|
| |
|
|
|
|
||||
| Home Help Feedback Subscriptions Archive Search Table of Contents | ||||
First published online February 20, 2004
Journal of Experimental Biology 207, 1163-1182 (2004)
Published by The Company of Biologists 2004
doi: 10.1242/jeb.00856
The anterior cardiac plexus: an intrinsic neurosecretory site within the stomatogastric nervous system of the crab Cancer productus
1 Department of Biology, University of Washington, Box 351800, Seattle,
Washington 98195-1800 USA
2 Friday Harbor Laboratories, University of Washington, 620 University Road,
Friday Harbor, Washington 98250 USA
* Author for correspondence at address 1 (e-mail: crabman{at}u.washington.edu)
Accepted 5 January 2004
 |
Summary |
|---|
 TOP Summary IntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Key words: incident light microscopy, transmission electron microscopy, laser scanning confocal microscopy, FLRFamide-related peptide, neurohormone, neuromodulation, crab, Cancer productus
|
Introduction |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
;
Passano, 1951). While many
definitions of a neuroendocrine site exist, in the broadest of terms, it is
generally accepted to be a region in which nerve terminals come in contact
with the circulatory system (Cooke and
Sullivan, 1982). Given this definition, many regions of the
crustacean nervous system are likely to contribute to the hormone complement
present in the hemolymph. Most crustacean neuroendocrine sites appear as
rather unorganized regions of secretory terminals within or just below the
sheath of nerves and ganglia (Friend,
1976; King, 1976;
Sullivan et al., 1977;
Livingstone et al., 1981;
Kobierski et al., 1987;
Kilman and Marder, 1996;
Kilman, 1998;
Skiebe et al., 1999;
Skiebe and Ganeshina, 2000).
Others, however, are more formally delimited, giving rise to what are often
termed neurosecretory organs (Bliss,
1951; Passano,
1951; Alexandrowicz and
Carlisle, 1953; Carlisle and
Knowles, 1959; Maynard,
1961a,b;
Maynard and Maynard, 1962;
Cooke and Sullivan, 1982).
Small molecule transmitters, biogenic amines, peptides, and diffusible
gases have been identified in crustacean neuroendocrine sites
(Evans et al., 1976;
Beltz and Kravitz, 1983;
Schwarz et al., 1984; Siwicki
et al., 1985,
1987;
Siwicki and Bishop, 1986;
Stangier et al., 1986,
1988;
Kobierski et al., 1987;
Dircksen, 1992;
Rodriguez-Sosa et al., 1994;
Christie et al., 1995,
2003;
Wood et al., 1996;
Chang et al., 1999; Skiebe et
al., 1999,
2002;
Yang et al., 1999;
Lee et al., 2000;
Wang et al., 2000). These
bioactive agents affect many target tissues and have been shown to directly
control or influence a variety of physiological processes such as molting,
metamorphosis, color change and the regulation of hemolymph glucose levels
(Huberman, 1990;
Keller, 1992;
Rao, 1992;
Rao and Riehm, 1993;
Chang, 1993;
Wainwright et al., 1996;
Fingerman, 1997;
Fingerman et al., 1998;
Soyez, 1997;
Chung et al., 1999;
Phlippen et al., 2000).
Circulating hormones have also been implicated in numerous aspects of nervous
system function, including modulation of the neural circuits and muscles
involved in feeding-related behavior (Turrigiano and Selverston,
1989,
1990;
Heinzel et al., 1993; Marder
et al., 1994,
1995; Christie and Nusbaum,
1995,
1998; Jorge-Rivera and Marder,
1996,
1997;
Jorge-Rivera, 1997;
Jorge-Rivera et al., 1998;
Weimann et al., 1997).
In crustaceans, the stomatogastric nervous system (STNS;
Fig. 1), an extension of the
central nervous system, controls the ingestion and movement of food through
the foregut (Selverston and Moulins,
1987; Harris-Warrick et al.,
1992). Four ganglia are contained within the STNS: the
stomatogastric ganglion (STG), the oesophageal ganglion (OG) and the paired
commissural ganglia (CoGs). A number of nerves connect these ganglia and/or
innervate the muscles of the foregut. Several distinct, but interacting,
neural circuits are present within the STNS, including one contained within
the STG that produces both the pyloric and gastric mill rhythms
(Selverston and Moulins, 1987;
Harris-Warrick et al., 1992).
Multiple forms of the pyloric and gastric mill motor patterns have been shown
to exist (Selverston and Moulins,
1987; Harris-Warrick et al.,
1992; Marder et al.,
1994,
1995,
1997;
Marder and Calabrese, 1996;
Skiebe, 2001). Research from
many laboratories has shown that much of this flexibility is imparted through
the actions of neuromodulators, including circulating hormones present in the
hemolymph (Turrigiano and Selverston,
1990; Heinzel et al.,
1993; Christie and Nusbaum,
1995,
1998;
Weimann et al., 1997).
|
In many genera, particularly those of the infraorder Astacidea (chelate
lobsters and freshwater crayfish), hormones released from both extrinsic and
intrinsic sites are likely to influence the STNS. The extrinsic sites include
some or all of the classically defined neuroendocrine organs, i.e. the sinus
glands (SGs) of the eyestalk, the pericardial organs (POs) present in the
venous cavity surrounding the heart, and the post-commissural organs (PCOs)
located within the posterior commissure near the oesophagus
(Cooke and Sullivan, 1982), as
well as sites located on the second roots of the thoracic ganglia and on the
ventral nerve cord (Livingstone et al.,
1981; Kobierski et al.,
1987).
