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The Journal of Neuroscience, December 15, 2001, 21(24):9572-9584
Schwann Cells Express Active Agrin and Enhance Aggregation of
Acetylcholine Receptors on Muscle Fibers
Jie-Fei
Yang,
Guan
Cao,
Samir
Koirala,
Linga V.
Reddy, and
Chien-Ping
Ko
Section of Neurobiology, Department of Biological Sciences,
University of Southern California, Los Angeles, California 90089-2520
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ABSTRACT |
To explore novel roles of glial cells in synaptic function and
formation, we examined the expression of agrin in frog Schwann cells
and tested their role in the aggregation of acetylcholine receptors
(AChRs). Using reverse transcription-PCR, we found that Schwann
cells along nerve fibers in tadpoles expressed only the inactive agrin
isoform B0 but began to also express active agrin isoforms B11 and B19
at approximately metamorphosis. During nerve regeneration in the adult,
the expression of these active agrin isoforms in Schwann cells was
upregulated, including the appearance of the most potent isoform, B8.
This upregulation was induced by regenerating axons but not by nerve
injury per se. In muscle cultures, the presence of adult Schwann cells
enhanced the number and the total area of AChR aggregates 2.2- and
4.5-fold, respectively, and this enhancement was eliminated by heparin
treatment. Furthermore, adult Schwann cells in culture expressed active
agrin isoforms and produced agrin protein. Using a novel technique to
selectively ablate perisynaptic Schwann cells (PSCs) at the
neuromuscular junction, we found that PSCs also expressed active agrin
isoforms B11 and B19, and these active isoforms were upregulated,
including the appearance of B8, during reinnervation. Observation
in vivo showed that extrajunctional AChR aggregates were
associated with PSC sprouts after nerve injury and subsequent
reinnervation. These results suggest that, contrary to the prevailing
view that only neurons express active agrin, glial cells also express
active agrin and play a role in the aggregation of AChRs both in
vitro and in vivo.
Key words:
acetylcholine receptors; agrin; complement; frog; glia; muscle; neuromuscular junction; nerve regeneration; Schwann cells
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INTRODUCTION |
Glial cells outnumber neurons and
are widely distributed throughout the nervous system, including at the
chemical synapse. However, our knowledge of the role of glial cells in
the synapse is rudimentary. As with other chemical synapses, the
neuromuscular junction (NMJ) is composed of three intimately juxtaposed
cellular elements: the presynaptic nerve terminal, the postsynaptic
specializations, and synapse-associated glial cells. Previous studies
on the NMJ, the best-understood synapse, have focused almost
exclusively on the role of the presynaptic nerve terminal and the
postsynaptic specializations (Sanes and Lichtman, 1999
). The role of
synapse-associated glial cells, which are called perisynaptic Schwann
cells (PSCs) (also known as terminal Schwann cells) at the NMJ, has
been overlooked until recently. One of the key findings regarding the
role of PSCs is that PSCs sprout profusely after nerve injury and lead regenerating axons and nerve terminal sprouts (Reynolds and Woolf, 1992; Son and Thompson, 1995a,b; O'Malley et al., 1999; Koirala et
al., 2000). Nerve terminals also grow along the preceding PSC sprouts
seen during synaptic remodeling in intact frog muscles (Chen et al.,
1991; Chen and Ko, 1994; Ko and Chen, 1996), as well as during
synaptogenesis in tadpole muscles (Herrera et al., 2000). These
studies suggest that PSCs play important roles in synaptic repair
and growth (Son et al., 1996). It has also been shown that glial cells
modulate synaptic function at the NMJ (Robitaille, 1998; Castonguay and
Robitaille, 2001) and the CNS synapse (Araque et al., 1999; Bacci et
al., 1999). Thus, glial cells should be viewed as an active partner of
the tripartite chemical synapse in both the PNS and the CNS.
To further explore synapse-glial interactions, the present study aimed
to test a hypothesis that glial cells play a role in the aggregation of
acetylcholine receptors (AChRs). It has been well established that
agrin plays a major role in the aggregation of AChRs and the
differentiation of the postsynaptic apparatus at the NMJ (McMahan,
1990; Ruegg and Bixby, 1998; Sanes and Lichtman, 1999). Motor neurons
express different isoforms of the agrin protein; three isoforms, B8,
B11, and B19, with inserts of 8, 11, and 19 amino acids, respectively,
at the B (for chick and frog)/Z (for rat) site are active in the
aggregation of AChRs, and one isoform, B0, without the inserts, is
inactive (Ferns et al., 1992, 1993; Ruegg et al., 1992; Tsim et al.,
1992). Although various non-neuronal tissues, including muscle fibers,
also express the inactive isoform of agrin (Ferns et al., 1992; Ruegg
et al., 1992; Tsim et al., 1992; Ma et al., 1994; Smith and O'Dowd,
1994), the prevailing view is that only neurons express active isoforms
of agrin and induce AChR aggregation (Ruegg and Bixby, 1998). However,
it has not been rigorously tested whether and how Schwann cells express active agrin isoforms and whether Schwann cells also play a role in the
aggregation of AChRs. The present study addressed these questions.
Parts of this work have been published previously in abstract form
(Qiang et al., 1998; Cao et al., 1999; Yang and Ko, 1999).
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MATERIALS AND METHODS |
Animals. Adult grass frogs (Rana pipiens)
(7-8 cm rump-to-nose length; weighing 25-35 gm) were obtained from
Charles Sullivan (Nashville, TN) and maintained in the laboratory for
at least 2 weeks before experiments. Frogs were kept at 24°C on a 12 hr light/dark cycle in individual tanks and fed with mealworm
(Tenebrio molitor) larvae twice a week. Adult Xenopus
laevis were obtained from Nasco (Fort Atkinson, WI), bred
following their methods, and embryos were staged according to the
system of Nieuwkoop and Faber (1994). Tadpoles and juveniles of
bullfrog (Rana catesbeiana) at different stages were
obtained from Charles Sullivan.
Adult Schwann cell culture. Adult Xenopus sciatic
nerves were dissected out, and epineurial membranes were removed.
Nerves were cut into small pieces (~2 mm) and digested with
0.3% collagenase and 0.25% trypsin-EDTA (Life Technologies,
Gaithersburg, MD). Dissociated cells were plated on
laminin-1-coated culture dishes with culture medium consisting of 45%
Leibovitz's L-15 medium (Life Technologies), 45% Ringer's solution
(in mM: 115 NaCl, 2 CaCl2,
2.5 KCl, and 10 HEPES, pH 7.4) and 10% fetal calf serum (Life
Technologies). Serum-free medium, L-15/Ringer's solution (1:1, v/v),
was used from the second week and subsequently changed once every week.
Because the sciatic nerve does not contain any neuronal cell bodies, it
is virtually impossible that the Schwann cell culture would be
contaminated with neurons. The identity of Schwann cells in culture was
verified by staining with monoclonal antibody (mAb) 2A12 (Astrow et
al., 1998) or anti-glial fibrillary acidic protein (GFAP) antibody
(Georgiou et al., 1994). Only cultures containing 90% or more Schwann
cells (the rest were fibroblasts) were used for coculturing with
muscle. After Schwann cells had been cultured for 3-4 weeks,
Xenopus muscle was added. Pure embryonic Xenopus
muscle cultures were prepared according to Tabti and Poo (1994).
Briefly, neural tubes and associated myotomal tissues of stage
21-23 Xenopus embryos were dissected, and myotomal
tissues were detached from the neural tube after 15 min 0.1%
collagenase treatment. The detached myotomal tissue was further
dissociated in Ca2+- and
Mg2+-free Ringer's solution. The
dissociated cells were then plated on either coverslips with adult
Xenopus Schwann cells grown on them or coverslips coated
with laminin-1. The culture medium contained 50% L-15 and 50%
Ringer's solution. For heparin treatment, 300 µg/ml heparin (H-3393;
Sigma, St. Louis, MO) was included in the muscle medium or Schwann
cell-muscle coculture medium. On day 7 in coculture, cultures were
fixed with 2% paraformaldehyde and stained with Texas Red-tagged
-bungarotoxin (-BTX) (0.3 µg/ml; Molecular Probes, Eugene, OR)
for AChR aggregates and mAb 2A12 for Schwann cells. Images were
captured with a Spot Digital Camera (Diagnostic Instruments, Sterling
Heights, MI), and the number and size of AChR aggregates were analyzed
using ImageTool (University of Texas Health Science Center at San
Antonio, San Antonio, TX).
