195
Neurogenesis in myriapods and chelicerates and its importance
for understanding arthropod relationships
Angelika Stollewerk1,2 and Ariel D. Chipman
Department of Zoology, University of Cambridge, Downing Street, Cambridge CB2 3EJ, UK
Synopsis Several alternative hypotheses on the relationships between the major arthropod groups are still being discussed.
We reexamine here the chelicerate/myriapod relationship by comparing previously published morphological data on neurogenesis in the euarthropod groups and presenting data on an additional myriapod (Strigamia maritima). Although there are
differences in the formation of neural precursors, most euarthropod species analyzed generate about 30 single neural
precursors (insects/crustaceans) or precursor groups (chelicerates/myriapods) per hemisegment that are arranged in a regular
pattern. The genetic network involved in recruitment and specification of neural precursors seems to be conserved among
euarthropods. Furthermore, we show here that neural precursor identity seems to be achieved in a similar way. Besides these
conserved features we found 2 characters that distinguish insects/crustaceans from myriapods/chelicerates. First, in insects and
crustaceans the neuroectoderm gives rise to epidermal and neural cells, whereas in chelicerates and myriapods the central area
of the neuroectoderm exclusively generates neural cells. Second, neural cells arise by stem-cell-like divisions of neuroblasts in
insects and crustaceans, whereas groups of mainly postmitotic neural precursors are recruited for the neural fate in chelicerates
and myriapods. We discuss whether these characteristics represent a sympleisiomorphy of myriapods and chelicerates that has
been lost in the more derived Pancrustacea or whether these characteristics are a synapomorphy of myriapods and chelicerates,
providing the first morphological support for the Myriochelata group.
Introduction
The relationships between and within the major
arthropod groups have not been consistently resolved.
Several alternative hypotheses are being discussed.
The so-called Mandibulata hypothesis suggests a
clade composed of insects, crustaceans, and myriapods
with various ideas as to the relationships within this
clade. The Pancrustacea hypothesis assumes a crustacean origin of insects or a sister group relationship
between both groups (Zrzavý and Štys 1997; Shultz
and Regier 2000; Dohle 2001; Mallatt and others
2004; Regier and others 2005), and the Atelocerata
hypothesis unites insects and myriapods as a clade
(Snodgrass 1938, 1950, 1951; Briggs and Fortey 1989;
Schram and Emerson 1991; Bergström 1992; Wheeler
WC and others 1993; Kraus O and Kraus M 1994, 1996;
Emerson and Schram 1998; Wheeler WC 1998; Wills
and others 1998; Bitsch C and Bitsch J 2004). Whereas
the Atelocerata hypothesis is mainly supported by
morphological evidence, the idea of Pancrustacea
was initially based on the phylogenetic analysis of
ribosomal-RNA sequence data in which crustaceans
and insects grouped together to the exclusion of
myriapods (Field and others 1988; Turberville and
others 1991; Ballard and others 1992; Friedrich and
Tautz 1995; Giribet and Ribera 1998). Recent data
on comparative developmental biology support this
molecular sister group relationship, although the
synapomorphies seem to be shared mainly by insects
and malacostracans (Dohle and Scholtz 1988; Patel and
others 1989; Whitington and others 1993; Osorio and
others 1995; Whitington 1996; Dohle 1998; Nilsson
and Osorio 1998; Duman-Scheel and Patel 1999;
Dohle 2001). Several independent phylogenetic analyses based on molecular data support a chelicerate/
myriapod sister group relationship, the so-called
Myriochelata hypothesis (Friedrich and Tautz 1995;
Hwang and others 2001; Kusche and Burmester
2001; Nardi and others 2003; Mallatt and others
2004; Pisani and others 2004), a link that had never
been considered by comparison of morphological
structures. However, recent comparative studies on
neurogenesis in the diplopod Glomeris marginata
and the chilopod Lithobius forficatus have shown that
the myriapods and the chelicerates share several features that cannot be found in homologous form in
From the symposium “The New Microscopy: Toward a Phylogenetic Synthesis” presented at the annual meeting of the Society for Integrative and
Comparative Biology, January 4–8, 2005, at San Diego, California.
1 E-mail: [email protected]
2 Present address: Johannes-Gutenberg University Mainz, Department of Genetics, Johann-Joachim-Becherweg 32, 55099 Mainz, Germany.
Integrative and Comparative Biology, volume 46, number 2, pp. 195–206
doi:10.1093/icb/icj020
Advance Access publication February 16, 2006
Ó The Society for Integrative and Comparative Biology 2006. All rights reserved. For permissions, please email: journals.permissions@
oxfordjournals.org.
