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A flexible genetic toolkit for arthropod
neurogenesis
rstb.royalsocietypublishing.org
Angelika Stollewerk
School of Biological and Chemical Sciences, Queen Mary University of London, Mile End Road, London E1 4NS, UK
Review
Cite this article: Stollewerk A. 2016 A flexible
genetic toolkit for arthropod neurogenesis.
Phil. Trans. R. Soc. B 371: 20150044.
http://dx.doi.org/10.1098/rstb.2015.0044
Accepted: 21 August 2015
One contribution of 16 to a discussion meeting
issue ‘Homology and convergence in nervous
system evolution’.
Subject Areas:
evolution, neuroscience
Keywords:
neurogenic potential, neural precursors,
neural progenitors, asymmetric division,
patterning genes, Notch signalling
Author for correspondence:
Angelika Stollewerk
e-mail: [email protected]
Arthropods show considerable variations in early neurogenesis. This
includes the pattern of specification, division and movement of neural precursors and progenitors. In all metazoans with nervous systems, including
arthropods, conserved genes regulate neurogenesis, which raises the question of how the various morphological mechanisms have emerged and
how the same genetic toolkit might generate different morphological
outcomes. Here I address this question by comparing neurogenesis
across arthropods and show how variations in the regulation and function
of the neural genes might explain this phenomenon and how they might
have facilitated the evolution of the diverse morphological mechanisms
of neurogenesis.
1. Introduction
During neurogenesis neurons and glia are formed and their number, types,
morphologies, final positions and connections shape the architecture of the
brain. Consequently, variations in the morphology and function of metazoan
nervous systems must originate from evolutionary changes of the developmental
processes that generate the neural cells and regulate their differentiation.
In order to understand the causes and sequence of evolutionary variations,
Volker Hartenstein and I, in a recent review [1], compared early neurogenesis
across the animal kingdom by subdividing the developmental processes of
early neurogenesis into distinct morphological modules. Neurogenesis starts
with patterning cells with neurogenic potential (module A) and further early
steps include the patterning (module B), proliferation (module C) and movement (module D) of neural progenitors. Variants of these modules can be
detected across the animal kingdom and allow conclusions to be drawn on
the mechanisms of neurogenesis in the urbilaterian [1]. For example, the
module ‘patterning cells with neurogenic potential’ (module A) is present as
three different variants. In animals like cnidarians that have a nerve net,
the whole ectoderm has neurogenic potential (‘generalized neurogenic
ectoderm’, module A1), whereas arthropods and vertebrates have restricted
ectodermal domains that generate the central nervous system (CNS) (‘neuroectoderm’, module A2). In some clades, the neurogenic potential is restricted
to subectodermal cells (module A3) such as the neoblasts in flatworms
(Platyhelminthes) [1].
Interestingly, the genetic toolkit of early neurogenesis is highly conserved in
all animal phyla despite significant variations in the morphological modules. In
this review, I will address the question of how the various morphological modules might have emerged and how the same genetic toolkit might generate the
different modules of neurogenesis, using arthropods as model organisms.
Arthropods are particularly well suited to analyse this question because of
the large size of the phylum, great variations in shapes and behaviours, and
the availability of comparative data on neurogenesis for all groups [1,2].
Furthermore, large-scale molecular phylogenies have greatly improved the resolution of arthropod relationships and thus facilitate evolutionary
investigations of arthropod neurogenesis (figure 1) [3]. It is now widely
accepted that the paraphyletic crustaceans group with the insects forming the
pancrustacean clade (also named Tetraconata clade). The pancrustaceans are
a sister group of the myriapods and are united with the latter in the
& 2015 The Author(s) Published by the Royal Society. All rights reserved.