Several neuroendocrine release zones within the STNS itself may also
contribute to its hormonal modulation in lobsters and crayfish. One such site
is an extensive plexus located within the sheath of the anterior portion of
the STNS (Maynard and Dando,
1974; Kilman,
1998; Skiebe et al.,
1999; Skiebe and
Wollenschläger, 2002;
Christie et al., 2003). While
the extent of this structure remains undetermined in most species, in the
American lobster Homarus americanus, and the Australian freshwater
crayfish Cherax destructor and Cherax quadricarinatus, it is
known to span the anterior portion of the stomatogastric nerve (stn)
and all or a portion of the superior oesophageal (son), oesophageal,
dorsal posterior oesophageal, inferior oesophageal and inferior ventricular
nerves (Skiebe and Wollenschläger,
2002; Christie et al.,
2003). In these species, transmission electron microscopy confirms
the ultrastructure of the site to be neuroendocrine in nature
(Kilman, 1998;
Skiebe and Ganeshina, 2000;
Christie et al., 2003). A
second intrinsic neuroendocrine site is present on each circumoesophageal
connective (coc) near the CoG
(Skiebe et al., 1999). Like
the plexus in the anterior portion of the STNS, this site is superficially
located and has been shown to possess an ultrastructure consistent with a
neuroendocrine release zone (Skiebe et
al., 1999). Thus far this plexus has been identified in only one
species, C. destructor (Skiebe et
al., 1999). Additionally, in several species, including H.
americanus and the California spiny lobster Panulirus
interruptus (Infraorder Palinura), neuroendocrine profiles have been
identified ultrastructurally in the sheath surrounding the STG and the nerves
immediately adjacent to it, i.e. the stn and the dorsal ventricular
nerve (Friend, 1976;
King, 1976). These sites too
may contribute to the hormonal control of the STNS.
As in lobsters and crayfish, the STNS of Brachyuran crabs is also modulated
by hormones released from extrinsic neuroendocrine sites (Christie and
Nusbaum, 1995,
1998;
Weimann et al., 1997). Unlike
the former groups, little is known about intrinsic neurosecretory zones in the
STNS of these animals. Using incident light microscopy, Maynard and Dando
(1974) identified an
iridescent region on each of the paired anterior cardiac nerves
(acns) in the blue crab Callinectes sapidus. They
interpreted this iridescence as indicative of a neurohemal release site.
Similarly, Skiebe and Wollenschläger
(2002), using antibodies to
vesicle-associated proteins, identified putative neuroendocrine sites on the
acns of the European edible crab Cancer pagurus. In neither
species is information available to confirm that the acn plexi are
ultrastructurally identifiable as neuroendocrine release zones.
In the present study, we used incident light microscopy to survey the STNS
of the red rock crab Cancer productus for putative neuroendocrine
sites. As with C. sapidus and C. pagurus
(Maynard and Dando, 1974;
Skiebe and Wollenschläger,
2002), the only putative neurosecretory sites identified were on
the acns. Using transmission electron microscopy, we confirmed that
these sites, which we have named the anterior cardiac plexi or ACPs, are
ultrastructurally identifiable as neuroendocrine release zones. All
innervation to the ACPs originates from four axons that project to the plexi
via the stn and sons. Modulator immunolabeling
shows that all four of the axons innervating the ACPs exhibit
FLRFamide-related peptide immunoreactivity. Our confirmation of the ACPs of
C. productus as neuroendocrine plexi shows that intrinsic
neurosecretory sites exist in the STNS of this species. This finding
strengthens the hypothesis that the acn sites previously identified
in C. sapidus (Maynard and Dando,
1974) and C. pagurus
(Skiebe and Wollenschläger,
2002) are also neuroendocrine in nature. Moreover, our results set
the stage for future biochemical and physiological studies of the ACPs, the
hormones contained within them and their actions on potential target tissues.
Some of this data has appeared previously in abstract form
(Christie et al., 2002).
|
Materials and methods |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
For tissue collection, crabs were anesthetized by packing in ice for 30–60 min, their foregut removed, and the STNS (Fig. 1) dissected from the foregut in chilled (approximately 4°C) physiological saline (440 mmol l–1 NaCl; 11 mmol l–1 KCl; 13 mmol l–1 CaCl2; 26 mmol l–1 MgCl2; 10 mmol l–1 Hepes acid, pH 7.4, adjusted with NaOH). Following dissection, tissue was pinned in a Sylgard 184 (KR Anderson, Santa Clara, California, USA)-lined Petri dish and subsequently processed as described below.
Incident light microscopy of living tissue
To examine fresh, unfixed tissue, the STNS was dissected as described
above, then pinned flat in a Sylgard-lined Petri dish containing chilled
(approximately 10°C) physiological saline. Preparations were viewed, and
maps of the putative location of each ACP drawn, using either a Wild M5A
(Heerbrugg, Switzerland) or Nikon SMZ1000 (Tokyo, Japan) stereomicroscope with
illumination provided by a Fiber-Lite Model 190 fiber optic illuminator
(Dolan-Jenner Industries, Inc., Woburn, Massachusetts, USA). Each preparation
was examined at multiple magnifications. The illuminating beam was adjusted
several times at each magnification so as to allow the tissue to be examined
at multiple illuminating angles. Incident light micrographs were taken using a
CoolSNAP camera system (Roper Scientific, Inc., Tucson, Arizona, USA) mounted
on the Nikon microscope.
