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Immunolocalization of a voltage-gated calcium Channel [beta] Subunit in the tentacles and cnidocytes of the Portuguese man-of-war, Physalia physalis.


Cnidocytes, the defining character of members of the phylum Cnidaria, are exceedingly complex cells. This complexity is evident in the elaborate architecture (Beckman and Ozbek, 2012) of the tubule that is everted from the cnidocyst found within the cells, in the venom (Calton and Burnett, 1973; Diaz-Garcia, 2012) released by the discharged tubule, and by the not fully understood mechanism that achieves discharge in a millisecond (Niichter et al., 2006). The presumed high cost of producing such complex cells is compounded by the fact that they can be used only once. Not surprisingly, therefore, cnidocyte discharge is tightly regulated, presumably to help ensure that these cnidocytes are used only when there is a high probability that discharge will achieve the desired results such as capture of prey, locomotion, or defense.

Discharge of cnidocytes involved in prey capture typically requires the close apposition of appropriate chemical and mechanical stimuli that are detected by receptors on the cnidocytes themselves, as well as on associated sensory cells. Receptors on cnidocytes are located in the ciliary complex found on the apical surface of the cell. This consists of a central kinocilium (or cnidocil) surrounded by an array of actin-rich stereovilli (Goltz and Thurm, 1991). Mechanical stimulation of this complex is transduced into membrane depolarizations (Brinkmann et al., 1996). The same structure also serves as a contact chemoreceptor (Thurm et al., 1998b; reviewed by Kass-Simon and Scappaticci, 2002). Chemoreceptors are also present elsewhere in the tentacle. Intracellular recordings from single cnidocytes in the tentacles of Physalia (Purcell and Anderson, 1995) and Cladonema (Price and Anderson, 2006) revealed bursts of synaptically-driven electrical activity in cnidocytes after perfusion of the tentacle with prey-specific chemical extracts. Furthermore, simultaneous recordings from pairs of cnidocytes revealed near synchrony in the timing of these electrical events in different cnidocytes, suggesting that they receive a common input. This input is presumably from a second population of chemoreceptors that serves to detect the "odor" of nearby prey, and may somehow prime the cnidocytes in anticipation of contact with the source of that odor. Thus, cnidocytes are postsynaptic to elements of the animal's nervous system.

Neural control of cnidocyte discharge is also present in a more global context inasmuch as the threshold for discharge is greatly elevated in Hydra fed to repletion (Smith et al., 1974; Thurm et al., 1998b), a response that is absent in nerve-free Hydra. Moreover, intracellular recordings have shown that mechanical stimuli applied to one cnidocyte in a capitate tentacle produce electrical responses in other cnidocytes in that same cluster (Oliver et al., 2008), suggesting that cnidocytes also serve as primary sensory receptors whose output is conveyed to other cnidocytes by way of the nervous system. This conclusion is supported by electron microscopic studies (Holtmann and Thurm, 2001) that reveal chemical synapses from cnidocytes to neurons.

The odor-evoked electrical events recorded from cnidocytes in the capitate tentacles of the hydroid Cladonema have all the hallmarks of synaptically-driven action potentials (Price and Anderson, 2006) (but see Oliver et al., 2008), and patch-clamp recordings from stenotele cnidocytes isolated from Cladonema show that these cells are capable of producing overshooting, [Na.sup.+]-dependent action potentials (Anderson and McKay, 1987). Thus, cnidocytes, at least the ones involved in prey capture, are not the independent effectors they were originally thought to be (Parker, 1919); rather they are closely integrated into the animal's nervous system where they function both pre- and post-synaptically.

A further elaboration of the concept that cnidocytes contain the machinery of chemical synapses is the understanding that cnidocyte discharge is an exocytotic process. This model derives from the observation that the cyst is structurally a vesicle inasmuch as it is enveloped by a membrane (Watson and Mariscal, 1984; Golz, 1994), and the assumption that one of the first steps in discharge would likely be fusion of the cyst membrane with the cell membrane, as occurs during synaptic transmission and other exocytotic processes (Lubbock et al., 1981). This model is given credence by the observation that discharge of a cnidocyte that has been impaled with a microelectrode does not always result in a loss of membrane potential (Brinkmann et al., 1996; Thurm et al., 1998b) as would undoubtedly occur if discharge simply ruptured the cell's membrane.

The facts that cnidocytes are capable of producing action potentials, function as pre-synaptic entities, and apparently undergo exocytosis during discharge prompted a search for ion channels in isolated cnidocytes (Bouchard et al., 2006). That study, which employed standard molecular cloning techniques to screen a cDNA library prepared from mRNA of isolated cnidocytes of Physalia physalis, revealed the presence of voltage-gated [Na.sup.+], [K.sup.+], and [Ca.sub.2+] channels, together with a [Ca.sup.2+] channel beta subunit. The K+ channel ([PpK.sub.V]1) and [Ca.sup.2+] beta subunit (Pp[Ca.sub.V][beta]) were cloned in their entirety and functionally expressed.

Voltage-gated calcium channels (VGCC) have a common basic structure consisting of [[alpha].sub.1], [[alpha].sub.2][delta], and [beta] subunits (Catterall, 2000). The [beta] subunit is a cytoplasmic protein involved in the trafficking to the membrane of the [[alpha].sub.1] subunit of the channel, and also modulates its kinetic of activation (Dolphin, 2003), current-voltage relationship, and recovery from inactivation (Jeziorski et al., 1999). Here, the cellular location of Pp[Ca.sub.V][beta] was determined using standard immunolocalization techniques with both isolated cnidocytes and cnidocytes in situ to determine if it is involved in the exocytotic events that underlie cnidocyte discharge, synaptic transmission to elements of the animal's nervous system, or both.


Generation of the Pp[Ca.sub.V][beta] antisera

Antisera against Pp[Ca.sub.V][beta] were produced commercially (21st Century Biochemicals, Marlboro, MA) as follows. Two peptides (# 1 -VDENDFKPLFEGNSNEPHC and #2-LAK RSAFQHPGKQPVIQKKG) corresponding to regions of the Pp[Ca.sub.V][beta] subunit sequence were synthesized, conjugated to keyhole limpet hemocyanin (KLH), and injected, together, into each of two rabbits. The anti-sera were affinity-purified using the two peptides. All experiments were performed with bleed #7 from one immunized rabbit.

