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847 The largest growth cones in the animal kingdom: an illustrated guide to the dynamics of Aplysia neuronal growth in cell culture Peter Lovell* and Leonid L. Moroz1,*,y *The Whitney Laboratory for Marine Bioscience, University of Florida, 9505 Ocean Shore Blvd., St Augustine, FL 32080, USA; yDepartment of Neuroscience and McKnight Brain Institute, University of Florida, Gainesville, FL 32611, USA Synopsis The marine mollusc, Aplysia californica is a powerful experimental model in cellular and systems neuroscience. Aplysia neurons are large, colored, and located at the ganglionic surface. Because of this, many neurons can be easily identified in terms of their physiological properties, synaptic connections, and behavioral roles. Simple networks can be reconstructed in cell culture and have been widely useful for cellular and molecular biological studies of neuronal growth, synaptogenesis, and learning and memory mechanisms. Here, we show that Aplysia neurons can form truly gigantic growth cones reaching up to 630 mm in diameter making them the largest growth cones ever reported in the animal kingdom. Second, using time-lapse video microscopy we have characterized the dynamics of neuronal outgrowth for 3 identified cell types (mechanosensory neurons, L7 motoneurons, and modulatory MCC neurons) representing 3 major functional classes of neurons. We show both cell-specific and neurite-specific growth characteristics and an irregular oscillatory rate of outgrowth ranging from 20 to 100 mm/h. Third, we characterized the dynamics of axotomy-induced neurite outgrowth as well as extrasomatic localization of b-tubulin mRNA in restricted regions of neuronal processes including growth cones and varicosities. The extrasomatically located mRNAs can be an important pool of neuronal transcripts supporting semiautonomous behavior of growth cones and localized synthesis of proteins in distinct and distant neuronal compartments. The reported data are compared with the existing literature from Lymnaea and Helisoma neurons as well as vertebrate preparations. Finally, our observations can provide an illustrated guide to complex behavior of neurons and glia in cell culture as well as their dependence upon various trophic factors and responses to neuronal injury. Introduction to neuronal growth cones Proper functioning of the adult nervous system in all animals is dependent on the formation of synaptic connections between specific presynaptic and postsynaptic neurons in the central nervous system, and between neurons and their targets (for example, muscle fibers or secretory cells), during development. Before synaptogenesis can occur, physical contact must be established even though the presynaptic and postsynaptic elements may be located in different regions of the brain, different ganglia in invertebrates, or different muscles throughout the body. To bridge this physical gap, neurons extend long processes that are capable of finding specific neuronal or muscle targets in the proverbial haystack of potential targets. This fundamental ability underlies all subsequent functions of a neuron and the nervous system. Neuronal outgrowth is the process whereby neurons establish long polarized processes extending from their somata to prospective synaptic partners. This process is accomplished by dynamic regulation of the actin and microtubule-based cytoskeleton within the motile growth cone at the distal end of the extending neurite. Growth cones are tipped at their distal edge by filopodia, narrow cylindrical extensions capable of extending tens of microns from the periphery of the growth cone (Dent and Gertler 2003). These filopodia are primarily composed of bundled F-actin oriented with the barbed (þ) end facing the distal membrane and the pointed end facing back into the core of the growth cone. Between the extended filopodia exists the lamellipodia, a flattened, veil-like region composed of an F-actin meshwork (Dent and Gertler 2003). Early work with dorsal root ganglion neurons (Bray and Chapman 1985) and later Aplysia neurons (Goldberg and Burmeister 1986) demonstrated that growth cones progress through 3 morphologically distinct stages to form new axon segments: protrusion, engorgement, and consolidation. Protrusion occurs by the elongation of filopodia and lamellipodia through the polymerization of actin 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. 1 E-mail: [email protected] Integrative and Comparative Biology, volume 46, number 6, pp. 847–870 doi:10.1093/icb/icl042 Advance Access publication October 3, 2006 The Author 2006. Published by Oxford University Press on behalf of the Society for Integrative and Comparative Biology. All rights reserved. For permissions please email: [email protected]. 848 filaments. Engorgement occurs when the lamellipodial veils become invested with vesicles and organelles, likely through both Brownian motion and directed microtubule-based transport. Consolidation occurs as the proximal part of the growth cone assumes a cylindrical shape and transport of organelles becomes bidirectional, thus adding a new distal segment of the axon (Dent and Gertler 2003). Subsequent studies have reinforced this 3-phase model of growth-cone movement during neuritogenesis in PC12 cells, rodent sympathetic (Aletta and Greene 1988), and cortical neurons (Halloran and Kalil 1994). Growth-cone motility, including extension, collapse, or turning in response to diffusible or substratebound cues, all occur due to an interaction between surface receptors on the growth cone and the underlying actin cytoskeleton. Binding of ligands to their specific receptors activates intracellular signal cascades (extensively reviewed by (Huber and others 2003) that converge on actin-associated proteins. More than 20 proteins bind directly to F-actin and/or G-actin and have been localized immunocytohemically to the growth cone including gelsolin (Tanaka and others 1993; Lu and others 1997), ADF/cofilin (Meberg and Bamburg 2000), Arp2/3 (Goldberg and others 2000), filamin (Letourneau and Shattuck 1989), fascin (Cohan and others 2001), and thymosin (Roth and others 1999; van Kesteren and others 2006). These actin-binding proteins regulate the dynamics of the actin cytoskeleton by (1) binding and/or sequestering actin monomers, (2) nucleating actin filaments, (3) capping the barbed or (4) pointed ends of actin filaments, (5) acting as barbed-end anticapping proteins, (6) severing F-actin, (7) bundling, cross-linking, or otherwise stabilizing F-actin, and (8) anchoring F-actin to membrane adhesions or specific regions of the membrane (Dent and Gertler 2003). To further complicate the issue, many of these actin-binding proteins are probably capable of multiple functions depending on the internal state of the growth cone, the area in which they are localized, and their phosphorylation state. While the localization of these binding proteins in the growth cone is well established, direct functional studies (that is, knockout and overexpression, GFP-labeling and dynamic localization, and determination of temporal and spatial activation patterns) are lacking for many of these proteins. While many elegant studies have examined neurite outgrowth and pathfinding in intact animals (Sabry and others 1991; Nyhus and Denburg 2000), the majority of experiments involved in elucidating fine details of growth-cone mechanics and dynamics have been conducted in primary cell culture. Along with vertebrate preparations (Lohof and others 1992; Hopker and P. Lovell and L. L. Moroz others 1999; Gomez and others 2001) and other invertebrate preparations, molluscan neurons grown in cell culture have long been a favorite model for research in this field (Burmeister and others 1991; Peter and others 1994; Goldberg 1998; Van Wagenen and others 1999; Koert and others 2001; Spira and others 2001; Geddis and others 2004; Trimm and Rehder 2004; Welshhans and Rehder 2005). Molluscan neurons have a number of significant advantages over other preparations for growth-cone research. First, their neurons are extremely large (up to 1 mm diameter cell bodies for R2 in Aplysia) and specific individual neurons can be visually identified accurately between animals. Second, many of these visually identifiable neurons have been functionally well characterized in the intact ganglia allowing accurate selection of presynaptic or postsynaptic targets for coculture experiments. Third, the identified neurons are extremely easy to individually remove from the intact ganglia and place in cell culture. Fourth, in simple primary cell culture conditions devoid of glial or other cellular contamination, molluscan neurons exhibit robust and rapid neurite outgrowth. Finally, the growth cones that form at the tip of the extending neurites are huge (often 20–40 mm wide but can be more than 500 mm wide) in comparison to those found in other preparations. Taken together, molluscan growth cones represent the simplest and most accessible model for investigating neurite outgrowth and growthcone dynamics in terms of formation of functional networks and structural changes associated with learning and memory (Kandel 2001). The use of Aplysia cell culture in neuroscience: overview The development of cell culture techniques for isolated, identified molluscan neurons allows researchers to examine neurons under controllable and experimentally malleable conditions. Theoretically, this means in cell culture the precise extracellular environment to which the neurons are exposed is known and therefore allows for the direct assessment of defined variables on various neuronal parameters. For example, trophic molecules or other factors can be added to the media (Munno and others 2000), substrate surfaces can be modified, or multiple identified neurons can be cultured together to examine interactions (Syed and others 1990; Lovell and others 2002) and their effects on any measurable element from the molecular level to learning and memory (Kandel 2001). In many cases, this direct control of experimental variables is not