Light and transmission electron microscopy
For light level and transmission electron microscopy, methods modified from
standard techniques were used (Dircksen,
1992; Kilman and Marder,
1996; Kilman,
1998; Webster et al.,
2000). Specifically, ACPs were identified using incident light
microscopy (see above) and subsequently isolated. These stretches of the
acn were fixed using one of two protocols. In the first protocol,
acns were fixed for 1–2 h at 4°C in freshly prepared
Karnovsky's fixative [2.5% glutaraldehyde (EM grade; Electron Microscopy
Sciences, Fort Washington, Pennsylvania, USA), 2% paraformaldehyde (EM grade;
Electron Microscopy Sciences), 0.1% CaCl2 and 5% sucrose in 0.2 mol
l–1 sodium cacodylate buffer, pH 7.2], rinsed twice (at 15
min intervals) in sodium cacodylate buffer and then post-fixed for 1 h with 1%
OsO4 in sodium cacodylate buffer. Following post-fixation, tissue
was rinsed twice in sodium cacodylate buffer (at 15 min) and subsequently
dehydrated in a graded ethanol series (40%, 60%, 80%, 95% and 100%).
Dehydrated tissue was passed through an ethanol/LX-112 epoxy resin (Ladd
Research Industries, Williston, Vermont, USA) series (3:1, 1:1, 1:3; 90 min
each) and then left in 100% LX-112 overnight. After this overnight
infiltration, tissue was transferred to embedding molds filled with fresh
LX-112 and polymerized at 60°C for 8 h.
In the second protocol, acns were fixed overnight in 4% paraformaldehyde, 0.1% glutaraldehyde in 0.1 mol l–1 sodium phosphate (P) buffer (pH 7.4) at 4°C. Following fixation, tissue was rinsed twice in P buffer (at 10 min intervals) and subsequently post-fixed in 1% OsO4 in 0.01 mol l–1 sodium phosphate buffer, pH 7.4. Following post-fixation, tissue was rinsed twice in distilled water (at 10 min intervals) and then dehydrated in graded ethanol series (see above). Some tissue fixed via this second protocol was embedded in LRWhite (LRW) resin (Electron Microscopy Sciences). Here, tissue was taken from ethanol into 100% activated LRW for 2 h. Tissue was subsequently transferred to fresh LRW overnight and then transferred again to fresh LRW for 2 h. Tissue was then embedded in fresh LRW using #1 gelatin capsules (Ted Pella, Redding, California, USA) and polymerized at 45°C for 48 h. The remaining acns fixed via the second protocol were taken from ethanol into a 1:1 mix of ethanol:propylene oxide for 30 min. Tissue was then rinsed twice in propylene oxide (at 15 min intervals), transferred into a 1:1 mix of propylene oxide:100% EMBed (EMB) resin (Electron Microscopy Sciences) for 2 h and then into fresh EMB for 2 h. After 2 h, the resin was replaced with fresh EMB and allowed to infiltrate overnight. After this overnight infiltration, tissue was again transferred to fresh EMB for 2 h and subsequently polymerized in fresh EMB for 48 h at 45°C.
Regardless of the resin used, polymerized blocks were sectioned for light
microscopy at 1.0 µm using glass knives and for transmission electron
microscopy at 70–90 nm with a diamond knife (Diatome, Fort Washington,
Pennsylvania, USA). All sectioning was done using a RMC MT6000 ultramicrotome
(Research and Manufacturing Company Inc., Tucson, Arizona, USA). Sections used
for light microscopic analysis were mounted on glass microscope slides and
subsequently stained with 1% Toluidine Blue, 1% borax in distilled water for
60 s at 60°C. Micrographs were taken with a Nikon CoolPix 4500 digital
camera mounted on a Nikon Eclipse E800 microscope using a PlanFluor 40x
1.35NA oil immersion lens. For transmission electron microscopy, sections were
mounted on copper mesh grids and stained with 4% uranyl acetate and Reynolds'
lead citrate (Reynolds, 1963)
for 1 h and 30 s, respectively. Tissue was examined and micrographs generated
using a Philips CM100 transmission electron microscope (Philips Electronic
Instrument Company, Mahwah, New Jersey, USA) at 60 kV.
The gross ultrastructure of the acn was the same regardless of the tissue processing used.
Wholemount immunocytochemistry
For wholemount immunocytochemistry, tissue was fixed overnight in freshly
made 4% paraformaldehyde in P buffer (pH 7.4; see above for composition).
Fixed tissue was rinsed five times over approximately 5 h in a solution of P
buffer containing 0.3% Triton X-100 (P-Triton). Incubation in primary
antibody (or antibodies) was done in P-Triton, with 10% normal goat serum
(NGS) added to diminish nonspecific binding. Following incubation in primary
antibody, tissues were again rinsed five times over approximately 5 h in
P-Triton and then incubated in secondary antibody (or antibodies). As with the
primary antibody, secondary antibody incubation was done in P-Triton with 10%
NGS. After secondary antibody incubation, each preparation was rinsed five
times over approximately 5 h in P buffer and then mounted between a glass
microscope slide and coverslip using either a solution of 80% glycerine, 4%
n-propyl gallate (pH 
9.0) or Vectashield mounting medium (Vector
Laboratories, Inc., Burlingame, California, USA). Fixation and incubation in
both primary and secondary antibody were done at 4°C. Incubation in both
primary and secondary antibody was done using gentle agitation. All rinses
were done at room temperature without agitation. Secondary antibody incubation
and all subsequent processing was conducted in the dark. Likewise, slides were
stored in the dark at 4°C until examined.