196
A. Stollewerk and A. D. Chipman
insects
crustaceans
chelicerates/myriapods
Fig. 1 Differences in the formation of neural precursors in
the arthropod groups. In insects and crustaceans, single
neural precursors (neuroblasts) are specified. Whereas
insect neuroblasts delaminate into the embryo shortly
after formation, crustacean neuroblasts remain in the
outer cell layer (neuroectoderm) and divide to give rise
to ganglion mother cells that are pushed into the
interior of the embryo by directed mitosis. In both
chelicerates and myriapods, groups of neural precursors
are selected and form invagination sites that eventually
detach from the apical surface and differentiate into
neural cells.
insects and crustaceans. The most distinctive difference
is that groups of neural precursors are singled out from
the neuroectoderm of the spider and the myriapods,
rather than individual cells (that is, neuroblasts) as in
insects or crustaceans (Fig. 1) (Cupiennius salei:
Stollewerk and others 2001; Limulus polyphemus:
Mittmann 2002; C. salei: Stollewerk 2002; Stollewerk
and others 2003; G. marginata: Dove and Stollewerk
2003; L. forficatus: Kadner and Stollewerk 2004).
Here we give an overview of the modes of neurogenesis in the major arthropod groups with special
focus on myriapods and chelicerates. Furthermore,
we present new data on the geophilomorph centipede
Strigamia maritima (Myriapoda) and discuss the data
in a phylogenetic context.
Neural precursor formation
in insects
Neurogenesis has been studied in detail in the insect
Drosophila melanogaster. The ventral neuroectoderm
of the Drosophila embryo gives rise to both neural
and ectodermal cells (Jiménez and Campos-Ortega
1979, 1990; Cabrera and others 1987). The competence
to adopt the neural fate depends on the presence of
the proneural genes achaete, scute, and lethal of scute.
These genes are expressed in clusters of cells in each
hemisegment at the beginning of neurogenesis. In a
second step, proneural gene expression is restricted
to a single cell of the cluster, the future neuroblast
(Cabrera and others 1987; Romani and others 1987;
Skeath and others 1992). This process is called lateral
inhibition and is mediated by the neurogenic genes
Notch and Delta (Simpson 1990; Martin-Bermudo
and others 1995; Heitzler and others 1996; Seugnet
and others 1997). It has been predicted that proneural
gene expression is higher in a particular cell of the
proneural cluster as a result of predetermination or
an extrinsic signal. Since the proneural genes activate
the expression of Delta, Delta is also up-regulated in
this cell. Delta binds to Notch and activates Notch in
the neighboring cells, which eventually leads to the
activation of the E(spl) genes. The gene products of
this complex repress proneural gene expression,
which in turn leads to a down-regulation of Delta in
neighboring cells (Nakao and Campos-Ortega 1996;
Ligoxygakis and others 1998). As a result of this feedback loop, proneural gene expression is maintained in
the neuroblast but down-regulated in the remaining
cells of the proneural cluster. Although this model
predicts a higher expression of Delta in single cells
(presumptive neuroblasts), it has not been demonstrated that Delta transcripts accumulate at higher
levels in individual cells within the proneural clusters.
Once a neuroblast is determined, it delaminates into
the embryo and divides asymmetrically to produce
ganglion mother cells (Goodman and Doe 1993). The
ganglion mother cells divide only once to give rise to
neural cells that differentiate into neurons and glia.
The neuroblasts do not delaminate all at once but
in 5 discrete waves. Each neuroblast has a distinct identity and gives rise to an invariant lineage of neural
progenies. The identity of the neuroblasts is specified
in the ventral neuroectoderm by segment polarity and
dorsoventral patterning genes (see review in Skeath
1999).
Neurogenesis has also been studied in insects
other than Drosophila. The pattern of neuroblasts is
similar in all insects analyzed: they are arranged in
7 anteroposterior rows with 3–6 neuroblasts each
(Bate 1976; Broadus and Doe 1995; Wheeler SR
and others 2003). It has been shown in Tribolium
castaneum and in Schistocerca americana that single
neuroblasts are selected in sequential waves, similar
to D. melanogaster (Broadus and Doe 1995; Wheeler
SR and others 2003). Within the insect group,
proneural genes have been identified in several
Diptera, a butterfly, and the flour beetle T. castaneum
(Precis coenia: Galant and others 1998; Ceratitis
capitata: Wülbeck and Simpson 2000; Calliphora
vicina: Pistillo and others 2002; Phormia terranovae:
197
Neurogenesis in myriapods and chelicerates
Skaer and others 2002; Anopheles gambiae: Wülbeck
and Simpson 2002; T. castaneum: Wheeler SR and
others 2003).