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Arthropoda
2
Onychophora
Myriapoda
Euchelicerata
Crustacea
Hexapoda
C3/D2
C3/D1
+C3/D1
C1/D3/D4
C1/D3/D4
B1/C2/D2
B2 (invariant neural
progenitors/precursors)
A2 (neuroectoderm)
Figure 1. Arthropod phylogeny and the evolution of the neurogenesis module. The arthropod relationships shown in the phylogenetic tree are the framework for
the discussions in this review. A2– D4 represent parts of the module variants recently described by [1]. Module A: patterning cells with neurogenic potential. In
contrast with animals with a generalized ectoderm (module A1, not shown), all arthropods have a restricted area with neurogenic potential (neuroectoderm) that
gives rise to the central nervous system (module A2). Module B: patterning of neural progenitors. Neural progenitors/precursor groups show a similar, invariant
pattern in all euarthropods (module B2), while the neural progenitors of onychophorans appear in random positions (module B1). The ancestral pattern of euarthropod neurogenesis is the selection of neural precursor groups. Modules C and D: proliferation and movement of neural progenitors, respectively. The neural
precursors of chelicerates and myriapods (mainly) do not divide (module C1) and directly differentiate into neurons and glia. The neural precursor groups are
internalized by ingression (module D3) or invagination (module D4). Both mechanisms are seen in chelicerates (including pycnogonids) and myriapods. In addition,
pycnogonids have asymmetrically dividing progenitors (module C3) that remain in the epithelium (module D1). By contrast, the neural progenitors of onychophorans
divide symmetrically (module C2) after their delamination (module D2). Insect and crustacean neuroblasts divide asymmetrically (module C3), however insect
neuroblasts delaminate (module D2), while crustacean neuroblasts remain in the epithelium (module D1). The scheme and description refer to the processes
in the ventral neuroectoderm.
Mandibulata group [3–6]. The chelicerates represent a basal
branch and are a sister group of the Mandibulata. Insects,
crustaceans, myriapods and chelicerates together compose
the euarthropods. The onychophorans (velvet worms) are
the sister group of the euarthropods and together with the
latter form the phylum Arthropoda (figure 1). The following
discussion of the neurogenesis modules is based on this
phylogenetic framework (but see [7– 10] for discussion).
2. Patterning regions with neurogenic potential
In arthropods, the procephalic and the ventral neuroectoderm
generate the tripartite brain and the ventral ganglia, respectively (module A2; figure 1) [11 –13]. The size of the
neuroectoderm differs from a few cell rows in crustaceans
and insects to wide areas containing many small cells in myriapods and chelicerates (figure 2) [14–19]. Data from insects
and a representative of a basal branch of the euarthropods,
the spider Parasteatoda tepidariorum (chelicerate), show that
dorsoventral (DV) cell fates, including the regions with neurogenic potential, are determined by morphogen gradients
involving Dpp/Sog [20–24]. The determination and patterning of the ventral neurogenic region by Dpp/Sog might,
therefore, be an ancestral feature of arthropod neurogenesis.
This is further supported by data from other Ecdysozoa,
Lophotrochozoa and Deuterostomia, which indicate that the
BMP(Dpp)/chordin(Sog) morphogen system is conserved
across bilaterians. However, there seem to be differences in
the degree of the contribution of the dpp/sog homologues in
patterning the neurogenic region in arthropods. In Drosophila
melanogaster sog null mutants, the DV extension is slightly
reduced [22], while in the flour beetle Tribolium castaneum
and in the spider (chelicerate) P. tepidariorum, sog RNAi results
in an (almost) complete loss of the ventral neuroectoderm
[20,23]. Data are not available for myriapods and data from
crustaceans are inconclusive because the only sog gene that
has been identified shows a restricted expression in the mandibular ventral midline cells [25]. The fact that the regulation
of DPP signalling has considerably diverged even within
insects [23,24] suggests that evolutionary variations in the
regulation of the DV patterning genes and their targets
might correlate with the different extensions of the neurogenic
regions in arthropods (figure 2). This assumption is further
supported by comparative studies of the DV extension of the
ventral midline in Diptera and Hymenoptera [26]. While in
D. melanogaster and in the mosquito Anopheles gambiae the ventral midline is 1–2 cells wide along the DV axis, it extends over
5–6 cells in the honeybee Apis mellifera. This difference is
because of variations in the regulation of single-minded—a transcription factor which is essential for the development of the
midline cells—by DV patterning genes [26].
