Integrative and Comparative Biology Advance Access originally published online on July 25, 2006
Integrative and Comparative Biology 2006 46(6):880-889; doi:10.1093/icb/icl020
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Neural regulation of a complex behavior: body patterning in cephalopod molluscs
Institute of Neuroscience, University of Oregon Eugene OR 97403, USA
Correspondence: 1E-mail: tublitz{at}uoneuro.uoregon.edu
 |
Synopsis |
|---|
Unshelled cephalopods have a remarkable ability to alter their appearance, using textural, postural, and chromatic elements to generate a myriad of body patterns. Of the unshelled cephalopods, it is generally acknowledged that cuttlefish express the most detailed and widest range of body patterns, including static and dynamic patterns. In this paper we present data on the neuronal mechanisms underlying this amazing behavior, focusing on the neuroregulation of the chromatic elements, the chromatophore organs, in the European cuttlefish Sepia officinalis. Cephalopod chromatophore organs, including those in Sepia, are unlike those in any other animal taxa; each consists of a pigment-containing chromatophore cell that expands in response to the coordinated activation of a set of radial muscles which are directly attached to the chromatophore cell. We show that the chromatophore muscles are regulated by 2 different excitatory transmitters, glutamate and the family of FMRFamide-related peptides (FaRPs). Glutamate mediates rapid and transient chromatophore cell expansion whereas the FaRPs are responsible for slower, more sustained responses. Using retrograde dye filling, immunocytochemical and in situ hybridization techniques, we demonstrate that the cell bodies of the glutamatergic and FaRPs-containing motoneurons innervating the fin chromatophore muscles are primarily localized to the posterior chromatophore and fin lobes in the posterior subesophageal mass of the Sepia brain. Data are also presented showing that some fin chromatophore motoneurons have multiple axons in different nerve branches, which accounts for overlapping chromatophore motor fields by adjacent peripheral nerves.
|
Introduction |
|---|
Understanding the mechanisms underlying behavioral plasticity—the ability of the nervous system to alter the performance of an individual behavior depending on changing internal or external conditions—remains one of the most important unresolved issues in modern neuroscience. Neural modulation of a behavior may be short or long term, ranging from milliseconds to a lifetime. Short-term modulations meet the animal's immediate needs while long-term neural changes are the result of experience or a normal part of development.
One well-known causal factor of short- and long-term behavioral changes is neuromodulators. The role of neuromodulators has been examined in numerous animal systems, including locusts, crustaceans, lamprey, and mammals to name but a few (Kettunen and others 2005
; Szucs and others 2005; Heidel and Pfluger 2006; Tryba and others 2006). These studies describe the effects of a host of neuromodulators and demonstrate that they can act centrally on the CNS, peripherally on target muscles, or both (for example, Edwards and Kravitz 1997; Ayali and Harris-Warrick 1998; Johnson and others 2003; Zilberstein and others 2004).
Studies on the role of neuromodulators in behavioral plasticity have focused on both vertebrate and invertebrate model systems. The former has utilized mammalian experimental models such as rats and monkeys to uncover a myriad of new neuromodulators regulating nearly every aspect of physiology and behavior and have identified novel mechanisms likely to underpin behavioral plasticity (for example, long-term potentiation; Xu and others 2006). Over the past 60 years invertebrate model systems have also yielded important new insights on the mechanisms underlying behavior at the systems cellular and molecular levels because of their highly tractable nervous systems, ease of experimental manipulation, and robust behavioral repertoires. Although these preparations, including molluscs, annelids, and insects, are amenable to a wide range of experimentation, they frequently lack the behavioral complexity seen in the simplest of vertebrates. One invertebrate taxon that combines a very rich behavioral repertoire with a tractable CNS is the cephalopod molluscs, a group that includes octopus, squid, and cuttlefish (Messenger 1988). The unique combination of properties found in this group of organisms make them excellent models for studies on the regulation of behavioral plasticity by neuromodulators.