possible in the undefined and extremely complicated environment of the intact ganglion. Therefore, while identified neurons can and should be studied in situ to determine Aplysia neurons in cell culture their functions in the intact brain, cell culture provides a powerful tool to uncover exactly how these neurons are able to accomplish these functions. Although neurons from multiple molluscan species have been examined in defined cell cultures, the vast majority of relevant work has been conducted on 3 species of gastropod molluscs: Helisoma trivolvis, Lymnaea stagnalis (2 species of freshwater snails, Pulmonata), and the sea slug Aplysia californica (Opisthobranchia). All 3 of these preparations have been used extensively for analysis of general biophysical properties of identified neurons, transmitter– receptor interactions (for example, Goldberg and others 1991; Price and Goldberg 1993; Spencer, Lukowiak and others 2000; Jacklet and Koh 2001; Koert and others 2001; Zhao and Klein 2002), and requirements for neurite outgrowth and regeneration (for example, Williams and Cohan 1994; Goldberg 1998; Spira and others 2001; Spira and others 2003; Vanmali and others 2003; Hislop and others 2004; van Diepen and others 2005; van Kesteren and others 2006) and thus extensive overlap exists in the literature allowing direct comparison across species. Each preparation, however, has been channeled into 1 of 3 distinct lines of research. Early embryonic development of the nervous system has been most extensively studied in Helisoma, including early pioneering work on culturing of embryonic molluscan neurons in order to study similarities and differences between neurons at early developmental stages and those of adults (for example, Goldberg 1998). Lymnaea has been used primarily to determine underlying regulatory mechanisms of neuron-specific formation of synapses (Syed and others 1990; Syed and Spencer 1994; Feng and others 1997; Lovell and others 2002; Meems and others 2003). Work on Aplysia, arguably the most studied of the 3 preparations, has focused mostly on synaptic plasticity following synapse formation and how this plasticity relates to learning and memory in the intact brain (for example, Lin and Glanzman 1996; Murphy and Glanzman 1996; Armitage and Siegelbaum 1998; Kandel 2001; Lee and others 2001). It is virtually impossible to overestimate the impact Aplysia cell culture has had on the synaptic underpinnings of learning and memory in the nervous system of all organisms (Kandel 2001). Starting from the classical conditioning of a simple defensive reflex and the identification in situ of the neurons responsible for this behavior (Castellucci and others 1970; Kupferman and others 1970, 1971; Pinsker and others 1970, 1973; Carew and others 1972), researchers were able to remove an individual synaptic component of this network and reform it in 849 appropriate cell cultures. Specifically, a single presynaptic sensory neuron was paired with an identified postsynaptic motoneuron (L7) where it formed an excitatory chemical synapse (Rayport and Schacher 1986). With this model synapse in cell culture, it was possible to demonstrate that changes in synaptic strength correlate directly with habituation or sensitization of the reflex behavior in situ. Moreover, this plasticity of synaptic strength was shown to be differentially modulated by the input of the neurotransmitter serotonin and the neuropeptide FMR Famide (Abrams and others 1984; Rayport and Schacher 1986; Sun and Schacher 1996; Sun and others 1996). Following from this, extensive research has been conducted on both presynaptic and postsynaptic mechanisms of synaptic plasticity (Lin and Glanzman 1996; Zhu and others 1997; Bao and others 1998; Kandel 2001; Jin and Hawkins 2003; Kim and others 2003; Li and others 2005), identification and characterization of the signaling cascades triggered within the neurons (Hu and others 2006), and, more recently, work at the molecular level has begun to identify dynamic changes in gene expression that both lead to and follow from these synaptic changes (Sun and others 2001; Kim and others 2006). The majority of the findings from the work on the reconstructed identified sensory-motor synapse of Aplysia have been directly correlated with similar data from other animal models from invertebrates to higher vertebrates, including humans. With such a large number of researchers using Aplysia and other molluscan neurons as experimental models, the methods used to place these cells in culture have become somewhat standardized. However, along with the many similar techniques utilized between the various preparations, 2 conspicuous technical differences are apparent. First, the physical method of removing neurons from the intact ganglia is different in most laboratories using Aplysia compared to those using Lymnaea or Helisoma. After enzymatic treatment to soften the sheaths of connective tissue surrounding the ganglia in all the preparations, Aplysia neurons are most often removed by penetrating the desired soma with a sharp glass electrode and then either pulling away the rest of the ganglion while holding the cell still or by gently moving the glass electrode away from the ganglion. When the individual neuron is free from the rest of the ganglion it is removed from the tip of the glass electrode and subsequently transferred to the final culture dish with a pipette. Lymnaea and Helisoma neurons, on the other hand, are generally removed from the intact ganglion via a suction pipette with an inner diameter slightly larger than the diameter of the neuronal 850 somata to be removed. This latter technique likely causes less damage to the neuron as the soma is not penetrated with a glass electrode. The sharp-electrode technique, however, is advantageous as the identity of the isolated neuron can be checked electrophysiologically as well as visually in the intact ganglion prior to removal. The second primary difference between Aplysia cell culture techniques and those used for Lymnaea or Helisoma is the nature of the additives to the cell culture media to promote robust neurite outgrowth. Molluscan neurons can survive in culture in the absence of trophic factors but they show little neurite outgrowth. Although the identity of these trophic factors is unknown they are added to the media in 2 different ways. For Aplysia, the trophic factors are added to the media by inclusion of up to 50% hemolymph. For Lymnaea and Helisoma, trophic factors are released into the media by incubating whole ganglia in the culture media for a period of time prior to addition of individual cells. Under these conditions the ganglia are thought to release unidentified soluble factors and factors that bind to the substrate that promote neurite outgrowth from cultured cells. Curiously, Aplysia hemolymph has been shown to promote neurite outgrowth in cultured Lymnaea neurons (Munno and others 2000) and media conditioned by Aplysia brain can induce neurite outgrowth from identified Aplysia neurons (Fejtl and Carpenter 1995). In this study, we will (1) illustrate several key steps in cell culture protocols used in our laboratory, and (2) present an overview centered around the initiation of neurite outgrowth and growth-cone motility in cultured identified neurons from the marine mollusc, A. californica. Although much of this work is an extension of early studies from the 1980s, we have revisited these topics with modern technological approaches and a consideration of new insights in neurobiology over recent years. Furthermore, with the advancement of molecular techniques over recent years (http://www.genome.gov/Pages/Research/ Sequencing/SeqProposals/AplysiaSeq.pdf) and the impending completion of the Aplysia genome (http:// www.genome.gov/13014443), we wish to further establish this model system as a basic tool for studying the underlying genomic control of neurite outgrowth and growth-cone dynamics. Methods: cell culture protocols and in situ hybridization in Aplysia Animals Adult A. californica were obtained from the National Resource for Aplysia (Miami, FL; http://www.rsmas. miami.edu/groups/sea-hares/) and maintained in P. Lovell and L. L. Moroz aquaria within flowing natural seawater. These animals were subjected to a 12 h dark/light cycle and fed red algae twice per week. Animal sizes varied depending on the identity of the neurons to be isolated: 2–5 g for motoneuron L7, 20–40 g for the serotonergic giant cell MCC, and 50–100 g for buccal and pleural sensory neurons. Cell culture Adult Aplysia were anaesthetized by injection of isotonic MgCl2 and pinned down in a dissection tray. Ganglia were removed via a dorsal midline incision, treated with enzyme for 1 h at 34 C in 1% protease IX (Sigma P-6141) in sterile, filtered, natural seawater, briefly rinsed in 4 C sterile seawater, then pinned down in defined media [DM: 1% w/v L-15 Leibovitz medium (Sigma L-4386) in sterile seawater with 20 mg/ml gentamicin sulfate (Sigma G-3632)] containing an additional 10 mM of D-glucose (Sigma G-7528). The sheaths of connective tissue surrounding the ganglia were mechanically removed with fine forceps. Neuronal somata of the desired neurons were visually identified with a dissecting microscope and subsequently removed from the ganglia via suction applied by a gilmont syringe (Gilmont, GS 110, Barrington, IL) attached to a sigmacoated (Sigma SL-2) glass capillary pipette (WPI 1B150-6) held with a micromanipulator (WPI KITE-R). As shown in detail in Figure 1, by carefully balancing suction pressure with axial movement of the pipette tip, individual neurons can be gently removed with a long section of their axon stump still attached to the soma. The glass pipettes were constructed with smooth firepolished tips that had inner opening diameters 10 mm larger than the cell body to be isolated. Cells were individually removed and transferred to falcon 3001 culture dishes with attached poly-L-lysine coated glass coverslips (Sigma P-1399) and maintained in 50% DM 50% filtered hemolymph at room temperature. Hemolymph, prepared fresh for each