In all arthropods the neurogenic potential differs along
the anterior –posterior (AP) axis. The (anterior) procephalic
Phil. Trans. R. Soc. B 371: 20150044
Pycnogonida
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Euarthropoda
Mandibulata
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(a)
pc
(b)
11
(f)
13
vne
Glomeris marginata msh
(c)
t1
4-1 4-2 4-3 4-4 3-5
5-5 5-6
5-3
5-1
5-4
5-2
6-4
6-1
6-2
7-1
7-3 7-4
MNB
7-2
t3
(g)
Glomeris marginata SoxB1
(d)
Tribolium castaneum msh
(e)
1-2
1-3
2-4
2-5
2-1 2-2 2-3
3-4 3-5
3-1 3-2 3-3
4-4
4-3
4-1 4-2
5-6
5-4
5-1 5-2 5-3
1-1
6-2 6-3 6-4
6-1
7-1 7-2 7-3 7-4
MNB
Glomeris marginata msh
Cupiennius salei msh
Tribolium castaneum
vnd, ind msh
Figure 2. Patterning of the neurogenic region in euarthropods. The dashed lines indicate the ventral midline of the neuroectoderm. (a) Flat preparation of a
Glomeris marginata embryo (millipede) showing the RNA expression pattern of the SoxB1 gene in all neuroectodermal cells. (b,c) Comparison of the msh RNA
expression pattern in flat preparations of three ventral neuromeres of G. marginata and Tribolium castaneum (beetle). Note the difference in the width of the
neuroectoderm in these species. The boxes frame the neuromere of the second leg segment in G. marginata and the second thoracic neuromere of T. castaneum,
respectively. (d,e) Schematic of the arrangement of the neural precursor groups in hemi-neuromeres of G. marginata and the spider Cupiennius salei. The precursor
groups expressing msh are shown in brown. ( f,g) Schematic drawings comparing the expression of the dorsoventral patterning genes vnd (blue), ind (red) and msh
(brown) in the neuroblasts of Drosophila melanogaster and T. castaneum. The neuroblasts 6-2 and 7-2 express both vnd and ind in D. melanogaster, while neuroblasts
4-2 and 5-3 express ind and msh in T. castaneum. l1–l3, leg segment 1 to 3; pc, procephalic neuroectoderm; t1–t3, thoracic segments 1 to 3; vne, ventral
neuroectoderm.
neuroectoderm not only generates a higher and more
diverse number of neurons than the ventral neuroectoderm
but also forms the complex brain centres such as the mushroom bodies that integrate sensory information and regulate
behaviour [27]. In vertebrates, the Wnt/b-catenin pathway
patterns the CNS along the AP axis and in some species
determines the overall AP cell fates (reviewed in [28]). An
involvement of this signalling pathway in patterning the
AP axis has also been demonstrated in lophotrochozoans,
in particular in regenerating planarians, where the
reduction of Wnt signalling results in radial hypercephalized
individuals [29].
In arthropods, Wnt signalling seems to regulate posterior
elongation and segmentation but is neither required for the
formation of anterior (head) segments nor the overall AP patterning of the CNS [28,30]; rather, the function of Wnt in
establishing AP fates in the arthropod CNS seems to be confined to neuromeres, the metameric units of the CNS [31– 34].
The mechanisms establishing different AP fates in the
anterior part of the embryo show considerable variations
within and between arthropod groups. The AP morphogen
Bicoid, for example, is not present outside of dipterans and
Hunchback, another head morphogen, does not regulate
the expression of AP patterning genes (e.g. Hox genes)
in the spider P. tepidariorum, in contrast with the case in
D. melanogaster [35,36]. If the overall AP patterning mechanisms are connected with the regionalization of the CNS,
these differences might contribute to existing variations of
the arthropod CNS. In D. melanogaster, AP axis formation is
in fact directly linked to the AP regionalization of the CNS.
AP morphogens (e.g. Bicoid, Hunchback) regulate the genes
that pattern the CNS along the AP axis (e.g. orthodenticle,
ems, Hox genes) (e.g. [37,38]). However, there are no data
available on how overall AP patterning is linked to CNS
regionalization in arthropods other than insects.
Thus, while the overall function of BMP in DV axis formation and patterning of the neurogenic region seems to be
conserved across the animal kingdom, the function of
Wnt in AP patterning of the CNS has been restricted to the
metameric units in arthropods. Furthermore, the upstream
mechanisms that set up the initial differences in CNS AP
fates have diverged in arthropods.
However, many of the genes downstream of the primary
AP/DV subdivision are conserved across the animal kingdom, such as the transcriptional regulators of the SoxB
family, which maintain the proliferative state of the neuroepithelium and at the same time inhibit neural
differentiation (figure 2) [39–42].