Reverse transcription-PCR. Adult frogs were anesthetized
with 15-30 min immersion in 0.1% tricaine (3-aminobenzoic acid ethyl ester; Sigma). For the short-term denervation study, the sciatic nerve
was transected and allowed to regenerate. For the long-term denervation
study, a 5 mm segment of the sciatic nerve was removed, and the severed
nerve was examined every 2 weeks to visually verify that the distal
stump was completely segregated from the proximal stump. At different
time points after axotomy, both distal and proximal nerve stumps were
collected and analyzed by reverse transcription (RT)-PCR. Segments of
these stumps were stained with anti-neurofilament 200 antibody (Sigma)
to verify the presence or the absence of axons.
Tissues including spinal cord, sciatic nerve trunk, and cutaneous
pectoris (CP) muscle of frog, or cultured cells were collected, frozen
in liquid nitrogen, and stored at 
70°C. The epineurial sheath,
which contains fibroblasts, was routinely removed. Total RNAs of these
tissues were isolated using QuickPrep Total RNA extraction kit
(Amersham Pharmacia Biotech, Arlington Heights, IL) and were reverse
transcribed using oligo-dT primer by First-Strand cDNA synthesis kit
(Amersham Pharmacia Biotech). PCR reaction mixtures were prepared with
cDNAs from reverse transcription using the PCR Supermix (Life
Technologies). The PCR reaction was performed using the Robocycler
Gradient 40 (Stratagene, La Jolla, CA) for 30 cycles of 94°C for 1 min, 57°C for 1.5 min, and 72°C for 1.5 min in a 50 µl volume
containing 0.8 mM dNTPs, 1× PCR buffer (20 mM
Tris-HCl, pH 8.4, and 50 mM KCl), 1.5 mM
MgCl2, and 0.25 U of Taq
DNA-polymerase. The PCR cycle numbers and composition of the PCR buffer
were optimized to fall in the linear range of signal. Primers flanking
the frog agrin alternative splicing site B were designed based on the
GenBank sequence under accession number AF096690 (Werle et al., 1999):
forward, 5' 574 TTT GAC GGA AAG ACT TAC CTG
594 3'; and backward, 5'
726 GGC TTC AGT CTT TAT GCT CAG CTC
702 3'. The PCR products were analyzed on
polyacrylamide gels following Sambrook and Russell (2001). After
electrophoresis, the gels were visualized by UV transilluminator and
imaged with a digital camera. The PCR fragments were subcloned into
pCR2 vector (Invitrogen, Carlsbad, CA), and the identity of each
fragment was confirmed by DNA sequencing (performed by Research
Genetics, Huntsville, AL). Nested PCR was used to further confirm that
the PCR products represented the frog agrin gene fragments. The
internal nest primers (forward, 5' 590 CCT GGA
GTA CCA CAA A 606 3'; and backward, 5'
707 AGC TCA AAT TCA TTG GT
690 3') were located within the PCR fragment
generated from the previous PCR reaction and thus were used to ensure
the identity of those PCR products. To reveal the relative abundance of
different agrin isoforms, bands of RT-PCR data were scanned, and the
number of pixels in each band was calculated and expressed as the
percentage of the total agrin isoforms within the same lane. To verify
the absence of neuronal mRNA contamination in our samples, primers for
Xenopus neurofilament were designed according to the GenBank
sequence under accession number U85969 (Gervasi and Szaro, 1997):
forward, 5' 298 TAC ATC GAG AAG GTC CAT
315 3'; and backward, 5'
1169 AAA AGT TTC CTG TAT GCA
1152 3'.
SDS-PAGE and immunoblotting. Schwann cells in culture
(5 × 106 cells per lane) were
collected by 0.25% trypsin-EDTA treatment and lysed in a buffer
containing 2% SDS and 62.5 mM Tris-HCl, pH 7.4. Conditioned media from Schwann cell and muscle cocultures or from pure
muscle cultures were concentrated using Microcon-30 (Millipore,
Bedford, MA). The concentrated media or the total lysate of Schwann
cells were collected and prepared for SDS-PAGE (Sambrook and
Russell, 2001).
After electrophoresis, proteins in the polyacrylamide gels were
transferred to a polyvinylidene difluoride membrane (Immobilon-P; Millipore) using a Mini Trans-Blot Cell (Bio-Rad, Hercules, CA). The
membrane was stained with Ponceau S [0.1% Ponceau S (w/v) in 5%
acetic acid (v/v)] to confirm the presence of proteins. The membrane
was blocked with 5% dry nonfat milk in Tris-buffered saline-Tween 20 (TBS-T) (20 mM Tris, 0.14 M NaCl, pH 7.6, and 0.1% Tween 20) for 1 hr at room temperature, followed by incubation with anti-agrin mAb C3 (a kind gift from Dr. Earl W. Godfrey, Eastern
Virginia Medical School, Norfolk, VA) (Godfrey et al., 1988) at
1:100 for 1 hr at room temperature. The membrane was thoroughly washed
with TBS-T before it was further incubated with alkaline
phosphatase-conjugated goat anti-mouse IgG secondary antibody for 1 hr
at room temperature. After TBS-T rinses, immunoreactivity was detected
using Alkaline Phosphatase Conjugate Substrate kit (Bio-Rad).
PSC ablation in the cutaneous pectoris muscle.
Affinity-purified mAb 2A12 (Astrow et al., 1998) [60 µg/ml in 100 µl of normal frog Ringer's solution (NFR)] was injected bilaterally
beneath the frog CP muscle. The CP muscles were dissected the next day and incubated in guinea pig complement at 30°C for 1 hr. The guinea pig complement was diluted with additional 40% distilled water to
maintain normal frog osmolarity. Immediately after the complement treatment, the CP muscle on one side was then taken for RT-PCR sampling
by separating the NMJ-rich and NMJ-poor regions according to the
innervation pattern visible under a dissecting microscope. The
contralateral muscle was used to confirm PSC cell death by staining
with ethidium homodimer-1 (EthD-1) (1:500 in NFR; Molecular Probes) and
FITC-peanut agglutinin (PNA) (1:100 in NFR) for 1 hr. EthD-1 labels
nuclei of lysed cells, whereas PNA recognizes the extracellular matrix
associated with PSCs (Ko, 1987).
Observation of extrajunctional AChR aggregates in the frog
muscle. Adult frogs were anesthetized by 15-30 min immersion in 0.1% tricaine, and the nerve to the CP muscle was transected 1-2 mm
from the muscle. This denervation procedure permits reinnervation of
the muscle beginning at ~14 d after axotomy. At 17-28 d after axotomy, animals were killed, and CP muscles were dissected.
Muscles in whole mount were treated with 3% normal goat serum in NFR
for 45-60 min, after which they were incubated overnight in 7.2 µg/ml mAb 2A12 in NFR. Biotinylated goat anti-mouse IgM µ-specific
secondary antibody (Sigma) was applied for 45 min at 1:50 in NFR, along with Texas Red-tagged -BTX at 0.5 µg/ml. After 30 min wash with NFR, the muscle was fixed in 2% paraformaldehyde for 30 min. In some cases, instead of -BTX, a monoclonal antibody against the 
subunit of the nicotinic AChR (Affinity BioReagents, Golden, CO)
was used at 1:400 after fixation.
7-amino-4-methylcoumarin-3-acetic acid-tagged streptavidin at 1:50 or
1:75 was applied to reveal mAb 2A12, and Texas Red-tagged goat
anti-mouse IgG at 1:400 revealed the anti-AChR antibody. Muscles
were rinsed again and then treated for 45-60 min with 5% goat serum
in frog PBS containing 0.5% Triton X-100 (PBS-T). Anti-synapsin I
(polyclonal antibody kindly provided by Dr. Bai Lu, National Institutes
of Health, Bethesda, MD) and anti-neurofilament 200 kDa (Sigma)
antibodies, diluted 1:300 and 1:400, respectively, in 0.5% PBS-T
containing 5% goat serum, were applied overnight. These antibodies
were revealed using 1:150 FITC-tagged goat anti-rabbit IgG and 1:400
FITC-tagged goat anti-mouse IgG 
-specific secondary antibodies,
respectively, in 0.3% PBS-T containing 5% goat serum. After rinsing
with PBS and stripping the muscles of excess connective tissue, the
muscles were post-fixed for 30 min in 2% paraformaldehyde.
