Download The largest growth cones in the animal kingdom

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. Two endogenous neuropeptides modulate the gill and
siphon withdrawal reflex in Aplysia by presynaptic facilitation involving camp-dependent closure of a serotoninsensitive potassium channel. Proc Natl Acad Sci USA
81:7956–60.
Aletta JM, Greene LA. 1988. Growth cone configuration and
advance—a time-lapse study using video-enhanced differential interference contrast microscopy. J Neurosci 8:
1425–35.
Alkon DL, Epstein H, Kuzirian A, Bennett MC, Nelson TJ.
2005. Protein synthesis required for long-term memory is
induced by PKC activation on days before associative
learning. Proc Natl Acad Sci USA 102:16432–7.
Armitage BA, Siegelbaum SA. 1998. Presynaptic induction and
expression of homosynaptic depression at Aplysia sensorimotor neuron synapses. J Neurosci 18:8770–9.
Bandtlow C, Zachleder T, Schwab ME. 1990. Oligodendrocytes
arrest neurite growth by contact inhibition. J Neurosci
10:3837–48.
Bao JX, Kandel ER, Hawkins RD. 1998. Involvement of
presynaptic and postsynaptic mechanisms in a cellular
analog of classical conditioning at Aplysia sensory-motor
neuron synapses in isolated cell culture. J Neurosci
18:458–66.
Bray D, Chapman K. 1985. Analysis of microspike movements
on the neuronal growth cone. J Neurosci 5:3204–13.
Buettner HM, Pittman RN. 1991. Quantitative effects of
laminin concentration on neurite outgrowth in vitro.
Dev Biol 145:266–76.
Burmeister DW, Rivas RJ, Goldberg DJ. 1991. Substrate-bound
factors stimulate engorgement of growth cone lamellipodia
during neurite elongation. Cell Motil Cytoskeleton 19:
255–68.
Carew TJ, Pinsker HM, Kandel ER. 1972. Long-term
habituation of a defensive withdrawal reflex in Aplysia.
Science 175:451–4.
Castellucci V, Pinsker H, Kupferman I, Kandel ER. 1970.
Neuronal mechanisms of habituation and dishabituation of
gill-withdrawal reflex in Aplysia. Science 167:1745–8.
Cohan CS, Kater SB. 1986. Suppression of neurite elongation
and growth cone motility by electrical activity. Science
232:1638–40.
Cohan CS, Welnhofer EA, Zhao L, Matsumura F, Yamashiro S.
2001. Role of the actin bundling protein fascin in growth
cone morphogenesis: localization in filopodia and lamellipodia. Cell Motil Cytoskeleton 48:109–20.
Dabrowski M, Aerts S, Van Hummelen P, Craessaerts K,
De Moor B, Annaert W, Moreau Y, De Strooper B. 2003.
867
Aplysia neurons in cell culture
Gene profiling of hippocampal
J Neurochem 85:1279–88.
neuronal
culture.
Dent EW, Gertler FB. 2003. Cytoskeletal dynamics and
transport in growth cone motility and axon guidance.
Neuron 40:209–27.
Fejtl M, Carpenter DO. 1995. Neurite outgrowth is enhanced
by conditioning factor(s) released from central ganglia of
Aplysia californica. Neurosci Lett 199:33–6.
Feng ZP, Klumperman J, Lukowiak K, Syed NI. 1997. In vitro
synaptogenesis between the somata of identified Lymnaea
neurons requires protein synthesis but not extrinsic growth
factors or substrate adhesion molecules. J Neurosci
17:7839–49.
Fundin BT, Rice FL, Ernfors P. 2002. Patterned gene programs
and target remodeling following axotomy at a major site for
sensory innervation. J Neurobiol 53:370–80.
Geddis MS, Tornieri K, Giesecke A, Rehder V. 2004. PLA(2)
and secondary metabolites of arachidonic acid control
filopodial behavior in neuronal growth cones. Cell Motil
Cytoskeleton 57:53–67.