Furthermore, many genes that translate the positional
information into neural/lineage subtype identity are
Phil. Trans. R. Soc. B 371: 20150044
Drosophila melanogaster
vnd, ind msh
t2
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Neural precursors/progenitors can appear in either random
(module B1) or fixed (module B2) positions within regions
of neurogenic potential (figure 3) [1]. Furthermore, single
(modules D1 and D2) or groups of contiguous precursors
can be selected (modules D3 and D4). In the arthropod
neuroectoderm, different types of founder cells of the
nervous system are generated over a period of time ranging
from hours (e.g. in the insect D. melanogaster) to months
(e.g. in the onychophoran Euperpatoides kanangrensis) [55]. In
chelicerates and myriapods groups of contiguous mainly
postmitotic neural precursors are selected in the neuroectoderm, while in onychophorans, crustaceans and insects
single neural progenitors are selected that divide either symmetrically or in a stem-cell like mode (see below) (figure 3)
[7,14,15,17,18,55,56]. In onychophorans, the sister group of
euarthropods (chelicerates, myriapods, crustaceans, insects),
neural progenitors appear in random positions in the
neuroectoderm and their number per hemi-neuromere is
variable (60– 100 progenitors; figure 3). By contrast, in all
euarthropods neural progenitors/precursors are generated
in fixed positions and show species-specific invariable
numbers within the same neuromeres (figures 2 and 3).
The morphological differences can be explained by
variations in the regulation of two classes of genes, the
achaete-scute homologues (ASH) and the Notch signalling pathway. These genes are used for the generation and regulation of
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3. Patterning of neural precursors/progenitors
neural cells throughout the animal kingdom; however, they
show variations in their spatial and temporal expression
(reviewed by [1]). In insects, chelicerates and myriapods the
invariant spatial–temporal expression of ASH genes sets up
the areas where neural progenitors/precursor groups form
[17,18,57,58]. The positions of the neural progenitors/precursors are refined by a transcriptional feedback loop between
the Notch signalling pathway and the ASH genes, which
results in the selection of regular arranged, spaced neural
precursor groups/progenitors [59,60]. In branchiopod crustaceans, the ASH genes are not expressed before neural
progenitors (neuroblasts) are formed and therefore are not
involved in positioning them (figure 3) [15]. However, an
invariant spatial pattern of ASH expression appears with a
time delay. Neuroblasts are also arranged in an invariant
pattern in another crustacean group, the malacostracans, but
molecular expression data are not available [14].
By contrast, in the onychophoran E. kanangrensis, ASH is
expressed at very low, homogeneous levels in the neuroectoderm [55]. High levels of expression can be detected and are
maintained in the neural progenitors after their delamination,
suggesting that Ek ASH is required for neural progenitor
maturation (figure 3).
Two mechanisms might contribute to the variations in patterning neural precursors/progenitors in the neuroectoderm
in arthropods. Firstly, the variations could be due to changes
in the prepatterning mechanisms regulating ASH expression.
Given the homogeneous low ASH expression in onychophorans, molecular changes in the achaete-scute homologues must
have occurred in the lineage leading to the euarthropods that
facilitated the spatio-temporal regulation of these genes allowing for the upregulation of transcripts in invariant positions.
However, data on ASH regulation in the embryonic CNS are
fragmentary in insects and not available at all in other arthropods. Skeath et al. [61] report that the D. melanogaster ASH
genes achaete, scute and lethal of scute are regulated by the
products of the pair-rule and DV axis patterning genes (e.g.
dpp) in the ventral neuroectoderm and their expression
patterns are maintained by the segment polarity proteins. In
the D. melanogaster procephalic neuroectoderm, lethal of scute,
which is the main proneural gene in the brain, is regulated
by the head gap genes tailless (tll), orthodenticle (otd), empty
spiracles (ems) and buttonhead (bhd) [62]. It can be speculated
that evolutionary changes in the spatial– temporal expression
of ASH genes are associated with variations in the expression
and interactions of the segmentation genes, which have
considerably diverged in arthropods, in particular regarding
the gap and pair-rule genes (e.g. [63 –69]).
However, segmentation genes might not be used at all for
positioning neural precursors/progenitors in some groups.
For example, in the neuroectoderm of the branchiopod
crustacean Daphnia magna, the default neural cell fate is suppressed in the central area of the hemi-neuromeres by the
activation of Notch signalling so that neural progenitors are
arranged in a ring-like structure (figure 3). This pattern
seems to be representative for branchiopods; however, in
malacostracan crustaceans the neuroblast arrangement is
not disrupted by a non-neural central area, suggesting that
the mechanisms of neuroblast positioning have diverged in
crustaceans [14].
Secondly, variations in the regulation of Notch activity
could underlie the variation in the pattern of neural
precursors/progenitors in arthropods.
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conserved in metazoans. Downstream of the primary AP and
DV patterning mechanisms, additional signalling pathways
as well as homeodomain proteins establish neuronal subtype
identity [43]. For example, the function of Hox genes in determining segmental identity is conserved across the animal
kingdom [43]. However, although these genes seem to be
an integral downstream part of the molecular processes that
pattern the neurogenic region along the AP axis of the CNS
in all arthropods (e.g. [44–47]), there are considerable functional variations even within individual arthropod groups.