Histological staining for acetylcholinesterase (AChE) was performed
according to the method of Karnovsky (1964). Finally, the muscles were
mounted on glass slides in Citifluor mountant (Ted Pella Inc., Redding,
CA) and viewed under fluorescence optics. Digital images were captured,
and measurements of length and area were made using the Scion Image
program (Scion Corp., Frederick, MD). Measurements are mean ± SEM.
To determine the properties of extrajunctional AChR aggregates and
their association with PSCs, four measurements were made: 1, size
(length along major axis) of AChR aggregate; 2, distance from closest
axon or nerve terminal; 3, distance from the closest PSC process; and
4, distance from the original synapse (marked by AChE stain).
Extrajunctional AChR "aggregates" were defined as -BTX- or
anti-AChR antibody-positive clusters that measured 10 µm or
greater in length and were at least 10 µm away from AChE-stained original sites. AChR aggregates were considered to be associated with
PSCs alone when the aggregates overlapped with 2A12 staining and were
at least 10 µm away from synapsin I- or neurofilament 200-stained
processes. All measurements were restricted to the area on each muscle
fiber extending 200 µm beyond either end of an NMJ; AChR aggregates
were very rare outside of this area.
It is important to rule out the possibility that AChR aggregates may be
associated with PSCs purely by chance. Our approach was to determine
the proportion of observed muscle area occupied by PSC sprouts and then
calculate the relative probability that randomly distributed AChR
aggregates would fall within this area. For muscle fibers with
PSC-associated AChR aggregates, the average PSC sprout area (without
nerve terminals or axons) was ~330
µm2. The total observed area per
muscle fiber was ~57350 µm2. After
excluding original junctional sites, as well as areas under axons and
nerve terminals, the total extrajunctional muscle area was ~52950
µm2. Within this extrajunctional area,
an AChR aggregate could randomly either fall within the area occupied
by PSC sprouts (330 µm2) or outside
(52620 µm2). The ratio of PSC sprout
area to surrounding muscle area is ~1:160 (330:52620). Therefore, if
extrajunctional AChR aggregates were randomly distributed over the
muscle surface, only 0.6% (1 of 160) would be expected to colocalize
with PSC sprouts alone. Our observed results were compared with these
expected results for random association using the
2 test.
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RESULTS |
Expression of agrin isoforms in developing and adult
Schwann cells
Previous molecular cloning studies have shown that motor neurons
express four isoforms of agrin that contain either 0, 8, 11, or 19 amino acids at the B (for chick and frog) or Z (for rat) site (Ferns et
al., 1992; Ruegg et al., 1992; Tsim et al., 1992; Werle et al., 1999).
Only B8, B11, and B19 isoforms are capable of inducing AChR aggregation
at the NMJ, whereas B0 is inactive in inducing AChR aggregation (Ferns
et al., 1992, 1993; Ruegg et al., 1992; Gesemann et al., 1995; Daggett
et al., 1996). To investigate expression of agrin isoforms in glial
cells, Schwann cells along the sciatic nerves of adult frogs
(Rana pipiens) were examined by RT-PCR. Although the
predominant cell type in the sciatic nerve is the Schwann cell, the
epineurial sheath, which is rich in fibroblasts, was routinely removed
to ensure that we used primarily Schwann cells for the RT-PCR study. We
found that frog Schwann cells along the sciatic nerve expressed not
only the inactive isoform B0, as shown previously (Werle et al., 1999), but also the active isoforms B11 and B19 (Fig.
1A, lane
6). To confirm the identity of agrin transcripts in Schwann
cells, each PCR fragment was subcloned and sequenced. PCR with nested
primers (see Materials and Methods) were also used to further confirm that these PCR products were frog agrin gene fragments (Werle et al.,
1999). Our results showed that agrin expression in adult Schwann cells
was similar to that in the spinal cord neurons, except for the absence
of the B8 isoform. In contrast to Schwann cells, the epineurial sheath,
which is rich in fibroblasts, of the frog sciatic nerve showed no agrin
expression at all (Fig. 1B, lane
6). To exclude the possibility of contamination with neuronal mRNAs in the sciatic nerve preparation, we performed RT-PCR
for neurofilament, a neuron-specific marker. As expected, spinal cord
tissue expressed neurofilament transcripts (Fig. 1C, lane 1, arrow). However, neurofilament
transcripts were expressed in neither Schwann cells along the sciatic
nerve (Fig. 1C, lane 2) nor muscle tissue (Fig.
1C, lane 3). Thus, it is unlikely that our
sciatic nerve sample is contaminated with neuronal mRNAs. It is most
likely that the active agrin isoforms seen in the sciatic nerve are
expressed by Schwann cells rather than by neurons or fibroblasts (see
Discussion).
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Figure 1.
Expression of agrin isoforms in developing and
adult Schwann cells. A, Expression of agrin isoforms in
Schwann cells along the sciatic nerve in adult and bullfrog tadpoles at
different developmental stages was examined by RT-PCR. Lane
1, The spinal cord of stage XXII bullfrog tadpoles showed all
four isoforms of agrin: B0, B8, B11, and B19. Lane 2,
Schwann cells at stage XXI expressed only the inactive B0 isoform.
Lane 3, A trace amount of the B11 isoform was detected
in Schwann cells at stage XXII. Lanes 4,
5, Schwann cells at stages XXIV and XXV expressed active
isoforms B11 and B19 besides B0. Lane 6, Adult Schwann
cells expressed three agrin isoforms, B0, B11, and B19, but did not
show B8. B, Fibroblasts in the epineurial sheath of the
sciatic nerve in bullfrog tadpoles at stages XXI-XXV (lanes
2-5) and in adult (lane 6) did not
express any agrin isoforms. As a positive control, the adult spinal
cord expressed all four agrin isoforms (lane 1).
C, RT-PCR showed neurofilament mRNA fragment (0.87 kb)
in the frog spinal cord tissue (lane 1,
arrow) but not in Schwann cells along the sciatic nerve
(lane 2) or muscle (lane 3).
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To determine the expression of active agrin isoforms in Schwann cells
during development, sciatic nerves from tadpoles of bullfrog
(Rana catesbeiana) at different developmental stages were
examined. Bullfrog tadpoles were chosen because their large size
provided ample developing Schwann cells for analysis. Before the onset
of metamorphosis (stage XXI), Schwann cells along the sciatic nerve
expressed only the inactive isoform B0 (Fig. 1A, lane 2). During and after metamorphosis (stages XXII to
XXV), Schwann cells began to express active isoforms B11 and B19, in addition to B0 (Fig. 1A, lanes 3-5).
Similar to adult Schwann cells (Fig. 1A, lane
6), developing Schwann cells did not express B8. In
contrast, spinal cord neurons of the above developmental stages, as
well as in adult, expressed all four isoforms (Fig. 1A,B, lane 1). Similar
to adult fibroblasts (Fig. 1B, lane
6), fibroblasts in the epineurial sheath of the sciatic
nerve in tadpoles at various stages expressed neither B0 nor active
agrin isoforms (Fig. 1B, lanes 2-5).
Expression of agrin isoforms in adult Schwann cells
after axotomy
To examine whether and how axons regulate the expression of adult
Schwann cell agrin, we compared Schwann cells along the frog
(Rana pipiens) sciatic nerve after short-term denervation, in which nerve regeneration was allowed, with Schwann cells after long-term denervation, in which nerve regeneration was prevented. The
result of the short-term denervation is shown in Figure
2, A and B. As
expected from the above study (Fig. 1A, lane
6), adult Schwann cells in intact frog sciatic nerve
expressed B0, B11, and B19 (Fig. 2A,
Intact). Two weeks after axotomy, Schwann cells along nerve
fibers (Fig. 2A, 2w) began to show
upregulation of active agrin isoforms in both proximal and distal nerve
segments. In addition to the upregulation of B11 and B19, the most
potent isoform in AChR aggregation, B8, now appeared. At this time
after the short-term denervation, axons began to reinnervate through the distal nerve stump, as revealed by anti-neurofilament 200 staining
(data not shown). To examine Schwann cell agrin upregulation after
axonal regeneration, each lane in Figure
2A (Distal) was analyzed based on
the area and intensity of each band (see Materials and Methods). This
information allowed us to reveal the relative abundance, and
changes in the expression pattern, of different agrin isoforms after
nerve injury. Figure 2B shows the percentage of all
active agrin isoforms combined (B8/B11/B19) relative to the total
isoforms (active plus inactive) within each individual lane at various
time periods after axotomy. The percentages of B8 and B0 in each lane
were also plotted individually. In Schwann cells along the intact
nerve, B11/B19 constituted <45% of total agrin isoforms. Two weeks
after axotomy, the active isoforms, including B8, constituted over 60%
of total agrin. This upregulation of active agrin became more prominent
as regeneration progressed and peaked ~6-7 weeks after nerve
transection. During this peak expression, there was a concomitant
decrease in the relative percentage of B0, and the pattern of Schwann
cell agrin expression closely resembled neuronal agrin expression (Fig.