|
Body patterning in cuttlefish: a richly detailed and plastic behavior |
|---|
One behavior unique to cephalopods is their remarkable ability to rapidly produce highly detailed coloration patterns extending across the entire body. Although all unshelled cephalopods manifest this behavior, the most information is known about the European cuttlefish Sepia officinalis because of its vast repertoire of patterns (Holmes 1940
; Packard and Hochberg 1977; Hanlon 1982; Hanlon and Messenger 1988). Sepia, in common with other cephalopods, are capable of adjusting their body coloration to merge with numerous different substrates, including many that are visually complex. Sepia also display specific body patterns in response to disturbances as well as to the appearance of predators, prey, and conspecifics (for example, during courtship). Many patterns are startlingly dynamic, for example, the "passing cloud" display, a set of several dark bands across the width of the body that move anteriorly at 5–10 Hz (Packard and Hochberg 1977). The complexity of these displays suggests a CNS origin, and most appear to be visually mediated since blinded cephalopods exhibit significantly fewer displays (Sanders and Young 1940). These detailed patterns are generated by a suite of chromatic elements, including iridiphores, leucophores, and chromatophores, the latter of which are responsible for the amazing ability of Sepia and other cephalopods to generate a body pattern in less than a second, much faster than any other metazoan (Messenger 1988).
The rapidity of cephalopod body pattern generation is owing to the unique structure of the chromatophore system. Cephalopod chromatophores are true multicellular organs (Fig. 1); at their core lays the chromatophore cell, a pigment-containing cell with a highly elastic plasmalemma. Attached to and radiating from the chromatophore cell are 6–20 striated muscles cells, the chromatophore muscles, which emanate from the chromatophore cell like the spokes of a bicycle wheel (Cloney and Florey 1968). Expansion (or retraction) of the chromatophore cell occurs as a result of the contraction (or relaxation) of the appropriate chromatophore muscles. Because chromatophore muscles produce graded contractions, many intermediate expansion states of the chromatophore cell are possible. Individual chromatophore cells also exhibit dynamic responses, for example, "flickering" behavior produced by rapid minicontraction/relaxation cycles of the chromatophore muscles. Ultimate control of body patterning in cephalopods lies within the CNS since most if not all chromatophore muscles are innervated (Reed 1995).
|
|
Chromatophore regulation by glutamate and FMRFamide-related peptides in S. officinalis |
|---|
Unlike vertebrates where regulation of striated muscular activity is controlled centrally in the spinal cord at the motoneuronal level, neural control of invertebrate muscles frequently occurs in the periphery. Many invertebrate striated muscles are polyinnervated by multiple types of motoneurons, each with their own array of transmitters and modulators. We have been investigating the neuroregulation of chromatophores and their associated muscles in the European cuttlefish S. officinalis. Our attention has focused on the cuttlefish fin, which has fewer cellular layers and is amenable to experimentation without impacting the overall health of the organism.
Using isolated pieces of fin, numerous putative bioactive substances were bioassayed to assess their effects on chromatophores. These investigations were limited to the dark brown chromatophores, 1 of the 3 chromatophore types in cephalopods, the others being yellow and reddish orange in color (Hanlon and Messenger 1988).
Of the classical transmitters tested on the isolated Sepia fin bioassay, glutamate was the only effective substance, causing chromatophore expansion (Fig. 2A). The response to glutamate was very rapid, within a few seconds after application. The threshold for glutamate was between 10–4 and 10–3 M. This high threshold concentration was likely the result of rapid glutamate uptake by other cells in the isolated fin preparation. All suprathreshold concentrations of glutamate always caused a complete and fully reversible chromatophore expansion with little or no desensitization.
|
In addition to glutamate, cuttlefish chromatophores are also regulated by the several members of the FRMFamide neuropeptide family. Like glutamate, FMRFamide application triggered chromatophore expansion (Fig. 2B). Unlike glutamate, there was a significant delay between peptide application and chromatophore expansion. The delay between peptide application and the chromatophore response was usually about 1 min or longer, substantially longer than that for glutamate. The FMRFamide-induced chromatophore expansion persisted for several minutes after the peptide was washed out of the bath, in contrast to the glutamate response which rapidly returned to the pre-application state shortly after glutamate was removed from the bath.