culture, was removed from large (100 g) animals, pooled, and centrifuged at 10 000 g for 10 min. Following this, the hemolymph was filtered through a 0.2 mm pore size, non-protein-binding syringe filter (acrodisc 32 mm syringe filter with 0.2 mm Supor membrane: Pall Co., UK) to sterilize it and yet maintain soluble proteinaceous trophic factors. Due to their small size (30–50 mm diameter cell body) sensory neurons were first transferred as a cluster onto a non-adhesive surface (uncoated plastic falcon 3001 culture dish), triturated with the sigmacoated pipette to separate individual sensory neurons, then transferred individually to the final culture conditions as above. Aplysia neurons in cell culture 851 Fig. 1 Isolation of individual neurons from the Aplysia CNS. The soma of individual neurons in the abdominal ganglion are visually identified and removed via a silicone-coated glass capillary manufactured to have an interior diameter 10 mm larger than the desired soma and fire polished to remove sharp edges. (A) The tip of the pipette is maneuvered to cover the soma of the selected cell using a micromanipulator and a slight amount of negative pressure is applied with a gilmont syringe. (B) This suction pulls the desired neuronal cell body into the tip of the pipette. At this point, if the suction is reduced and the pipette is pulled away (C) the neuron being extracted will hang loosely from the ganglion but still be attached via the axon stump. (D) The neuron is pulled from the ganglion by applying just enough negative suction to hold the cell body in the tip of the pipette while slowly moving the pipette away from the ganglion along its axis in a smooth, pulling motion. (E) By balancing the suction pressure with movement, the cell can be drawn slightly up into the pipette and the axon severed as it exits the ganglion. (F) At this stage, the neuron is completely removed from the ganglion along with a long segment of axon and is ready for transfer to the desired cell culture. White scale bar ¼ 100 mm. In situ hybridization For fluorescent in situ hybridization (FISH), biotinlabeled antisense b-tubulin RNA probes were obtained by in vitro transcription of a 1374 bp DNA according to the manufacturer’s instructions (Boehringer Mannheim, Indianapolis, IN). Their sense probes were also designed for use as nonspecific controls. For further details see Jezzini and colleagues (2005). Cultured cells were rinsed briefly in artificial seawater (ASW) and fixed in 3.7% formaldehyde in PBS for 30 min at 4 C. The cells were washed 3 times with 0.1 M PBS at room temperature and treated with 1% Triton X100 in PBS. Following this, cells were rinsed in 0.1 M triethanolamine (pH 8.0, plus 0.25% acetic anhydride) for 10 min at room temperature, equilibrated in 50% deionized formamide in 5· saline sodium citrate (SSC, pH 7.2) for 20 min at 42 C, and then hybridized overnight at 42 C in hybridization buffer (50% deionized formamide, 5· SSC, 0.02% SDS, 2% blocking reagent) containing 1.5 mg/ml of the biotin-labeled probes. The following day, unbound probe was removed with two 15 min washes of 2· SSC at 42 C and with 2 washes of 0.1· SSC at 45 C. The cultured neurons were then equilibrated in buffer I (0.1 M Tris–HCl and 0.15 M NaCl) for 2 min followed by buffer II (0.1 M Tris–HCl with 0.15 M NaCl, 0.3% Triton X-100, and 2% normal goat serum) for 30 min at room temperature. Biotin-labeled mRNA 852 was fluorescently tagged by incubation in streptavidin–FITC (Invitrogen, Gaithersburg, MD; 1:200, diluted in buffer II) for 4 h at 4 C. Unbound antibody was washed out via 3 washes with PBS at room temperature. mRNA distribution in cultured neurons was also analyzed using digoxigenin (DIG)-labeled RNA probes. Briefly, DIG-labeled antisense b-tubulin (GB#: AAP13560) RNA probes were reversed-transcribed from linearized cDNA clones (P-GemT vector) using the DIG RNA Labeling Kit (SP6/T7) from Roche Diagnostics (Mannheim, Germany). Cells were processed as above up until the initial overnight hybridization, at which point the 2 techniques diverge. Cells were incubated overnight at 42 C in hybridization buffer containing 1.5 mg/ml of the DIG-labeled RNA probes. Unbound probe was subsequently removed via two 15 min washes in 2· SSC and 2 washes in 0.1· SSC at 42 C. Cells were washed 3 times in PBT (1· PBS, 0.1% Triton X100, 2 mg/ml BSA) at room temperature and then placed in 10% normal goat serum in PBT at 4 C for 30 min. Following this, cells were incubated at 4 C for 1 h in PBT containing 1% goat serum and a 1:2000 dilution of alkalinephosphatase-conjugated DIG-antibodies (Roche Diagnostics). Cells were subsequently washed 3 times in PBT and then twice in detection buffer (100 mM NaCl, 50 mM MgCl2, 0.1% Tween-20, 1 mM levamisol, 100 mM Tris–HCl, pH 9.5). The cultured neurons were then stained by addition of 20 ml/ml NBT/BICP solution from the DIG nucleic acid detection kit (Roche Diagnostics) to the detection buffer in the dark. Development of staining was checked every 15 min and stopped after peak staining was seen (1.5 h). The staining reaction was stopped by incubating the neurons in 4% paraformaldehyde in methanol for 10 min followed by a final wash in 100% ethanol. Immunocytochemistry Cultured Aplysia neurons were labeled using immunocytochemistry to examine b-tubulin and actin cytoskeletal organization in their growth cones. Cultured neurons were washed with Ca2þ-free, low-ionicstrength artificial seawater (100 mM NaCl, 10 mM KCl, 5 mM MgCl2, 15 mM HEPES, 60 g/l glycine, pH 7.9) and then fixed with 3.7% formaldehyde in cytoskeletal stabilization buffer (CBS: 80 mM Pipes, 5 mM EGTA, 1 mM MgCl2, 4% PEG MW 35 000) at 4 C for 20 min. Neurons were subsequently washed 3 times in CBS, extracted with 1% Triton X100, washed 3 times in CBS and left in CBS at 4 C overnight. The next day, cells were washed twice in PBS P. Lovell and L. L. Moroz then incubated in 1:200 diluted monoclonal anti b-tubulin antibody (mAB Tub.21: Sigma) in PBS for 45 min. Following this, cells were washed 4 times in PBS then incubated in AlexaFluor-488 goat antimouse secondary antibody (1:200) and AlexaFluor594 Phalloidin (1:10, Molecular Probes) in PBS for 45 min in the dark. Cells were washed 3 times in 4 C PBS, put in bleach prevention buffer (50% glycerol, 49.8% PBS, 0.212% n-propyl gallate) and imaged. Imaging Cultured neurons were imaged using a Nikon TE2000-E inverted microscope with phase contrast, Nomarski DIC optics, and epi-fluorescence (X-Cite 120 Fluorescence Illuminator system, Exfo.). Images were captured using a Retiga EXi (Q-imaging) digital camera controlled by a PC computer running Metamorph software (version 6.2r4, Universal Imaging Corp.). This system allows high-resolution time-lapse analysis of cultured neurons. Moreover, the integrated morphometric analysis tools included in the imaging software enabled rapid and accurate measurement of neurite outgrowth from the acquired digital images and time-lapse videos. Neurite length was measured at each frame of the time-lapse image stacks from where they exit the soma to their leading edge. Growth-cone movement was measured by tracking changes in position of the center of the growth cone from frame to frame in the time-lapse video stack. Axotomy Neurites of cultured Aplysia neurons were transected using a glass capillary drawn into a fine point (1 mm) mm) held in a micromanipulator. Cultured neurons were first imaged on an inverted Nikon TE2000-E microscope to select neurites for axotomy. The tip of the glass capillary was then positioned next to the selected neurite and adjusted so that it gently contacts the substrate. This tip was then drawn in 1 smooth motion across the neurite perpendicular to the axis of the neurite. Both the proximal (attached to the cell body) and distal (severed) parts of the neurite were then examined using time-lapse video microscopy for subsequent growth-cone formation and initiation of neurite outgrowth. Aplysia neuronal cell culture: an illustrated guide Glial cells Neurons and glia are 2 major classes of cells in the nervous system. Although glial cells significantly outnumber neurons and perform multiple functions, they are investigated less often. There is very little 853 Aplysia neurons in cell culture known about invertebrate glia and even a classification of glial cell types in invertebrates, including mollusks, has hardly begun (Radojcic and Pentreath 1979; Roots 1981; Sonetti and others 1994). Only 1 specific glial marker, a secretory protein Ag, has been described in Aplysia (Lockhart and others 1996) although its functions remain unknown. Indeed, in addition to neurons, the central nervous system of Aplysia contains many other cell types, including glial cells. These cells are often attached to the surface of the neuronal somata and may be transferred to the culture dish along with the neuron. We have found this to be especially true for large abdominal motoneurons and for MCC (see Fig. 2); however, sensory neurons generally have very few, or no, visible glial cells attached to their somata. The presence of glial cells attached to the surface of the identified neurons in cell culture is important and yet often overlooked. When cultured or freshly isolated neurons are fixed, the glial cells will also be fixed and subsequent molecular analysis will be contaminated with glial mRNA. In this sense, all expressed sequence tag (EST) collections from identified molluscan neurons probably also contain sequences from unidentified glial cells. Within a few days in cell culture conditions on a glass substrate coated with poly-L-lysine, the glial cells migrate from the neuronal cell body and attach to the substrate forming a halo around the neuron (Fig. 2A and B). These glial cells are easy to differentiate from the neurons due to their very small relative size (20–30 mm diameter) and their characteristic microtubular organization. Whereas neurons typically have parallel bundles of microtubules (see Fig. 5B), immunocytochemical staining for b-tubulin in glial cells reveals radiating patterns of microtubules extending outward from a common