For example, the expression of the Hox gene labial in the
head segment that gives rise to the tritocerebrum is conserved in all arthropods [48]; however, studies in insects
show divergent functions of Labial in brain development.
While in D. melanogaster labial mutants the tritocerebrum
and additional brain structures are affected, labial RNAi
does not show a phenotype in the milkweed bug Oncopeltus
fasciatus and the functional defects are restricted to the
tritocerebrum in the flour beetle T. castaneum [49–51].
The genes patterning the neurogenic region along the DV
axis are also highly conserved across the animal kingdom
[52]. Yet again, detailed comparative analyses of the
expression of the DV patterning genes muscle segment homeobox (msh), intermediate neuroblasts defective (ind) and ventral
nervous system (vnd) defective in arthropods revealed considerable variations in the expression of these genes relative
to the neural progenitors/precursors in the respective
domains (figure 2) [53,54]. Furthermore, evolutionary
changes have occurred in the neuronal subtype specific
genes that are regulated by the DV patterning genes. For
example, msh regulates the expression of the motor- and
interneuronal marker islet in the spider Cupiennius salei,
whereas this is not the case in D. melanogaster [53].
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onychophorans
chelicerates
insects
crustaceans
intermediate
neural precursor
neuron/glia
neural
precursor
neural
progenitor
neuro/glia
ganglion
mother
cell
neuroblast
symmetrically
dividing
neuroblast
Figure 3. Pattern of neural precursor/progenitor specification and division. Small sections of the neuroectoderm (white squares: neuroectodermal cells) are shown
for each group. (a) Onychophorans: ASH shows a low homogeneous expression in the neuroectoderm (light grey) and is upregulated in the neural progenitors (dark
grey) after their delamination. Chelicerates: ASH is upregulated in large domains and becomes restricted to groups of cells. A similar pattern is seen in myriapods
(not shown). Insects: ASH is expressed in small proneural clusters. Expression becomes restricted to single neuroblasts. Crustaceans (branchiopods): ASH is upregulated after neuroblast specification. Over time more neuroblasts express ASH and the gene is expressed at different levels indicated by lighter and darker grey.
(b) Division pattern and movement of neural precursors/progenitors. Onychophorans: single neural progenitors delaminate and divide symmetrically to produce
intermediate neural precursors, which divide again. The expression of snail and prospero has not been studied in onychophorans. Chelicerates: most neural precursors
are postmitotic. Neural precursor groups express snail (orange). Neural precursors co-express prospero (red) before detaching from the group. Insects: neuroblasts
express snail and prospero and divide asymmetrically to produce GMCs, which divide again to produce neurons and glia. Crustaceans show the same division pattern
but snail is expressed long before prospero. Snail-positive neuroblasts can divide symmetrically.
The core mechanism of the Notch signalling pathway is
the activation of the Notch receptor by the ligand Delta,
which results in the intramembrane proteolysis of the receptor and the release of an intracellular fragment that acts as a
transcriptional regulator [70]. The interactions between the
transmembrane proteins Notch and Delta are complex,
and different regulatory mechanisms are used in different
developmental processes such as ligand– receptor trans- and
cis-activation, receptor cis-inhibition by the ligand or
mutual receptor– ligand cis-inhibition [70,71].
Biological and mathematical models show that the selection pattern of precursors can be modulated by regulating
Notch signalling in different ways [71,72]. The selection of
regular arranged, spaced neural precursor groups/progenitors can be explained by the ‘lateral inhibition model’
where Notch signalling and the ASH genes are linked in a
transcriptional feedback mechanism [70]. In insects, the
ASH genes are expressed in small clusters in the ventral
neuroectoderm. One of the cells adopts the neural fate and
expresses higher levels of ASH, which in turn leads to an
increase of Delta expression [59]. Delta trans-activates the
Notch receptor in the neighbouring cells. The products of
the Notch target genes, the hairy/Enhancer of Split (HES)
genes, downregulate ASH and inhibit the neighbouring
cells from adopting the neural fate. In chelicerates and
myriapods, lateral inhibition operates between the neural
precursors and the neuroectoermal cells surrounding the
selected precursor group [18,60]. However, the neural precursor group itself seems to be insensitive to Notch signalling.
