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Department of Molecular Biology and Biochemistry, Simon Fraser University, Burnaby, British Columbia V5A 1S6, Canada
* To whom correspondence should be addressed. Email: brandhor{at}sfu.ca
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Abbreviations: D-NAME, D-nitroarginine-methyl-ester • GA, geldanamycin • GBD, geldanamycin binding domain • HSP90, heat shock protein 90 • L-NAME, L-nitroarginine-methyl-ester • NO, nitric oxide • NOS, nitric oxide synthase • ODQ, 1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one • RD, radicicol • sGC, soluble guanyl cyclase • SNAP, S-nitroso-N-acetylpenicillamine
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Introduction |
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TOPAbstractIntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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Competent echinoid larvae will settle and initiate metamorphosis if provided with a hard surface covered with an appropriate organic film, particularly a microbial film (reviewed by Strathmann, 1987; Pearse and Cameron, 1991; also see Discussion). In the absence of such cues, some species delay metamorphosis (Caldwell, 1972; Cameron and Hinegardner, 1974). When placed in clean glass or plastic dishes with fresh seawater, L. pictus larvae rarely metamorphose. This allows experimental investigation of the induction of metamorphosis. The mechanism by which external cues are detected and transduced into the initiation of metamorphosis remains poorly understood, but apparently involves a neurosensory response. Further, it is not clear whether larval or juvenile sensory perception (or both) is responsible for transducing external signals under natural conditions. Evidence for the involvement of neural responses from both the larva and the juvenile has been reported. Electrical stimulation of the oral ganglion or the apical neuropile of Dendraster excentricus larvae induced metamorphosis (Burke, 1983). In contrast, observation of settling behaviors and the prevention of settling (and consequently of metamorphosis) in the presence of inducers clearly demonstrates a role for the juvenile sensory apparatus in L. pictus (Cameron and Hinegardner, 1974; Burke, 1980; our observations). Investigations of the molecular and anatomical basis of signaling events that regulate echinoid metamorphosis can thus be placed in this historical context.
Nitric oxide synthase (NOS) catalyzes the conversion of L-arginine to L-citrulline with the production of the gas nitric oxide (NO). NOS expression and NO function have been documented in both nervous and non-nervous tissues alike across a range of eukaryotic organisms, indicating their antiquity and importance in regulating many cellular processes (Schulte et al., 1998; Cueto et al., 1996; Kuzin et al., 1996; Czar et al., 1997). That NO is diffusible through biological membranes suggests that it may have served as a primitive signaling system between cells before more elaborate mechanisms of cell adhesion and receptor-based signaling evolved. In mammalian cells, NOS activity in vivo requires interaction with heat shock protein 90 (HSP90) (Garcia-Cardena et al., 1998; Bender et al., 1999). We recently reported that metamorphosis of two species of ascidian tadpole larvae is induced by drugs that inhibit the activity of the protein chaperone HSP90, NOS, or soluble guanylyl cyclase (sGC) (Bishop et al., 2001). Among larval tissues, NOS activity is concentrated in the tail muscle cells of the ascidian tadpole Cnemidocarpa finmarkiensis. Removal of the tail stimulates metamorphosis of the head, consistent with there being a signal, probably NO, from the tail that represses metamorphosis. NOS produces NO, a gaseous signaling molecule whose most common effector is sGC (Garthwaite et al., 1995; Salter et al., 1996; Hebeiss and Kilbinger, 1998). Thus, inhibition of NOS often results in a corresponding reduction of cGMP (McDonald and Murad, 1996, for review). Metamorphosis of the marine gastropod Ilyanassa obsoleta was also reported to be induced by inhibition of NOS activity (Froggett and Leise, 1999), indicating that NO may repress metamorphosis in a variety of animals.
To further test the idea that NO-mediated repression of metamorphosis occurs widely within the bilaterian clade, we used the sea urchin L. pictus. We report that NO/cGMP signaling is an important regulator of the events surrounding the transition of form from the larva to the juvenile in L. pictus, that it is downstream from a natural inductive cue, and that this regulation may be dependent upon HSP90 function. NOS was detected in both larval and juvenile organs; such organs may be involved in sensing or transducing the response to natural inductive cues.