Neural precursor formation in
malacostracan crustaceans
Neuroblasts have also been described in malacostracan
crustaceans and exist perhaps also in branchiopods
(Leptochelia spp.: Dohle 1972; Diastylis rathkei: Dohle
1976; Neomysis integer: Scholtz 1984; Peracarida: Dohle
and Scholtz 1988; Gammarus pulex: Scholtz 1990;
Cherax destructor: Scholtz 1992; Leptodora kindti:
Gerberding 1997; Decapoda: Harzsch and others
1998; Harzsch 2001; Scholtz and Gerberding 2002;
Harzsch 2003). However, there are several differences
from insect neuroblasts. Neuroblasts in malacostracan
crustaceans are generated by so-called ectoteloblasts,
specialized stem cells that are located in the posterior
region of the germ band anterior to the proctodeum
(with the exception of Amphipoda; Scholtz 1990).
Furthermore, crustacean neuroblasts do not delaminate into the embryo but remain in the outer surface.
Similar to insects, crustacean neuroblasts divide asymmetrically to give rise to smaller ganglion mother cells
that are pushed into the embryo by directed mitosis
(Scholtz 1992). The ganglion mother cells also divide
once to produce 2 neurons. In contrast to insects,
crustacean neuroblasts can generate epidermal cells
after budding off ganglion mother cells (Scholtz
and Gerberding 2002). Two achaete-scute homologues
have been identified in the branchiopod crustacean
Triops longicaudatus (Wheeler SR and Skeath 2005).
The expression pattern of these genes is similar to
the distribution of transcripts of the Achaete-Scute
Complex genes in Drosophila.
Neural precursor formation in
chelicerates and myriapods
In a few classical accounts, neuroblasts have been
described in 3 chelicerate species, but it is possible
that the data were partly misinterpreted owing to
technical limitations at the time (Yoshikura 1955;
Mathew 1956; Winter 1980). Apart from these studies,
the literature suggests that neurogenesis occurs by
a generalized inward proliferation of neuroectodermal cells to produce paired segmental thickenings in chelicerates and myriapods (Anderson 1973).
However, recent analyses of neurogenesis in 2 chelicerates (both spiders) and 2 myriapods (a diplopod and
a chilopod) have revealed that in contrast to insects
and crustaceans, groups of neural precursors are specified for neural fate (Fig. 1) in both myriapods and
chelicerates (Stollewerk and others 2001; Mittmann
1
2
3
4
5
6
7
Glomeris marginata
1
2
3
4
5
6
7
Cupiennius salei
Fig. 2 The invaginating neural precursor groups show
a similar arrangement in chelicerates and in myriapods.
Neural precursor groups are arranged in 7 rows with
3–6 invagination sites each, in both chelicerates and
myriapods. The order of formation is different in the
spider C. salei and in the centipede L. forficatus compared
with the diplopod G. marginata. In both Lithobius
(not shown) and Cupiennius, the first invagination sites
(black) arise in a coherent anterior-lateral region of
each hemisegment, whereas in Glomeris the first neural
precursor groups (black) are distributed over the
hemisegment. Subsequent invagination sites (white, gray,
striped) also arise at different positions in Cupiennius
and Glomeris.
2002; Stollewerk 2002; Dove and Stollewerk 2003;
Kadner and Stollewerk 2004). Although the neural precursors arise in 4 sequential waves in regions that are
prefigured by proneural genes, similar to Drosophila,
the precursor groups form invagination sites that
persist in the ventral neuroectoderm during the entire
course of neurogenesis. Approximately 30 invagination
groups per hemisegment detach from the apical surface
at about the same time, 3 days after the beginning of
neurogenesis. Interestingly, the invagination groups
show a similar pattern in the myriapods and the
spider (Fig. 2): they are arranged in 7 transverse rows
with 3–6 invagination sites each (Stollewerk and others
2001; Dove and Stollewerk 2003; Kadner and
Stollewerk 2004). Although the final pattern of invagination sites is similar in both groups (Fig. 2), the
order of formation of the individual invagination
groups is different. In both the spider C. salei and
the chilopod L. forficatus, the first invagination sites
arise in the anteriolateral region of each hemisegment,
whereas in the diplopod G. marginata, the first invaginating groups are visible in the middle of each
hemisegment (Stollewerk and others 2001; Dove and
Stollewerk 2003; Kadner and Stollewerk 2004). One can
be speculate that this difference in timing has an impact
on the identity of the neural precursors in Glomeris
compared with the spider and the chilopod, as genes
that are involved in neural diversity might be expressed during different time windows in the ventral
neuroectoderm.