Antibodies
To determine whether FLRFamide-related peptides were present in the ACPs,
we used a rabbit polyclonal antibody generated against FMRFamide (catalog
#20091; Immunostar Inc., Hudson, Wisconsin, USA). This antibody was chosen as
it has been used previously for mapping the distribution of this peptide
family in several crustacean species (Schmidt and Ache,
1994a,b;
Tierney et al., 1997;
Fénelon et al., 1998;
Blitz et al., 1999;
Kilman et al., 1999;
Meyrand et al., 2000). As all
known crustacean FMRFamide-related peptides contain the carboxy-terminal amino
acid sequence FLRF rather than FMRF
(Trimmer et al., 1987;
Krajniak, 1991;
Mercier et al., 1993;
Weimann et al., 1993), in this
paper we will refer to the FMRF antibody and its immunolabeling as FLRFamide
antibody and immunolabeling, respectively. In our study the FLRFamide antibody
was used at a final dilution of 1:300 with an incubation time of 48–72
h.
As a general marker for electron-lucent vesicles (ELVs), an antibody
generated against the synaptic vesicle-associated protein synapsin was used.
This antibody is a mouse monoclonal antibody generated against a glutathione
S-transferase fusion protein, which includes a portion of a
Drosophila synapsin homolog (SYNORF1;
Klagges et al., 1996; provided
for this study by E. Buchner). This antibody has been used previously to
identify the location of putative synaptic neuropil and neurosecretory sites
in several crustaceans (Skiebe,
2000; Skiebe and Ganeshina,
2000; Skiebe and
Wollenschläger, 2002). In our study, the synapsin antibody
was used at a final dilution of 1:20 with an incubation time of approximately
72 h.
The secondary antibodies used in our study were goat anti-rabbit IgG labeled with Texas Red (catalog #111-075-144; Jackson ImmunoResearch Corporation, West Grove, Pennsylvania, USA) and goat anti-mouse IgG labeled with FITC (catalog #115-095-146; Jackson ImmunoResearch Corporation). Both antibodies were used at final dilutions of 1:300 with incubation times of 12–24 h.
Preadsorption controls
To strengthen our confidence that the FLRFamide immunoreactivity seen in
the ACP was due to the presence of FLRFamide-related peptides, we conducted a
series of preadsorption controls. In these experiments, TNRNFLRFamide
(American Peptide Company, Sunnyvale, California, USA), SDRNFLRFamide
(American Peptide Company), Cancer borealis tachykinin
related-peptide Ia (APSGFLGMRamide; synthesized by the Protein Chemistry
Laboratory, University of Pennsylvania School of Medicine, University of
Pennsylvania, Philadelphia, Pennsylvania, USA, and provided for this study by
M. Nusbaum), or proctolin (RYLPT; Peninsula Laboratories, Belmont, California,
USA) were used as blocking agents. In each experiment, the FLRFamide antibody
was incubated with a blocking agent for 2 h at room temperature prior to
applying the solution to the tissue. Immunoprocessing was then performed as
described above. TNRNFLRFamide and SDRNFLRFamide were chosen for these
experiments as they are the only FLRFamide-related peptides thus far isolated
from crabs of the genus Cancer
(Weimann et al., 1993).
Cancer borealis tachykinin related-peptide Ia and proctolin were
chosen as they too are known to be present in species of the genus
Cancer in their native form
(Marder et al., 1986;
Christie et al., 1997b).
Lucifer Yellow nerve backfilling
In some experiments, one acn was backfilled with Lucifer Yellow-CH
dilithium salt (LY; Sigma; Saint Louis, Missouri, USA or Molecular Probes,
Eugene, Oregon, USA) prior to fixation and FLRFamide immunoprocessing. In
these experiments, a Vaseline well was built around the acn and the
saline within the well subsequently replaced with distilled water. After
several minutes, the distilled water was removed and replaced with a solution
of 10–20% LY in distilled water and the nerve was transected within the
well. Following transection, the preparation was incubated at 10°C for
18–72 h in the dark. Dye was subsequently removed from the well and the
preparation fixed and immunolabeled as described above. All immunoprocessing
of LY backfilled preparations was done in the dark.