Gioio AE, Lavina ZS, Jurkovicova D, Zhang H, Eyman M,
Giuditta A, Kaplan BB. 2004. Nerve terminals of squid
photoreceptor neurons contain a heterogeneous population
of mRNAs and translate a transfected reporter mRNA. Eur J
Neurosci 20:865–72.
Giuditta A, Kaplan BB, van Minnen J, Alvarez J, Koenig E.
2002. Axonal and presynaptic protein synthesis: new insights
into the biology of the neuron. Trends Neurosci 25:400–4.
Goldberg DJ, Burmeister DW. 1986. Stages in axon
formation—observations of growth of Aplysia axons in
culture using video-enhanced contrast-differential interference contrast microscopy. J Cell Biol 103:1921–31.
Goldberg DJ, Foley MS, Tang DJ, Grabham PW. 2000.
Recruitment of the Arp2/3 complex and Mena for the
stimulation of actin polymerization in growth cones by nerve
growth factor. J Neurosci Res 60:458–67.
Haydon PG, Cohan CS, McCobb DP, Miller HR, Kater SB.
1985. Neuron-specific growth cone properties as seen in
identified neurons of Helisoma. J Neurosci Res 13:135–47.
Hermann PM, van Kesteren RE, Wildering WC, Painter SD,
Reno JM, Smith JS, Kumar SB, Geraerts WPM, Ericsson LH,
Smit AB and others. 2000. Neurotrophic actions of a novel
molluscan epidermal growth factor. J Neurosci 20:6355–64.
Hislop J, Dyer JR, Scott D, van Kesteren RE, Sossin WS. 2004.
Characterization of a novel molluskan tyrosine kinase
receptor that inhibits neurite regeneration. J Neurobiol
60:127–36.
Hopker VH, Shewan D, Tessier-Lavigne M, Poo MM, Holt C.
1999. Growth-cone attraction to netrin-1 is converted to
repulsion by laminin-1. Nature 401:69–73.
Hu JY, Wu F, Schacher S. 2006. Two signaling pathways
regulate the expression and secretion of a neuropeptide
required for long-term facilitation in Aplysia. J Neurosci
26:1026–35.
Huber AB, Kolodkin AL, Ginty DD, Cloutier JF. 2003.
Signaling at the growth cone: ligand-receptor complexes
and the control of axon growth and guidance. Annu Rev
Neurosci 26:509–63.
Huff T, Muller CSG, Otto AM, Netzker R, Hannappel E. 2001.
Beta-thymosins, small acidic peptides with multiple functions. Int J Biochem Cell Biol 33:205–20.
Jacklet JW, Koh HY. 2001. Nitric oxide as an orthograde
cotransmitter at central synapses of Aplysia: responses of
isolated neurons in culture. Am Zool 41:282–91.
Jezzini SH, Bodnarova M, Moroz LL. 2005. Two-color in situ
hybridization in the CNS of Aplysia californica. J Neurosci
Methods 149:15–25.
Jin I, Hawkins RD. 2003. Presynaptic and postsynaptic
mechanisms of a novel form of homosynaptic potentiation
at Aplysia sensory-motor neuron synapses. J Neurosci
23:7288–97.
Goldberg JI. 1998. Serotonin regulation of neurite outgrowth in
identified neurons from mature and embryonic Helisoma
trivolvis. Perspect Dev Neurobiol 5:373–87.
Jontes JD, Buchanan J, Smith SJ. 2000. Growth cone and
dendrite dynamics in zebrafish embryos: early events in
synaptogenesis imaged in vivo. Nat Neurosci 3:231–7.
Goldberg JI, McCobb DP, Guthrie PB, Lawton RA, Lee RE,
Kater SB. 1988. Characterization of cultured embryonic
neurons from the snail Helisoma. In: Beadle, Lees, Kater,
editors. Cell culture approaches to invertebrate neuroscience.
p. 85–108.
Kandel ER. 2001. Neuroscience—the molecular biology of
memory storage: a dialogue between genes and synapses.
Science 294:1030–8.