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Materials and Methods |
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Pharmacological inhibition
L-NAME (L-nitroarginine-methyl-ester) and its enantiomer D-NAME, radicicol (RD), and ODQ (1H-[1,2,4]oxadiazolo[4,3-a]quinoxalin-1-one) were obtained from Sigma Chemical Corp. (St. Louis, MO). Because there is variation in the rate of development of the juvenile rudiment, individual L. pictus plutei were selected by examination under a stereomicroscope and transferred to wells of 24-well plastic culture dishes (Flow Labs, McLean, VA). Larvae were selected for experiments based on the presence of a large, pigmented rudiment having well developed spines and tube feet. Each well contained about 10 larvae in 2 ml MFSW or experimental solutions in MFSW. To quadruplicate sets of these selected larvae were added L-NAME, D-NAME, RD, ODQ, or MFSW in 1- or 2-ml final volumes. Metamorphosis was monitored using a stereomicroscope; it was scored if the larval epithelium had collapsed on top of an everted juvenile. The activity of tube feet was used as an indicator of larval vitality. The concentrations of L-NAME, RD, and ODQ used in the experiments reported here were chosen because they elicited a metamorphic response in ascidian larvae (Bishop et al., 2001). L-NAME and D-NAME were prepared as 1 M stocks in water and diluted to a final concentration of 1-10 mM with MFSW. ODQ was prepared as a 100 mM stock in DMSO and diluted into MFSW to 50 µM. RD was prepared as a 5 mM stock in DMSO and diluted into MFSW to 5µM. SNAP (S-nitroso-N-acetylpenicillamine) was prepared as a 100 mM stock in DMSO and diluted to 0.1 mM in MFSW. For RD, ODQ, and SNAP treatments, experimental and control wells all contained a final concentration of 0.1% DMSO; this concentration of DMSO did not have any inductive properties. Unless significant metamorphosis was observed sooner, experiments were scored at 24, 48, and sometimes 72 h. A low frequency of spontaneous metamorphosis was observed for larvae placed in plastic dishes; this response tends to occur shortly after the assessment of a larvae and its transfer into a well. If such a response was observed before the addition of drugs, juveniles were removed.
To create a natural inductive cue, glass syracuse dishes were submerged for several days in recirculating tanks containing natural seawater. Ten larvae were exposed to the substrate in MFSW either in the presence or absence of 0.1 mM SNAP. Results shown are from a single experiment.
Microsurgical removal of oral hoods and pre-oral hoods was accomplished using a fine-edged stainless steel pin (Fine Science Tools, Vancouver, BC) fused to a glass pipette. Dissected oral and pre-oral hoods retained their capacity to swim. Quadruplicate sets of 5 larvae or hoods per well (a total of 20 operations) were used for each experiment.
All experiments were tested for statistical significance by performing a one-tailed Student’s t test with the assumption of homoscedastic variance. In all graphs (made using Microsoft Excel 97), the asterisks denote statistical significance; P values are provided in the figure legends. Specific statistical comparisons are described in the figure legends.
NADPHd histochemistry and NOS immunohistochemistry
The NADPH diaphorase staining protocol described by Weinberg et al. (1996) was used with modifications. Larvae were fixed in 2% glutaraldehyde and 1% formaldehyde in sodium phosphate buffer for 1 h at room temperature. Formaldehyde was freshly prepared by dissolving paraformaldehyde (EM grade, Ted Pella, CA) in MFSW, adjusting the pH to 7.4, and then diluting in PB to 1%. After rinsing with PB, fixed larvae were incubated in 0.4 mg/ml nitrotetrazolium blue substrate with 2 mg/ml NADPH from 2 to 16 h at 37 °C. As a negative control, specimens were incubated in 50% ethanol for 2 h and then incubated in nitrotetrazolium blue in the absence of NADPH; no staining was observed under these conditions. Under the fixation conditions used, NOS is the only diaphorase expected to be active (Weinberg et al., 1996). Stained larvae were examined as whole mounts by microscopy or were dehydrated in a graded ethanol series, embedded in polyester wax (BDH Laboratory Supplies, Poole, England), and sectioned at 8 µm. Sectioned larvae were examined using an Olympus Vanox microscope, and images were captured using a Sony DXC-950 3CCD camera.