The tetrapeptide FMRFamide is a member of the FMRFamide peptide family whose signature structure is a C-terminus F/LxRFamide. We isolated the FMRFamide coding gene in Sepia using standard molecular biological techniques. The gene structure is shown in Figure 3A. The single open reading frame of the Sepia FMRFamide gene codes for 14 FMRFamide-related peptides (FaRPs). There are 11 copies of FMRFamide and single copies each of FLRFamide, FIRFamide, and the novel decapeptide ALSGDAFLRFamide. Each of the 4 FaRPs caused expansion of brown chromatophores when applied to the isolated fin preparation. Figure 3B shows that all 4 FaRPs elicited similar responses in terms of rate of expansion after peptide application and rate of relaxation after peptide washout but they had different thresholds. FLRFamide was the most potent (threshold concentration 10–9 M), followed by the decapeptide ALSGDAFLRFamide (10–8 M), and FMRFamide and FIRFamide (10–7 M).
|
Sepia also contains a second FaRP gene that codes for 3 additional FaRPs: 2 hexapeptides, NSLFRFamide and GNLFRFamide, and the pentapeptide TIFRFamide (Fig. 3C). Each hexapeptide caused chromatophore expansion when applied to the isolated fin preparation (Fig. 3D). The pentapeptide TIFRFamide was ineffective on the chromatophore bioassay at all concentrations. The latency and duration of the effect of the 2 FaRP hexapeptides were similar to that for FMRFamide. The threshold for each hexapeptide was 10–6 M.
|
Expression of glutamate and FaRPs in the periphery and in the CNS of S. officinalis |
|---|
To determine if glutamate and FaRPs are expressed in chromatophore motoneurons, immunocytochemical and in situ hybridization techniques were used. Using an FMRFamide polyclonal antibody, we demonstrated that FMRFamide-like immunoreactivity was present in the nerve cells associated with chromatophore muscles. A dense meshwork of FMRFamide-immunopositive fibers was found throughout the dermal chromatophore layer in the cuttlefish fin. Many FMRFamide-immunopositive fibers traversed across or along the length of individual chromatophore muscles (Fig. 4A). FMRFamide-like immunoreactivity in peripheral nerves was punctate, particularly when the nerve was closely apposed to the chromatophore muscle (Fig. 4B).
|
FaRP-like immunoreactivity was also present centrally in the Sepia brain. The region of the brain with the highest percentage of FMRFamide-immunopositive cells was the posterior chromatophore lobes (PCL), the location of chromatophore motoneurons (Fig. 5A; Boycott 1953
, 1961; Young 1976; Dubas and others 1986a; Dubas and others 1986b; Miyan and Messenger 1995). The PCL is part of the posterior subesophageal mass (PSEM), a region that also includes the palliovisceral and fin lobes (FLs). An adult Sepia contains 
30,000 neurons in the PCL (Hanlon and Messenger 1988), with cell bodies of several size classes ranging from 20 to over 60 µm in diameter. The PCL is immediately adjacent to the pallial nerve, a nerve thought to contain the axons of the PCL chromatophore motoneurons (Boycott 1961; Young 1976; Dubas and others 1986a; Dubas and others 1986b). FMRFamide immunostaining was present in a subset of PCL somata located in the most posteriolateral region, where the pallial nerve exits the lobe (Fig. 5A and C). Only the smallest PCL cells, those with soma diameters of 20 µm, expressed FMRFamide-like immunoreactivity, and only 20% of the cells in this size class were FRMFamide immunopositive. FMRFamide-like immunoreactivity was also present in cells in adjacent lobes including the palliovisceral and FLs as well as in several other regions of the brain.
|
Confirmation of the presence of FaRPs in a subset of cells in the PCL was obtained using in situ hybridization methods to identify the cells that express the FMRFamide gene. The FMRFamide gene was expressed in the same 20 µm diameter PCL cells that contain FMRFamide-like immunoreactivity (Fig. 5C). This was confirmed using double labeling techniques (Fig. 5A and B). Occasionally, there were a few cells positive for one staining procedure but not the other (Fig. 5A and B).