198
To obtain more data on diverse myriapod groups,
we analyzed neurogenesis in the geophilomorph
centipede S. maritima. In contrast to Lithobius and
Glomeris, Strigamia undergoes so-called epimorphic
development. Myriapods showing this kind of development generate all segments during embryogenesis,
whereas in Lithobius and Glomeris further segments
are added during posthatching larval stages. Since
Strigamia does not have a considerably longer period
of embryogenesis (approximately 30 days [Arthur and
Chipman 2005], compared with approximately 15 days
in Cupiennius, Lithobius, and Glomeris), the 50 or so
segments must arise and differentiate in quick succession. This raises the question of whether neural precursor formation is altered in adaptation to an
accelerated development of individual segments.
In Strigamia segments arise from a posterior
undifferentiated disc (Chipman and others 2004b).
As segments are added sequentially, the older segments
begin to differentiate, with the first signs of neurogenesis becoming apparent approximately 5–6
segments anterior to the undifferentiated area that is
the fifth or sixth youngest segment (Chipman and
Stollewerk 2006). As the segments arise from the
posterior disc, they are broad in their mediolateral
extent and anteroposteriorly compressed (Fig. 3A
and B). Shortly after their first appearance, they
separate into clear left and right hemisegments.
Morphogenetic movements cause individual segments
to broaden along the anteroposterior axis while the
mediolateral extent is reduced (Fig. 3C). Later in
development, after all of the segments have been
generated, the left and right halves of the germ band
drift apart in a process known as lateral migration
(Kettle and others 2003; Chipman and others 2004b).
Throughout development, there is an anterior to
posterior gradient in the degree of differentiation
of individual segments, spanning a wide range of
stages in the neurogenic process. This allows the
whole course of neurogenesis to be observed in a
small number of specimens.
Similar to the spider and the other myriapods,
Strigamia has approximately 30 invagination sites
per hemisegment (based on counts of invagination
sites at different axial positions in multiple embryos).
In the narrow posterior segments, they arise at
stereotypical positions (see below) and are eventually arranged in 3 rows (Fig. 3B). Interestingly, the
morphogenetic movements that reduce the mediolateral extent of the segments lead to an arrangement
of the invagination sites that is similar to the other
myriapods and the spider: they are arranged in 7
rows with 3–6 invagination sites each (Fig. 3C;
compare with Glomeris Fig. 3D).
A. Stollewerk and A. D. Chipman
Fig. 3 Pattern of invagination sites in the geophilomorph
centipede S. maritima. Flat preparations of stage 5a
embryos (for staging, see Chipman and Stollewerk 2006)
stained with phalloidin-FITC; anterior is toward the top,
medial to the left in (B–D). (A) In the posterior region
of the germ band, the segments are broad in their
mediolateral extent and anteroposteriorly compressed.
(B) Strigamia has about 30 invagination sites per
hemisegment, similar to the spider and the other
myriapods. All invagination sites are already present in
the narrow posterior segments. They are arranged in
3 rows and arise at stereotypical positions.
(C) Morphogenetic movements that reduce the
mediolateral extent of the segments lead to an
arrangement of the invagination sites that is similar to
that in the other myriapods and the spider. In each
anterior hemisegment, the neural precursor groups are
arranged in 7 rows with 3–6 invagination sites each.
(D) Similar pattern of invagination sites in a hemisegment
of the diplopod G. marginata. ant, antennal segment;
ic, intercalary segment; md, mandibular segment;
mx1, maxillary segment 1; mx2, maxillary segment 2;
mxp, maxillipede segment; l1 to l4, trunk segments
corresponding to leg pairs 1–4; l15 to l17, trunk
segments corresponding to leg pairs 15–17; 1–7,
row of invagination sites 1–7.
Proneural genes in the spider and
the myriapods
In Drosophila the proneural genes are essential for
neural fate. The genes of the so-called Achaete-Scute
Complex achaete, scute, and lethal of scute are expressed
prior to formation of the neuroblasts in the ventral
neuroectoderm (Jiménez and Campos-Ortega 1979,
1990; Cabrera and others 1987; Martin-Bermudo
and others 1991). Mutations in these genes lead to
Neurogenesis in myriapods and chelicerates
the absence of neuroblasts. Homologues of achaetescute have been identified in the spider C. salei
and the myriapods G. marginata and L. forficatus
(Stollewerk and others 2001; Dove and Stollewerk
2003; Kadner and Stollewerk 2004). As in Drosophila,
these homologues can be detected in regions of the
neuroectoderm where neural precursors will be
generated hours later. Expression is up-regulated in
the neural precursor groups, and transcription is
down-regulated in the surrounding cells. This expression pattern is different from that in Drosophila,
where proneural transcripts become restricted to single
cells (neuroblasts). Functional studies in the spider
have shown that the achaete-scute homologues
are essential for neural fate, similar to the case of
Drosophila (Stollewerk and others 2001).