2A, 4w-8w, compare with Fig.
1A,B, lane 1). From 8 weeks after axotomy, the active isoforms began to show downregulation
(Fig. 2B). However, 12 weeks after a single nerve
transection, when nerve regeneration was complete, the active agrin
expression level in Schwann cells remained slightly higher than that in
Schwann cells along intact nerves, and a trace amount of B8 (3%) could
still be detected (Fig. 2A,B).
Whether or not the normal expression pattern would be totally restored beyond 12 weeks after axotomy was not examined.
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Figure 2.
Upregulation of active agrin isoforms in adult
Schwann cells induced by nerve regeneration. Expression of agrin
isoforms in the Schwann cells along the frog sciatic nerve after
short-term denervation (A, B) and
long-term denervation (C, D) was examined
by RT-PCR. A, Schwann cells in the intact sciatic nerve
expressed only three agrin isoforms: B0, B11, and B19. However, after a
single nerve transection that allowed nerve regeneration, Schwann cells
in both the proximal and the distal nerve stumps upregulated the
expression of active isoforms, including the appearance of B8. The
number of weeks after axotomy is denoted on the top of
each lane. B, A plot shows changes in the
percentage of B0 (filled circles), B8
(open triangles), and all active isoforms (B8/B11/B19;
filled triangles) relative to the total isoforms (see
Materials and Methods) at different time points after short-term
denervation (n = 3 experiments; mean ± SEM).
The increase in the relative expression of active agrin and a
concomitant decrease in B0 began at ~2 weeks and peaked at ~6-7
weeks after short-term denervation. A trace amount of B8 (3%) was
still detected 12 weeks after axotomy. C, Schwann cells
in the sciatic nerve segment proximal to the transection site 2-6
weeks after long-term denervation upregulated the active agrin isoforms
compared with the intact nerve (top left panel).
The same Schwann cell samples along the intact nerve and the proximal
nerve segment 2-6 weeks after axotomy did not show any neurofilament
mRNA, in contrast to the spinal cord (bottom left
panel). The proximal segment contained regenerating
axons, as revealed by positive anti-neurofilament staining (top
right panel). In contrast, the distal segment, which was
chronically severed from the proximal segment, was absent of
anti-neurofilament staining (bottom right panel).
Schwann cells in the distal segment showed neither upregulation of B11
and B19 nor appearance of B8 (middle left panel).
D, A plot shows an increase in the relative expression
of all agrin isoforms (B8/B11/B19;
filled triangles) and B8 (open
triangles) in the proximal segment after long-term denervation.
However, in the chronically segregated distal segment, the relative
expression of the total active isoforms (B8/B11/B19; filled
squares) remained unchanged, and no B8 (open
squares) was detected up to 6 weeks after axotomy
(n = 3 experiments; mean ± SEM).
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To confirm that the upregulation of active agrin was triggered by
axonal regeneration instead of simply by nerve injury per se, we also
examined agrin expression in Schwann cells after long-term denervation
(Fig. 2C,D). At 2 weeks after axotomy, the
proximal nerve stump showed positive neurofilament staining (Fig.
2C, top right panel), indicative of nerve
regeneration, but the disconnected distal stump was absent of
neurofilament staining (Fig. 2C, bottom right
panel). As expected for regenerated nerves (Fig.
2A,B), Schwann cells in the
proximal nerve segment expressed all four isoforms of agrin (Fig.
2C, top left panel). In contrast, Schwann cells in the chronically denervated distal segment expressed only B0,
B11, and B19 and not B8 (Fig. 2C, middle left
panel). At 4-6 weeks after chronic denervation, the
contrast between the upregulation of active agrin in the proximal stump
and its absence in the distal stump became even more prominent. Similar
to the intact sciatic nerve (Fig. 1C), Schwann cells in the
regenerated proximal stump 2-6 weeks after axotomy did not express
neurofilament mRNA (Fig. 2C, bottom left
panel). Thus, the appearance of B8 isoform in the proximal
Schwann cells after nerve regeneration is likely not a contamination of
neuronal agrin mRNA. The difference in the relative expression of
active versus inactive isoforms between Schwann cells in the proximal
and distal segments is further shown in Figure 2D.
Schwann cells along the proximal nerve stump showed upregulation of all
active agrin isoforms from 45% to over 65% and B8 from 0% to over
17%, relative to the total agrin isoforms at 4-6 weeks after axotomy.
In contrast, Schwann cells in the distal nerve segment showed a total
absence of B8 and no upregulation of the other active isoforms, which
remained ~44-48% up to 6 weeks after long-term denervation. These
results suggest that upregulation of active agrin isoforms in Schwann
cells is not induced by nerve injury per se but instead by nerve regeneration.
Adult Schwann cells enhance AChR aggregation on muscle
in culture
Because adult Schwann cells express active agrin isoforms, we
investigated whether these glial cells play a role in the aggregation of AChRs on muscle fibers. To address this question, Xenopus
cultures were used for functional assay. Schwann cells obtained from
adult Xenopus sciatic nerves were cocultured with primary
Xenopus myotubes, which were prepared from stage 21-23
embryos. In culture, these Schwann cells de-differentiate to a
nonmyelinating phenotype (Brockes et al., 1979), similar to PSCs. All
cultures contained similar density of Schwann cells at ~15,000 cells
per dish (35 mm in diameter). The identity of Schwann cells in
vitro was confirmed by positive staining with mAb 2A12 that
recognizes the Schwann cell membrane (Astrow et al., 1998) or with
anti-GFAP antibody (Georgiou et al., 1994). Embryonic Schwann cells
were not used because they expressed only the inactive B0 isoform as
shown above. To analyze the effect of Schwann cells on AChR
aggregation, we compared AChR clusters on muscle fibers grown in the
presence of adult Schwann cells (SC+M) with those on muscle grown alone
(M) in culture. Neurons were not added to either of these cultures, and
the absence of neuronal contamination was further verified by the
absence of staining with anti-neurofilament antibody in these cultures (data not shown). Figure 3 shows an
example of an SC+M culture (A, A') and an M
culture (B, B') 7 d in culture. The general
cellular morphology in both cultures appeared similar under
phase-contrast optics (A, B). In addition, there
was no significant difference in muscle fiber length between SC+M
culture (241.8 ± 51.2 µm; mean ± SD; n = 96) and M culture (221.6 ± 33.2 µm; n = 98).
However, as shown with fluorescence staining, there were more and
larger AChR clusters (arrowheads in A' and
B') labeled with -BTX in SC+M cultures than in M cultures
(compare A' with B'). The average number of AChR
clusters per 100 µm muscle length in the SC +M culture was 2.2-fold
(p < 0.001) of that in the M culture (Fig. 3C). The average area of individual AChR "hotspots" in
the SC+M culture was 43.4 ± 7.3 µm2, which was significantly larger
(p < 0.02) than the 22.7 ± 3.7 µm2 in the M culture. Thus, the total
area of AChR clusters per 100 µm muscle length in the SC+M culture
was also significantly enlarged (p < 0.001) to
4.5-fold of that in the M culture (Fig. 3D).
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Figure 3.
Adult Schwann cells enhance AChR aggregation on
muscle in culture. Aggregation of AChRs on embryonic
Xenopus SC+M cells for 7 d was compared with
that in M. A, B, Phase-contrast images of
an SC+M culture (A) and an M culture
(B) show similar muscle morphology.
A', B', Fluorescence images of the same
cultures labeled with Texas Red-conjugated -BTX show more AChR
hotspots in SC+M than M. C, A plot shows a significant
increase in the number of AChR hotspots per 100 µm muscle length in
SC+M, which was 2.2-fold of that in M. Treatment with heparin (300 µg/ml) eliminated this increase in the coculture (SC+M+H), but the
treatment did not show effect in pure muscle culture (M+H).