Glutamate was also found in the PCL. About 90% of PCL cells expressed glutamate-like immunoreactivity (Fig. 5D). Some cells were more intensely stained than others. Although many brain regions also exhibited glutamate immunostaining, all contained a much lower percentage of glutamate immunopositive cells than the PCL. Because 40% of the cells in the PCL expressed FMRFamide-like immunoreactivity (Fig. 5C) and 90% were glutamate positive (Fig. 5D), it was not surprising that some PCL cells were immunopositive for both glutamate and FRMFamide (Fig. 5E). Ninety percent of the FMRFamide-expressing cells were also positive for glutamate immunostaining. It is clear that many chromatophore motoneurons in the PCL co-express both glutamate and FaRPs and that both act as excitatory neurotransmitters on the chromatophore motoneurons. Because of their presence in the PCL and their differential rate of activation of chromatophore muscles, we propose that glutamate acts as the fast excitatory neurotransmitter at the chromatophore neuromuscular junction (NMJ) mediating rapid changes in body patterns, whereas the slower acting FaRPs are the slow excitatory neurotransmitter at the chromatophore NMJ responsible for sustained body patterns (Loi and others 1996; Loi and Tublitz 1997, 2000).
|
Peripheral innervation patterns of chromatophore motoneurons |
|---|
Top
SynopsisIntroductionBody patterning in cuttlefish:...Chromatophore regulation by...Expression of glutamate and... Peripheral innervation patterns... Central location of fin...Concluding remarksREFERENCES |
|---|
The peripheral innervation patterns of chromatophore motoneurons are not well described in any cephalopod. We have begun to address this issue in S. officinalis using a previously developed neuroanatomical nomenclature for the branches of the fin nerve (Gaston and Tublitz 2004
). In particular, we have been interested in determining the extent of the peripheral motor fields of the chromatophore motoneurons. Previous electrical stimulation studies demonstrated extensive overlap of motor fields across adjacent fin nerve branches (Gaston and Tublitz 2004). To investigate this further, we assessed the extent of chromatophore motor fields before and after transection of adjacent fin nerve branches (Gaston and Tublitz 2004). Stimulations of the distal ends of transected nerves (n = 6 branches from 4 animals) and their immediately adjacent neighbor nerves (n = 8) revealed changes in the extent of the motor field and in the number of chromatophores active in that field (Fig. 6). Decreases in both percent chromatophore activity (Fig. 6A) and extent of activated region (Fig. 6B) were typically seen for both transected and adjacent nerves. The transection experiments often revealed primary (1°) and secondary (2°) fields within the region of chromatophore activity (Fig. 6C). Where 1° and 2° fields of stimulation existed, the decreases in activity resulting from transection were most apparent for 2° fields, with little or no change in 1° fields (Fig. 6C).
|
Figure 7 summarizes nerve transection experiments. In transected nerves, changes in the extent of the activated region (Fig. 7A, A1) and in the percentage of chromatophore activity (Fig. 7A, A2) following a cut were small for 1° fields of stimulation (mean decrease of 2 and 11%, respectively) but markedly decreased for 2° fields (mean decrease of 88 and 98%, respectively). These decreases were significant in 2° fields but not 1° fields for both region of activity (F(1,7) = 114, P = 0.00001) and percent chromatophore activity (F(1,7) = 42.9, P = 0.003; 2-way mixed design ANOVA). In immediately adjacent nerves (Fig. 7B, B1 and B2), changes in both 1° and 2° stimulation fields following a cut varied; in most instances, the region of activity and the percent chromatophore activity within the region either decreased or remained the same for adjacent nerves. Because of the high degree of variability, changes in immediately adjacent nerves were not statistically significant for either region or percent chromatophore activity (F(1,10) = 0.13, n.s., and F(1,10) = 0.07, n.s., respectively; 2-way mixed design ANOVA).