Goldberg JI, Mills LR, Kater SB. 1991. Novel effects of
serotonin on neurite outgrowth in neurons cultured from
embryos of Helisoma trivolvis. J Neurobiol 22:182–94.
Goldberg JL. 2003. How does an axon grow? Genes Dev
17:941–58.
Kaplan BB, Lavina ZS, Gioio AE. 2004. Subcellular compartmentation of neuronal protein synthesis: new insights into
the biology of the neuron. Ann N Y Acad Sci 1018:244–54.
Khalsa SBS, Whitmore D, Block GD. 1992. Stopping the
circadian pacemaker with inhibitors of protein synthesis.
Proc Natl Acad Sci USA 89:10862–6.
Gomez TM, Robles E, Poo MM, Spitzer NC. 2001. Filopodial
calcium transients promote substrate-dependent growth
cone turning. Science 291:1983–7.
Kim H, Lee SH, Han JH, Lee JA, Cheang YH, Chang DJ,
Lee YS, Kaang BK. 2006. A nucleolar protein ApLLP induces
ApC/EBP expression required for long-term synaptic
facilitation in Aplysia neurons. Neuron 49:707–18.
Halloran MC, Kalil K. 1994. Dynamic behaviors of growth
cones extending in the corpus callosum of living cortical
brain-slices observed with video microscopy. J Neurosci
14:2161–77.
Kim JH, Udo H, Li HL, Youn TY, Chen M, Kandel ER,
Bailey CH. 2003. Presynaptic activation of silent synapses
and growth of new synapses contribute to intermediate and
long-term facilitation in Aplysia. Neuron 40:151–65.
868
P. Lovell and L. L. Moroz
Knobel KM, Jorgensen EM, Bastiani MJ. 1999. Growth cones
stall and collapse during axon outgrowth in Caenorhabditis
elegans. Development 126:4489–98.
Meberg PJ, Bamburg JR. 2000. Increase in neurite outgrowth
mediated by overexpression of actin depolymerizing factor.
J Neurosci 20:2459–69.
Koert CE, Spencer GE, van Minnen J, Li KW, Geraerts WPM,
Syed NI, Smit AB, van Kesteren RE. 2001. Functional
implications of neurotransmitter expression during axonal
regeneration: serotonin, but not peptides, auto-regulate axon
growth of an identified central neuron. J Neurosci
21:5597–606.
Meems R, Munno D, van Minnen J, Syed NI. 2003. Synapse
formation between isolated axons requires presynaptic soma
and redistribution of postsynaptic AChRs. J Neurophysiol
89:2611–19.
Kupferman I, Castellucci V, Pinsker H, Kandel E. 1970.
Neuronal correlates of habituation and dishabituation of the
gill-withdrawal reflex in Aplysia. Science 167:1743–5.
Kupferman I, Pinsker H, Castellucci V, Kandel ER. 1971.
Central and peripheral control of gill movements in Aplysia.
Science 174:1252–6.
Moccia R, Chen D, Lyles V, Kapuya EEY, Kalachikov S,
Spahn CMT, Frank J, Kandel ER, Barad M, Martin KC. 2003.
An unbiased cDNA library prepared from isolated Aplysia
sensory neuron processes is enriched for cytoskeletal and
translational mRNAs. J Neurosci 23:9409–17.
Lee JA, Kim HK, Kim KH, Han JH, Lee YS, Lim CS, Chang DJ,
Kubo T, Kaang BK. 2001. Overexpression of and RNA
interference with the CCAAT enhancer-binding protein on
long-term facilitation of Aplysia sensory to motor synapses.
Learn Mem 8:220–6.
Moroz LL, Edwards JR, Puthanveettil SV, Kohn AB, Ha T,
Heyland A, Knudsen B, Sahni A, Yu F, Liu L, Jezzini S,
Lovell P, Iannucculli W, Chen M, Nguyen T, Sheng H,
Shaw R, Kalachikov S, Panchin YV, Farmerie W, Russo JJ,
Ju J, Kandel ER. 2006. The neuronal transcriptome of Aplysia
californica: A platform for the neurogenomics of defined
neurons, neuronal compartments and neuronal circuitry.