D, A plot shows the total area (in square micrometers) of AChR clusters per
100 µm of muscle length in SC+M, M, and after heparin treatment in
SC+M+H and M+H. Similar to C, the area of AChR clusters
in SC+M was significantly enlarged to 4.5-fold of that in M, and this
increase was eliminated by heparin treatment. In C and
D, n denotes the number of muscle fibers
observed, and all values are mean ± SD. E-H, SC+M
cocultures were examined to determine the spatial relationship between
AChR aggregates and Schwann cell-muscle contacts. The contacts
(arrows) could be observed with phase contrast optics
(E) and confirmed with staining of Schwann cells
with mAb 2A12 (F). The same culture
double-labeled with -BTX (G) showed that the
majority of these contacts were not colocalized with AChR clusters
(arrowheads). Only in rare cases were AChR clusters
colocalized with the Schwann cell-muscle contact
(asterisk). The spatial relationship between AChR
clusters and the contacts is further shown in H, which
is a merged image of E-G.
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To test whether AChR aggregates colocalized with contacts between
Schwann cell processes and muscle fibers, we double-labeled the SC+M
culture with mAb 2A12 for Schwann cells (Fig. 3F) and -BTX for AChRs (Fig. 3G). As shown in Figure 3E, contacts
(arrows) between Schwann cell processes and muscle fibers
were visualized with phase-contrast optics. Schwann cell processes were
verified with mAb 2A12 staining (F). Although there
were numerous AChR clusters (Fig. 3G,H,
arrowheads) on the muscle fibers, most of them were not
associated with Schwann cell processes. Only in rare cases were AChR
clusters colocalized with Schwann cell-muscle contacts (Fig.
3E-H, asterisks). In 138 muscle fibers from nine cultures observed, 98.5% of AChR aggregates were not colocalized with
Schwann cell-muscle contacts. This contrasts with nerve-muscle contacts, in which AChRs are clustered (Cohen et al., 1979; Frank and
Fischbach 1979). Our results suggest that the increase in AChR
aggregation by Schwann cells is likely mediated by soluble factors,
such as agrin, released into the tissue culture medium.
Is the increase in AChR aggregation by Schwann cells mediated
by agrin?
To determine whether the enhancement of AChR aggregation by
Schwann cells might be mediated by agrin, we first verified with RT-PCR
the expression of active agrin transcripts in cultured Schwann cells.
As shown in Figure 4A,
lane 3, Schwann cells grown alone in culture expressed not
only B0 but also the active isoforms B11 and B19. The expression of
these three isoforms was also seen in SC+M culture (Fig.
4A, lane 4) throughout the
coculture period. Similar to the expression pattern in adult Schwann
cells after chronic denervation in vivo (Fig.
2C,D), B8 was not expressed in these
"denervated" Schwann cells, either grown alone or with muscle, in
culture. The absence of neurons was verified by the absence of
neurofilament mRNA in these cultures (data not shown). Muscle cells
grown alone in culture (M) expressed only the inactive isoform B0 (Fig.
4A, lane 2), whereas spinal neurons in
culture (Fig. 4A, lane 1) expressed all
four isoforms. Thus, Schwann cells, muscles, and neurons in
vitro retain their characteristic expression pattern of agrin
isoforms as seen in vivo.
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Figure 4.
Agrin expression in adult Xenopus
Schwann cells in vitro. A, Agrin isoform
expression in cultured Schwann cells was examined by RT-PCR.
Lane 1, As a positive control, spinal neurons in culture
showed all four agrin isoforms. Lane 2, Muscle in
culture displayed only the B0 isoform. Lane 3, Schwann
cells in culture expressed B0, B11, and B19 isoforms. Lane
4, SC+M for 7 d also showed B0, B11, and B19 isoforms.
Lane 5, As a negative control, the lane showed no bands
when no RNA samples were used for RT-PCR. B, Agrin
protein in cultured Schwann cells and conditioned media was detected by
Western blot using the anti-agrin antibody C3. Lane 1,
Native agrin protein over 200 kDa was detected in the total cell lysate
of cultured Schwann cells. Lane 2, As a positive
control, brain tissues showed similar immunoreactivity over 200 kDa.
Conditioned media of Xenopus Schwann cells cultured for
6 weeks (SC-CM; lane 3) and
Xenopus muscle conditioned media from day 7 in culture
(Mu-CM; lane 4) showed a positive
band at ~70 kDa (arrow), but bands over 200 kDa were
absent. The 70 kDa band may be an inactive degradation product of
native agrin protein.
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To confirm that the expression of active agrin isoforms resulted in the
production of agrin protein, lysate of cultured Schwann cell bodies was
examined with Western blot using anti-Xenopus agrin mAb C3
(Godfrey et al., 1988). Similar to brain tissues (Fig.
4B, lane 2), cultured Schwann cells (Fig.
4B, lane 1) expressed native agrin protein
at over 200 kDa. To determine whether agrin protein was released into
the culture medium, Western blot of conditioned medium from Schwann
cells cultured for 6 weeks was examined. As shown in Figure
4B, lane 3, the Schwann cell-conditioned medium did contain the agrin protein (arrow), as also
observed in the muscle-conditioned medium (lane 4) as
a control. However, in contrast to the native agrin protein over 200 kDa seen in Schwann cell bodies, the 70 kDa band seen in the
conditioned medium likely belongs to an inactive degraded product of
the agrin protein, as shown previously in basal lamina extracts of the
Torpedo electric organ (Nitkin et al., 1987). Consistent
with the absence of native agrin protein bands above 200 kDa, we found
that Schwann cell-conditioned medium added to M culture alone did not
cause an increase in AChR aggregates (data not shown). This is contrary
to the effect seen in SC+M coculture (Fig. 3), in which muscle fibers
were probably exposed to a continuous supply of native agrin protein
from cocultured Schwann cells (see Discussion).
To further determine whether Schwann cell-derived agrin might be
involved in the enhancement of AChR aggregation, Schwann cell and
muscle cocultures were treated with heparin (300 µg/ml), which has
been shown to inhibit agrin-mediated AChR clustering on muscle (Hopf
and Hoch, 1997). As shown in Figure 3, C and D, the number and the area of AChR hotspots in SC+M cultures were significantly reduced by heparin treatment (SC+M+H). However, spontaneous AChR aggregation, which is independent of agrin, was not
affected in M cultures by heparin treatment. Although the direct
evidence is lacking (see Discussion), the present data are consistent
with the hypothesis that the enhancement of AChR aggregation seen in
SC+M cultures is mediated by Schwann cell-derived agrin.
Agrin expression in the perisynaptic Schwann cell at the NMJ
To investigate whether Schwann cells at the NMJ are similar to
Schwann cells along the nerve with respect to expression of active
agrin isoforms, we used a novel technique to selectively ablate PSCs
in vivo (Reddy et al., 1999) (the detailed procedure of this
technique will be described in a future paper). To ablate PSCs in
vivo, we took advantage of mAb 2A12, which specifically labels the
PSC surface membrane (Astrow et al., 1998). Labeling of PSCs with mAb
2A12 followed by treatment with guinea pig complement results in the
formation of membrane-attack complexes on PSCs. Membrane-attack
complexes form pores on the cell membrane and cause the
antibody-labeled cells to lyse (Howard and Hughes-Jones, 1988). To
verify PSC lysis, EthD-1, which stains the nuclei of cells with damaged
membranes, was applied after the complement treatment. As shown in
Figure 5A, EthD-1 labeling (in
red, arrows) colocalized with the soma of PSCs,
which were revealed by mAb 2A12 immunofluorescent staining (in
green). Over 80% of PSCs were ablated using this treatment.
In contrast to PSCs, Schwann cells along axons are not labeled with mAb
2A12 in whole mount because mAb 2A12 does not penetrate the perineurium
surrounding nerve fibers (Astrow et al., 1998) and thus are not lysed.
We did not observe any morphological damage to nerve terminals or
muscle fibers after mAb 2A12 and complement treatment (L. V. Reddy, S. Koirala, Y. Sugiura, and C. P. Ko, unpublished
observations). In control experiments using complement treatment
alone, without 2A12 application, over 95% of PSCs were not lysed as
indicated by the absence of EthD-1 labeling (Fig. 5B,
arrows). Because mAb 2A12 was not used for the control, PSCs
were revealed with FITC-conjugated PNA (in green), which
recognizes the extracellular matrix associated with PSCs (Ko, 1987).