Neurogenic genes in the spider
and the myriapods
It has been shown in Drosophila that the neurogenic
genes Notch and Delta are responsible for the restriction of proneural gene expression to a single cell of a
cluster (Simpson 1990; Martin-Bermudo and others
1995; Heitzler and others 1996; Seugnet and others
1997). However, a dynamic expression of Delta
(mRNA and protein) that correlates with the specification of neuroblasts has not been observed in the
central nervous system of fly embryos, although it is
assumed that within a proneural cluster the cell
expressing the highest level of Delta is selected for
the neural fate. Similarly, Notch seems to be expressed
at homogeneous levels in all ventral neuroectodermal cells, indicating that Notch transcripts are
not excluded from neural precursors (Heitzler and
Simpson 1993).
Two Delta homologues, CsDelta1 and CsDelta2,
have been identified in the spider C. salei, and 1
Delta homologue each in the myriapods G. marginata
and L. forficatus (Stollewerk 2002; Dove and Stollewerk
2003; Kadner and Stollewerk 2004). In contrast to
the case of flies, expression of the spider CsDelta2
and the myriapod Delta genes can be correlated with
the formation of neural precursors. Delta transcripts
can be detected in all neuroectodermal cells but
accumulate in the invaginating neural precursors.
Furthermore, the spider and myriapod Notch homologues (1 in each species) show a heterogeneous
expression pattern throughout neurogenesis. The
up-regulation of Notch in distinct regions in the spider
and the myriapods might correlate with the formation of invagination sites, but this has to be analyzed
in more detail. Functional studies in the spider revealed
that Notch and Delta mediate lateral inhibition,
199
similar to the case of Drosophila, although groups of
neural precursors, rather than single cells, are selected.
We have identified 1 Delta and 1 Notch homologue
in S. maritima (Chipman and Stollewerk 2006).
StmNotch shows a heterogeneous expression pattern
similar to the spider and the other myriapods (data
not shown). However, the expression pattern of the
Strigamia Delta gene is different from that in the
other euarthropod groups (Fig. 4). First, Delta expression reveals that invagination sites are added continuously during neurogenesis. In the most posterior
segment of the Strigamia embryo that exhibits neurogenesis (representing the earliest stages of neurogenesis), 2 invagination sites are visible (Fig. 4A). In
the next anterior (developmentally older) segment,
an additional invagination site has been generated,
and in the next anterior segment a further 2 invagination sites have been added. This pattern suggests that
Fig. 4 Expression pattern of Strigamia Delta. Flat
preparations (A and B) and sagittal section (C) of
embryos (stage 5b) stained for a DIG-labeled StmDelta
probe. Anterior is toward the top in (A) and (B), and
to the left in (C). (A) Invagination sites are added
continuously in Strigamia. In the most posterior segment
of the Strigamia embryo that exhibits neurogenesis
(representing the earliest stages of neurogenesis),
2 invagination sites are visible. In the next anterior
(developmentally older) segment, an additional
invagination site has been generated (3) and in the next
anterior segment 2 further invagination sites have been
added (4 and 5). (B and C) Flat preparation (B) and
sagittal section (C) through the anterior region of the
germ band (maxillary segment 2 to trunk segment 4).
StmDelta transcripts seem to accumulate at higher levels
in single cells within the invagination groups (arrows).
The single cells are surrounded by cells expressing
lower levels of Delta.
200
invagination sites are added continuously during
neurogenesis, rather than in several distinct waves as
in the spider and the other myriapods.
Furthermore, StmDelta transcripts seem to
accumulate at higher levels in single cells within the
invagination groups (arrows in Fig. 4B and C). The
single cells are surrounded by cells expressing lower
levels of Delta (Fig. 4). To understand this expression
pattern, we further analyzed the morphology of
the invaginating cell groups by staining Strigamia
embryos with phalloidin-FITC, a dye that stains the
actin cytoskeleton. An accumulation of actin around
single cells in the ventral neuroectoderm was observed
by confocal microscopy (Fig. 5A). A detailed analysis
of the morphology of the invagination groups revealed
that this staining is due to the cell processes of
the cells of individual invagination groups that are
attached to a single cell of the group (Fig. 5B
and C). These data suggest that StmDelta transcripts
are not present at higher levels in single cells but
accumulate around single cells within invagination
groups as a result of this distinct morphological
arrangement.
Fig. 5 F-actin accumulates around single cells in the
ventral neuroectoderm of Strigamia. Flat preparations
of embryos (stage 5a) stained with phalloidin-FITC
(A and B). Anterior is toward the top. (A) An
accumulation of F-actin can be seen at different
apicobasal levels in the ventral neuroectoderm of
Strigamia (arrow). (B) A higher magnification of
invagination groups reveals that this staining is due to
the cell processes of the cells of individual invagination
groups that are attached to a single cell of the group.