Cell. Forthcoming.
Leong CCW, Syed NI, Lorscheider FL. 2001. Retrograde
degeneration of neurite membrane structural integrity of
nerve growth cones following in vitro exposure to mercury.
Neuroreport 12:733–7.
Munno DW, Woodin MA, Lukowiak K, Syed NI,
Dickinson PS. 2000. Different extrinsic trophic factors
regulate neurite outgrowth and synapse formation between
identified Lymnaea neurons. J Neurobiol 44:20–30.
Letourneau PC, Shattuck TA. 1989. Distribution and possible
interactions of actin-associated proteins and cell adhesion
molecules of nerve growth cones. Development 105:505–19.
Murphy GG, Glanzman DL. 1996. Enhancement of sensorimotor connections by conditioning-related stimulation in
Aplysia depends upon postsynaptic Ca2þ. Proc Natl Acad
Sci USA 93:9931–6.
Li Q, Roberts AC, Glanzman DL. 2005. Synaptic facilitation
and behavioral dishabituation in Aplysia: dependence on
release of Ca2þ from postsynaptic intracellular stores,
postsynaptic exocytosis, and modulation of postsynaptic
AMPA receptor efficacy. J Neurosci 25:5623–37.
Lin XY, Glanzman DL. 1996. Long-term depression of Aplysia
sensorimotor synapses in cell culture: inductive role of a rise
in postsynaptic calcium. J Neurophysiol 76:2111–14.
Lockhart ST, Levitan IB, Pikielny CW. 1996. Ag, a novel
protein secreted from Aplysia glia. J Neurobiol 29:35–48.
Lohof AM, Quillan M, Dan Y, Poo MM. 1992. Asymmetric
modulation of cytosolic Camp activity induces growth cone
turning. J Neurosci 12:1253–61.
Lovell P, Jezzini SH, Moroz LL. 2006. Electroporation of
neurons and growth cones in Aplysia californica. J Neurosci
Methods 151:114–20.
Lovell P, McMahon B, Syed NI. 2002. Synaptic precedence
during synapse formation between reciprocally connected
neurons involves transmitter-receptor interactions and AA
metabolites. J Neurophysiol 88:1328–38.
Nilsson A, Moller K, Dahlin L, Lundborg G, Kanje M. 2005.
Early changes in gene expression in the dorsal root ganglia
after transection of the sciatic nerve; effects of amphiregulin
and PAI-1 on regeneration. Mol Brain Res 136:65–74.
Nyhus JK, Denburg JL. 2000. A paradoxical gradient of a basal
lamina-associated repellent is essential for pathfinding by the
Ti1 pioneer axons in cockroach embryos. Mol Cell Neurosci
16:481–98.
Peter N, Aronoff B, Wu F, Schacher S. 1994. Decrease in
growth cone neurite fasciculation by sensory or motor cells
in vitro accompanies down-regulation of Aplysia celladhesion molecules by neurotransmitters. J Neurosci
14:1413–21.
Pinsker H, Kupfermann I, Castellucci V, Kandel E. 1970.
Habituation and dishabituation of the gill-withdrawal reflex
in Aplysia. Science 167:1740–2.
Pinsker HM, Hening WA, Carew TJ, Kandel ER. 1973. Longterm sensitization of a defensive withdrawal reflex in Aplysia.
Science 182:1039–42.
Lu M, Witke W, Kwiatkowski DJ, Kosik KS. 1997. Delayed
retraction of filopodia in gelsolin null mice. J Cell Biol
138:1279–87.
Price CJ, Goldberg JI. 1993. Serotonin activation of a cyclicAMP-dependent sodium current in an identified neuron
from Helisoma trivolvis. J Neurosci 13:4979–87.
Lyles V, Zhao YL, Martin KC. 2006. Synapse formation and
mRNA localization in cultured Aplysia neurons. Neuron
49:349–56.