Universal anti-NOS (Affinity Bioreagents, Golden, CO) polyclonal rabbit antibody was used to detect NOS in growing and mature larvae. Larvae were fixed for 2 h at room temperature in 4% formaldehyde (prepared as outlined above). Fixed larvae were blocked with PB saline containing 5% bovine serum albumin and 0.1% Triton-X-100 and then incubated in 1:100 anti-NOS overnight at 4 °C. Larvae were incubated in secondary antibody (goat anti-rabbit-Alexa 568, Molecular Probes, Eugene OR) for 2 h at room temperature and then rinsed, mounted, and viewed on a Zeiss LSM 410 confocal microscope. Images were processed using Adobe Photoshop 5.5 or 6.0.
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Dendraster excentricus pluteus larvae contain cells that express catecholamines in the lower lip of the mouth; this region was thus termed an oral ganglion (Burke, 1983). Removal of the oral hood (OH), which includes the oral ganglion, induced metamorphosis (Burke 1983). That observation and the expression of NOS in the oral ganglion cells of L. pictus larvae led to the hypothesis that these cells repress metamorphosis via their production of NO. To test this idea, we microsurgically removed either the entire OH or the pre-oral hood (PH) from mature larvae and scored the frequency of metamorphosis. This operation did not induce metamorphosis of L. pictus after 6 h (not shown), so L-NAME was added to see if larvae lacking the OH or the PH had retained their capacity to undergo metamorphosis. Neither postoperative larvae nor the OH and PH were responsive to L-NAME at concentrations that induced metamorphosis in intact control larvae (Fig. 6A). To further test if the postoperative larvae and the dissected tissues had retained the capacity to metamorphose, we added 50 µM ODQ after 14 h of incubation in L-NAME. This resulted in a very rapid metamorphic response (Fig. 6A). The OH and PH did not initially undergo epithelial collapse typical of intact metamorphosing larvae, although they did so within 24 h (data not shown). Microscopic analysis indicated that the OH and PH were not necrotic, but rather they had undergone genuine cellular rearrangements characteristic of the epithelium of metamorphosing larvae. Therefore, microsurgical removal of the OH or the PH did not lead to metamorphosis of postoperative larvae, and apparently decreased their capacity to respond to NOS inhibition but not sGC inhibition. Dissected OH and PH tissues underwent rearrangements typical of metamorphosing larvae, but only after a protracted period in drug.
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Frequently, some of the treated larvae did not respond by initiating metamorphosis, even after longer incubations (72 h in some cases). In similar experiments, we have found that a fraction of selected larvae do not respond to dishes coated with microbial film. The fraction of resistant larvae in such experiments was variable (not shown), but similar to the fraction resistant to potent drug treatments such as ODQ (Fig. 4). Variation in response to inducers, whether natural or otherwise, may represent variation in sensitivity of sensory perception, levels of NO repression, or response to a reduction of NO signaling (or a combination thereof) among larvae of a clutch and among clutches. Perhaps the resistant larvae had not achieved competence to undergo metamorphosis, despite the morphological similarity of their rudiments to those that did metamorphose. In fact, in some cases, larvae containing less well developed rudiments were responsive to drugs, whereas those with large, highly pigmented rudiments were not. It is clear that competence does not strictly correspond to the presence of a fully formed rudiment within the larva. Assessing competence is problematic in that one does not know whether a lack of response is due to lack of competence or failure to respond to an inductive cue.