The white square borders an invagination group. (C) The
schematic drawing shows the distinct morphological
arrangement of an invagination group in Strigamia. ant,
antennal segment; ic, intercalary segment; md,
mandibular segment; mx1, maxillary segment 1; mx2,
maxillary segment 2; mxp, maxillipede segment; l1, trunk
segment corresponding to leg pair 1; ml, ventral midline.
A. Stollewerk and A. D. Chipman
Specification of neuroblast identity
in arthropods
In Drosophila, segment polarity genes and dorsoventral
patterning genes are expressed during neurogenesis
in the ventral neuroectoderm (see review in Skeath
1999). These genes subdivide the neuroectoderm
into a gridlike pattern, so that each proneural cluster
shows a different gene expression profile. The neuroblasts maintain the specific expression pattern of the
proneural cluster from which they delaminate and give
rise to an invariant lineage of distinct neural progenies.
Thirty neuroblasts delaminate from the neuroectoderm
of each hemisegment. They are arranged in 7 rows
with each row expressing a different subset of segment
polarity genes. It has been shown in Drosophila that the
function of the segment polarity genes is specifically
required in neuroblasts. Mutations in these genes lead
either to the absence of specific neuroblasts or to
changes in the identity of neuroblasts (Skeath 1999).
The specification of neuroblast identity has not
been analyzed in any detail in arthropods other than
Drosophila. Although segment polarity genes have
been identified in other insects, in crustaceans, in
myriapods, and in a spider, their function during
neurogenesis has not been studied except for engrailed
(Patel and others 1992; Brown and others 1994, 1997;
Dawes and others 1994; Patel 1994; Damen and others
2000; Telford 2000; Davis and others 2001; Damen
2002; Dearden and others 2002; Hughes and
Kaufman 2002; Mouchel-Vielh and others 2002;
Copf and others 2003; Kettle and others 2003;
Chipman and others 2004a, 2004b; Eckert and others
2004; Janssen and others 2004; Peel 2004). However,
Patel and coworkers (1989) investigated the expression
pattern of the segment polarity gene engrailed in several
insects and crustaceans and showed that engrailed
expression in neuroblasts is conserved. In all species
analyzed, engrailed is expressed in neuroblast rows 6
and 7, and 1 neuroblast of row 1.
We have analyzed engrailed expression in the
spider Cupiennius and the geophilomorph centipede
Strigamia. In both the spider and the centipede,
engrailed is expressed in segmental stripes in the posterior region of the germ band (Fig. 6A and D) (Damen
2002; Kettle and others 2003; Chipman and others
2004b). In more anterior, developmentally advanced
segments that are undergoing neurogenesis, engrailed
expression covers a broader region at the posterior
border of the segments. In addition, in the central area
of the ventral neuroectoderm the engrailed expression
domain extends into the anterior region of the next
posterior segments, whereas the engrailed stripe lateral
to the limb buds is still restricted to a few cell rows at
201
Neurogenesis in myriapods and chelicerates
Fig. 6 Comparison of engrailed expression in the spider
Cupiennius and the geophilomorph centipede Strigamia.
Flat preparations (A and B) and sagittal sections
(C and D) of embryos stained for DIG-labeled engrailed
probes. Anterior is toward the top in (A) and (B), and
to the left in (C) and (D). (A) In Cupiennius, engrailed is
expressed in small dorsoventral stripes in newly formed
segments in the posterior region of the germ band
(arrowhead). In the anterior region of the germ band
(190 h after egg laying), engrailed expression covers a
broader region at the posterior border of the segments
and extends into the anterior region of the next
posterior segments (arrows). The anterior expression
domain is restricted to the mediocentral region of each
hemisegment. The black lines indicate the segmental
borders. (B) In Strigamia (stage 5b) engrailed expression
extends into the groove (arrow). In contrast to the
spider, engrailed is also expressed in the ventral midline.
The black lines indicate the segmental borders.
(C) The sagittal section through a ventral neuromere of
the spider shows that engrailed is expressed not only in
the outer neuroectodermal cell layer but also in the
basally located invaginating neural precursors.
The arrowhead indicates the segmental border. (D) In
Strigamia the segments have a peak and trough structure.
As in Cupiennius, engrailed is expressed in a small stripe
in the posterior region of the germ band that coincides
with the posterior border of the segments
(arrowheads). In more anterior segments that are
undergoing neurogenesis, engrailed expression also
covers the anterior region of the segments (arrows).
We have compared the pattern of engrailed expression with the pattern of invagination sites in the spider.
Single-color double-staining with engrailed and antihorseradish peroxidase, which is exclusively expressed
in the cell processes of the invaginating neural precursor groups at this time, revealed that engrailed
is expressed in the invagination groups of rows 6
and 7 and row 1 in the spider (Fig. 7A–D).