This is reflected in the high levels of Delta expressed by the
precursor groups suggesting that the Notch receptor is
cis-inactivated [60,71].
In onychophorans, the homogeneous, low expression of
ASH and the strong expression of Notch and Delta in the ventral neuroectoderm suggest that Notch signalling keeps ASH
expression at low levels in all cells (figure 3) [55]. Random
changes in the sensitivity to Notch signalling, possibly by
cis-inactivation of the Notch receptor by the ligand Delta,
could allow for individual cells to adopt the neural fate and
delaminate. This mechanism could explain the seemingly
random positions and great range in progenitor numbers
per hemi-neuromere.
The pattern of neural precursors/progenitors is much
more complex in the developing brain. In chelicerates and
myriapods, the same types of founder cells (neural precursor
groups) are visible in the procephalic and ventral neuroectoderm [73]. In both neurogenic areas, the ASH genes are
expressed in large domains from which several neural precursor groups arise (although the underlying morphological
processes are more complex in the procephalic neuroectoderm, see below). By contrast, in insects, ASH genes are
expressed in large domains (rather than small clusters) in
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Arthropods show also variations in the developmental modules following neural determination. Selected cells can either
differentiate directly into neural cells (neurons/glia) (module
C1), as is the case for many of the neural precursors in chelicerates and myriapods, or they can undergo symmetric (e.g.
onychophorans) or asymmetric divisions (insects, crustaceans)
(modules C2 and C3; figure 1). They can also show combinations of these variants (e.g. sea spiders; figure 1). The
associated movements either of the neural precursors/progenitors themselves and/or of their progeny lead to the
formation of internal cell layers, basal to the neuroectoderm
(figure 3). The movements show variations that are partially
linked to the differences in the specification patterns of neural
precursors/progenitors. Insect neuroblasts start to divide
asymmetrically to produce ganglion mother cells (GMCs)
after segregating individually from the neuroectoderm (delamination; modules C3 and D2), while crustacean neuroblasts
remain in the neuroectoderm and generate GMCs at the basal
side by directed asymmetric mitosis (module C3 and D1;
figures 1 and 3) [14–16,77]. Onychophoran neural progenitors delaminate before they divide symmetrically to
produce intermediate neural precursors (modules C2 and
D2; figures 1 and 3) [7,55]. By contrast, there are no cell divisions within the neural precursor groups of chelicerates and
myriapods (module C1) and only few neural precursors
divide after detaching from the precursor groups (ingression;
module D3) or the internalization of the whole group (invagination; module D4; figures 1 and 3).
However, in myriapods single and groups of mitotic cells
seem to prefigure the areas where neural precursor groups
are selected, suggesting a link between the formation of
neural precursors and cell proliferation [18]. However, the
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4. Proliferation and movement of neural
precursors
division pattern has not been analysed in detail and it is
not known if the neural precursors within a group are related
by lineage. Interestingly, sea spiders, which are most
probably the sister group of all other chelicerates (euchelicerates), show a combination of asymmetrically dividing
neural progenitors and neural precursor groups [78]. However, in this case, neural precursor groups are generated
before the neural progenitors appear.
Despite these differences, committed neural progenitors/
precursors express a set of conserved factors that were shown
to be involved in asymmetric division and neural cell fate
determination in D. melanogaster [79]. This raises the question
of how these genes can support the various morphological
outcomes in all euarthropod groups (data are not available
for onychophorans).
In the following, I will discuss a few of the factors for
which comparative data exist in euarthropods. In D. melanogaster, Asense activates many genes in neuroblasts, among
others those that are required for asymmetric division [80].
The neuroblasts immediately delaminate after their formation and start to divide (figure 3). The neural
differentiation factor Prospero is asymmetrically distributed
into the GMCs where it enters the nucleus, acts antagonistically to Asense and promotes cell-cycle exit [81–83]. The
Snail family members are required for the mitotic activity
of the neuroblast and the spindle rotation, which occurs
prior to the production of GMCs [79]. The asymmetric distribution of Prospero to GMCs is affected in mutants
carrying a deletion of three Snail family genes [79]. Furthermore,
the Snail family members might be directly involved in triggering the delamination of neuroblasts by downregulating
E-cadherin [84,85].