centrally located microtubule-organizing center (Fig. 2C and D). Neuronal morphology and requirements for growth Fig. 2 Glial cells detach from the neuronal cell body and spread out on the substate in cell culture. Large neurons, including MCC and abdominal motoneurons, are often covered with glial cells following removal from the intact ganglion. These glial cells spread out around the neuronal soma (A and B) and attach to the poly-L-lysine substrate within a few days in vitro. Three putative glial cells are marked (*) in panel (B). Immunocytochemical staining for b-tubulin (C and D, green staining) reveals a distinct morphology of the glial cells characterized by microtubules radiating out from a central point (arrow). At least 4 glial cells are fully shown in panel (D). Note a neuronal processes between 2 glial cells in panel (C). White scale bar ¼ 100 mm (A and B), 8 mm (C), 20 mm (D). Although Aplysia neurons (Schacher and Proshansky 1983), along with those of other molluscan species (Wong and others 1981; Wong and others 1984; Feng and others 1997; Lovell and others 2002), can survive several days in cell culture in the absence of extrinsic trophic factors they generally show very little neurite outgrowth. To this end, a long standing and still incompletely answered question in the field of molluscan cell culture is what precise factors are required to induce neurite outgrowth? The general answer to this question today is that to induce robust neurite outgrowth, molluscan neurons need to be exposed to an unknown set of trophic factors. This is accomplished either by adding filtered hemolymph to the media or by creating brain-conditioned media whereby whole ganglia are incubated for a period of time during which they release an unknown set of trophic factors into the media. This media is then filtered and used in subsequent cell cultures. Since the constituents from different media can induce differential growth from the same neurons (Wong and others 1981; Schacher and Proshansky 1983; Hermann and others 2000) as do different physical substrates (Syed and others 1999), all neurons grown in subsequent experiments in this study will use defined media supplemented with hemolymph such that differences can be attributed to intrinsic neuronal properties, not extrinsic variables. When placed in defined media supplemented with hemolymph, identified Aplysia neurons tend to grow in dissimilar ways. Following removal from the intact cerebral ganglion, the giant serotonergic neuron MCC 854 P. Lovell and L. L. Moroz Fig. 3 Aplysia neurons display highly variable morphology in cell culture depending on the identity of the neuron. (A) In cell culture, the large serotonergic neuron MCC typically extends a great number of thin, curving neurites from the distal end of the severed axonal stump resulting in a dense meshwork of new growth. In contrast (B and C) pleural sensory neurons extend far fewer processes directly from their soma; these processes may be straight or branched. (D) Growth cones from MCC neurons are usually wide, flat structures that extend well back into the shaft of the extending neurite while growth cones at the distal ends of pleural sensory neurons (E and F) are more compact structures that taper abruptly into the shaft of their respective neurite. White scale bars ¼ 100 mm in each panel. (Weiss and Kupfermann 1976) typically has a long axonal stump still attached to the cell body (Fig. 3A). New neurite outgrowth generally occurs from the distal end of this axonal stump; however, fine processes are often also seen extending out from the sides of the axon stump. These new processes are typically fine, highly branched, very rarely straight, and contain many small varicosities giving them a bumpy appearance in cell culture (Fig. 3A). When MCC neurons are placed in culture without the long axon stump, they most often die. Those that do survive, however, grow similar thin tangles of neurites extending directly outward from the cell body. In both cases, the growth cones at the leading tips of these processes are wide and long resulting in a long, tapered junction between the end of the new neuritic shaft and the growth cone (Fig. 3D). In contrast, pleural sensory neurons assume a much simpler morphology in cell culture typified by only a few straight processes (between 1 and 3 primary neurites) (Fig. 3B and C). These primary processes often branch away from the cell body; however, they rarely show the complex web of branching produced by MCC neurons. The growth cones at the tip of pleural sensory neurites are small and discrete structures that taper sharply into the shaft of the extending neurite (Fig. 3E and F). Although the final morphology achieved in cell culture is dependent on the identity of the neuron and details of the culture conditions, is neurite outgrowth initiated in a consistent manner? Once initiated, does growth continue indefinitely at a constant rate or does it progress through different stages over time? In our laboratory, we routinely observe that neurite outgrowth from Aplysia neurons in cell culture progresses through a consistent set of phases in L-15 media supplemented with hemolymph as follows: (1) lag (“silent”) phase (first 10–12 h); (2) initiation of growth (10–18 h); (3) phase of intensive growth (18 h to 3 or 4 days); (4) stationary phase (4–12 days); (5) degeneration phase (>10–12 days). Following removal from the intact ganglia and placement in cell culture, Aplysia neurons have a round cell body and, to a greater or lesser degree, an axon stump (Fig. 4). During the first 10–12 h in culture, the axon stump is partially retracted, if present, and little new neurite outgrowth occurs. Around the 12–18 h period, new neurites emerge from the cell body or the remnants of the axon stump (mean time to emergence of first neurite ¼ 17.1 – 5.7 h; n ¼ 10 pleural sensory neurons). We consistently see roughly synchronized sprouting from individual neurons in a single-cell culture dish even if they are plated far apart to reduce the likelihood of signaling between the neurons. From 18 h to 3 or 4 days in culture, Aplysia neurons grow robustly with many active growth-cone-tipped neurites. By Day 4 or Day 5, growth cones disappear to be replaced by blunt, round, phase light varicosities and little new growth will occur. Neurons will stay in this state of arrested growth for up to 2 weeks after which they will start to deteriorate. These phases of neurite 855 Aplysia neurons in cell culture Fig. 4 Neurite outgrowth in cell culture is initiated following a consistent time delay. Following removal from the intact ganglia and placement under appropriate conditions of cell culture, Aplysia sensory neurons maintain a spherical morphology composed of their somata and vestiges of axon stumps. Neurite outgrowth is initiated from the cell body (see inserts at time ¼ 9 h 20 m and 10 h 50 m) following a delay generally occurring 10 h after plating in culture. Once initiated, neurite outgrowth proceeds rapidly with a mean growth rate of 50 mm/h for each neurite tipped by a growth cone. White scale bar ¼ 100 mm. outgrowth are generally consistent between different neurons. In addition, we have found that if neurons are maintained in DM for 24 or 48 h in the absence of trophic factors derived from hemolymph or brain, they fail to initiate neurite outgrowth. Upon addition of hemolymph to the dish, these neurons that had failed to grow in DM alone will sprout robustly after 12 h (P. Lovell, unpublished observations) demonstrating that it takes a consistent time period to respond to extrinsic trophic factors. We have further determined that this hemolymph-induced growth after 48 h in DM can be blocked by the addition of actinomycin D (10 mM, n ¼ 3) to the culture dish indicating that the trophic factor-induced neurite outgrowth requires a transcription-dependent step. Taken together, Aplysia neurons in cell culture appear to initiate growth after a consistent time in cell culture and progress through rapid phases of neurite outgrowth followed by an extended period characterized by little new outgrowth. This initial outgrowth is dependent on the presence of extrinsic trophic factors in the cell culture media. The precise identities of these factors are unknown. Finally, different neurons achieve completely different morphologies when exposed to the same culture conditions. This may be indicative of different inherent growth programs in the different neurons or it may be due to different responses to the same extrinsic growth factors. Growth-cone morphology Although the size and shape of growth cones are highly variable between preparations, between identified neurons from a single animal (for examples from Helisoma see Haydon and others 1985), and even between growth cones extending from a single cell, they often share some common morphological features. Generally, growth cones in Aplysia have thick central (C) domain (Fig. 5A) containing organelles and the distal ends of microtubules (Fig. 5B). At the leading edge, this central domain is fringed by an active motile peripheral (P) domain composed of a veil-like lamellipodium strung between finger-like filopodia. The filopodia and the lamellipodia are distinct structures from one another with the straight ridges formed by the filopodia clearly visible extending through the flat lamellipodium (Fig. 5A). This distinction becomes more evident following immunocytochemical staining of the growth cone for b-tubulin and actin (Fig. 5B). b-Tubulin is packed into tight parallel microtubules in the shaft of the growth cone and splay out in the C domain, although it is rarely 856 seen entering the P domain. The P domain is composed of a mesh of actin in the lamellipodia with straight parallel bundles of actin forming the filopodia (Fig. 5B). Fig. 5 Morphology of Aplysia growth cones in cell culture. (A) Growth cones are generally considered to be composed of 2 functionally distinct domains: a central (C) domain containing organelles and the distal ends of microtubules and a dynamic peripheral domain composed of motile