Interestingly, there are 7 invagination sites in rows 6
and 7 and this number is identical to the number of
neuroblasts that are engrailed positive in rows 6 and 7
in insects and crustaceans (Patel and others 1989;
Duman-Scheel and Patel 1999).
We have also compared engrailed expression with
the position of invagination sites in the geophilomorph
centipede. In Strigamia the segments have a peak and
trough structure that is most obvious in sagittal sections (Figs. 6D and 8D). In the posterior region of
the germ band the engrailed stripe divides the trough
into 2 halves (Fig. 6D). Based on this expression
pattern and the fact that the posterior border of the
engrailed domain coincides with the posterior border
of segments in all arthropods analyzed, we conclude
that the posterior half of the trough belongs to the next
posterior segment. In more anterior segments that
exhibit neurogenesis, engrailed is expressed throughout
the trough, indicating that it is not only expressed at
the posterior border of the segments but also in neural
precursors in the anterior region of each segment.
Analysis of phalloidin-FITC-stained embryos revealed
that the neural precursor groups that belong to
the anterior row of invagination sites extend into the
groove (Fig. 8B and C). Similarly, the groups that
belong to the posterior rows extend into the groove
from the other site (Fig. 8B). Double-stainings with
Delta and engrailed suggest that engrailed is expressed
in the first anterior row of invagination groups and
both in rows 6 and 7 (Fig. 8E).
Conclusions
the posterior border of the segments (Fig. 6A and B,
arrows; Fig. 7A; Damen 2002). In spiders and
myriapods, all cells of the ventral neuroectoderm
give rise to neural cells. The epidermis arises lateral
to the neuromeres only after invagination of the neural
precursors (Stollewerk 2002; Dove and Stollewerk
2003; Stollewerk 2004). Therefore, it can be concluded
that during neurogenesis, engrailed is specifically
expressed in neural precursors.
In Strigamia, engrailed is expressed in the ventral
midline (Fig. 6B). Further analysis will show whether
this expression corresponds to an accumulation of
transcripts in neural cells.
We have presented comparative morphological and
molecular data on neurogenesis in the euarthropod
groups. Although there are differences in the formation
of neural precursors, most arthropod species analyzed
generate approximately 30 single neural precursors
(insects/crustaceans) or precursor groups (chelicerates/
myriapods) per hemisegment, which are arranged in
regular rows. Homologues of achaete-scute are necessary for the formation of neural precursors, and the
neurogenic genes Notch and Delta restrict the proportion of cells that adopt a neural fate at a certain
time. In insects, chelicerates, and 2 of the 3 myriapods
analyzed, neural precursors are produced in several
202
A. Stollewerk and A. D. Chipman
Fig. 7 Comparison of the pattern of engrailed expression with the pattern of invagination sites in the spider C. salei.
Flat preparations of embryos stained for DIG-labeled engrailed probes (A–C), anti-horseradish peroxidase (B and C),
and with phalloidin-FITC (D). The black and white lines indicate the segmental borders. (A) Expression domain of
engrailed in segments that exhibit neurogenesis (190 h after egg laying). (B and C) Single-color double-staining with
engrailed and anti-horseradish peroxidase. At this stage (190 h of development) the horseradish peroxidase antigen is
exclusively expressed in the cell processes of the invaginating neural precursors, as seen on the apical view of the
ventral neuroectoderm (B). On the basal view of the same area, engrailed expression is visible in the neural precursors
of rows 6, 7 and 1 (C). (D) Pattern of invagination sites in an embryo of the same stage stained with phalloidin-FITC.
Note that there are 7 invagination sites in rows 6 and 7. The segmental border is clearly visible because of the distinct
shape of the border cells: they are mediolaterally elongated. 1, 6, 7: row of invagination sites 1, 6, 7.
Fig. 8 Comparison of engrailed expression with the position of invagination sites in the geophilomorph centipede
Strigamia (stage 5b). Flat preparations of embryos stained with phalloidin-FITC (A–C), flat preparation of an embryo
double-stained for a DIG-labeled Stmengrailed probe (blue) and a Fluorescein-labeled StmDelta probe (red) (Chipman
and others 2004a) (E), and sagittal section of an embryo stained for a DIG-labeled Stmengrailed probe (D). (A–B) Apical
and basal optical section, respectively, of the same anterior region of the ventral neuroectoderm (2 hemisegments).