Confocal and epifluorescense microscopy
All fluorescent preparations, regardless of the type of processing, were
viewed and data collected using one of two Bio-Rad MRC 600 laser scanning
confocal microscopes (Bio-Rad Microscience Ltd., Hemel Hempstead, UK), a
Bio-Rad Radiance 2000 laser scanning confocal microscope or a Nikon Eclipse
E600 epifluorescense microscope. The Bio-Rad MRC 600 system located at the
University of Washington (Department of Biology) is equipped with a Nikon
Optiphot upright microscope and a krypton/argon mixed gas laser. Nikon Fluor
10x 0.5NA dry, PlanApo 20x 0.75NA dry and PlanApo 60x 1.4NA
oil immersion lenses were used for imaging. Bio-Rad supplied YHS or K1/K2
filter sets and Comos software were used for imaging all preparations on this
system (filter specifications are as described in
Christie et al., 1997a). The
Bio-Rad MRC 600 system located at Friday Harbor Laboratories is equipped with
a Nikon Optiphot inverted microscope and uses the same laser, filters and
software as the MRC 600 system located at the University of Washington. With
the addition of a Nikon Fluor 40x 0.85NA dry lens, the objective lenses
were also the same as those located on the system at the University of
Washington. The Bio-Rad Radiance 2000 laser scanning confocal microscope is
equipped with a modified Nikon Eclipse E600FN microscope and a krypton/argon
mixed gas laser. Nikon PlanApo 10x 0.45NA DIC dry, PlanApo 20x
0.75NA DIC dry and PlanApo 60x 1.4NA DIC oil immersion lenses were used
for imaging. Bio-Rad supplied LaserSharp 2000 software and 560 DCLP dichroic
and HQ 515/30 and E600LP emission filters were used for imaging tissue on this
system. The Nikon Eclipse E600 epifluorescense microscope is equipped with
Nikon PlanFluor 10x 0.30NA dry, PlanFluor 20x 0.50NA dry and
PlanFluor 40x 0.75NA dry lenses and B-2E/C FITC (EX, 465–495 nm;
DM, 505 nm; BA, 515–555 nm) and G-2E/C TRITC (EX, 528–553 nm; DM,
565 nm; BA, 600–660 mn) filter sets.
Figures were produced using a combination of Photoshop (version 7.0; Adobe Systems Inc., San Jose, California, USA) and Canvas (version 8.0; Deneba Systems Inc., Miami, Florida, USA) software. Contrast and brightness were adjusted as needed to optimize the clarity of the printed figures.
|
Results |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
).
This visual effect is attributed to the high density of
peptide/amine-containing neurosecretory granules contained within these
structures (Maynard and Maynard,
1962). To survey the STNS of C. productus for putative
neurosecretory sites, we conducted an incident light examination of the entire
STNS of this species. This included all ganglia, their interconnecting nerves
and the major motor nerves of this system
(Fig. 1). In each of the
preparations examined (N=47 preparations), we observed a well
delimited area of punctate, bluish-white iridescence on each of the paired
acns (Fig. 2A). On
each acn, the iridescent region started close to the junction of this
nerve and the stn and extended distally for up to several mm. The
iridescent profiles present at this site appeared superficially located,
within or just below the sheath of the nerve. In no other regions of the STNS
did we consistently observe any distinct areas of iridescence within the
sheath (Fig. 2B).
|
Light and transmission electron microscopy
To characterize the ultrastructure of the iridescent structures just
described, we isolated several pieces of the acn (N=9
acns) and subjected them to light and transmission electron
microscopy. In each of these preparations, the iridescent region of the
acn could be roughly divided into two parts: the central core and the
peripheral zone. The central core of the acn was characterized by the
presence of large (approximately 10 µm) diameter axons
(Fig. 3A–C). All axons in
the central core contained filamentous axoplasm and were tightly ensheathed by
a thick wrapping of glia (Fig.
3D). Dense-core vesicles (DCVs) and ELVs were occasionally evident
in these axons. At the end of the acn closest to the stn,
five large diameter axons were present
(Fig. 3A). Within the center of
each acn segment, smaller diameter neurites emanated from the large
fibers just described, radiating toward the periphery of the nerve
(Fig. 3B). These neurites also
showed glial wrapping and contained filamentous axoplasm as well as DCVs and,
occasionally, ELVs. At the distal end of each acn segment, only one
large diameter fiber was seen, suggesting that the other four axons terminate
in the iridescent portion of the acn
(Fig. 3C).
|
|
|
The nerve terminals present at the periphery of the acn ranged in
size from <1 µm to approximately 10 µm in major cross-sectional
diameter and contained numerous DCVs as well as mitochondria and,
occasionally, ELVs (Figs 4,
5). In no terminals were any
conventional synapses seen. In many terminals, however, morphological
correlates of hormone secretion were evident (Figs
4B,
5). These ultrastructural
features included DCVs docked to the plasma membrane and omega
(
)-figures (Fig. 5).
Based on their location and ultrastructure, we have named the above described neurosecretory region of each acn the anterior cardiac plexus or ACP.
FLRFamide labeling and sources of innervation of the anterior cardiac plexi
FLRFamide labeling
FLRFamide-related peptides have been shown to be present in many crustacean
neuroendocrine sites (Kobierski et al.,
1987; Krajniak,
1991; Mercier et al.,
1993; Christie et al.,
1995; Kilman,
1998; Skiebe et al.,
1999; Pulver and Marder,
2002). To determine whether the ACP contains FLRFamide-related
peptides, we conducted wholemount immunolabeling of the acns
(N=62 acns). In each acn, intense FLRFamide
immunoreactivity was seen in the portion of the nerve that contained the ACP.
This label consisted both of immunopositive fibers and varicosities (Figs
6,7,8,9).
In the region of the acn closest to stn, four
FLRFamide-immunopositive fibers were present. After travelling distally for
several hundred µm, these axons arborized, producing a large number of
smaller diameter processes which radiated from the central core of the nerve,
giving rise to a dense plexus of fine neurites studded with varicosities in
the periphery (Figs 6,
7). The immunopositive
varicosities varied widely both in shape and size
(Fig. 7). These varicosities
showed pronounced clustering in the perineural sheath region, which gives rise
to a bark-like appearance of the immunolabel, particularly in the portion of
the ACP closest to the stn (Fig.