Heterologous expression

cDNA for Pp[Ca.sub.V][beta] was ligated into the oocyte expression vector EGFP-PXOOM (Jespersen et al., 2002) (a gift from Dr. D. Boudko, Rosalind Franklin University) to generate a chimeric [beta] subunit fused with EGFP at the N-terminus. The construct was linearized with Not1, and cRNA for oocyte injection was generated by in vitro transcription with T7 polymerase using the mMessage mMachine (Ambion Inc., Carlsbad, CA). [beta] subunit cRNA was co-expressed with cRNA of the [[alpha].sub.1] subunit of the voltage-gated [Ca.sup.2+] channel from the scyphozoan jellyfish Cyanea capillata (Cc[Ca.sub.V][[alpha].sub.1]) (Jeziorski et al., 1998). Cc[Ca.sub.v][[alpha].sub.1] expresses well in Xenopus oocytes and can form functional complexes with a variety of [beta] subunits, including those from mammals (Jeziorski et al., 1998, 1999). cDNA for Cc[Ca.sub.V][[alpha].sub.1] was ligated into the pXENEX vector (provided by Dr. R. Greenberg, University of Pennsylvania), linearized with NgoM1, and transcribed into cRNA as before. The integrity and quantity of the transcripts was evaluated by agarose gel electrophoresis. Collagenase-treated stage V and VI X. laevis oocytes were injected with cRNA at a ratio of 3 parts of Pp[Ca.sub.V][beta] cRNA to 1 part of Cc[Ca.sub.v][[alpha].sub.1] cRNA as described elsewhere (Bouchard et al., 2006). Oocytes were incubated for 3-6 days at 17 [degrees]C in sterile ND96 supplemented with 2.5 mmol P1 sodium pyruvate, 100 units/ml penicillin, 100 mg/ml streptomycin, and 5% horse serum.

Western blotting

Only oocytes that displayed GFP fluorescence at the membrane were processed for protein extraction. Proteins from these oocytes were prepared using the method of Stebbins-Boaz et al. (2004). Specifically, fluorescent oocytes were placed in a 1.5-ml Eppendorf tube and solubilized on ice with a 1.5-pellet pestle (Kimble/Kontes) in 10 ml/oocyte of homogenization buffer [0.1 mol [1.sup.-1] KCl, 1 mmol [1.sup.-1] Mg[Cl.sub.2], 50 mmol [1.sup.-1] Tris-HCl (pH 7.5), 1 mmol [1.sup.-1] DTT, 80 mmol [1.sup.-1] [beta]-glycerophosphate (MP Biochemicals, Inc., Solon, Ohio) with 1 ml/ml of protease inhibitor cocktail (cat# P8340 Sigma-Aldrich). The homogenate was centrifuged at 10,000 X g for 10 min at 4[degrees]C, and the supernatant was divided into 20-ml aliquots and stored at -20[degrees]C. All samples were prepared by boiling in a Laemmli buffer (12.5 mmol [1.sup.-1] Tris-base pH 6.5, 3% sodium dodecyl sulfate (SDS), 0.05% bromophenol blue, 5% glycerol, 25 mmol [1.sup.-1] DTT, 2% [beta]-ME) for 10 min. Proteins (two oocytes/well) and molecular weight marker mix (Pierce 3-Color Prestained Protein, ThermoScientific, Fisher, Rockford, IL) were separated by sodium dodecyl sulfate-polyacrylamide electrophoresis (SDS-PAGE) using 4%-12% Bis-Tris gels and 1 X NuPAGE MES SDS running buffer. After electrophoresis, the samples were transferred onto a nitrocellulose membrane Protran (Whatman, Dassel, Germany) for 2 h at 30 V in 1 X Tris-Glycine transfer buffer. Reagents for protein fractionation and electro-transfer were obtained from Invitrogen (Carlsbad, CA). Subsequently, nitrocellulose membranes were blocked for 2 h at room temperature in 1 X TBS containing 5% nonfat milk, 0.2% Tween-20; they were then incubated overnight in a cold room with primary antibodies at the following concentrations: rabbit polyclonal anti-[PpCav.sub.[beta]] whole immune serum or pre-immune serum, 1: 2,000; peptide 1 and peptide 2 affinity-purified sera, 1: 300; and rabbit polyclonal anti-GFP (NB 600-310, Novus Biologicals, Littleton, CO), 1: 5,000. Following the primary antibody incubation, membranes were washed six times, 5 min each, with TBS. Blots were incubated for 2 h at 37[degrees]C with goat anti-rabbit IgG conjugated to alkaline phosphatase (Jackson ImmunoResearch Laboratories, Inc.,) diluted 1:2000 in blocking solution. Blots were washed again in 1 X TBS 10 times for 5 min each. The Alkaline Phosphatase Conjugate Substrate Kit (Bio-Rad Laboratories Hercules, CA) was used for colorimetric reactions.

Absorption of [beta] subunit antibody with antigens for the Western blotting

A dilution of 1:333 of whole serum in PBS was incubated overnight at 4[degrees]C with 1 mg/ml of each peptide #1 and #2. The aliquot was centrifuged and the supernatant was further diluted to bring the antibody concentration to 1:2000 before it was applied to the Western blots. Blocking conditions were the same for the affinity-purified antibody with the following difference: 12-15 mg of affinity-purified antibodies were diluted in 100 ml of PBS in presence of 100 mg of the corresponding antigenic peptide before dilution to a working concentration (1:300). The amount of homogenated proteins loaded per well for the blocking experiment corresponded to 1 oocyte/well.