|
The changes in region of activity and in percent chromatophore activity in the above described transection experiments suggest that at least some axons innervating fin chromatophores bifurcate and project down multiple fin nerve branches. To further demonstrate the presence of bifurcating axons, single nerve stimulations (n = 4 nerves from 4 animals) were performed while sequentially cutting immediately adjacent nerves. A fin nerve diagram for 1 single nerve transection experiment appears in Figure 8A, while data from all experiments are summarized in Figure 8B–D. In all cases, decreases in region of activity and/or in percent chromatophore activity for the stimulated nerve were evident following the transection of both immediately adjacent nerves. The results from these data are consistent with results described in previous transection experiments presented here (Figs 6 and 7).
|
Overlapping regions of chromatophore activity observed here are in agreement with previous reports of overlapping chromatophore motor fields in other cephalopods (Rowell 1963
; Packard 1974; Dubas 1987; Ferguson and others 1988). In octopus, Dubas (1987) suggested that the overlap is owing to innervation of chromatophores by multiple axons, while Ferguson and colleagues (1988) further showed that the same chromatophore in overlapping regions could be innervated by motoneurons traveling in neighboring stellar nerves of the squid. Buhler and colleagues (1975) reported overlapping chromatophore motor fields as well; however, these authors attributed their result to current spread. Here, changes in activated regions and percent chromatophore activity through nerve transection experiments suggest overlapping is due, at least in part, to motoneuron axon bifurcation and projection down multiple fin nerve branches. This does not rule out the presence of nonbifurcating axons, as both possibilities could and likely do exist because during some transections there was no change in the activated region or in percent chromatophore activity, indicating no shared axons between neighboring branches. Spread of current to neighboring chromatophores is a possible but unlikely explanation for the lack of change, for if current spread was the case, the activated region or percent chromatophore activity would have remained the same following all transections instead of only a subset. |
Central location of fin chromatophore motoneurons |
|---|
One longstanding unanswered question about chromatophore motoneurons is the location of their somata in the Sepia brain. To address this question, individual branches of the fin nerve were backfilled with Texas Red Dextran using the procedures described in Gaston and Tublitz (2004)
. The 3 branches were backfilled: the most anterior and posterior primary branches (named "A1" and "P1" using the nomenclature in Gaston and Tublitz 2004) and the most anterior secondary branch of A1 (named "A1a"). The vast majority of the cell bodies labeled in these backfills were localized to the regions of the PSEM, including the PCL, the palliovisceral lobe (PVL), the FL, and the posterior posterior chromatophore lobe (PPCL; Loi and Tublitz 2000). A brain section of the PSEM with a few labeled cells is shown in Figure 9A. Figure 9B depicts the distribution of cells labeled by the 3 backfills. The majority of cells were localized to the PCL and FL. A significantly lower number of labeled cells were found in the PVL, and only a few were located in the PPCL. Although the 3 backfilled nerve branches undoubtedly contained a mix of motor and sensory axons, and the former almost certainly included nonchromatophore motoneurons, these data strongly suggest that the PCL and FL are the primary locations of the chromatophore motoneurons innervating the fin in Sepia. This conclusion is supported by studies in Sepia and in other cephalopods (Sereni and Young 1932; Boycott 1961; Dubas and others 1986a; Dubas and others 1986b; Gaston and Tublitz 2004).