Although crustacean neuroblasts do not delaminate, they
show the same division pattern as insect neuroblasts, and it is
therefore conceivable to assume that the function of Snail and
Asense in asymmetric cell division is conserved in both
groups. Furthermore, prospero is asymmetrically distributed
into the GMCs in crustaceans and its expression coincides
with the onset of asymmetric division of the neuroblasts,
suggesting that it acts as a neural cell fate determinant similar
to the case in insects [15,86]. However, Snail must have an
additional earlier function in crustacean neurogenesis
because it is the earliest gene to be expressed in neuroblasts
and there is a considerable time delay between neuroblast
formation and the start of asymmetric divisions [15,86]. The
fact that snail is expressed before ASH suggests that the
former rather than the latter gene is required for neural cell
fate determination. This potential function of Snail is supported by data from D. melanogaster showing that Snail can
activate neuroblast fate genes such as grainyhead, numb,
prospero and seven-up [80]. Furthermore, Snail could be
involved in the symmetric divisions of crustacean neuroblasts, which occur before the start of GMC production
(figure 3) [15]. The function of Snail in regulating cell-cycle
genes and the proliferative divisions of stem cells has
been demonstrated both in D. melanogaster and in
vertebrates [80,87 –89].
The delayed progression of crustacean neuroblasts to
asymmetric divisions compared with insects can again be
explained by changes in the use of the participating genes
[86]. Notch signalling is used not only for binary cell fate
decisions in the hemi-neuromeres between the central presumptive epidermal cells and the surrounding neuroblasts,
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some areas of the procephalic neuroectoderm, from which
several neuroblasts delaminate [74]. Furthermore, in the
dorsal midline of the procephalic neuroectoderm of
D. melanogaster, groups of neural precursors form within a
large lethal of scute domain [74]. Yet again, different patterns
and regulations of ASH and Notch signalling can be seen in
different ontogenetic stages (e.g. larvae) and in the sensory
nervous system (e.g. [75,76]). The mechanisms seen in the
developing insect brain show similarities to the ancestral
patterns of neurogenesis described in onychophorans and
chelicerates/myriapods.
These data have led to the hypothesis that the neural precursor groups of chelicerates and myriapods have evolved
from areas of massive, random neural progenitor delamination due to changes in the regulation of the ASH genes and
in the regulation of Notch signalling [55]. There is strong evidence that neural precursor groups were present in the last
common ancestor of euarthropods and thus represent the
ancestral pattern of euarthropod neurogenesis [55].
Additional changes in the regulation of the participating
genes have led to the evolution of neuroblasts from the
neural precursor groups in the last common ancestor of
insects and crustaceans. Thus, changes in the prepatterning
mechanisms and the versatility of the Notch signalling pathway might explain the various patterns of neural precursors/
progenitors in arthropods.
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5. Conservation of structure despite changes
of gene expression
Phil. Trans. R. Soc. B 371: 20150044
So far, I have discussed how the expression of conserved genes
in neural precursors/progenitors can lead to different morphological outcomes. However, in arthropod neurogenesis
the reverse mechanism also exists: the expression of conserved
genes in different patterns resulting in the same morphological
outcome. This phenomenon might contribute to internal buffering mechanisms ensuring that evolutionary modifications
do not result in destabilization and functional breakdown of
neuronal structures. In the following, I will give two examples
relating to the axonal scaffold of euarthropods.
In all euarthropods, axonal tracts are established in the
developing ventral nerve chord in a highly conserved pattern
[53]. Two longitudinal tracts connect the brain and the ventral
ganglia. In each ventral neuromere, two transverse tracts, the
anterior and posterior commissures, connect the longitudinal
fascicles. During later development this arrangement can be
obscured by the fusion of ganglia. The conserved axonal scaffold seems to be related to the equally highly conserved
arrangement of the neural precursor groups/progenitors in
euarthropods. In all four groups, they are arranged in seven
rows and three to six columns in each hemi-neuromere
(figure 2) [53]. Spatial and temporal cues within the neuroectoderm lead to the expression of a unique set of genes in
the neural precursors/progenitors, which in turn determines
the subtype identity of the progeny [43,53,54,98 –100]. The
neural precursors/progenitors produce early sets of pioneer
neurons that establish the axonal scaffold [101]. One set of
pioneer neurons is characterized by the expression of the
motor and interneuronal marker even-skipped [54,102,103].
The position of eve-positive neurons is highly conserved in
insects and crustaceans. However, a recent comparison of
the neuroblast expression patterns in D. melanogaster and in
the flour beetle T. castaneum revealed that despite their conserved positions the expression profiles of the neuroblasts
have diverged considerably, including the profiles of those
generating the eve-positive pioneer neurons [54,99]. Interestingly, these evolutionary changes include the expression of
patterning genes which are highly conserved throughout
the animal kingdom, such as the DV patterning genes msh,
ind and vnd (figure 2). These data show that the early neuronal programme, which generates the pioneer neurons,
tolerates considerable molecular variations, thus ensuring
the formation of the conserved axonal scaffold.