Specimens of Physalia physalis L. were collected from the beach in Marineland, Florida, at high tide, and maintained in flowing seawater at the Whitney Laboratory for Marine Bioscience. Immunocytochemistry was performed on intact tentacles and on isolated cnidocytes. In the former case, small pieces of tentacle were excised from the animal with scissors and anesthetized in isotonic Mg[Cl.sub.2] diluted 1:1 with normal seawater. Fully relaxed tentacles were lightly stretched and pinned to the base of a Sylgard (Dow Corning, Midland, MI)-coated petri dish, then fixed with 4% paraformaldehyde in 0.1 mol [1.sup.-1] phosphate buffer (PB) containing 24 g/1 NaCl overnight at 4[degrees]C or for 4 h at room temperature, then rinsed (6 X 1 h) in PBS containing 0.25% Triton-X 100 (PBS-T) followed by a single rinse in PBS-T containing 10% goat serum (PBS-T-G). The tentacle pieces were then cut either into short 1-2 cm lengths suitable for mounting on a microscope slide, or into single cnidosacs that could be viewed axially. Tentacles were incubated overnight at 4[degrees]C in PBS-T containing 10% NGS and the 7th bleed of the rabbit antisera diluted 1:4000. Once rinsed (6 X 1 h) in PBS-T, tentacles were incubated in the secondary antibody, Alexa Fluor 488 goat anti-rabbit (Invitrogen, Molecular Probe, Eugene, OR) diluted 1:200 in PBS-T overnight at 4[degrees]C. Unbound secondary antibody was washed off and cells' nuclei were labeled with 4',6-diamidino-2-phenylindole (DAPI) (Invitrogen, Molecular Probes, Eugene, OR), rinsed, then mounted on a microscope slide with either Fluoromount (SouthernBiotech) or 60% glycerol in PBS containing 1 mg/ml o-phenylenediamine HCl to minimize fading. Edges of coverslips were sealed with nail polish for glycerol-mounted samples. The experiment was replicated using 8 [micro]g/ml of the affinity-purified antibody #2. The same affinity-purified antibody concentration was used for the blocking experiment. For this experiment, 16 [micro]g/ml antiserum was incubated in PBS overnight at 4[degrees]C in the presence of 3.7 mg/ml of peptide 2. The following day, the solution was centrifuged 5 min at 10,000 X g, and the supernatant was diluted in an equal volume of PBS containing 10% NGS. This solution was then used in immunochemistry on tentacles, following the protocol described above.

Isolated cnidocytes were obtained by heat shock (McKay and Anderson, 1988; Bouchard et al., 2006). Briefly, pieces from fishing tentacles from P. physalis were placed in Eppendorf tubes containing 500 [micro]l of [Ca.sup.2+] -and [Mg.sup.2+] -free artificial seawater (ASW), then warmed to 45[degrees]C for 15 min, vortexed for 5 s, and plunged into an ice bath for 30 s. Residual fragments of tentacle were removed, and the remaining liquid was layered on chilled (4[degrees]C) Percoll (Sigma, St. Louis, MO) diluted 1:1.5 with concentrated ASW (mmol [1.sup.-1]) NaCl, 1120 mmol [1.sup.-1]; KCl, 22 mmol [1.sup.-1]; Ca[Cl.sub.2], 20 mmol [1.sup.-1]; Mg[Cl.sub.2], 65 mmol F1; NaHC[O.sub.3], 2 mmol [1.sup.-1]. The samples were then spun at 5000 rpm (2987 X g) in a Sorval RC 5B centrifuge for 15 min at 4[degrees]C. The pellet was rinsed

three times with [Ca.sup.2+]/[Mg.sup.2+] -free ASW. Small aliquots of the cnidocyte suspension were placed directly onto gelatin-coated microscope slides or charged slides (Fisher Scientific). After 5 min of settling time, excess fluid was removed with a fine pipette and replaced with 3% paraformaldehyde in PBS containing 0.1% saponin (PBS-S). Isolated cnidocytes were fixed for 45 min at room temperature, rinsed (6 X 15 min) in PBS-S and then in PBS-S-G, incubated overnight at 4[degrees]C in primary antisera, rinsed (6 X 15 min), incubated in secondary antisera for 2 h at 37[degrees]C, rinsed, and mounted as before. Preimmune serum was tested on isolated cnidocyte using the above protocol.

Except for the experiment using pre-immunized rabbit serum, immune serum and affinity-purified antibody #2 were preabsorbed with KLH at a final concentration of 1 mg/ml overnight at 4[degrees]C. Images were collected using a Leica SP5 confocal microscope.


A polyclonal antibody specific to peptides from [PpCa.sub.v][beta] was generated and characterized by Western immunoblotting of protein homogenates of Xenopus oocyte expressing [PpCa.sub.v][beta]. Immunochemistry experiments were performed using the immune serum and pre-immune serum on whole tentacles and isolated cnidocytes. In addition, a fraction of the immune serum affinity-purified with the peptide immunogen #2 was tested on whole tentacles.

Characterization of the [PpCa.sub.v][beta] antisera

The co-expression of the [PpCa.sub.v][beta] subunit protein with a jellyfish [[alpha].sub.1] subunit, [CcCa.sub.v][[alpha].sub.1], ensured traffic of the voltage-gated calcium channel (VGCC) complex to the oocyte membrane. EGFP fused to the N-terminus of the [PpCa.sub.v][beta] enabled oocytes expressing [PpCa.sub.v][beta] at the membrane to be selected on the basis of a cell's peripheral fluorescence (Fig. 1a), and was used as a tag on [PpCa.sub.v][beta] in Western blots to identify the chimeric [PpCa.sub.v][beta]/EGFP protein using an anti-GFP antibody. A band of 109 kDa was stained by the anti-GFP antibody and the immune serum (Fig. 1b left and right panels). Although the electrophoretic mobility of [PpCa.sub.v][beta]/EGFP is less than expected (Mr of [PpCa.sub.v][beta] = 59.2; Mr of EGFP = 26: expected total = 86.0), labeling of a band of the same molecular weight by both anti-[PpCa.sub.v][beta] and anti-GFP (Fig. 1b, right and left panel) indicated that they likely targeted the same protein. The immune serum produced no specific staining on blots of lysates from uninjected oocytes (Fig. 1b, left panel, lane c). Together, these findings indicate that the immune serum contains antibodies specific for [PpCa.sub.v][beta].

In a second immunoblotting experiment, we were interested to know whether the whole immune serum contained antibodies against both peptides used to immunize the rabbit. Two fractions of the immune serum were affinity-purified using peptides #1 and #2. Each affinity-purified serum immunostained a single 109-kDa band on blots of electrophoresed oocyte homogenates that contain the recombinant [PpCa.sub.v][beta]/EGFP protein (Fig. 1c).

In a third immunoblotting experiment, the specificity of the immune serum was further tested by pre-incubation with the two [PpCa.sub.v][beta] peptides used to immunize the rabbit (Fig. Id). Identical amounts of the same oocyte lysate (1 oocyte/ well) containing the chimeric [PpCa.sub.v][beta]/EGFP were electrophoresed and blotted together on nitrocellulose and incubated with the same immune serum except that in one case, the immune serum was preabsorbed against the immunogenic peptides. No staining was observed using the preabsorbed serum, while the normal serum still labeled a band of 109 kDa.