|
|
Concluding remarks |
|---|
Body patterning behavior in unshelled cephalopods is truly remarkable. Cephalopods are the fastest color-changing organisms in the animal kingdom, producing a myriad of complex patterns used for camouflage as well as interspecific and intraspecific communication (Hanlon and Messenger 1988
). Cephalopod body patterning behavior is arguably the most complex behavior among invertebrates and is clearly one of the most fascinating. The studies presented here provide but a glimpse into the complicated neural mechanisms underlying this behavior. A complete understanding of the neuronal control of body patterning in cephalopods will require a multidisciplinary approach, ranging from molecular to behavioral studies. With the advent of modern neurobiological techniques, what once was a mystery is beginning to be understood. Future studies on body patterning behavior in cephalopods should reveal important new principles on the neuronal regulation of behavior that transcends species differences. |
Acknowledgements |
|---|
The work described in this paper was supported by the National Science Foundation, the American Heart Association, the Medical Research Foundation of Oregon, and NIGMS T32GM07257.
Conflict of interest: None declared.
|
Footnotes |
|---|
From the symposium "Recent Developments in Neurobiology" presented at the annual meeting of the Society for Integrative and Comparative Biology, January 4–8, 2006, at Orlando, Florida.
|
REFERENCES |
|---|
Ayali, A and RM Harris-Warrick. 1998. Interaction of dopamine and cardiac sac modulatory inputs on the pyloric network in the lobster stomatogastric ganglion. Brain Res 794:155–61.[CrossRef][Web of Science][Medline]
Boycott, BB. 1953. The chromatophore system of cephalopods. Proc Linnean Soc London 164:235–40.
Boycott, BB. 1961. The functional organization of the brain of the cuttlefish Sepia officinalis. Proc R Soc Lond B 153:503–34.
Buhler, A, D Froesch, K Mangold, H Marthy. 1975. On the motor projection of the stellate ganglion in Octopus vulgaris. Brain Res 88:69–72.[CrossRef][Web of Science][Medline]
Cloney, RA and E Florey. 1968. Ultrastructure of cephalopod chromatophore organs. Z Zellforsch Mikrosk Anat 89:250–80.[CrossRef][Web of Science][Medline]
Dubas, F. 1987. Innervation of chromatophore muscle fibres in the octopus Eledone cirrhosa. Cell Tissue Res 248:675–82.[CrossRef]
Dubas, F, RT Hanlon, GP Ferguson, HM Pinsker. 1986a. Localization and stimulation of chromatophore motoneurons in the brain of the squid, Lolliguncula brevis. J Exp Biol 121:1–25.
Dubas, F, RB Leonard, RT Hanlon. 1986b. Chromatophore motoneurons in the brain of the squid, Lolliguncula brevis: an HRP study. Brain Res 374:21–9.[CrossRef][Web of Science][Medline]
Edwards, DH and EA Kravitz. 1997. Serotonin, social status and aggression. Curr Opin Neurobiol 7:812–9.[CrossRef][Web of Science][Medline]
Ferguson, GP, FM Martini, HM Pinsker. 1988. Chromatophore motor fields in the squid, Lolliguncula brevis. J Exp Biol 134:281–95.
Gaston, MR and NJ Tublitz. 2004. Peripheral innervation patterns and central distribution of fin chromatophore motoneurons in the cuttlefish Sepia officinalis. J Exp Biol 207:3089–98.
Hanlon, RT. 1982. The functional organization of chromatophores and iridescent cells in the body patterning of Loligo plei (Cephalopoda: Myopsida). Malacologia 23:89–119.[Web of Science]
Hanlon, RT and JB Messenger. 1988. Adaptive coloration in young cuttlefish (Sepia officinalis L.): the morphology and development of body patterns and their relation to behavior. Phil Trans R Soc Lond B 320:437–87.[CrossRef]
Heidel, E and HJ Pfluger. 2006. Ion currents and spiking properties of identified subtypes of locust octopaminergic dorsal unpaired median neurons. Eur J Neurosci 23:1189–206.[CrossRef][Web of Science][Medline]
Holmes, W. 1940. The color changes and color patterns of Sepia officinalis L. Proc Zool Soc Lond 110:17–35.