8). Blister-like protuberances of the acn sheath were
often seen in this region of the ACP. These protrusions commonly contained
large numbers of tightly clustered FLRFamide-immunopositive varicosities
(Fig. 9). No FLRFamide-like
immunoreactivity was seen in any of the acns past the plexi, which
suggests that the four FLRFamide immunolabeled acn fibers terminate
in the ACPs (Figs 6,
7).
|
|
|
|
Preadsorption controls
As our study is the first using the Immunostar FLRFamide antibody in C.
productus, and the first time that any modulator staining has been
reported in the ACP, we conducted experiments to confirm that the staining we
report is suppressed specifically by preadsorption of the antibody with
extended FLRFamide peptides. Incubation of the FLRFamide antibody with
TNRNFLRFamide or SDRNFLRFamide (10–6 mol
l–1) completely abolished immunolabeling in the ACPs
(N=6 ACPs for each peptide; data not shown). Staining in these
structures after the FLRFamide antibody had been preincubated with either
Cancer borealis tachykinin-related peptide Ia or proctolin
(10–3 mol l–1; N=6 ACPs for each
peptide) was no different from antibody preincubations at room temperature
with no blocking agent present (N=6 ACPs; data not shown).
Sources of FLRFamide innervation to the anterior cardiac plexi
As just described, four axons are the sole source of FLRFamide innervation
to each ACP. In several preparations, including the one shown in
Fig. 6A, the projection pathway
of these fibers could be traced with relatively little ambiguity using the
immunolabeling alone. In these preparations, two large diameter immunopositive
axons in each son could be seen to project into the stn (for
a total of four large diameter, FLRFamide-immunopositive axons in this nerve;
note that a number of small FLRFamide-immunopositive axons were also present
in the stn) and bifurcated just anterior to the insertion point of
the acns. At the acns, one branch derived from the
bifurcation of each axon entered the right acn with the other branch
of each axon entering the left acn. The result of this branching
pattern is the four axons seen per acn
(Fig. 6B).
To confirm the projection pathway of the axons just described, and to attempt to locate the somata of these axons, we conducted a series of experiments in which we backfilled one acn with LY and subsequently processed the nerve backfill for FLRFamide immunolabeling (N=5 preparations). In these experiments, the backfill site on the acn was just distal to the insertion point of this nerve into the stn. This location ensured the presence of the FLRFamide-immunopositive axons at the site of nerve transection.
In all backfilled acns, LY-filling was evident in five axons. Four of the five axons were double-labeled by the FLRFamide antibody and could be seen to project into the contralateral acn as well as into the stn, traveling toward the anterior ganglia (data not shown). At the junction of the stn and sons, two of the four double-labeled axons entered the right son with the remaining pair entering the left son (Fig. 10A). Thus, these backfills do confirm the projection pathway of the FLRFamide-containing fibers described above. It should be noted that additional single-labeled FLRFamide immunopostive axons were also present in each son (Fig. 10A). These axons too appeared to project into the stn. Unlike the double-labeled axons, these FLRFamide immunolabeled fibers bypassed the acns and projected into and innervated the neuropil of the STG (data not shown).
|
The source of most somata that project from the anterior portion of the
STNS to the posterior nerves and STG is the paired CoGs
(Coleman et al., 1992). As the
CoGs contain numerous FLRFamide-immunopositive cell bodies (approximately 40
somata per ganglion; data not shown), we felt confident that the somata of the
FLRFamide axons innervating the ACPs would be found to reside here. While LY
routinely reached the CoGs (N=8 of 10 CoGs; data not shown), in no
preparation were we able to dye-fill any somata in these ganglia.
Interestingly, in one CoG, two very weakly dye-filled,
FLRFamide-immunopositive axons could be seen to project out of the neuropil
toward the coc, which connects the STNS to both the supraoesophageal
and the fused thoracic ganglia (data not shown). The direction in which these
axons projected within the coc could not be determined in this
ganglion.
The LY-filled acn axon not labeled by the FLRFamide antibody also
projected into both the contralateral acn and the stn, but
here the fiber traveled posteriorly, toward the STG
(Fig. 10B). Though
unconfirmed, it appears likely that this fiber is the axon of the anterior
median (AM) neuron, a neuron whose soma is located within the STG and is known
in numerous decapods to project axons through both acns to innervate
the muscles of the cardiac sac region of the foregut
(Maynard and Dando, 1974;
Selverston and Moulins, 1987;
Harris-Warrick et al.,
1992).
The FLRFamide-immunopositive axons appear to be the sole source of innervation to the anterior cardiac plexi
As described above, four axons projecting via the sons
and stn provide all of the FLRFamide innervation to the ACPs.
Moreover, our ultrastructural and LY backfill data suggest that only one other
neuron, likely to be the AM neuron, projects through acns, which
contain the ACPs. To assess whether the putative AM neuron contributes to the
innervation of the ACPs, we conducted double-immunolabels of these structures
pairing the FLRFamide antibody with an antibody generated against the
vesicle-associated protein synapsin. Synapsins are expressed on ELVs and as
the AM neuron is known to contain glutamate (contained in ELVs; Selverston and
Moulins, 1986), this antibody should be a general marker for its terminals.