The few EthD-1-positive cells not colocalized with PSC cell bodies in
the control were likely blood cells that are inevitably damaged during
muscle dissection. The above results indicate that PSCs in
situ can be selectively ablated using mAb 2A12 followed by
complement treatment (Reddy, Koirala, Sugiura and Ko,
unpublished observations).
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Figure 5.
Agrin expression in perisynaptic Schwann cells at
the neuromuscular junction. A, PSCs at the NMJ were
selectively ablated in vivo by labeling the PSC membrane
with mAb 2A12 and subsequent application of guinea pig complement,
which forms membrane attack complexes and lyses cells. PSC lysis was
revealed by ethidium homodimer-1, which enters cells with damaged
membranes and stains their nuclei (in red,
arrows). PSCs (in green) were revealed by
labeling with FITC-conjugated secondary antibody to mAb 2A12. Over 80%
of PSCs were ablated by this technique. B, A control
muscle treated with complement alone did not show dead PSCs
(arrows mark nuclei of living PSCs). On average, <5%
of PSCs were ablated in the absence of mAb 2A12. PSCs in the control
group were revealed with FITC-conjugated peanut agglutinin (in
green). C, The acute effect of PSC
ablation on agrin expression in normal and reinnervated muscles was
examined by RT-PCR. Within the NMJ-rich region of intact CP muscle
(PSC+), agrin expression typical of adult Schwann cells
was observed (lane 1, compare with Fig.
2A, Intact), whereas only the B0
isoform was found in the NMJ-poor region (NMJ,
lane 2), which consists mainly of muscle tissue. When
PSCs were killed, active agrin bands (B11 and B19) became much weaker
(lane 3). With PSCs intact, 2 weeks after axotomy, nerve
regeneration induced the upregulation of B11/B19 and the appearance of
B8 (lane 5), similar to Schwann cells along the
regenerated sciatic nerve (lane 5, compare with Fig.
2A). This upregulation of active isoforms was
significantly weaker after PSC ablation (lane 7).
NMJ-poor regions showed no response to PSC ablation either with or
without denervation (lane 2 vs lane
4, lane 6 vs lane 8).
Lane 9, As a negative control, the lane showed no bands
when no RNA samples were used for RT-PCR.
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To test whether PSCs express active agrin isoforms, intact and
PSC-ablated muscles were examined by RT-PCR (Fig. 5C). The central region of the CP muscle enriched in NMJs (NMJ+) was
separated from the remaining NMJ-poor (NMJ) region. In
innervated muscles without PSC ablation (Fig. 5C), agrin
isoforms B0, B11, and B19 were detected in the NMJ-rich region
(NMJ+, PSC+, lane 1). This expression
pattern was similar to that of adult Schwann cells along intact nerve
(Fig. 2A). In contrast, only the B0 isoform was found
in the NMJ-poor region (Fig. 5C, NMJ,
lane 2), which consists mainly of muscle tissue. After acute
ablation of PSCs in innervated muscle, active agrin bands (B11 and B19)
became much weaker (Fig. 5C, lane 3; Table
1). Thus, the reduction in B11 and B19 is
correlated with the absence of PSCs. The weaker bands of active
isoforms in lane 3 after PSC ablation may originate, in
part, from the unablated Schwann cells along intramuscular nerves (see
Discussion). These axonal Schwann cells constitute ~20% of the total
number of Schwann cells as counted by nuclear staining (Hoechst 33342)
in the CP muscle.
To investigate whether and how agrin expression in Schwann cells
responds to nerve injury, we performed RT-PCR of reinnervated muscles
(Fig. 5C). In reinnervated muscles 2 weeks after nerve transection, upregulation of B11/B19 and the appearance of B8 were
found in the NMJ-rich tissue with intact PSCs (lane 5;
compare with lane 1). This result is similar to the
upregulation of active agrin isoforms in Schwann cells along
regenerating sciatic nerves (Fig. 2A). After acute
PSC ablation, the active agrin bands in the reinnervated NMJ-rich
tissue were significantly weaker (compare lane 7 with
lane 5), indicating that the active agrin mRNA was present
in PSCs. These changes in agrin expression were quantified by comparing
the relative intensity of various bands within the same lane (Table 1).
After PSC ablation, the relative expression of active agrin isoforms in
the NMJ-rich tissue in innervated muscles was decreased from 37 to 14%
of total agrin isoforms and in reinnervated muscles from 63 to 26%
(Table 1). NMJ-poor regions showed no response to PSC ablation either
with or without axotomy (Fig. 5C, lane 2 vs
lane 4, lanes 6 vs lane 8). Thus, PSC
ablation does not affect the agrin expression pattern in muscle.
Extrajunctional AChR clusters are associated with PSC sprouts
in vivo
The expression of active agrin isoforms and their upregulation in
PSCs after nerve regeneration suggested that PSCs might also play a
role in the aggregation of AChRs in vivo. Because of
the close apposition between PSCs and nerve terminals, it is impossible
to distinguish the contribution of PSCs to AChR aggregation from that
of nerve terminals in normal muscles. To circumvent this difficulty, we
took advantage of the fact that PSCs sprout, often tens of micrometers,
beyond the tips of regenerating nerve terminals during reinnervation
(Koirala et al., 2000). This allowed us to examine in vivo
whether PSCs play a role in AChR aggregation independent of nerve
terminals. CP muscles of adult frog (Rana pipiens) were
excised 17-28 d after axotomy, when many NMJs bore PSC sprouts longer
than corresponding regenerating nerve terminals (Koirala et al., 2000)
and then were fluorescently stained for AChRs, PSCs, axons, and nerve
terminals. Figure 6 shows an example of
an NMJ with extrajunctional AChR aggregation associated with PSC
sprouts 4 weeks after axotomy. A prominent PSC sprout labeled with mAb
2A12 (Fig. 6A, arrowhead) extended over
100 µm beyond the boundary of the original synaptic site (Fig.
6A,B,D,
arrows), which was marked by AChE staining (Fig.
6D). Large and diffuse AChR aggregates labeled with
-BTX staining (Fig. 6B, arrowhead) were
associated with the PSC sprout. Although regenerating nerve terminals
were observed in neighboring junctional branches (Fig. 6C,
asterisk), nerve terminals and axons were absent along these extrajunctional AChR aggregates (Fig. 6C,
arrowhead), suggesting that PSCs directly induce such
extrajunctional AChR aggregates. In contrast to the junctional AChRs,
which appeared as bright bands and were sharply colocalized with the
original junctional site, the extrajunctional AChRs appeared less dense
and were not strictly confined by the boundary of the PSC sprouts
(compare the region marked by the arrowhead in A
with that in B). This lack of precise colocalization between
the extrajunctional AChRs and PSC sprouts indicates that the AChR
clusters are not on the PSC membrane but rather on muscle. Furthermore,
some extrajunctional AChR clusters show a staining pattern that
resembles sarcomeres. Figure 6E shows an example of
this "sarcomeric" pattern revealed in a high-magnification view of
the extrajunctional AChR cluster marked by the arrowhead in
Figure 6B. The brighter stripes of AChR staining
match with the light bands of sarcomeres in the same region of the
muscle fiber (Fig. 6F). The sarcomeric pattern is
not an artifact of intracellular staining of sarcomeres with -BTX
because the pattern was also seen in muscle that was freshly dissected and labeled with -BTX without any previous fixation, membrane permeabilization, or other staining procedures (Fig. 6G). The more pronounced sarcomeric pat- tern
seen in Figure 6G again matches with the light bands of
sarcomeres (Fig. 6H). PSC-associated AChRs with the
sarcomeric pattern and weaker staining than the junctional AChRs were
also observed with an antibody to the subunit of the nicotinic AChR
(data not shown). Besides PSC-associated extrajunctional AChR clusters,
small speckles of AChRs (~1-5 µm) randomly distributed over the
muscle surface were also seen with -BTX or anti-AChR staining in
denervated and reinnervated muscles (data not shown). These speckles
were too small to reveal the sarcomeric pattern and were not further
examined in the present study.
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Figure 6.