The absence of bands below and above 109 kDa on blots treated with the GFP-antibody but their presence on blots treated with the affinity-purified antibodies as well as on blots incubated with preimmune serum (Fig. 1b, c) suggests that the unspecific labeling is caused by cross-reaction with the rabbit's endogenous antibodies.

Despite the observation that the apparent molecular weight of [PpCa.sub.v][beta]/EGFP exceeded the calculated molecular weight by 23 kDa, the following experimental evidence supported the possibility that the band at 109 kDa corresponded to the [PpCa.sub.v][beta]/EGFP: both [PpCa.sub.v][beta] antibodies and anti-GFP antibody labeled a band of the same molecular weight on blots of oocyte homogenates expressing [PpCa.sub.v][beta] EGFP. Immune serum that had been preincubated with immunogens did not bind the [PpCa.sub.v][beta]/EGFP band transferred on blots.


Antibody labeling of the tentacles with anti-[PpCa.sub.v][beta] showed a widely distributed punctate labeling in the ectoderm of the cnidosac, which contains cnidocytes and other cells (Figs. 2, 3). As expected, similar immunochemistry results were obtained when affinity-purified antibody #2 (AP2) was used (Fig 2a). When AP2 was preabsorbed with peptide 2, the antibody lost all ability to bind its antigen in the tentacle (Fig. 2b), which confirmed the specificity of the AP2 antibody. Because the fluorescent labeling in the cnidosac's ectoderm is localized in a small area of the cell, it is difficult to identify the cell types labeled with the antibody.

Cnidocytes isolated from the tentacles of Physalia were exclusively either large or small holotrichous isorhizas with dimensions consistent with those reported by Bardi and Marques (2007). Both typically bore a small cytoplasmic extension at the basal end of the cell, which was most obvious in the smaller holotrichous isorhizas.

Immunostaining of isolated cnidocytes with anti-[PpCa.sub.v][beta] revealed the presence of small (1.20 SE 0.07 [micro]m, n = 13) plaques that appeared to be cell membrane-associated around the base of the cnidocyte (Fig. 4 a, b). These plaques appeared to be restricted to a belt that encircled the cnidocyte basal to the cyst. In whole-mount immunochemistry, similar staining was most evident when a cnidosac was viewed side on (i.e., axially). Occasional random specific staining was observed elsewhere on isolated cnidocytes but, most notably, was absent from the apical surface of the cell. Pre-immune serum did not stain isolated cnidocytes (Fig. 4c).


The findings presented here confirm that the voltage-gated [Ca.sup.2+] channel [beta] subunit [PpCa.sub.v][beta] previously cloned from the cnidocytes of the tentacles of Physalia physalis is expressed in those cnidocytes, presumably as part of heteromeric voltage-gated [Ca.sup.2+] channel complexes.

The specificity of the staining was confirmed by extensive characterization of the [PpCa.sub.v][beta] antibody by Western immunoblotting on protein homogenates from Xenopus oocytes expressing a [PpCa.sub.v][beta] cDNA fused to an EGFP gene (Fig. 1). The immunoblotting experiment demonstrated that antibodies against two distinct parts of [PpCa.sub.v][beta] reacted to a single band on Western immunoblots of protein homogenates from Xenopus oocytes expressing [PpCa.sub.v][beta]/EGFP (Fig. 1c). The same band reacted to the GFP antibodies (Fig. lb). The binding to this band as well as immunostainings in the cnidosac (Fig. 2a, affinity-purified antibody #2--AP2) using the [PpCa.sub.v][beta] sera were blocked by preabsorbtion of the antisera with the peptides used to generate them (Fig. 1d, whole serum preabsorbed with peptides #1 and #2; and Fig. 2b, AP2 preabsorbed with peptide #2). The discrepancy between the apparent molecular weight of the stained band and the Mr of the [PpCa.sub.v][beta]/EGFP should not be taken as a matter of concern because anomalous migration of polypeptides on SDS-PAGE has been reported. Post-translational modifications, hydrophobicity, and the presence of acidic or basic amino acids in polypeptides are some factors that can result in anomalous protein mobility on SDS-PAGE gels (Matagne et al., 1991; Righetti et al., 2001).

Cnidocytes are polarized cells, which in Physalia are grouped into bulbous clusters named cnidosacs. The apex of the cnidocyte with its cnidocil apparatus projects from the surface of the tentacle, while the basal pole, embedded in the ectoderm of the cnidosac, is inserted into the mesoglea by its basal stalk and basal pad of the fibrillar system (Cormier and Hessinger, 1980). No systematic study has yet been undertaken to identify the cellular composition of the cnidosac in Physalia. The ultrastructure of neighboring cells, which are the counterpart of the Hydra's accessory cell, and mucus cells in the cnidosac were reported by Flessinger and Ford (1988) and Cormier and Hessinger (1980), respectively. More recently, antibodies specific to the RFamide neuropeptide family revealed an assemblage of neurites distributed at the base of cnidocytes, and sensory cells that project to the surface of the cnidosac in Physalia (Anderson et al., 2004). In this species, electrophysiological analysis of the cnidocytes' responses, recorded in situ, showed that application of various fractions of fish mucus triggered electrical activity in the cnidocytes without discharge of the cyst (Purcell and Anderson, 1995). This chemosensory signal is presumably driven by neurons acting as intermediates between the cnidocytes and the chemoreceptor-bearing sensory cells. In respect to these observations, sensory signals regulating the discharge likely derive from neuronal terminals that make synaptic contacts onto cnidocytes. Immunostaining with anti-[PpCa.sub.v][beta] reveals that [PpCa.sub.v][beta] is distributed throughout the ectoderm of cnidosacs of Physalia and localized in small plaques, which are distributed at different depths (Fig. 3a-c). Plaques that are localized deep in the ectoderm of the cnidosac might be associated with cnidocytes but also with presynaptic nerve terminals of the peptidergic network identified previously at the base of the cnidocyte assemblage (Anderson et al., 2004). Plaques at superficial depths appear to be linearly arranged (Fig. 3c) as might be expected if they represent synapses along the length of a neuron. This network of fine processes that spread close to the surface of the cnidosac was not labeled by the peptidergic antibodies (Anderson et al., 2004).