Since the AM neuron is not FLRFamide immunopositive (no intrinsic STG somata
show evidence of FLRFamide labeling in C. productus; data not shown),
we would expect that if it contributes to the innervation of the ACPs,
synapsin-positive, FLRFamide-negative terminals should be present in the
double-labeled preparations.
In all double-labeled preparations (N=18 ACPs), we found that both the FLRFamide and synapsin immunolabels are coincident in the ACP (Fig. 11). In each ACP, most FLRFamide-immunopositive varicosities were found to exhibit some degree of synapsin labeling. In no ACP did we observe any terminals that contained only synapsin immunoreactivity. Thus, all inputs to the ACPs can be accounted for by the four FLRFamide-immunopositive input axons.
|
|
Discussion |
|---|
|
TOPSummaryIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
;
Fingerman and Aoto, 1959;
Knowles, 1965;
Bunt and Ashby, 1967;
Andrews et al., 1971;
Smith, 1974;
Silverthorn, 1975;
Andrews and Shivers, 1976;
Strolenberg et al., 1977;
Nordman and Morris, 1980; Weatherby,
1981; Kobierski et al.,
1987; Dircksen,
1992; Skiebe et al.,
1999; Skiebe and Ganeshina, 2002;
Christie et al., 2003). All
appear to be separated from the hemolymph space by only an epineurium, often
termed the basement membrane or basal lamina. This structure is acellular and
appears to be composed of an amorphous, collagen-like material, which is often
fenestrated with minute open spaces. Nerve terminals, often in aggregates, lie
beneath the epineurium, as do glia and other support cells. The nerve
terminals are often densely packed with large, membrane-bound vesicles of
varying electron density and, in some neurosecretory sites, small ELVs are
also present. The ACPs of C. productus exhibit all of these
ultrastructural characteristics; compare
Fig. 4 of this study and fig. 1
of Strolenberg et al. (1977),
fig. 5 of Weatherby (1981),
fig. 8 of Kobierski et al.
(1987), fig. 2 of Dircksen
(1992) or fig. 10 of Skiebe et
al. (1999). As we have shown
in this study, each ACP consists of nerve terminals present directly under a
well-defined epineurium. The terminals are densely packed with DCVs and also
contain a small population of ELVs. Thus, based on ultrastructural morphology,
we believe the ACPs are neurosecretory in nature.
In addition to a conserved ultrastructure, many anatomical investigations
of crustacean neurosecretory sites show morphological evidence of the
exocytosis of DCVs (Bunt and Ashby,
1967; Smith, 1974;
Silverthorn, 1975;
Andrews and Shivers, 1976;
Nordmann and Morris, 1980;
Weatherby, 1981;
Dircksen, 1992;
Christie et al., 2003). As
defined by Normann (1969,
1976), an exocytotic event can
be seen via transmission electron microscopy to consist of several
steps: (1) a slight invagination of the plasma membrane of a nerve terminal
towards a DCV, (2) direct contact, or docking, of the DCV membrane with the
plasma membrane of the terminal, (3) fusion of the terminal and the DCV
membranes resulting in the formation of an -figure and (4) extrusion of
the electron-dense vesicle core. In our study we have shown that these same
exocytotic steps can be seen in transmission electron micrographs of the
C. productus ACPs (compare Fig.
5 of this study and fig. 11 of
Weatherby, 1981). While
generally considered a rare occurrence
(Strolenberg et al., 1977), we
found -figures common on the nerve terminals that comprise the ACPs. In
fact, on many terminals, multiple -figures were evident (Figs
4B,
5). The presence of these
morphological correlates of neurosecretion further strengthens our belief that
the ACPs of C. productus are active neuroendocrine signalling
centers.
Source of innervation and hormone complement of the anterior cardiac plexi
Nerve backfilling, in combination with light and transmission electron
microscopy, shows that all innervation to the ACPs is provided by four axons,
which project to these structures via the sons and
stn. These axons terminate in the ACPs and do not innervate the
muscles of the cardiac sac region of the foregut as does the other axon
(presumably that of the AM neuron) present in each acn. Our goal with
the nerve backfills was to identify the location of the somata innervating the
ACPs. While our LY backfills routinely traveled to each of the paired CoGs,
which are the source of most of the somata projecting to the posterior portion
of the STNS (Coleman et al.,
1992) and the location of approximately 40
FLRFamide-immunopositive somata (A. E. Christie, unpublished observations),
they failed to fill any cell bodies in these ganglia. Thus, it appears
possible that the somata responsible for the innervation of the ACPs may
reside outside the STNS. In support of this hypothesis is one double-labeled
CoG. In this ganglion, weakly dye-filled, FLRFamide-immunopositive axons
appeared to project from the CoG neuropil toward the coc. Since this
nerve connects the CoG with both the supraoesophageal ganglion and the fused
thoracic ganglia, both are potential locations for the cell bodies innervating
the ACPs. At present, we are conducting cobalt chloride backfills from the
acns to these ganglia in an attempt to determine if somata in one or
both of these sites are the source of the axons innervating the ACPs.
Our ultrastructural analysis of the ACPs shows that these neurohemal release sites contain both DCVs and ELVs. This finding suggests that the structures could contain a diverse complement of hormonal modulators including peptides, amines and classical small molecule transmitters. In this paper, we have used immunocytochemistry to show that each of the four axons that innervate the ACPs exhibit FLRFamide-related staining. We are continuing to survey the ACPs for additional modulators and as this study continues, it will be interesting to see what hormone complement is present at these sites and if the neurons that innervate the plexi have conserved or distinct cotransmitter phenotypes.