PSC sprouts may induce AChR aggregation in
vivo. NMJs 4 weeks after axotomy were fluorescently labeled
with mAb 2A12 for PSCs (A), -BTX for AChRs
(B), and antibodies to neurofilament 200 kDa and
synapsin I for axons and nerve terminals, respectively
(C). A, A prominent PSC sprout
(arrowhead) extended beyond the boundary of the original
junction delineated by AChE (arrow in D;
corresponding arrows in A and
B). B, A large extrajunctional AChR
aggregate (arrowhead) colocalized with the PSC sprout
(arrowhead in A). C, Axons
and nerve terminals were absent from extrajunctional AChR aggregates
(arrowhead) and PSC sprouts, suggesting that PSC sprouts
may directly induce AChR clustering in vivo.
Asterisk marks the closest extent of regenerating nerve
terminals. D, The original synaptic sites were labeled
with Karnovsky's AChE stain. The scale bar in D applies
to A-D. E, A high-magnification view of
the AChR aggregate marked by the arrowhead in
B. F, Bright-field view of the same
region of the muscle fiber as in E. The AChR
aggregate showed sarcomeric staining pattern with stripes of
brighter AChRs (arrowheads in E) matching
with the light bands (arrowheads in
F) of sarcomeres. G, An
extrajunctional AChR aggregate in a reinnervated muscle (3 weeks after
axotomy), which was freshly dissected and labeled with -BTX without
previous fixation, membrane permeabilization, or other staining
procedures. H, Bright-field view of the same region of
the muscle fiber as in G. The sarcomeric pattern also
showed brighter stripes of AChRs (arrowheads in
G) matching with the light bands
(arrowheads in H) of sarcomeres.
The scale bar in H applies to E-H.
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To further test whether PSC sprouts play a role in the aggregation of
extrajunctional AChRs as exemplified by Figure 6, the degree of
association between extrajunctional AChR aggregates and PSC sprouts in
reinnervated muscles was quantified 17-28 d after axotomy
(n = 225 aggregates, 8 muscles). We found that 88% of
extrajunctional AChR aggregates were associated with PSC sprouts, whereas 12% were distributed over the remaining muscle surface. Some
of the latter aggregates also showed the sarcomeric pattern. We showed
previously that Schwann cell sprouts are induced by regenerating nerve
terminals (Koirala et al., 2000). Consistent with this finding, most of
the PSC sprouts associated with extrajunctional AChR aggregates were
also accompanied by regenerating nerve terminal sprouts. The 88% of
PSC-associated aggregates mentioned above included 74% that showed
overlying nerve terminal sprouts and 14% that were devoid of nerve
terminals. To exclude extrajunctional AChR aggregation probably caused
by nerve terminal sprouts, we focused solely on the 14% (32 of 225) of
aggregates that were colocalized with PSC sprouts alone and, on
average, 17 ± 5 µm away from the closest nerve terminals or axons.
To exclude the possibility that the association between PSC sprouts and
extrajunctional AChR aggregates was by chance, a statistical analysis
was performed (see Materials and Methods). If randomly distributed,
only 0.6% of the 58 extrajunctional AChR aggregates not associated
with nerve terminals would colocalize with PSC sprouts alone, and
99.4% would be distributed over the remaining muscle surface
(excluding areas under axons and nerve terminals). However, we found
that 55.2% (32 of 58) of extrajunctional AChR aggregates were
associated with PSC sprouts alone, which is significantly greater than
random association (p < 10
5; 2
test). In addition, PSC-associated extrajunctional AChR aggregates were
significantly larger (p < 0.05; Student's
t test) than randomly distributed aggregates (31 ± 9 vs 14 ± 4 µm in length. Furthermore, we found that 52% of PSC
sprouts without accompanying nerve terminals (n = 56, 8 muscles) showed associated AChR aggregates. Together, these in
vivo results suggest that PSC sprouts may induce AChR aggregation
after nerve injury.
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DISCUSSION |
The present study demonstrates that adult Schwann cells express
active agrin isoforms and that this expression is upregulated by axonal
regeneration. Adult Schwann cells enhance the aggregation of AChRs on
muscle fibers in vitro. Although the evidence is indirect, our results are consistent with the hypothesis that the enhancement of
AChR aggregation by cultured Schwann cells is mediated by agrin. Using
complement-mediated lysis to selectively ablate PSCs at the NMJ
in vivo, we show that these synapse-associated glial cells also express active agrin isoforms. Finally, we present evidence that
PSCs may induce AChR clusters in vivo. These novel findings suggest that, in addition to neurons, glial cells also play a role in
the aggregation of postsynaptic receptors. Receptor aggregation by glia
may complement the role of neurons and may be particularly important in
ensuring rapid restoration of synaptic function during regeneration.
Adult Schwann cells express active agrin isoforms
Werle et al. (1999) have reported that Schwann cells in the frog
sciatic nerve express only the inactive B0 isoform. However, using the
same preparation and the same primers for RT-PCR, we found that frog
Schwann cells do express the active agrin isoforms B11 and B19, besides
the inactive B0. One possible explanation for the conflicting result
may be attributed to different amount of RNA, or different temperature
and number of cycles, used for our RT-PCR. To exclude the possibility
of false-positive bands in our samples, we subcloned each PCR fragment
and confirmed its sequence as agrin. The expression of active agrin
isoforms is not an artifact of contamination by neuronal mRNA that
might be present in axons or nerve terminals, because the sciatic nerve samples used for our RT-PCR study did not contain any neurofilament mRNA. In addition, we showed that Schwann cells cocultured with muscles, but devoid of neurons, also express active agrin isoforms. Furthermore, if our RT-PCR samples were contaminated with neuronal agrin mRNA, we would detect all four agrin isoforms, including B8, as
found in the spinal cord. However, the B8 isoform was not observed in
Schwann cells along normal sciatic nerve, in nerve terminals, or in
cultured Schwann cells. Moreover, B11 and B19 isoforms were also
expressed in the chronically segregated distal segment of the sciatic
nerve. Because the half-life of mRNAs in animal cells is usually <24
hr (Lewin and Siliciano, 1996), the active agrin isoforms in the distal
nerve segment seen even at 6 weeks after long-term denervation must be
actively transcribed, most likely by Schwann cells along the nerve.
Contamination of active agrin from other cell types is also unlikely;
Schwann cells are the predominant cell type along the nerve fiber, and
we found that fibroblasts in the epineurial sheath do not express any
agrin isoforms. Together, these results demonstrate that adult Schwann cells express active agrin isoforms. Anti-agrin staining has also been
observed at frog PSCs (Werle et al., 1999). However, it is not possible
to distinguish between the active and inactive forms of agrin proteins
using the currently available antibodies.
Expression of Schwann cell agrin during development and
axonal regeneration
Similar to embryonic Schwann cells in the chick sciatic nerve,
which express only B0 (Ruegg et al., 1992), developing Schwann cells in
tadpoles do not express active agrin. Although it is not known how the
expression of B11 and B19 is triggered in Schwann cells during
metamorphosis, it is clear that the expression of these active isoforms
occurs long after synaptogenesis. Thus, the expression of B11 and B19
isoforms by Schwann cells during development is not induced by
innervation per se. The absence of active agrin expression in tadpole
Schwann cells also indicates that Schwann cell-derived agrin is
unlikely to play a role in the aggregation of AChRs during initial
synaptogenesis. Consistent with this view, there is no AChR clustering
along Schwann cell sprouts that extend beyond developing nerve
terminals during synapse formation in tadpole muscles (Herrera et al.,
2000). This is in contrast to adult muscles, in which aggregates of
AChRs are formed along PSC sprouts after nerve injury and reinnervation
(see below).
In adult animals, expression of all active agrin isoforms in Schwann
cells was upregulated after nerve injury. However, the upregulation is
not attributable to nerve injury per se because no upregulation occurs
in Schwann cells along chronically severed nerve fibers. Rather, the
upregulation of active agrin isoforms coincides with axonal
regeneration, suggesting that regenerating nerves play a role in the
upregulation. This upregulation of active agrin isoforms is reminiscent
of the induction of PSC sprouting, which also occurs in response to
nerve regeneration, but not nerve injury alone, at the frog NMJ
(Koirala et al., 2000). The mechanism underlying activation of Schwann
cells by regenerating axons and nerve terminals is unknown.