In the case of staining on isolated cnidocytes, plaques (Fig. 4) are distributed in a belt-like fashion that encircles the basal region of the cnidocyte. This is the area occupied by the nucleus and other organelles (Hessinger and Ford, 1988). Curiously, a circular pad at the base of the cyst of the large cnidocyte of Physalia was described in a scanning electron microscopy study (see fig. 11 in Cormier and Hessinger, 1980). The fringe of the circular pad is covered with small projections, which are in continuity with the rods that form the fibrillar basket, which surrounds the cyst. We do not know whether there is a structural connection between the [PpCa.sub.v][beta] labeling localized at the cnidocyte membrane and the cyst's circular pad identified in the Cormier and Hessinger study.

Occasional and randomly distributed lateral labeling above the basal ring or below the apical pole of cnidocytes was easily identified as cells or clusters of cells attached to the surface of the cnidocyte membrane (result not shown). This labeling likely originated from cells belonging to the immunopositive [PpCa.sub.v][beta] network dispersed at the surface of the ectoderm of the cnidosac. Sporadic [PpCa.sub.v][beta] punctate labeling was also observed in the distal region of the foot of isolated cnidocytes of Physalia (result not shown). However, we do not know whether this labeling belonged to the cnidocyte itself or to an immunopositive presynaptic contact from another cell attached to the cnidocyte foot.

The immunoreactive plaques on the basal hemisphere of isolated cnidocytes likely correspond to clusters of VGCC associated with presynaptic terminals within the cnidocyte. As previously mentioned, ultrastructural and physiological studies that demonstrate the presence of presynaptic terminals at the base of the cnidocyte are lacking for Physalia. In various other hydrozoan species, however, physiological studies have shown that cnidocytes receive synaptic input from neurons and sensory cells in the tentacle (Thurm et al, 1998b, 2004; Price and Anderson, 2006; Oliver et al, 2008) and, moreover, function as primary sensory receptors for mechanical (Brinkmann et al., 1996) and chemical (Oliver et al, 2008) signals and communicate that information to other cell types (Holtmann and Thurm, 2001; Oliver et al., 2008; Scappaticci et al, 2010). In these capacities cnidocytes serve both pre- and post-synaptic functions. Neurocnidocyte synapses have long been recognized, and they resemble typical cnidarian synapses in terms of both the size and number of the synaptic vesicles (Westfall et al., 1998). Morphological evidence for afferent synapses from cnidocytes to other cell types is less clear. Holtmann and Thurm (2001) described in Coryne tubulosa synapses from cnidocytes to neurons and adjacent rootlet cell that are characterized by the presence of one or two exceptionally large (up to 1100 nm in diameter) vesicles per cnidocyte. They termed these magno-vesicles. Given the strength of the physiological evidence supporting a presynaptic role for cnidocytes, and no evidence for more conventional synapses, it is likely that magno-vesicle synapses constitute the afferent pathway. It is too early to say whether this unconventional synapse is generalized in the phylum. Given that Physalia is also a member of the class Hydrozoa and that its cnidocytes seem capable of integrating sensory information from multiple sources (Purcell and Anderson, 1995), there is no reason to assume that cnidocytes in Physalia do not function as pre-synaptic elements. As such, they must possess the molecular machinery for chemical synaptic transmission, including the various proteins involved in producing and packaging the neurotransmitter into the vesicles, and the various proteins that dock those vesicles to the terminal membrane for exocytosis, together with the heteromeric voltage-gated [Ca.sup.2+] channel complexes that underlie the voltage-gated [Ca.sup.2+] currents that trigger transmitter release.

Interestingly, the aspect and distribution of the immunoreactive [PpCa.sub.v][beta] plaques in cnidocytes mirror the expression pattern of some subtypes of VGCC complexes in hair cells of vertebrates (Layton et al., 2005). Vertebrate hair cells from the cochlea, the vestibular system, and the fish lateral line share many structural and functional resemblances with cnidocytes (Thurm et al., 1998a). These cells express VGCCs on their basolateral membranes, and hair cells involved in hearing possess the L-type channel [Ca.sub.v]1.3 that mediates Ca-triggered exocytosis of transmitters, which activate post-synaptic neurons of the auditory pathway for further processing in the brain. The importance of [Ca.sub.v]1.3 in hearing was demonstrated by the generation of [Ca.sub.v]1.3 knockout mice, which were completely deaf (Platzer et al., 2000; Dou et al., 2004; reviewed; Rutherford and Pangrsic, 2012). Similarly, in zebrafish and human, specific mutations of [Ca.sub.v]1.3 were identified and associated with deafness (Baig et al., 2011). In the inner hair cells of mouse, it appears that among the multiple types of [beta] subunit that are expressed in these cells, the b2 plays the most important role in synaptic function. Knockout mice for the [beta]2 resulted in decrease of [Ca.sub.v]1.3 expression at the membrane, which was functionally correlated to hearing impairment (Neef et al., 2009).

We believe that [PpCa.sub.v][beta] forms part of VGCC complexes that underlie synaptic transmission of sensory information and efferent events detected by cnidocytes to postsynaptic cells such as neurites and other cnidocytes, in a manner similar to that observed in other hydrozoans (Thurm et al., 1998b, 2004; Holtmann and Thurm, 2001; Oliver et al., 2008; Scappaticci et al., 2010).

Although nematocytes were studied among a limited number of species, they are found to be highly diverse. Even within a single species, different types of nematocytes play distinct roles and consequently are modulated differently (Scappaticci and Kass-Simon, 2008, 2010). The cnidocytes present in the cnidosac of the feeding tentacles of Physalia are penetrant. They target soft-bodied prey but, unlike many hydrozoan models used to study the physiology of nematocytes, they do not target crustaceans. Nevertheless, there is no reason to believe that the cnidocyte of Physalia is different from the ones in other hydrozoans (Purcell and Anderson, 1995; Anderson et al., 2004). In fact, we recently cloned a putative metabotropic glutamate receptor and localized it in plaques at the base of the isolated cnidocyte (unpubl. data). We still do not know whether this receptor is associated with areas positive for [PpCa.sub.v][beta]. Still, there is ample evidence that the afferent synapse of the hydrozoan nematocyte is glutamatergic (Sieger and Thurm, 1997) as is thought to be the case at the afferent synapse of the vertebrate hair cell (Meza, 2008). Glutamate was localized to nematocytes (Kass-Simon and Scappaticci, 2002), and ionotropic receptors for glutamate were also identified in these cells (Scappaticci et al., 2004). Physalia cnidocytes most likely share a glutamatergic synapse with other hydrozoan nematocytes.