Johnson, BR, P Kloppenburg, RM Harris-Warrick. 2003. Dopamine modulation of calcium currents in pyloric neurons of the lobster stomatogastric ganglion. J Neurophysiol 90:631–43.
Kettunen, P, A Kyriakatos, K Hallen, A El Manira. 2005. Neuromodulation via conditional release of endocannabinoids in the spinal locomotor network. Neuron 45:95–104.[CrossRef][Web of Science][Medline]
Loi, PK, RG Saunders, DC Young, NJ Tublitz. 1996. Peptidergic regulation of chromatophore function in the European cuttlefish Sepia officinalis. J Exp Biol 199:1177–87.[Abstract]
Loi, PK and NJ Tublitz. 1997. Molecular analysis of FMRFamide- and FMRFamide-related peptides (FaRPs) in the cuttlefish Sepia officinalis. J Exp Biol 200:1483–9.[Abstract]
Loi, PK and NJ Tublitz. 2000. Roles of glutamate and FMRFamide related peptides at the chromatophore neuromuscular junction in the cuttlefish, Sepia officinalis. J Comp Neurol 420:499–511.[CrossRef][Web of Science][Medline]
Messenger, JB. 1988. Why octopuses? Have we learned anything from studying their brains? Sci Prog 72:297–320.[Medline]
Miyan, JA and JB Messenger. 1995. Intracellular recordings from the chromatophore lobes of Octopus. In Abbott, NJ, WilliamsonR , MaddockL (Eds.). Cephalopod neurobiology.London Oxford University Press pp. 415–30.
Packard, A. 1974. Chromatophore fields in the skin of the octopus. J Physiol 238:38P–40P.[Medline]
Packard, A and FG Hochberg. 1977. In Nixon, M and MessengerJB (Eds.). Skin patterning in octopus and other genera. The biology of cephalopods. Symp Zool Soc Lond 38:191–231.
Reed, CM. 1995. Dye coupling in the muscles controlling squid chromatophore expansion. J Exp Biol 198:2631–4.
Rowell, CHF. 1963. Excitatory and inhibitory pathways in the arm of Octopus. J Exp Biol 40:257–70.[Abstract]
Sanders, FK and JZ Young. 1940. Learning and other functions of the higher nervous centres of Sepia. J Neurophysiol 3:501–26.
Sereni, E and JZ Young. 1932. Nervous degeneration and regeneration in Cephalopods. Pubbl Staz Zool Napoli 12:173–208.
Szucs, A, HD Abarbanel, MI Rabinovich, AI Selverston. 2005. Dopamine modulation of spike dynamics in bursting neurons. Eur J Neurosci 21:763–72.[CrossRef][Web of Science][Medline]
Tryba, AK, F Pena, JM Ramirez. 2006. Gasping activity in vitro: a rhythm dependent on 5-HT2A receptors. J Neurosci 26:2623–34.
Xu, NL, CQ Ye, MM Poo, XH Zhang. 2006. Coincidence detection of synaptic inputs is facilitated at the distal dendrites after long-term potentiation induction. J Neurosci 26:3002–9.
Young, JZ. 1976. The nervous system of Loligo II. Subesophageal centres. Phil Trans R Soc Lond B 274:101–67.[Web of Science][Medline]
Zilberstein, Y, E Fuchs, L Hershtik, A Ayali. 2004. Neuromodulation for behavior in the locust frontal ganglion. J Comp Physiol 190:301–9.
CiteULike 
Connotea 
Del.icio.us What's this?
This article has been cited by other articles:
|
|
 |
|
  L. M. Mathger, C.-C. Chiao, A. Barbosa, K. C. Buresch, S. Kaye, and R. T. Hanlon Disruptive coloration elicited on controlled natural substrates in cuttlefish, Sepia officinalis J. Exp. Biol., August 1, 2007; 210(15): 2657 - 2666. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||