In addition to the pioneer neurons, a sophisticated system
of axonal guidance molecules at the ventral CNS midline
plays an important role in generating the axonal scaffold in
euarthropods [104–106]. For example, Netrin, a conserved
chemoattractant, attracts commissural axons to the ventral
midline [107]. In insects and crustaceans, specialized midline
cells consisting of neurons and glia are the source of Netrin
expression [106,107]. In spiders, however, neuronal midline
cells are absent and the midline consists of epithelial cells
[108]. Furthermore, during embryogenesis, spiders undergo
a process called inversion whereby the germband splits
into two halves at the ventral side along the AP axis. The
two halves move apart and during this process a ventral
midline epithelium appears which extends between the
germband halves. After dorsal closure the germband halves
come together again ventrally and the midline epithelium
7
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but also for keeping neuroblasts in the transition phase which
is characterized by the expression of neural genes but the
absence of asymmetric divisions. Neuroblasts individually
leave the transition phase showing strong expression of
ASH, asense and prospero and start producing GMCs [86].
Both ASH and prospero expression are derepressed when
Notch activity is lowered, leading to premature production
of GMCs. Thus in this context, ASH seems to be required
for the maturation of the neuroblasts, rather than the
specification of the neural fate as is the case in insects.
Furthermore, in embryos with reduced Notch activity
the epithelial morphology is disturbed, indicating that
Notch signalling might have an additional function in keeping neuroblasts within the neuroectoderm [86]. This
function of Notch has been described both in vertebrates
and in arthropods. For example, inactivation of the Notch
effector genes Hes 1, 3 and 5 in mice results in reduction
of tight and adherens junctions at the apical end feed of
neural stem cells and loss of these cells from the spinal
cord neuroepithelium [90]. In the spider C. salei (chelicerate)
neural precursor groups disintegrate prematurely in the
CNS and peripheral nervous system (PNS), showing that
this function of Notch can also support the formation and
maintenance of neural precursor groups [17,91].
In spiders (chelicerates), most of the neural precursors
expressing snail and prospero are postmitotic and the few precursors that proliferate divide symmetrically, thus excluding
a function of these panneural factors in asymmetric division
[92]. However, Prospero seems to act as neural differentiation factor and might promote cell-cycle exit similar to the
case in insects and crustaceans because it is expressed in
the nuclei of neurons and their precursors in the CNS and
PNS [92,93]. Snail, on the other hand, must have a different
function. Interestingly, it has been shown in vertebrates and
recently also in D. melanogaster that members of the Snail
family are sufficient to induce cell-shape changes leading to
the delamination of neural precursors from the neuroepithelium [94–96]. It is, therefore, conceivable that Snail is
required for the morphological changes that occur during
invagination/ingression of the neural precursors both in chelicerates and myriapods, where snail is expressed in the same
pattern [35].
Asense is neither present in chelicerates nor in myriapods;
however, the spider C. salei has two ASH genes, which arose
by independent duplication in the spider lineage [97]. While
CsASH1 is initially expressed in large proneural domains,
CsASH2 is exclusively expressed in neural precursors and is
able to rescue asense-specific defects in D. melanogaster
mutants [97]. This suggests that there is yet another mechanism that could account for the morphological differences of
neurogenesis in arthropods—the convergent evolution of
genes promoting the neural programme.
To summarize, these explanations show that the emergence of the various neurogenesis modules might have
been facilitated by the diverse functions of the participating
genes as well as their flexibility of acting in different morphological contexts. Furthermore, the modularity of early
neurogenesis has allowed for combining various modules
so that neurogenesis could be adapted to different requirements not only in the individual arthropod groups/species
but also in different areas of neurogenesis in the same
organism.
Downloaded from http://rstb.royalsocietypublishing.org/ on June 12, 2017
Molecular genetic changes that positively influenced the flexible use of the gene product, for example by introducing
Competing interests. I declare we have no competing interests.
Funding. I received no funding for this study.
Acknowledgements. I am grateful to Nick Strausfeld, Frank Hirth and the
Royal Society for inviting me to participate in the March 2015 discussion meeting ‘Origin and evolution of the nervous system’. I also
gratefully acknowledge the participants of the discussion meeting
for thought-provoking and helpful comments.
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