Although a careful confocal microscopic observation suggested that the plaques are associated with the cnidocyte membrane, it is possible that some of these plaques are localized in the presynaptic terminals of neurons whose processes had remained attached to the cnidocytes during the isolation procedure. However, [PpCa.sub.v][beta] was cloned from a few, well-rinsed isolated cnidocytes by RT-PCR (Bouchard et al., 2006), indicating that [PpCa.sub.v][beta] is indeed present in cnidocytes. Thus, even though the number of plaques revealed by anti-[PpCa.sub.v][beta] may be more than one might expect given the apparent rarity of synapses in electron microscopy studies of cnidocytes, some of those plaques, at least, will mark the location of pre-synaptic termini within the cnidocyte. Ultrastructural analysis coupled to immunochemistry should help determine how [PpCa.sub.v][beta] plaques are organized within other structural components of the cnidocyte.

Implication of the results on the cyst exocytosis hypothesis

The absence of anti-[PpCa.sub.v][beta] staining at the apical surface of the cell is noteworthy. One of the first steps in cnidocyte discharge is thought to be a [Ca.sup.2+]-dependent exocytotic event whereby the membrane at the apical pole of the cnidocyst fuses with the cell membrane. In this model, exocytosis is mediated by voltage-gated [Ca.sup.2+] currents presumably carried by heteromeric voltage-gated [Ca.sup.2+] channels that include a [beta] subunit. This model is supported by the finding that intracellular microelectrode recordings from cnidocytes are not always terminated by cnidocyte discharge (Brinkmann et al., 1996; Thurm et al., 1998b), as would be the case if discharge were brought about simply by membrane rupture. Moreover, cnidocyte discharge in Hydra has been shown to be voltage- and [Ca.sup.2+]-dependent, antagonized by [Mg.sup.2+], and inhibited by inorganic calcium-channel blockers (Gitter et al, 1994), implying that a heteromeric voltage-gated channel located at the apical pole of the cnidocyte should be involved in the discharge. However, no specific staining by [PpCa.sub.v][beta] antibodies was ever observed at the location of the cyst exocytosis. It is possible that a second [Ca.sub.v][beta] is present in the Physalia cnidocyte and constitutes part of the [Ca.sup.2+] channel involved in discharge. While the number of [beta] subunits present in Physalia is unknown, only a single [Ca.sub.v][beta] has been identified in the genome of Hydra (Y. Moran, Hebrew University of Jerusalem; pers. comm.), another hydrozoan, suggesting that a second [Ca.sub.v][beta] in Physalia is unlikely. If this is so, then one must assume either than the VGCC complex that underlies exocytosis of the cyst does not include a [beta] subunit, or that the [Ca.sup.2+] required for exocytosis enters the apical end of the cell through another type of voltage-sensitive pathway (Zhou et al., 2004; Kass-Simon and Scappaticci, 2004; Liebeskind et al, 2011; Zhang et al., 2011; Gur Barzilai et al., 2012). Indeed, it has been argued (Gitter et al., 1994) that because organic antagonists of [Ca.sup.2+] channels do not block cnidocyte discharge in Hydra, the [Ca.sup.2+] channels that underlie discharge may be structurally different from those present elsewhere in the animal. However, [CcCa.sub.v][alpha]1, a voltage-sensitive [Ca.sup.2+] channel that has been cloned from the jellyfish Cyanea capillata and well-characterized by heterologous expression in Xenopus oocytes (Jeziorski et al., 1999), is only minimally sensitive to organic [Ca.sup.2+] channel blockers, suggesting that the pharmacological evidence for an unconventional [Ca.sup.2+] channel involved in discharge may not be relevant.

The [Ca.sup.2+] required for exocytosis may enter the cnidocyte by way of the voltage-gated channels that underlie the action potential in these cells (Anderson and McKay, 1987; Brinkmann et al., 1996; Oliver et al., 2008). These action potentials are Na+-dependent (Anderson and McKay, 1987; Oliver et al., 2008), but the [Ca.sup.2+]-permeability of the underlying channels has not been rigorously assessed. Importantly, the action potential in cnidocytes can trigger cnidocyte discharge (Brinkmann et al., 1996). The amount of [Ca.sup.2+] required to dock a single vesicle, the cyst, is likely to be very small, raising the distinct possibility that sufficient [Ca.sup.2+] might enter the cell during the action potential to trigger discharge, albeit in the absence of true, voltage-gated [Ca.sup.2+] channels.

In conclusion, this work suggests that the Physalia [PpCa.sub.v][beta] subunit is widely distributed in the animal's tentacles where it could function as a component of voltage-gated [Ca.sup.2+] channels at chemical synapses. The distribution of [PpCa.sub.v][beta] is best revealed with isolated cnidocytes, where it is confined to a few plaques that created a belt-like arrangement on the basal hemisphere of the cell. Labeled areas are presumed to be clusters of VGCC complexes, but only electron microscopy coupled to immunolabeling using the [PpCa.sub.v][beta] antibody would confirm whether these channel complexes are associated with presynaptic densities. The absence of antibody binding at the apical end of the cnidocyte, together with the fact that another [Ca.sub.v][beta] expressed in the cnidocyte of Physalia is unlikely, suggests that the [Ca.sup.2+] required for exocytosis of the cyst enters the cell by a pathway other than a conventional VGCC, perhaps by way of the voltage-gated [Na.sup.+] channel present in these cells (Bouchard et al, 2006).


This research was supported by National Science Foundation grant IOS1021769.

Literature Cited

Anderson, P. A. V., and M. C. McKay. 1987. The electrophysiology of cnidocytes. J. Exp. Biol. 133: 215-230.

Anderson, P. A. V., L. F. Thompson, and C. G. Moneypenny. 2004. Evidence for a common pattern of peptidergic innervation of cnidocytes. Biol. Bull. 207: 141-146.

Baig, S. M., A. Koschak, A. Lieb, M. Gebhart, C. Dafinger, G. Nurnberg, A. Ali, I. Ahmad, M. J. Sinnegger-Brauns, N. Brandt, et al. 2011. Loss of Cav1.3 (CACNA1D) function in a human channelopathy with bradycardia and congenital deafness. Nature Neurosci. 14: 77-84.