To our knowledge, the concept of competence does not describe a specific biological state in any marine invertebrate having planktotrophic larvae and benthic adults. Larvae with no rudiments or abnormal rudiments do not respond to inducers of metamorphosis (Cameron and Hinegardner, 1978; CDB, unpubl. obs.), so competence in urchins represents a discrete change in the physiological state of the larva that is related to the growth and development of the juvenile. Competence is a phenomenon that requires further investigation and should be considered in all studies on the regulation of metamorphosis. The acquisition of competence coincides with the initiation of metamorphosis in some animals, but not in others (Birkeland et al., 1971; Degnan et al., 1997; Bishop et al., 2001). This indicates that the fitness consequences associated with the timing of, and substrate choice during, settling and metamorphosis vary. What other signaling systems may be contributing to the timing events surrounding life cycle transformations? Studies on thyroxine in echinoids suggest its involvement in the evolutionary loss of larval feeding. The addition of exogenous thyroxine leads to a reduction of larval structures and the time to metamorphosis in D. excentricus (J. Hodin, Friday Harbor Laboratories, and A. Heyland, University of Florida; pers. comm.). It will be interesting to know if and how NO/cGMP and hormonal signals interact to regulate the timing of life cycle transformations in echinoids.
Between different clutches, we have observed a striking difference in the response of larvae to NOS inhibition. Increased sensitivity is manifested as a more rapid response given that identical concentrations of L-NAME and D-NAME were used. We cannot rule out other variations in culturing conditions, such as larval densities and food. The data on NO signaling presented here are from the clutch that was the most sensitive to NOS inhibition. This clutch often responded to NOS inhibition within 2 h, whereas another clutch often took 24-48 h to show a significant effect. The source of this variation is not clear.
We have shown that D-NAME has inductive properties that are suppressed by SNAP, indicating that application of D-NAME also leads to a decrease in NO. Although D-NAME is often used as an inactive negative control for L-NAME treatment, we propose that it does inhibit NOS, but less effectively than L-NAME; others have also noted this activity (Babal et al., 2000). Therefore, D-NAME should be used as a less active enantiomer of L-NAME, not an inactive enantiomer. The extent to which D-NAME is useful as a control for L-NAME treatment will depend on the sensitivity of the experimental system to NO reduction and the concentration of drug used.
There was a lag in the response to radicicol after the beginning of treatment. Radicicol competes with ATP for binding to HSP90, thereby inhibiting its function in binding and folding proteins (Schulte et al., 1998; Sharma et al., 1998). As a protein chaperone, HSP90 interacts with members of several signal transduction pathways (reviewed by Pratt, 1998). In concert with accessory proteins, HSP90 promotes the folding and maintenance of the active state of several known client proteins (Aligue et al., 1994; Whitesell et al., 1994; Nathan and Lindquist, 1995; reviewed by Caplan, 1999). NOS activity in some mammalian cells, including neurons, requires an interaction with HSP90; all three vertebrate isoforms of NOS are degraded in the presence of geldanamycin (GA), another HSP90 inhibitor (Joly et al., 1997; Garcia-Cardena et al., 1998; Bender et al., 1999). Like RD, this agent inhibits HSP90 function by competing with ATP for binding (Promrodou et al., 1997). When the folding function of HSP90 is impaired by inhibitory drugs such as RD and GA, its client proteins (which are often in complexes including HSP90) may be caught in a partially folded state that is then recognized by the ubiquitin-proteasome protein degradation machinery (reviewed by Pratt, 1998; Caplan, 1999). Thus, some client proteins are expected to be degraded or lose activity after HSP90 activity is inhibited. In this circumstance, a response to inhibition of HSP90 would not be expected until its activity had become limiting and its critical client proteins had lost activity or decayed. Such a lag in response was observed for three HSP90 inhibitors that induced metamorphosis when applied to ascidian larvae (Bishop et al., 2001). Thus, we consider this lag to be a consequence of the mechanism by which RD probably leads to a decline in NOS activity. However, a direct demonstration of interaction between HSP90 and NOS in urchins is warranted.