fingerlike filopodia (F) and a veil-like lamellipodia (L). White scale bar ¼ 20 mm. (B) Fluorescent immunocytochemical labeling of the cytoskeletal architecture reveals parallel bundles of b-tubulin (green) in the shaft of the axon that splay out in the C domain (*) with only limited entrance into the P domain (yellow arrow). The P domain is composed almost entirely of actin (red), especially the thin, finger-like filopodial projections (white arrow heads). P. Lovell and L. L. Moroz Aplysia neuronal growth cones in vitro typically range in size from a few microns in diameter for small sensory neurons up to 20 mm diameter for larger motoneurons. Occasionally, giant growth cones are formed from the severed end of the axon stump from large neurons growing in DM supplemented with hemolymph (Fig. 6). These growth cones can be an order of magnitude larger in diameter than normal growth cones and may reach up to 630 mm in diameter (see Fig. 6B). These are the largest growth cones ever reported in the animal kingdom. Moreover, these growth cones are physically large enough to allow direct physiological and molecular access to this specialized cellular compartment. In spite of this size difference, these growth cones are still clearly fringed with an active P domain containing motile lamellipodia and many individual filopodial extensions (Fig. 6A and D). Over time, these giant growth cones have been observed to retract the leading edge leaving behind individual thin neurites tipped with normal-sized growth cones (Fig. 6B,C and F). Following this retraction over the next 24 h, the giant growth cones disappear leaving a dense network of individual neurites extending from the end of the axon stump, a morphology typically seen in older Aplysia cell culture preparations (Schacher and Proshansky 1983; Schacher 1985). Fig. 6 Growth cones of Aplysia neurons can reach giant size in cell culture. Neurite outgrowth from the distal end of an axon stump is occasionally initiated by the formation of a giant growth cone that reaches up to 545 mm in diameter in this cell. These giant growth cones are morphologically similar ones and have a clearly defined core region surrounded by motile filopodia and lamellipodia (A and D). The width of the lamellipodia surrounding the core region is between 8 and 10 mm and is rather constant all around the growth cone. After reaching a critical size, the edges of these giant growth cones frequently pull back leaving thin neurites tipped with small growth cones (white arrows in B and D). Time-lapse ¼ 2.5 h (A to B), 3 h (B to C), 1 h (D to E), and 1 h (E to F). White scale bar ¼ 100 mm in each panel. 857 Aplysia neurons in cell culture Growth-cone dynamics Although we analyzed the morphology of Aplysia growth cones in cell culture (see preceding section) their primary function in situ is to traverse the physical gap between presynaptic and postsynaptic neurons. Therefore, the dynamics of their movement is arguably more important than their overall morphology. Moreover, we asked the following questions: Do growth cones from different identified Aplysia neurons move at the same rate? Do they move at a consistent or variable rate? Do trophic factorinduced and axotomy-induced growth cones move at the same rate? Does growth-cone movement correlate exactly with extension of the neurite? Once growth has begun, we have measured the rate of growth-cone movement of Aplysia neurons in culture using digital time-lapse microscopy (see also Table 1 Dynamics of growth-cone movement in Aplysia and other models Neuron Rate of growth-cone advance (mm/h) Reference Aplysia californica Pleural sensory 51.0 – 18.8 Current study MCC 56.8 – 16.6 Current study Table 1). From 17 neurons (10 sensory, 4 MCC, and 3 L7 motoneuron), we calculated that the mean rate at which the leading edge of the growth cone moves across a poly-L-lysine substrate is 51.0 – 19.1 mm/h at room temperature (Fig. 7A) with short-term maximal growth rates up to 100 mm/h. Looking at different identified neurons, we found that growth cones of sensory neurons and MCC neurons move at a similar rate (52.7 – 18.8 mm/h, n ¼ 10 and 56.8 – 16.6 mm/h, n ¼ 4, respectively; P ¼ 0.7111, unpaired t-test) while growth cones from L7 motoneurons advance at a slightly lower rate (34.2 – 14.5 mm/h, n ¼ 3; P ¼ 0.1487 and P ¼ 0.1199 sensory versus L7 and MCC versus L7, respectively; unpaired t-test). Also, sensory neurons that were held for 48 h in DM alone and subsequently supplemented with 50% hemolymph for 24 h started to grow with growth cones that moved at 50.4 – 15.9 mm/h (n ¼ 2). The large standard deviation in all these measurements is reflective of the way in which these growth cones move along the substrate. We find that growth cones advancing along a smooth unobstructed path on the poly-L-lysine substrate of the culture dish will grow at a rapid rate for a short period of time and intermittently pause. During this pause, filopodia will extend and retract A 250 L7 34.2 – 14.5 Current study Axotomy induced in seawater 65.4 – 22.7 Current study Axotomy induced in hemolymph 69.6 – 33.4 Current study Fasiculating 71.8 – 33.2 Current study 50 4.5 – 2.8 Current study 0 Helisoma trivolvis 8 12 Cohen and Kater (1986) Lymnaea stagnalis 28 Leong and colleagues (2001) Caenorhabditis elegans 20–55 Knobel and colleagues (1999) Rat Dorsal root ganglion 12 Bandtlow and colleagues (1990) Cortical 80 Halloran and Kalil (1994) Sympathetic 90 Buettner and Pittman (1991) FISH Mauthner cells Distance (m) B 100 Jontes and colleagues (2000) *See Figure 13 for additional details; please also note the significantly lower rate of particle retrograde transport in neurites as compared to the measured growth-cone advance in all preparations. Distance (µm) Retrograde particle transport rate in neurite* 200 150 R2 = 0.9792 100 0 50 100 0 20 40 150 200 250 300 350 100 120 140 180 160 140 120 100 80 60 40 20 0 60 80 Time (minutes) Fig. 7 Growth cones move at a high and variable rate. (A) Growth cones from 17 Aplysia neurons were imaged with time-lapse video microscopy and the distance traveled from a fixed starting point was measured. Although highly variable, Aplysia growth cones were found to move at a mean rate of 51.0 – 19.1 mm/h. (B) This variability in rate of neurite extension can even be seen when 2 growth cones from the same neuron are measured (black box with dotted line and white circle with solid line, each represent movement of an individual growth cone from the same cell). 858 P. Lovell and L. L. Moroz mean length of the single process in those cells with only 1 primary process was 712.2 – 193.2 mm. Interestingly, the total length of all neurites in a single cell divided by the number of primary processes in that cell produces a mean growth of 641.0 – 256.3 mm/ neurite. Taken together, this means that regardless of the number of primary processes exiting the cell body, each individual process must grow at approximately the same rate. Since the energy and resources required to grow 2 or 3 processes of equal length must be about 2 or 3 times more than that required to grow only 1 process of that length, the rate of neurite outgrowth must not be limited by energy requirements or other limitations of resources in the cell body and instead must be a function inherent to growth cones themselves. Since the primary function of growth cones is to extend neurites to their specific targets, we closely examined the rate of neurite extension independently from growth-cone movement. Using time-lapse video analysis, we measured the precise length of a neurite from the point at which it left the neuronal cell body to its distal tip at 15 min intervals for 12 h. This neurite exited the cell body (Fig. 8, white arrow), extended for a short distance, looped back on itself, from the P domain of the growth cone and the entire growth cone will move from side to side. Following this pause, the growth cone will begin to advance at a rate similar to that seen prior to the pause although not necessarily in the same direction. The frequency of the pausing behavior is highly variable both between cells and between individual growth cones extending from the same cell body (Fig. 7B). The growth cone represented by the dotted line (Fig. 7B) pauses less frequently than the growth cone represented by the solid line. Thus, although the rate at which they move is similar when they are both moving maximally, the total distance covered by the growth cone represented by the dotted line is far greater after 2 h. In addition to calculating precise rates of growthcone movement with time-lapse video microscopy, we took a random sample of 14 pleural sensory neurons, grew them in DM supplemented with hemolymph for 48 h, and measured the total length of all the neurites formed. From these 14 neurons, 6 neurons had 1 primary neurite extending out from the soma, 6 neurons had 2 primary neurites extending from the soma, and 2 neurons had 3 primary neurites. The mean length of the longest process extending from each neuron was 625.9 – 147.7 mm while the Length of neurite (µm) 350 300 250 200 150 100 50 0 0 100 200 300 400 500 600 700 800 Time (min) Fig. 8 Neurite extension from the cell body proceeds through phases of rapid growth interrupted by extended periods of slower growth. Tracking a single neurite as it emerges from the cell body (white arrow) reveals a complex behavior in terms of both angle and rate of extension. This neurite initially grows away from the cell body, loops back, and then grows away rapidly in a new direction. When the total length of the neurite is plotted against time, 2 distinct phases of extension emerge: a rapid phase of relatively fast and constant extension and a slow phase of little active extension. The 2 dotted lines placed on the graph are parallel to the rapid phases of growth and to each other. White scale bar ¼ 100 mm. 859 Aplysia neurons in cell culture 500 450 400 350 300 250 200 150 100 50 0 C 0 of this neurite and measured the distance it traveled to see how it correlated with neurite extension. This neurite, as mentioned above, exits to the left