Radojcic T, Pentreath VW. 1979. Invertebrate glia. Prog
Neurobiol 12:115–79.
Martin KC. 2004. Local protein synthesis during axon guidance
and synaptic plasticity. Curr Opin Neurobiol 14:305–10.
Rayport SG, Schacher S. 1986. Synaptic plasticity in vitro—cell
culture of identified Aplysia neurons mediating short-term
habituation and sensitization. J Neurosci 6:759–63.
Aplysia neurons in cell culture
Roots BI. 1981. Comparative studies on glial markers. J Exp
Biol 95:167–80.
Roth LWA, Bormann P, Wiederkehr C, Reinhard E. 1999. BetaThymosin, a modulator of the actin cytoskeleton is increased
in regenerating retinal ganglion cells. Eur J Neurosci
11:3488–98.
Sabry JH, Oconnor TP, Evans L, Toroianraymond A,
Kirschner M, Bentley D. 1991. Microtubule behavior during
guidance of pioneer neuron growth cones in situ. J Cell Biol
115:381–95.
Schacher S. 1985. Differential synapse formation and neurite
outgrowth at 2 branches of the metacerebral cell of Aplysia in
dissociated cell culture. J Neurosci 5:2028–34.
Schacher S, Proshansky E. 1983. Neurite regeneration by
Aplysia neurons in dissociated cell culture—modulation by
Aplysia hemolymph and the presence of the initial axonal
segment. J Neurosci 3:2403–13.
Scheibenstock A, Krygier D, Haque Z, Syed N, Lukowiak K.
2002. The soma of RPeD1 must be present for long-term
memory formation of associative learning in Lymnaea.
J Neurophysiol 88:1584–91.
Sonetti D, Ottaviani E, Bianchi F, Rodriguez M, Stefano ML,
Scharrer B, Stefano GB. 1994. Microglia in invertebrate
ganglia. Proc Natl Acad Sci USA 91:9180–4.
Spencer GE, Lukowiak K, Syed NI. 2000. Transmitter-receptor
interactions between growth cones of identified Lymnaea
neurons determine target cell selection in vitro. J Neurosci
20:8077–86.
Spencer GE, Syed NI, van Kesteren E, Lukowiak K,
Geraerts WPM, van Minnen J. 2000. Synthesis and
functional integration of a neurotransmitter receptor in
isolated invertebrate axons. J Neurobiol 44:72–81.
Spira ME, Oren R, Dormann A, Gitler AD. 2003. Critical
calpain-dependent ultrastructural alterations underlie the
transformation of an axonal segment into a growth cone
after axotomy of cultured Aplysia neurons. J Comp Neurol
457:293–312.
Spira ME, Oren R, Dormann A, Ilouz N, Lev S. 2001. Calcium,
protease activation, and cytoskeleton remodeling underlie
growth cone formation and neuronal regeneration. Cell Mol
Neurobiol 21:591–604.
Sun ZY, Kauderer B, Schacher S. 1996. Differential distribution
of functional receptors for neuromodulators evoking shortterm heterosynaptic plasticity in Aplysia sensory neurons.
J Neurosci 16:7540–9.
Sun ZY, Schacher S. 1996. Development of short-term
heterosynaptic facilitation at Aplysia sensorimotor synapses
in vitro is accompanied by changes in the functional expression of presynaptic serotonin receptors. J Neurophysiol
76:2250–61.
Sun ZY, Wu F, Schacher S. 2001. Rapid bidirectional
modulation of mRNA expression and export accompany
long-term facilitation and depression of Aplysia synapses.
J Neurobiol 46:41–7.
869
Syed NI, Bulloch AGM, Lukowiak K. 1990. In vitro
reconstruction of the respiratory central pattern generator
of the mollusk Lymnaea. Science 250:282–5.
Syed NI, Zaidi H, Lovell PJ. 1999. In vitro reconstruction
of neuronal networks: a simple model system approach. In:
Windhorst and Johansoon, editors. Modern techniques in
neuroscience. p. 361–376.