All of the biochemical characterizations concerning the inhibitory properties of anti-HSP90 drugs have been conducted with vertebrate cells. It is relevant to assess whether RD is likely to have the same effect on L. pictus HSP90 as it does on vertebrate HSP90. The crystal structure of a geldanamycin-HSP90 complex has been determined (Stebbins et al., 1997). The geldanamycin binding domain (GBD) is 43% conserved at the amino acid level between vertebrates and E. coli; the aspartic acid residue (Asp93) is absolutely conserved among all HSP90 homologs from 35 species. A hydrogen bond network between Asp93 and the carbamate group of GA has been suggested by structural and functional studies to play the most critical role in the binding of HSP90 to GA (Schnur et al., 1995; Stebbins et al., 1997). Thus, it is probable that GA has similar inhibitory properties on HSP90 from all organisms. RD and GA share no structural similarities, but RD can compete with GA for binding to the N-terminal portion of HSP90 that contains the GBD (Schulte et al., 1998). Moreover, like GA treatment, RD depletes cells of known HSP90 client proteins (Schulte et al., 1998). It is reasonable then to expect a set of highly conserved intermolecular interactions between the GBD of HSP90 of different organisms and RD and hence, a highly conserved mechanism of inhibition of HSP90 by RD. Consistent with this conclusion, GA and RD had similar effects on the initiation of ascidian metamorphosis (Bishop et al., 2001) and morphogenetic movements during sea urchin embryonic development (CB, unpubl. obs.).
Under natural circumstances, the initiation of metamorphosis by competent L. pictus larvae results from a sensory response to appropriate environmental cues. Minimally, this is a biochemical cue, although a hard surface is usually required (Cameron and Hinegardner, 1974). It is not clear what cells or organs are involved in transducing this chemo- and mechanosensory perception into a metamorphic response. The rate of biphasic potentials recorded from the larval body or near the rudiment increases more in response to a substrate "conditioned" with a microbial film than to an unconditioned substrate (Satterlie and Cameron, 1985). This suggests that both the larval and juvenile neural systems are responsive to environmental stimuli. We have not tested whether the drugs used herein can induce metamorphosis in the absence of contact with a hard surface, but the suppression by SNAP of the inductive properties of biofilm demonstrate that NO signaling is downstream of sensory perception leading to metamorphic events.
Various experiments have attempted to address how metamorphosis is initiated and coordinated. The results can differ among echinoid species. Although many species require a hard surface for settlement before metamorphosis, larvae of the sand dollar D. excentricus suspended in seawater can be induced to metamorphose by a heat-labile, low-molecular-weight compound extracted from the sand of a bed of adults (Highsmith, 1982; Burke, 1983, 1984). Low-voltage electrical stimulation of the oral ganglion on the lower lip of the larval mouth or the apical neuropile between the preoral and anterolateral arms on the preoral hood region of the D. excentricus larva induced metamorphosis (Burke, 1983). These sensitive larval areas have axonal connections (Burke, 1983), and there is a ciliary patch on the pre-oral hood that may have a sensory function (Nakajima, 1986). Electrical stimulation of the oral ganglion has been reported to induce metamorphosis in several echinoids, including L. pictus (Burke and Gibson, 1986), although Cameron and Hinègardner (1974) reported otherwise for L. pictus. The difference in these results may be methodological. Recently, Beer et al. (2001) reported that cells in the lower lip of the larval mouth of the sea urchin Psammechinus miliaris develop immunoreactivity to a serotonin antibody. We found staining for NOS protein and NOS activity in cells that appear to be neurons in the lower lip of the mouth, corresponding to the region of the oral ganglion (Burke, 1983), and in cells of the preoral hood, perhaps corresponding to the apical neuropile (Burke, 1983). When Burke (1983) excised the oral hood of D. excentricus—including the oral ganglion and apical neuropile—both fragments of the larva rapidly began metamorphosis, but this did not occur when only the preoral hood—lacking the oral ganglion—or larval arms—lacking both sites—were excised. The excised preoral hood and remaining larva were able to respond to a chemical cue for metamorphosis, but isolated larval arms did not (Burke, 1983). Isolated larval arms of some species, including D. excentricus, can be induced to contract by treatment with divalent ionophores or the neural transmitters adrenalin, noradrenalin, and dopamine (Burke, 1982, 1983). Dopamine induced only a few whole D. excentricus larvae to initiate metamorphosis, suggesting the local response of arms can be inhibited centrally.