of the cell body, does a large loop, then extends away at an angle 90 different from the original direction. Using digital analysis of the time-lapse videos (1 frame per 10 min for 12 h), we placed a red “X” at the distal tip of the growth cone in each frame then overlaid the track with the first and last images of the video (Fig. 9A and B). Interestingly, even though this track clearly reveals that the growth cone at the leading tip of the neurite travels over a long serpentine path, the final morphology of the neurite is relatively straight (Fig. 9B). By comparing the distance traveled by the growth cone with the length of the neurite (Fig. 9C) we found that the growth cone traveled over almost twice the distance of the total length of the neurite it was extending. Furthermore, we measured the velocity of the growth cone at each point in time over the 12 h period (Fig. 9D). We found that the point-topoint velocity of the growth cone was highly variable, almost punctate, with maximal rates >100 mm/h interspersed with much slower rates. D 140 Velocity (µm/h) Distance (m) and then extended away from the cell body (Fig. 8). When we graphed the total length of the neurite at each time interval, it became immediately apparent that neurite extension did not occur at a steady state over the entire 12 h period (Fig. 8). Instead, we found that the neurite grew at a fast constant rate for the first 90 min, then grew at a much slower rate for the next 7.5 h. Following this, the neurite resumed a rapid growth rate essentially similar to that seen in the first 90 min. In Figure 8, 2 dotted lines have been placed on the graph. These 2 lines are exactly parallel to each other and their slope (velocity) approximately matches the initial and late phases of neurite extension. While the drop in neurite extension rate at 90 min occurs when the growth cone is in a relatively empty part of the dish, the subsequent re-establishment of extension occurs soon after an interaction between the growth cone at the tip of the measured neurite and a second growth cone on the tip of a different neurite of the same cell. Because neurite extension does not proceed at a constant rate over a period of 12 h (Fig. 8), we examined the precise path traveled by the growth cone 100 200 300 400 500 600 700 800 Time (min) 120 100 80 60 40 20 0 0 100 200 300 400 500 600 700 800 Time (min) Fig. 9 The track of an individual growth cone may be entirely different from the final morphology of the extended neurite. A growth cone that emerges from the left side of a neuron (A) extends over 800 min to a final, relatively straight, position 90 from the initial extension (B). The actual path the growth cone takes to accomplish this growth is indicated by red Xs. (C) To achieve this morphology the actual distance traveled by the growth cone (black squares) is far greater than the length of the neurite that it is extending (white circles). (D) When the precise velocity of the growth cone is calculated for each 10 min interval a highly variable rate of growth is revealed. 860 P. Lovell and L. L. Moroz Overall, we found that Aplysia growth cones are capable of rapid rates of movement across a polyL-lysine substrate with appropriate conditions of cell culture. This rate was similar between the different identified neurons examined. However, the rate of growth-cone movement is not consistent over time. Instead, it is highly variable with long periods of little movement across the substrate interspersed with periods of rapid movement at a relatively consistent rate. An unexpected finding was that neurite extension and growth-cone movement are not somewhat independent. We found that the growth cone at the leading tip of a neurite can traverse long distances without actually leading to lengthening of the neurite. Axotomy-induced neurite outgrowth The rapid formation of new growth cones following axotomy of a neurite in cell culture is a well described phenomenon both in Aplysia and in other preparations. We sought to find out whether these growth cones exhibit growth dynamics similar to those of growth cones that extend out from the cell body and to what extent the formation and movement of these growth cones are dependent on trophic factors in the media. Pleural sensory cells, chosen because they generally have fewer and straighter processes in cell culture as compared to other neuronal types, were grown for 48 h in DM supplemented with hemolymph. A sharp glass electrode was drawn perpendicularly across a single neurite at the level of the substrate thus effectively severing the chosen neurite from the cell body (Fig. 10A and B). Within 5 min postaxotomy new growth cones are formed on both sides of the neurite: on the distal tip of the proximal neurite still attached to the cell body and on the proximal tip of the severed neurite (Fig. 10C). These growth cones continue to elaborate (Fig. 10D) and produce new neurites (Fig. 10E). Although new growth cones are formed on both sides of the cut site, they appear qualitatively distinct from one another in the many dozens of axotomy preparations we have performed. The growth cone formed at the tip of the neurite still attached to the cell body is morphologically similar to those found at the leading tip of most extending neuritis. The growth cone formed by the severed neurite, however, is typically smaller and composed of many fine extensions with little maintenance of active lamellipodial structures. Moreover, with the formation of the growth cone distal to the cut site on the severed neurite, we often see a collapse of the growth cone at the far end of the severed neurite (Fig. 10C and D). While the initiation of neurite outgrowth appears to be dependent on extrinsic trophic support when Fig. 10 New growth cones form rapidly after axotomy. (A) Aplysia pleural sensory neurons normally grow with a relatively simple morphology in cell culture facilitating easy axotomy of individual neurites. Immediately after axotomy with a glass microelectrode: (B) the severed ends, both proximal and distal to the site of the cut, retract slightly. (C) Five minutes postaxotomy, new growth cones appear flanking the cut site. (D) Ten minutes postaxotomy, the new growth cones flanking the site of the cut continue to enlarge and send out new filopodia while the growth cone on the distal end of the severed neurite has retracted its filopodia and has started to collapse. At 30 min postaxotomy (E), the growth cone proximal to the site of the cut is extending along the dish while the activity of the growth cone distal to the site of the cut has resulted in a highly branched morphology. White arrows in each panel point to the same immobile particle on the substrate. Black scale bar in (A) ¼ 100 mm. neurons are first placed in culture (see above), the formation of growth cones and subsequent neurite outgrowth following axotomy are not dependent on trophic factors. Pleural sensory neurons were grown in DM supplemented with hemolymph for 48 h following which time the media was replaced with filtered seawater. After an hour in seawater, neurites were selected, axotomized, and the resulting formation of growth cones assayed. All pleural sensory neurons in seawater formed growth cones both on the attached neurites and on severed neurites similar to those on neurites axotomized in the presence of hemolymph. We calculated mean growth rates of 861 Aplysia neurons in cell culture Fig. 11 Severed neurites can survive and grow in cell culture in the absence of the cell body. After removal from the intact ganglion and placement in cell culture, the cell body of an abdominal ganglia motor neuron was removed leaving behind a long piece of neurite attached to the substrate. (A) After 18 h in cell culture in the absence of the somata (represented by the dotted circle), new growth is clearly seen from the distal ends of the severed neurite. Little new growth is seen from the end of the neurite, however, where it was previously attached to the cell body. (B) With an additional 48 h in cell culture, new growth (white arrows) continues to emerge from the isolated neurite. White scale bar ¼ 100 mm. axotomy-induced growth cones on the poly-L-lysine substrate in seawater and hemolymph and found them to be nearly identical (65.4 – 22.7 mm/h, n ¼ 7 and 69.6 – 33.4 mm/h, n ¼ 3, respectively; P ¼ 0.8194, unpaired t-test). The growth cones that extend from the neurite attached to the cell body often traverse the gap created by the axotomy, fasiculate onto the severed neurite, and grow along the surface of the neurite. In this situation we calculated mean growth rates of 71.7 – 18.7 mm/h for neurons in seawater (n ¼ 5) and 71.8 – 33.2 mm/h (n ¼ 5) for neurons in DM supplemented with hemolymph. Along with overcoming the dependence of neurite outgrowth on extrinsic factors, axotomy also appears to be able to promote new growth in older cultures of neurons that had ceased active growth. Pleural sensory neurons were maintained in cell culture for 5 days upon which time they achieved a typical morphology, exhibiting few active growth cones, many phase-bright varicosities, and little evidence of active neurite outgrowth. Growth-cone formation following axotomy in these preparations (n ¼ 3) appeared similar to those in cultures only 48 h old. Growth cones formed within 5 min and rapid neurite outgrowth was observed for the next 2 h. To complement the above experiments, we isolated neurites in the absence of their cell bodies and placed them in DM supplemented with hemolymph. These neurites not only survived for up to a week but also exhibited new neurite outgrowth (Fig. 11). Taken as a whole, we found that axotomy of Aplysia neurites in cell culture leads to the rapid formation of new growth cones both proximal and distal to the site of the cut. In addition, this formation of new growth cones, and the movement of these growth cones and the subsequent extension of the neurites did not require extrinsic trophic factors or de novo protein synthesis. Lastly, axotomy was found to induce new outgrowth in neurons that had previously ceased to grow