Syed NI, Spencer GE. 1994. Target cell selection and specific
synapse formation by identified Lymnaea neurons in vitro.
Netherlands J Zool 44:327–38.
Tanaka J, Kira M, Sobue K. 1993. Gelsolin is localized in
neuronal growth cones. Dev Brain Res 76:268–71.
Trimm KR, Rehder V. 2004. Nitric oxide acts as a slow-down
and search signal in developing neurites. Eur J Neurosci
19:809–18.
van Diepen MT, Spencer GE, van Minnen J, Gouwenberg Y,
Bouwman J, Smit AB, van Kesteren RE. 2005. The molluscan
RING-finger protein L-TRIM is essential for neuronal
outgrowth. Mol Cell Neurosci 29:74–81.
van Kesteren RE, Carter C, Dissel HMG, van Minnen J,
Gouwenberg Y, Syed NI, Spencer GE, Smit AB. 2006. Local
synthesis of actin-binding protein beta-thymosin regulates
neurite outgrowth. J Neurosci 26:152–7.
Van Wagenen S, Cheng S, Rehder V. 1999. Stimulationinduced changes in filopodial dynamics determine the action
radius of growth cones in the snail Helisoma trivolvis. Cell
Motil Cytoskeleton 44:248–62.
Vanmali BH, Romanova EV, Messner MC, Singh M,
Maruniak J, Sweedler J, Kirk MD. 2003. Endogenous
neurotrophic factors enhance neurite growth by bag cell
neurons of Aplysia. J Neurobiol 56:78–93.
Verma P, Chierzi S, Codd AM, Campbell DS, Meyer RL,
Holt CE, Fawcett JW. 2005. Axonal protein synthesis and
degradation are necessary for efficient growth cone
regeneration. J Neurosci 25:331–42.
Weiss KR, Kupfermann I. 1976. Homology of the giant
serotonergic neurons (metacerebral cells) in Aplysia and
pulmonate molluscs. Brain Res 117:33–49.
Welshhans K, Rehder V. 2005. Local activation of the nitric
oxide cyclic guanosine monophosphate pathway in growth
cones regulates filopodial length via protein kinase G, cyclic
ADP ribose and intracellular Ca2þ release. Eur J Neurosci
22:3006–16.
Wildering WC, Hermann PM, Bulloch AGM. 2001. Lymnaea
epidermal growth factor promotes axonal regeneration in
CNS organ culture. J Neurosci 21:9345–54.
Williams DK, Cohan CS. 1994. The role of conditioning factors
in the formation of growth cones and neurites from the axon
stump after axotomy. Dev Brain Res 81:89–104.
Willis DE, Twiss JL. 2006. The evolving roles of axonally
synthesized proteins in regeneration. Curr Opin Neurobiol
16:111–18.
Wong RG, Barker DL, Kater SB, Bodnar DA. 1984. Nerve growth
promoting factor produced in culture media conditioned by
specific CNS tissues of the snail Helisoma. Brain Res 292:81–91.
870
Wong RG, Hadley RD, Kater SB, Hauser GC. 1981. Neurite
outgrowth in molluscan organ and cell cultures—the role of
conditioning factor(s). J Neurosci 1:1008–21.
Wu KY, Hengst U, Cox LJ, Macosko EZ, Jeromin A,
Urquhart ER, Jaffrey SR. 2005. Local translation of RhoA
regulates growth cone collapse. Nature 436:1020–4.
Zhao YL, Klein M. 2002. Modulation of the readily
releasable pool of transmitter and of excitation-secretion
P. Lovell and L. L. Moroz
coupling by activity and by serotonin at Aplysia
sensorimotor synapses in culture. J Neurosci 22:
10671–9.
Zhu H, Wu F, Schacher S. 1997. Site-specific and sensory
neuron-dependent increases in postsynaptic glutamate
sensitivity accompany serotonin-induced long-term facilitation at Aplysia sensorimotor synapses. J Neurosci
17:4976–86.