On the basis of his experiments, Burke (1983) proposed that there is a mutually inhibitory control of metamorphosis between the oral hood and remainder of the D. excentricus larva that is switched off in response to an appropriate cue (or electrical stimulation). The inhibitory region of the oral hood appears to be localized to the larval mouth (Burke, 1983), while the preoral and remaining regions of the larva must have sensory receptors for the chemical cue that induces metamorphosis. Data from histological sectioning and optical reconstructions of the L. pictus oral epithelium stained for NOS suggest that nitrergic neurons may reside within this epithelium, possibly performing a sensory role related to feeding or metamorphosis. These NOS-expressing cells were considered as candidate NO-signaling centers. We removed the pre-oral hood or the entire oral hood. In the former case, most of the oral cells remained with the larva; in the latter, they were removed (see Fig. 6B). In direct contrast to D. excentricus, L. pictus did not metamorphose in response to the removal of the oral hood, a basic distinction between these two species. Moreover, both classes of L. pictus postoperative larvae were less sensitive to NOS inhibition than were the intact controls, but they apparently retained their sensitivity to inhibition of sGC. In Figure 6A, ODQ was added directly to wells containing postoperative larvae that had been treated with L-NAME for 14 h; there may have been an additive effect of the two drugs. Accordingly, when tested separately, a five-fold excess of L-NAME or ODQ is required to induce metamorphosis of larvae lacking the oral hood over concentrations that induce metamorphosis of control larvae (CDB, unpubl. obs.). These experiments are difficult to interpret with respect to Burke’s model of mutual inhibition, but they do suggest the involvement of cells in the oral hood of L. pictus in a pathway that regulates metamorphosis by NO/cGMP signaling.
The regulatory role of these and other NOS-expressing cells in larvae or juveniles may be additive. In L. pictus, the tube feet of the rudiment appear to have sensory receptors that may be involved in inducing metamorphosis (Burke, 1980). We found intense staining for NOS activity in the nerve plexus lining the outer epithelial layer of the tube feet of the rudiment. NO has been implicated in the relaxation of adult tube feet (Billack et al., 1998). The presence of NOS in cells associated with other structures that may have a sensory role (the pre-oral hood, the tips of the anterolateral arms and epaulettes) suggests that the drugs we used act on one or more of these groups of cells to inhibit their production of NO. Indeed, microsurgical and expression data indicate that multiple larval structures and perhaps juvenile structures transduce sensory information, by NO/cGMP signaling, which leads to the initiation of metamorphosis. The frequency of metamorphosis of larvae of L. variegatus was increased by excess potassium or calcium ions (Cameron et al., 1989). Metamorphosis of Strongylocentrotus purpuratus larvae was induced by treatment with calcium ionophore A23187 or quercetin, an inhibitor of a [Ca,Mg]-ATPase (Klein et al., 1985). Ionic fluxes may play a role, perhaps in coordinating cellular responses (Burke, 1983; Pearse and Cameron, 1991). Some mammalian isoforms of NOS (endothelial and neuronal) are dependent on Ca2+ for their activation (Mayer et al., 1998). The inductive properties of Ca2+ flux may relate to the role of Ca2+ in the regulation of NOS activity.
With this report, there is now evidence that NO plays a repressive role in regulating the initiation of metamorphosis in a protostome (Ilyanassa) and three deuterostomes (two ascidians and an echinoid) (Froggett and Leise, 1999; Bishop et al., 2001). NO is involved in metamorphosis of larvae that do not grow before metamorphosis and retain much of the larval tissue (ascidians), larvae that grow as swimming veliger larvae but do not undergo profound changes upon metamorphosis (Ilyanassa), and larvae that undergo extensive growth and catastrophic metamorphosis in which most larval tissues are degraded and replaced by a radically different juvenile (echinoids). NO, a universal and ancient signaling molecule in eukaryotes, may have a role in regulating metamorphosis in a wide diversity of animals.
Sea urchin larvae are optically clear and can easily be cultured in large numbers. This fact, and a rich experimental literature on settling and metamorphosis, make echinoids a useful system with which to investigate the neuroanatomical basis for the regulation of metamorphosis. These features and our findings provide a basis for a more focused experimental effort to identify which cells or organs repress metamorphosis by NO production in L. pictus.
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Acknowledgments |
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Footnotes |
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Literature Cited |
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TOPAbstractIntroductionMaterials and MethodsResultsDiscussionLiterature Cited |
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0.01; PRD48 