after 5 days in culture. Extrasomatic localization of mRNA Over the past decade, the emerging concept of extrasomatic protein synthesis in polarized cells, such as neurons, has revolutionized modern neurobiology (Kaplan and others 2004; Martin 2004; Verma and others 2005; Wu and others 2005; Lyles and others 2006; van Kesteren and others 2006; Willis and Twiss 2006). For protein synthesis to occur in cellular compartments far removed from the cell body, such as growth cones, mRNA must be present in these compartments to act as the template for translation. Furthermore, it is expected that only a specific subset of all mRNA transcripts in a cell will be located extrasomatically. We investigated the extrasomatic subcellular location of b-tubulin mRNA in cultured 862 P. Lovell and L. L. Moroz Fig. 12 Extrasomatic mRNA is localized in specific morphologically distinct subcompartments outside the cell body. Although mRNA localization and protein synthesis is generally considered to be limited to the cell body, recent evidence suggests that this may not be the case for highly polarized cells such as neurons. A certain degree of diffusion of mRNA into neurites from big somata might be expected. If present, such passive diffusion should be associated with a clear positive gradient in the distribution of certain mRNAs, with the highest mRNA concentration near the somata. In contrast, a negative gradient, patched distribution or even uniform distribution will be indicators of specific mRNA transport. Experiments using FISH on isolated sensory neurons and MCC neurons have shown that the mRNAs for b-tubulin are not uniformly distributed along neurites in dissociated cell culture with a clear negative gradient, predominant localization of transcripts in the base of the growth cones, and somewhat patchy distribution in distant processes. (A) and (B) In situ hybridization with DIG-labeled probes against b-tubulin in cultured Aplysia buccal sensory neurons (A and B) and random abdominal motoneurons (E and F) reveals dark blue staining, indicative of the localization of b-tubulin mRNA, at the end of the axon stump (red arrow). Similar dark staining, above the level of background staining, can be seen in varicosities along the shaft of neurites (E, black arrows). Discrete patches of darker staining can be seen in areas along the shaft of smooth, thick neurites (F, black arrows). High magnification FISH [the antisense probes labeled with biotin–streptavidin–FITS, panel (C)] against b-tubulin reveals punctate staining in growth cones. (D) In situ hybridization with sense (control) FISH probes reveals no labeling in neurites and growth cones (white line outlines the end of the neurite). Aplysia neurons using in situ hybridization. We found that b-tubulin mRNA is not diffusely distributed throughout the extrasomatic cytosol. Instead, it is found clustered in discrete sites. Strong staining for b-tubulin mRNA was found at the distal end of the axon stump (Fig. 12A and B), in varicosities along the shaft of thin neurites (Fig. 12E), and in isolated patches along the shaft of large, smooth neurites (Fig. 12F). Although the comparatively dark staining in varicosities as opposed to the thin intervening neurite (see Fig. 12E) may be considered an artifact due to the thicker diameter of varicosities, we feel it represents true differences in mRNA localization since discrete darker areas are also seen along the shaft of a neurite of constant diameter (see arrows Fig. 12F). At higher magnification, FISH revealed punctate staining in distal tips of neurites with some clustering at bifurcation points along the neurite (Fig. 12C). Control preparations exposed to sense b-tubulin probes show little background staining (Fig. 12D). Aplysia neurons in cell culture 863 Fig. 13 Retrograde movement of vesicles along a neurite. Time-lapse video analysis coupled with Nomarski DIC optics allows for precise measurement of visible particles and vesicles as they are transported along a neurite. From a section of neurite attached to a pleural sensory neuron (A), the location of a single vesicle can be determined over time [see arrows in (C)]. The distance traveled by 4 visible particles over time (B) demonstrate a relatively consistent rate of retrograde transport at 4.5 – 2.8 mm/h. White scale bar ¼ 50 mm (A), 20 mm (C). Discussion and comparison with other species Growth-factor dependence A generally accepted concept in neurobiology is that adult neurons need extrinsic trophic factors in order to initiate neurite outgrowth. For vertebrate neurons this requirement is extended even further in that extrinsic trophic factors are necessary both for neurite outgrowth and for overall neuronal survival (Goldberg 2003). Molluscan neurons, on the other hand, can survive adequately in the absence of extrinsic trophic factors but produce little in the way of new outgrowth. Moreover, what little growth is produced from molluscan neurons in the absence of extrinsic trophic factors is morphologically quite different from the long thin processes produced under trophic factorrich conditions. The wide veil-like processes produced by neurons devoid of trophic factor support are composed of tangled arrays of microtubules as opposed to the parallel bundles of microtubules found in the axonal shafts of normally growing neurons (P. Lovell, unpublished observations). Hemolymph is the most common source of trophic factors traditionally used for Aplysia neurons (Schacher and Proshansky 1983) while other molluscan preparations typically use brain-derived trophic factors (Syed and others 1990; Syed and Spencer 1994; Goldberg 1998; Munno and others 2000; Geddis and others 2004). In either case, the identity of the trophic factors within the respective media has remained elusive despite many efforts in various laboratories to identify them. For example, epidermal growth factor (EGF) has been cloned from L. stagnalis and been shown to be sufficient to induce neurite outgrowth in cultured neurons (Wildering and others 2001). Careful chemical analysis, however, has been unable to demonstrate its presence in brainconditioned media. While this does not conclusively mean that EGF is not present in the media, it is suggestive that it is not the primary growth-inducing component. More likely, both media are composed of a cocktail of factors that work either synergistically or may serve overlapping redundant functions, thus making the interpretation of fractionation experiments difficult. Interestingly, it has been shown that trophic factors isolated from dialyzed Aplysia hemolymph are sufficient to induce robust neurite outgrowth from cultured Lymnaea neurons (Munno and others 2000), indicating the presence of similar factors 864 in both media that are conserved across species. The specific trophic factor(s) in media derived from hemolymph or brain still require identification before an understanding is reached of exactly how neurite outgrowth is induced in adult molluscan neurons. Although the identity of the trophic factors is unknown, their addition to the cell culture media consistently produces neurite outgrowth. This initiation of neurite outgrowth follows a strict temporal sequence. Aplysia neurons placed in media containing 50% hemolymph show little or no neurite outgrowth for 10–12 h but then undergoes rapid growth. If the neurons are held for 48 h in the absence of trophic factors and then supplied with hemolymph, they show the same 10–12 h time lag before neurite outgrowth is initiated. We have observed a similar delay in the initiation of neurite outgrowth from freshly isolated Lymnaea neurons placed in brain-derived media (P. Lovell, unpublished observations). At first, we thought that this delay might reflect a refractory period, possibly related to recovery from damage after removal from the intact ganglion, during which the neuron was incapable of growth. However, since a similar time lag between application of trophic factors and the outgrowth of neurites is seen in neurons that are held for 48 h without trophic factor support, it seems doubtful that the delay is related to the initial damage. More likely, the delay in the initiation of neurite outgrowth may be related to the time it takes to turn on a cellular program of active neurite outgrowth in an adult neuron. This idea is supported by the finding that transcription is necessary for trophic factor-induced neurite outgrowth. Furthermore, adult molluscan neurons are known to differ from those of embryonic molluscs in that the embryonic neurons do not need extrinsic trophic support to grow in cell culture (Goldberg and others 1988). Thus, while embryonic neurons may be in a default “grow” cellular program, adult neurons that have formed synapses with their respective targets would likely be in a completely different physiological state. A manifestation of this may be seen in cell culture in that by Day 4 or Day 5 molluscan neurons typically cease to grow. It remains to be seen whether Aplysia neurons, which have stopped growing on Day 5 and then removed from the primary culture dish and re-plated in a second dish, are capable of subsequent outgrowth and if so whether this follows the same temporal sequence. Axotomy In contrast to trophic factor-induced neurite outgrowth, axotomy-induced formation of new growth P. Lovell and L. L. Moroz cones and subsequent outgrowth shows little lag in time. Within minutes after transection of a neurite in vitro, a large, active growth cone forms proximal to the cut site and initiates new outgrowth. Since the times taken to induce the formation of a new growth cone, via extrinsic trophic factors and via axotomy, are so different, it is likely that 2 completely separate signaling pathways converge on the same cellular program, namely growth-cone formation and neurite outgrowth. Extensive work has shown that local Ca2þ influx at the site of the cut triggers a calpaindependent breakdown and reorganization of the local cytoskeleton to transform the neurite shaft into a functional growth cone (Spira and others 2001; Spira and others 2003). In