Bardi, J., and A. C. Marques. 2007. Taxonomic redescription of the Portuguese man-of-war, Physalia physalis (Cnidaria, Hydrozoa, Siphonophorae, Cystonectae) from Brazil. Iheringia Ser. Zool. 97: 425-433.

Beckmann, A., and S. Ozbek. 2012. The nematocyst: a molecular map of the cnidarian stinging organelle. Int. J. Dev. Biol. 56: 577-582.

Bouchard, C., C. G. Moneypenny, R. B. Price, L. F. Thompson, L. Stalheim, and P. A. V. Anderson. 2006. Cloning, functional expression and localization of voltage-gated calcium channel subunits from cnidocytes of the Portuguese Man O'War, Physalia physalis. J. Exp. Biol. 209: 2979-2989.

Brinkmann, M., D. Oliver, and U. Thurm. 1996. Mechanoelectric transduction in nematocytes of a hydropolyp (Corynidae). J. Comp. Physiol. A 178: 125-138.

Calton, G. J., and J. W. Burnett. 1973. The purification of the Portuguese man-of-war nematocysts toxins by gel diffusion. Comp. Gen. Pharmacol. 4: 267-270.

Catterall, W. A. 2000. Structure and regulation of voltage-gated [Ca.sup.2+] channels. Annu. Rev. Cell Dev. Biol. 16: 521--555.

Cormier, S. M., and D. A. Hessinger. 1980. Cellular basis for tentacle adherence in the Portuguese man-of-war (Physalia physalis). Tissue Cell 12: 713-721.

Diaz-Garcia, C. M. 2012. Toxins from Physalia physalis (Cnidaria) raise the intracellular [Ca.sup.2+] of beta-cells and promote insulin secretion. Curr. Med. Chem. 19: 5414-5423.

Dolphin, A. C. 2003. [beta]-subunits of voltage-gated calcium channels. Pharmacol. Rev. 55: 607-627.

Dou, H., A. E. Vazquez, Y. Namkung, H. Chu, E. L. Cardell, L. Nie, S. Parson, H. S. Shin, and E. N. Yantoah. 2004. Null mutation of alpha1D [Ca.sup.2+] channel gene results in deafness but no vestibular defect in mice. J. Assoc. Res. Otolaryngol. 5: 215-226.

Gitter, A. H., D. Oliver, and U. Thurm. 1994. Calcium- and voltage-dependence of nematocyst discharge in Hydra vulgaris. J. Comp. Physiol. A 175: 115-122.

Golz, R. 1994. Apical surface of hydrozoan nematocytes: structural adaptations to mechanosensory and exocytotic functions. J. Morphol. 222: 49-59.

Golz, R., and U. Thurm. 1991. Cytoskeleton-membrane interactions in the cnidocil complex of the hydrozoan nematocytes. Cell Tissue Res. 263: 573-583.

Gur Barzilai, M., A. M. Reitzel, J. E. M. Kraus, D. Gordon, U. Technau, M. Gurevitz, and Y. Moran. 2012. Convergent evolution of sodium ion selectivity in metazoan neuronal signaling. Cell Rep. 2: 242-248.

Hessinger, D. A., and M. T. Ford. 1988. Ultrastructure of the small cnidocyte of the Portuguese Man-O-War (Physalia physalis) tentacles. Pp. 75-94 in The Biology of Nematocysts, D. A. Hessinger and H. M. Lenhoff, eds. Academic Press, San Diego.

Holtmann, M., and U. Thurm. 2001. Mono- and oligo-vesicular synapses and their connectivity in a cnidarian sensory epithelium (Coryne tubulosa). J. Comp. Neurol. 432: 537-549.

Jespersen, T., M. Grunnet, K. Angelo, D. A. Klaerke, and S. P. Olesen. 2002. Dual-function vector for protein expression in both mammalian cells and Xenopus laevis oocytes. BioTechniques 32: 536-540.

Jeziorski, M. C., R. M. Greenberg, K. S. Clark, and P. A. V. Anderson. 1998. Cloning and functional expression of a voltage-gated calcium channel alphal subunit from jellyfish. J. Biol. Chem. 273: 22792-22799.

Jeziorski, M. C., R. M. Greenberg, and P. A. V. Anderson. 1999. Cloning and expression of a jellyfish calcium channel beta subunit reveal functional conservation of the alphal-beta interaction. Recept. Channels 6: 375-386.

Kass-Simon, G., and A. A. Scappaticci, Jr. 2002. The behavioral and developmental physiology of nematocysts. Can. J. Zool. 80: 1772-1794.

Kass-Simon, G., and A. A. Scappaticci, Jr. 2004. Glutamatergic and GAB Anergic control in the tentacle effector systems of Hydra vulgaris. Hydrobiologia 530/531: 67-71.

Layton, M. G., D. Robertson, A. W. Everett, H. A. M. W. Mulders, and G. K. Yates. 2005. Cellular localization of voltage-gated calcium channels and synaptic vesicle-associated proteins in the guinea pig cochlea. J. Mol. Neurosci. 27: 225-244.

Liebeskind, B. J., D. M. Hillis, and H. H. Zakon. 2011. Evolution of sodium channels predates the origin of nervous systems in animals. Proc. Natl. Acad. Sci. USA 108: 9154-9159.

Lubbock, R., B. L. Gupta, and T. A. Hall. 1981. Novel role of calcium in exocytosis: mechanism of nematocyst discharge as shown by X-ray microanalysis. Proc. Natl. Acad. Sci. USA 78: 3624-3628.

Matagne, A., B. Joris, and J. M. Frere. 1991. Anomalous behaviour of a protein during SDS/PAGE corrected by chemical modification of carboxylic groups. Biochem. J. 280: 553-556.

McKay, M. C., and P. A. V. Anderson. 1988. Preparation and properties of cnidocytes from the sea anemone Anthopleura elegantissima. Biol. Bull. 174: 47-53.

Meza, G. 2008. Modalities of GABA and glutamate neurotransmission in the vertebrate inner ear vestibule. Neurochem. Res. 33: 1634-1642.

Neef, J., A. Gehrt, A. V. Bulankina, A. C. Meyer, D. Riedel, R. G. Gregg, N. Strenzke, and T. Moser. 2009. The [Ca.sup.2+] channel subunit [beta]2 regulates [Ca.sup.2+] channel abundance and function in inner hair cells and is required for hearing. J. Neurosci. 29: 10730-10740.