Structures homologous to the anterior cardiac plexi are likely present in other Brachyuran species
In decapod crustaceans, some neurosecretory organs appear conserved across
phylogeny. In all species thus far examined, structures homologous to the SGs
and the POs have been identified (Cooke
and Sullivan, 1982). While little work has been done on the PCOs,
it appears that these structures are lacking in some species
(Cooke and Sullivan, 1982).
Whether structures homologous to the ACPs are present in other crabs remains
to be determined. There is preliminary evidence that in the genus
Cancer, the physical structure of the ACPs is conserved. Skiebe and
Wollenschläger (2002),
using antibodies to vesicle-associated proteins, have shown that structures,
apparently homologous to the ACPs, are present on the acns of C.
pagurus. Likewise, using incident light microscopy, we found iridescent
regions on the acns of Cancer antennarius, Cancer anthonyi,
Cancer borealis, Cancer irroratus and Cancer magister
(Christie et al., 2002). In
these latter species, antibodies to the same vesicle-associated proteins used
by Skiebe and Wollenschläger
(2002) label the iridescent
sites. Moreover, this labeling is identical to the vesicle-associated protein
staining seen in the ACPs of C. productus (A. E. Christie,
unpublished observations). At present, we are conducting transmission electron
microscopy on the putative ACPs of these Cancer species to confirm
that the ultrastructure of these sites is consistent with that of a
neuroendocrine release zone.
In addition to crabs of the genus Cancer, there is also evidence
suggesting that ACP-like structures are present in species from other
Brachyuran genera. In their treatise on the organization of the stomatogastric
neuromuscular system, Maynard and Dando
(1974) reported the presence
of an opaque bluish-white region on each acn of the blue crab
Callinectes sapidus. While no detailed description of the site was
undertaken, the location of this opaque area is the same as that of the ACP we
report here for C. productus. We have recently begun an incident
light examination of species from a number of other Brachyuran genera
(including Callinectes, Carcinus, Chionoecetes and
Telmessus) and have found iridescent areas on the acns of
these animals as well (A. E. Christie and J. M. Edwards, unpublished
observations). As more data are collected, it will be interesting to see how
broadly conserved ACP-like structures are in Brachyuran species.
It is unclear whether the ACPs, at least as they appear in C.
productus, will be found in the non-Brachyuran decapods, e.g. Palinura
(spiny lobsters) and Astacidea (chelate lobsters and freshwater crayfish).
This is due to the fact that the location and branching structure of the
acns vary between Brachyurans and these other animals
(Maynard and Dando, 1974). In
their extensive descriptions of the STNS of both the Caribbean spiny lobster
Panulirus argus and the American lobster H. americanus,
Maynard and Dando (1974) make
no mention of a putative neurosecretory site on nerves considered homologous
to the crab acns. Likewise, vesicle-associated protein labeling in
several Palinuran and Astacidean species shows no evidence of immunopositive
plexi on the acn homologs
(Skiebe, 2000;
Skiebe and Ganeshina, 2000;
Skiebe and Wollenschläger,
2002).
It is interesting to note that in H. americanus, as well as in the
California spiny lobster P. interruptus and the Australian freshwater
crayfish C. destructor and C. quadricarinatus, there is
evidence of a collection of neurosecretory profiles in the nerves of the
anterior portion of the STNS (Kobierski et
al., 1987; Kilman and Marder,
1996; Kilman,
1998; Skiebe,
2000; Skiebe and Ganeshina,
2000; Skiebe and
Wollenschläger, 2002;
Christie et al., 2003). In
these animals, this neurosecretory region overlies the same general region of
the foregut as the portion of the acn containing the ACP.
Transmission electron microscopy shows that in these species the sheath of
nerves in the anterior portion of the STNS is replete with neurosecretory-type
profiles that are similar in their ultrastructure to those found in the ACPs
of C. productus (Kobierski et
al., 1987; Kilman,
1998; Skiebe,
2000; Skiebe and Ganeshina,
2000, Skiebe and
Wollenschläger, 2002;
Christie et al., 2003).
Whether this site is the Astacidean and Palinuran homolog of the Brachyuran
ACPs remains unclear. The common ultrastructure of the sites and their general
location within the STNS suggest this may be the case. Recently, however, we
have found that the source of innervation to the respective sites is not
conserved. As we show in this paper, the ACPs are innervated by only four
axons. Our backfilling experiments suggest that the CoGs are not the location
of the somata of these fibers. In the freshwater crayfish, C.
quadricarinatus, we have found that at least 24 CoG neurons do contribute
to the innervation of the plexus (Christie
et al., 2003). The likely differing locations of the somata
innervating the C. productus ACPs versus the C.
quadricarinatus plexus suggest that the neurons projecting to the two
sites could receive different synaptic input resulting in quite different
secretory output. Clearly additional study will be needed to determine the
degree of homology between the Brachuran and the Astacidean and Palinuran
plexi.
Physiological functions of the anterior cardiac plexi
While the physiological role(s) of the ACPs within the STNS remains
unknown, several functions have been generally ascribed to circulating
hormones in this system. Many substances have been demonstrated to have
neuromodulatory capabilities in the STNS. The effects of many of these
modulators have been shown to be highly concentration dependent, particularly
on the ci