vertebrate preparations, the formation of a growth cone after axotomy requires de novo protein synthesis (Verma and others 2005). Pilot experiments in our laboratory, however, indicate that this may not be the case for Aplysia neurons. When anisomycin, a potent protein synthesis blocker commonly used in molluscan preparations (Khalsa and others 1992; Alkon and others 2005) is added to the culture media 1 h prior to neurite transection, new growth cones still form reliably at the site of the cut (P. Lovell, unpublished observations). Furthermore, transcription of new mRNA is clearly not necessary for axotomy-induced growth-cone formation since growth cones form on the severed neurite distal to the site of the cut. Finally, axotomy can induce growthcone formation and neurite outgrowth in the absence of extrinsic trophic factors and is equally effective on neurons that have ceased to grow after 5 days in culture. Potentially, this highlights the difference between regenerative neurite outgrowth after damage and induction of neurite outgrowth during development, 2 processes that share a common output but are regulated in completely different ways, a concept that recently has also arisen in regard to vertebrate preparations (Goldberg 2003). Growth rate We measured the rate of movement of growth cones growing along a poly-L-lysine substrate to be 50 mm/h (for a summary see Table 1). This growth rate is faster than previously reported for other molluscs [8–12 mm/h for H. trivolvis (Cohan and Kater 1986), 28 mm/h for L. stagnalis (Leong and others 2001)] cultured under similar conditions. Neurons from Caenorhabditis elegans have been shown to grow at 20–55 mm/h in vivo (Knobel and others 1999). Work with vertebrate neurons have revealed growth rates from 12 mm/h (rat dorsal root ganglion) (Bandtlow and others 1990) up to 80 mm/h 865 Aplysia neurons in cell culture (cortical neurons) (Halloran and Kalil 1994), 90 mm/h (sympathetic neurons) (Buettner and Pittman 1991), and 100 mm/h (fish: Mauthner neurons) (Jontes and others 2000). While it is difficult to compare our measured growth rates to those seen in vertebrates due to vast differences in culture conditions, especially the higher temperature used in vertebrate preparations, our measurements are notably higher even than those seen in other molluscs cultured under similar conditions. This difference may be partially due to the fact that different neurons grow at different rates or it may reflect a measurement artifact arising from the time interval at which growth rate was calculated. We have observed that Aplysia growth cones in cell culture grow in a highly phasic fashion with periods of constant rapid growth randomly interrupted by periods of no measurable forward growth. During these pauses, the Aplysia growth cones will often meander slightly from side to side and filopodia will be extended and retracted as though the growth cone is looking for guidance cues in the environment. With this in mind, while the growth cone is actively moving it will travel at rates exceeding 50 mm/h, although, the total distance between the initial and final points after 1 h may only be 20 mm. Thus, earlier studies that calculated growth rates from a lower sampling frequency may have underestimated how fast growth cones can move when they are actually moving. This is further supported by our observation that when Aplysia growth cones fasciculate on a neurite that already exists in the dish, and thus tend to pause and search for guidance cues less frequently, we measure growth rates of 70 mm/h. When we measured movement rates for multiple growth cones growing out from a common neurite branch point we found little difference from those growth cones at the tip of an unbranched neurite. Additionally, when we let pleural sensory neurons grow for 48 h and subsequently calculated the amount of outgrowth per primary neurite we found little difference in length between those neurons with 1 primary neurite and those with 2 or 3. Starting from the spherical cell body, growing 3 primary neurites each at the same rate must cost 3 times the metabolic energy of growing a single neurite at that same rate. Therefore, the overall cellular energy demands of the neuron must not be a rate-limiting step that determines either the speed of growth-cone movement or the subsequent rate of neurite extension. More likely, the rate of growth cone advance is an inherent characteristic of the growth cone itself and possibly dependent on such factors as maximum rate of actin assembly/disassembly in the filopodia and lamellipodia, microtubule extension, membrane insertion, and rate of anterograde transport of required molecules along the neurite. Supporting this idea is the fact that the leading edge of the lamellipodia of the giant (>500 mm diameter) growth cones we have observed advance at a rate of 60 mm/h. These growth cones are somewhat unique in that they only occasionally form at the tip of the axon stump of large neurons, they expand up to a giant size and they retract, leaving behind fine processes tipped with normally sized growth cones. The filopodia and lamellipodia of these growth cones must use the same molecular machinery (that is, actin dynamics) to move. Thus the 60 mm/h forward movement of filopodia/lamellipodia may be the default speed for this system under these cell culture conditions. Isolated neuronal processes and extrasomatic mRNAs Isolated neurites from a variety of invertebrate preparations have been shown to survive for long periods of time in vivo despite the absence of a functional cell body (Scheibenstock and others 2002). This situation is paralleled in cell culture where severed neurites can survive for days, exhibit new outgrowth, and have a limited capacity for synapse formation (Meems and others 2003). This autonomous behavior of the severed neurites and the growth cones emanating from them raises an important fundamental neurobiological question challenging a central dogma of cell biology: given the short half-life of most proteins, how can severed neurites grow and survive in the absence of the cell body if de novo protein synthesis can only occur in the cell body? Recently, it has been shown that extrasomatic regions of neurons, including growth cones, contain mRNA (Giuditta and others 2002; Moccia and others 2003; Gioio and others 2004; Moroz and others 2006) and are capable of localized de novo protein synthesis (Spencer, Syed and others 2000). While it is tempting to hypothesize that local protein synthesis in the growth cone is an important regulatory step in control of neurite outgrowth and synapse formation [for review see Martin (2004)], direct evidence demonstrating a functional contribution of locally synthesized proteins to neuronal outgrowth is just starting to emerge. mRNA encoding for b-thymosin, a small protein that binds and sequesters monomeric actin (Huff and others 2001) has been localized in extrasomatic regions of retinal ganglion cells (Roth and others 1999), Lymnaea pedal neurons (van Kesteren and others 2006), and Aplysia neurons (Moccia and others 2003). Knockdown of b-thymosin in cultured neurons, as well as in isolated neurites, using RNAi significantly enhanced neurite 866 outgrowth (van Kesteren and others 2006). Moreover, local translation of RhoA, a small GTPase that regulates the actin cytoskeleton upstream of actinbinding proteins, has been shown to be necessary for Sema3A-mediated growth-cone collapse in Xenopus retinal axons (Wu and others 2005), and regeneration of growth cones following axotomy requires localized de novo protein synthesis (Verma and others 2005). It is clear that mRNA is both present and locally translated in growth cones. However, an understanding of how this mRNA pool is selected from the somatic mRNA pool, how it is transported to the growth cone, how translation of this extrasomatic mRNA into protein is regulated, and what roles these mRNAs play in neurite outgrowth, growth-cone guidance, synaptic target selection, and synaptogenesis, is lacking. In part, this is due to the lack of broad-scale genomic tools that are applicable to model systems with individually identifiable neurons. Commercially obtainable microarrays are available for various vertebrate model systems and have been used to investigate gene-expression profiles during neuronal regeneration (Fundin and others 2002; Dabrowski and others 2003; Nilsson and others 2005). However, the small size of vertebrate neurons and the lack of individually identifiable cells limit the applicability of this technique. The heterogeneity of the neurons used in these studies, and thus potential transcriptome differences due to neuronal type, potentially overshadows transcriptome changes specifically related to neurite outgrowth. Conversely, progress with invertebrate preparations with large identified neurons has been limited to extrasomatic EST collection (Moccia and others 2003; Moroz and others 2006), differential display PCR (Gioio and others 2004) and conventional library-screening techniques (van Kesteren and others 2006). This bottleneck, however, has currently been overcome by obtaining 200 000 ESTs from the CNS of Aplysia (Moroz and others, 2006), the development of new tools for introducing charged molecules such as mRNA into subcellular compartments of cultured Aplysia neurons (Lovell and others 2006) and ongoing Aplysia genome sequencing (http://www.genome.gov/Pages/ Research/Sequencing/SeqProposals/AplysiaSeq.pdf). Acknowledgments Our work is supported by NSF and National Institutes of Health Center of Excellence in Genomic Science Grants P50 HG002806 and R01 NS39103 and in part by McKnight Brain Research Foundations and UF Opportunity Funds. We also would like to thank Mrs E. Bobkova, Drs A. Kohn, N. Alieva for help in P. Lovell and L. L. Moroz molecular biology techniques, in situ hybridization protocols, immunohistochemistry, and confocal microscopy. Conflict of interest: None declared. References Abrams TW, Castellucci VF, Camardo JS, Kandel ER, Lloyd PE. 1984. 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