Nuchter, T., M. Benoit, U. Engel, S. Ozbek, and T. W. Holstein. 2006. Nanosecond-scale kinetics of nematocyst discharge. Curr. Biol. 16: R316-318.

Oliver, D., M. Brinkmann, T. Sieger, and U. Thurm. 2008. Hydrozoan nematocytes send and receive synaptic signals induced by mechano-chemical stimuli. J. Exp. Biol. 211: 2876-2888.

Parker, G. H. 1919. The elementary nervous system. Pp. 83-84 in Monographs of Experimental Biology, L. Jacques, T. H. Morgan, and W. J. V. Osterhout, eds. J. B. Lippincott, Philadelphia.

Platzer, J., J. Engel, A. Schrott-Fischer, K. Stephan, S. Bova, H. Chen, H. Zheng, and J. Striessnig. 2000. Congenital deafness and sinoatrial node dysfunction in mice lacking class D L-type [Ca.sup.2+] channels. Cell 102: 89-97.

Price, R. B., and P. A. V. Anderson. 2006. Chemosensory pathways in the capitate tentacles of the hydroid Cladonema. Invert. Neurosci. 6: 23-32.

Purcell, J. E., and P. A. V. Anderson. 1995. Electrical responses to water-soluble components of fish mucus recorded from the cnidocytes of a fish predator, Physalia physalis. Mar. Freshw. Behav. Physiol. 26: 149-162.

Righetti, P. G., A. Stoyano, and M. Y. Zhukov. 2001. Sodium dodecyl sulphate polyacrylamide gel electrophoresis. Chapter 13, pp. 217-274 in The Proteome Revisited: Theory and Practice of All Relevant Electrophoretic Steps. Journal of Chromatography Library Series, Vol. 63, P. G. Righetti, A. Stoyano, and M. Y. Zhukov, eds. Elsevier Science, Amsterdam.

Rutherford, M. A., and T. Pangrsic. 2012. Molecular anatomy and physiology of exocytosis in sensory hair cells. Cell Calcium 52: 327-337.

Scappaticci, A. A., and G. Kass-Simon. 2008. NMDA and [GABA.sub.B] receptors are involved in controlling nematocyst discharge in hydra. Comp. Biochem. Physiol. A 150: 415-422.

Scappaticci, A. A., R. Jacques, J. E. Carroll, L. A. Hufnagel, and G. Kass-Simon. 2004. Immunocytochemical evidence for an NMDA 1 receptor subunit in dissociated cells of Hydra vulgaris. Cell Tissue Res. 316: 263-270.

Scappaticci A. A., Jr, F. Kahn, and G. Kass-Simon. 2010. Nematocyst discharge in Hydra vulgaris: differential responses of desmonemes and stenoteles to mechanical and chemical stimulation. Comp. Biochem. Physiol. A 157: 184-191.

Sieger, T., and U. Thurm. 1997. Glutamatergic properties of synaptic transmission of the hair-cell-analog nematocytes of Corynidae polyps and their modulation by choline-derivatives. P. 736 in From Membrane to Mind: Proceedings of the 25th Gottingen Neurobiology Conference, N. Eisner and H. Wassle, eds. G. Thieme, Stuttgart. (Abstract).

Smith, S., J. Oshida, and H. Bode. 1974. Inhibition of nematocyst discharge in Hydra fed to repletion. Biol. Bull. 147: 186-202.

Stebbins-Boaz, B., K. Fortner, J. Frazier, S. Piluso, S. Pullen, M. Rasar, W. Reid, K. Sinclair, and E. Winger. 2004. Oocyte maturation in Xenopus laevis is blocked by the hormonal herbicide, 2,4-dichlorophenoxy acetic acid. Mol. Reprod. Dev. 67: 233-242.

Thurm, U., M. Brinkmann, P. Golz. P. Lawonn, and D. Oliver. 1998a. The supra-molecular basis of the mechanoelectric transduction studied in concentric hair bundles of invertebrates. Pp. 228-236 in From Structure to Information in Sensory Systems, A. Taddei-Ferretti and C. Musio, eds. World Scientific Publishing, Singapore.

Thurm, U., M. Brinkmann, M. Holtmann, P. Lawonn, D. Oliver, and T. Sieger. 1998b. Modulation of the output of a mechanosensory cell by chemosensory and synaptic inputs. Pp. 237-253 in From Structure to Information in Sensory Systems, A. Taddei-Ferretti and C. Musio, eds. World Scientific Publishing. Singapore.

Thurm, U., M. Brinkmann, R. Golz, M. Holtmann, D. Oliver, and T. Sieger. 2004. Mechanoreception and synaptic transmission of hydrozoan nematocytes. Hydrobiologia 530/531: 97-105.

Watson, G. M., and R. N. Mariscal. 1984. Ultrastructure and sulfur cytochemistry of nematocyst development in catch tentacles of the sea anemone Haliplanella luciae (Cnidaria: Anthozoa). J. Ultrastruct. Res. 87: 159-171.

Westfall, J. A., D. D. Landers, and J. D. McCallum. 1998. Different nematocytes have different synapses in the sea anemone Aiptasia pallida (Cnidaria, anthozoa). J. Morphol. 238: 53-62.

Zhang, T., Z. Liu, W. Song, Y. Du, and K. Dong. 2011. Molecular characterization and functional expression of the DSC1 channel. Insect Biochem. Mol. Biol. 41: 451-458

Zhou, W., I. Chung, Z. Liu, and A. L. Goldin. 2004. A voltage-gated calcium-selective channel encoded by a sodium channel-like gene. Neuron 42: 101-112.


(1) Whitney Laboratory for Marine Bioscience and (2) Dept. of Physiology and Functional Genomics, University of Florida, 9505 Ocean Shore Blvd., St. Augustine, Florida 32080

Received 9 June 2014; accepted 26 August 2014.

* To whom correspondence should be addressed, at University of South Florida Sarasota-Manatee, 8350 N. Tamiami Trail, Sarasota, FL 34243. E-mail:

Abbreviations: Pp[Cav.sub.V][beta], voltage-gated calcium channel [beta] subunit of Physalia physalis; VGCC, voltage-gated calcium channel.
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Author:Bouchard, Christelle; Anderson, Peter A.V.
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Date:Dec 1, 2014
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