Schwann cells enhance AChR aggregation in vitro
Koenig et al. (1998) suggested that chick Schwann cells have the
capacity to enhance AChR aggregation on cultured myotubes. The present
study has shown that cultured adult Schwann cells express active agrin
mRNAs and produce native agrin protein. In addition, adult Schwann
cells increase the number and area of AChR hotspots on
Xenopus muscle in culture, and the increase in AChR
aggregation is eliminated by heparin. These findings are consistent
with the hypothesis that the enhancement of AChR aggregation by Schwann
cells is mediated by agrin. However, besides disrupting agrin function
(Hopf and Hoch, 1997), heparin has been shown to also interfere with
the function of neuregulin, laminin, and other molecules that affect
AChR synthesis and clustering (Yarden and Wen, 1994). Thus, we cannot
exclude the possibility that other soluble molecules released by
Schwann cells may also play a role in the enhancement of AChR
aggregation. Unfortunately, the lack of antibodies that perturb the
function of Xenopus agrin prevented us from testing the
direct involvement of agrin in SC+M cocultures.
The absence of enhancement of AChR aggregation by Schwann
cell-conditioned medium also precluded our attempt to use
immunoprecipitation to directly test the role of Schwann cell-derived
agrin. In the Schwann cell-conditioned medium, we only detected a 70 kDa band, which likely represents a degradation product of the agrin
protein and is probably not active in AChR aggregation (Nitkin et al., 1987). The absence of protein bands above 200 kDa in the conditioned medium may be attributable to the fact that the conditioned medium was
collected from long-term cultures, and thus most native agrin protein
was degraded. This is in contrast to the increase in AChR aggregation
observed in SC+M cocultures (Fig. 3). In the cocultures, Schwann cells
may continuously release native agrin protein, which presumably would
have an immediate and cumulative effect on AChR aggregation before
being degraded.
Expression of active agrin in the PSC
The present study has applied a novel technique using
complement-mediated lysis to selectively ablate PSCs (Reddy et al., 1999) (a full-length paper will be published in the future). Ablation of PSCs caused a reduction in the amount of active agrin isoforms, indicating that, similar to axonal Schwann cells, PSCs also express active agrin genes. As shown in Astrow et al. (1998), Schwann cells
along axons are not labeled by mAb 2A12 in whole-mount preparations and
thus are not ablated by the complement treatment. Because ~20% of
the total Schwann cells in the CP muscle belong to these axonal Schwann
cells and typically ~20% of PSCs were not ablated by the mAb 2A12
plus complement treatment, these remaining Schwann cells might
contribute to the active agrin isoforms still seen after the ablation
treatment in intact and reinnervated muscles (Table 1). Our approach
using PSC ablation and RT-PCR has provided evidence, albeit indirect,
that is consistent with the idea that PSCs also express active agrin
isoforms and the expression is upregulated by reinnervation.
Extrajunctional AChRs are associated with PSC sprouts
in vivo
The present study has shown that, during muscle reinnervation,
clusters of AChRs form outside of original synaptic sites and that
these clusters colocalize with PSC sprouts. Because the density of
AChRs falls sharply within a few micrometers of nerve terminals at the
NMJ (Matthews-Bellinger and Salpeter, 1978), it is unlikely that
neuronal agrin would have a diffusible effect on the aggregation of
extrajunctional AChRs located tens of micrometers away as observed in
the present study. It is also unlikely that the extrajunctional AChR
aggregation is caused by nerve terminals that extended and then
retracted before our observations. Using repeated observation of
identified NMJs in living frogs, we showed previously extension of PSC
sprouts tens or hundreds of micrometers longer than nerve terminals; in
many cases, PSC sprouts continued to extend even when nerve terminals
showed no change in length during a period of 2-3 months of multiple
observations (Chen et al., 1991; Chen and Ko, 1994; Ko and Chen, 1996).
Thus, frog nerve terminals do not extend and then retract through such
long distances. Furthermore, we showed recently that, at early stages
of reinnervation (2-4 weeks after axotomy), there is substantial
growth of nerve terminals but virtually no observed retraction (Koirala
et al., 2000). Thus, the clusters of extrajunctional AChRs were most
likely caused by PSC sprouts rather than by nerve terminals that
extended and then retracted.
The association of AChR aggregates and PSC sprouts is not random. In
stark contrast to the predicted value of 0.6% if the association were
purely by chance, we showed that 55.2% of extrajunctional AChR
clusters (without accompanying nerve terminals) are located at PSC
sprouts. Furthermore, 52% of PSC sprouts (without accompanying nerve
terminals) showed the presence of colocalized extrajunctional AChRs,
consistent with the suggestion that PSC sprouts play a role in the AChR aggregation.
The mechanism of PSC-induced AChR aggregation in vivo is not
known. Because PSCs express active agrin, it is tempting to speculate that the aggregation of extrajunctional AChRs is mediated by agrin released by PSCs. The colocalization between AChR clusters and PSC
sprouts in vivo would suggest that agrin molecules, if
released by PSC sprouts, probably bind basal lamina and are
concentrated locally, as is the case for neuronal agrin at the
NMJ. In contrast to the in vivo finding, Schwann
cell-enhanced AChR aggregation in vitro is independent of
cell contact, which may be attributed to the paucity of basal lamina on
embryonic muscle fibers in culture.
As in the present study, discrete patches of extrajunctional AChRs with
different shape and lower density than the junctional AChRs have been
reported in denervated mammalian muscles (Ko et al., 1977). Clusters of
AChR with various densities have also been observed in muscles treated
with agrin in vitro (Wallace, 1992) or in vivo
(Bezakova et al., 2001). In fact, the sarcomeric pattern can also be
discerned in some extrajunctional AChR clusters induced by agrin
application to rat muscle (Bezakova et al., 2001, their Fig. 4).
Because the sarcomeric pattern of AChR staining matches with the light
bands of the sarcomere, it is likely that AChRs are located in and
around the T tubules, which have membrane invaginations near the Z-line
in the light bands. Sheikh et al. (2001) have shown that sodium
channels cluster selectively around the mouths of the T tubules in the
frog skeletal muscle fiber. It is possible that AChRs also cluster only
around the mouths of the T tubules. Alternatively, AChRs may be evenly
distributed along the entire length of the T tubules, but their
invaginations give rise to a brighter signal of AChR staining in the
en face view.
The novel role of glial cells in receptor aggregation described in this
study may be important in laying the groundwork for rapid and
successful restoration of synaptic function during regeneration. In
addition to the presynaptic guidance of regenerating nerve terminals as
shown previously (for review, see Son et al., 1996), PSCs now appear to
play a postsynaptic role in inducing AChR aggregation ahead of
regenerating terminals. There is indirect evidence that PSC-induced
AChR aggregates may constitute sites of subsequent synaptogenesis. PSCs
sprout soon after regenerating nerve terminals arrive at endplates, and
nerve terminals grow along PSC sprouts (Koirala et al., 2000). Because
AChR aggregates are present along PSC sprouts ahead of nerve terminals,
these aggregates would very likely be "innervated" by the
regenerating nerve terminals. Consistent with this scenario, we observe
that, at later stages of reinnervation, all extrajunctional AChR
aggregates colocalize with overlying nerve terminals (Koirala et al.,
2000). Apart from a preparatory role in reinnervation, our results
suggest that PSCs could also play a role in the maintenance of AChRs at
the NMJ. Studies are underway using our novel PSC ablation technique to
determine whether PSCs play a role in the maintenance of AChR
clustering at NMJs, particularly after nerve injury.
| |
FOOTNOTES |
Received Aug. 7, 2001; revised Sept. 19, 2001; accepted Oct. 1, 2001.
This work was supported by National Institutes of Health Grant NS17954
and a Muscular Dystrophy Association research grant. We thank Dr. Earl
W. Godfrey of Eastern Virginia Medical School for the generous gift of
anti-agrin C3 monoclonal antibody and Dr. Bai Lu of National Institutes
of Health for the generous gift of the anti-synapsin I antibody. We are
grateful to Drs. John H. Caldwell, Earl W. Godfrey, and Karl W. K. Tsim for their critical comments. We also thank H. Qiang, T. Ma, Z. Feng, and C. David for their expert technical support.
Correspondence should be addressed to Dr. Chien-Ping Ko, Section of
Neurobiology, Department of Biological Sciences, University of Southern
California, Los Angeles, CA 90089-2520. E-mail: cko{at}mizar.usc.edu
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ABSTRACT
INTRODUCTION