The brackets indicate the extension of the groove. The neural precursors extend into the groove from anterior and
posterior (arrows in [B]). (C) Three hemisegments of the posterior region of the germ band. The arrow indicates neural
precursor groups that belong to the first row of invagination sites and extend into the groove. (D) engrailed expression
in neural precursors (arrows). (E) engrailed is expressed in rows 6 and 7 and throughout the groove, indicating that
transcripts also accumulate in neural precursors of row 1 (arrow). The white lines indicate the segmental borders.
sequential waves. Neural precursor formation has
been analyzed in only a limited number of crustacean
species (Dohle 1972, 1976; Scholtz 1984, 1990, 1992;
Harzsch and Dawirs 1994, 1996; Harzsch and others
1998; Gerberding and Scholtz 1999; Harzsch 2001,
2003). Based on the small amount of data available,
it can be assumed that neuroblasts in crustaceans
are continuously added during neurogenesis, rather
than being generated in several waves. The continuous
addition of neural precursor groups in the
centipede Strigamia might be an adaptation to the
distinct embryonic development of this species.
Approximately 50 segments are generated during
embryogenesis and differentiate in quick succession.
Gene expression studies and morphological analyses
revealed that each segment exhibits a different differentiation state along the anterior-posterior axis during
neurogenesis. Therefore, it can be concluded that each
segment initiates neurogenesis on its own, rather than
being synchronized with several segments, as seen in
203
Neurogenesis in myriapods and chelicerates
the spider and in the other myriapods. As has been
shown previously, neurogenesis occurs simultaneously
in the prosoma of the spider and in the head and first
trunk segments of G. marginata and L. forficatus, and at
least 2 to 3 segments show the same stage of neurogenesis in the posterior region of the germ band
(Stollewerk and others 2001; Dove and Stollewerk
2003; Kadner and Stollewerk 2004). Despite the differences in neural precursor formation in the euarthropod
group, neural precursor identity seems to be achieved
in a similar way. In all species analyzed, engrailed is
expressed in neural precursor rows 6, 7, and 1.
In addition to these conserved features, we found
2 characteristics that distinguish insects/crustaceans
from myriapods/chelicerates. First, in insects and
crustaceans the neuroectoderm gives rise to epidermal
and neural cells. In contrast, there is no decision
between epidermal and neural fate in the central region
of the ventral neuroectoderm of chelicerates and
myriapods. The epidermis arises lateral to the neuromeres and overgrows the ventral nerve cord after
formation of neural precursors. However, this characteristic seems to be ancestral (plesiomorphic), as it
has been shown in onychophorans (a group that is
assumed to be basal to the arthropods) that the
whole central regions of the hemisegments sink into
the embryo and thus give only rise to neural cells
(Eriksson and others 2003). The second distinguishing
characteristic is the presence of neuroblasts in insects/
crustaceans as opposed to neural precursor groups in
myriapods/chelicerates. Invaginating cell groups have
not been found in onychophorans or in tardigrades
(another potential outgroup to the euarthropods
(Eriksson and others 2003; Hejnol and Schnabel
2005). However, only 2 species have been analyzed,
which might be derived and thus do not represent
the ancestral state. The fact that approximately 30
neural precursors/precursor groups per hemisegment
are arranged in a strikingly similar pattern in most
euarthropod species analyzed suggests that a similar
pattern was present in the last common ancestor of
the arthropods. The formation of neural precursor
groups could be a sympleisiomorphy of myriapods
and chelicerates that has been lost in the more derived
Pancrustacea. However, it is also possible that this
characteristic represents a synapomorphy of myriapods
and chelicerates, providing the first morphological
support for the Myriochelata group (Friedrich and
Tautz 1995; Hwang and others 2001; Kusche and
Burmester 2001; Nardi and others 2003; Mallatt
and others 2004; Pisani and others 2004). At present,
we cannot distinguish between these 2 scenarios,
but the data presented in this review are clearly
inconsistent with the Atelocerata hypothesis, which
unites myriapods and insects, to the exclusion of
crustaceans. These data are intriguing and warrant
further research into neurogenesis in putative
arthropod sister groups. Additional data will allow a
polarization of the characteristic state changes and
help resolve the question of the relationships between
the major arthropod groups.
Acknowledgments
We thank the organizers of the symposium for the
opportunity to present our work. We are grateful to
Michael Akam and Pat Simpson for providing lab
space and for helpful discussions. Thanks to Pat
Simpson for critical reading of the manuscript. The
Deutsche
Forschungsgemeinschaft
(A.S.)
and
Federation of European Biochemical Societies
(A.D.C.) supported this research.
References
Anderson D. 1973. Embryology and phylogeny in annelids and
arthropods. Oxford: Pergamon Press.
Arthur W, Chipman AD. 2005. The centipede Strigamia
maritima: what it can tell us about the development and
evolution of segmentation. BioEssays 27:653–60.
Ballard JWO, Olsen GJ, Faith DP, Odgers WA, Rowell DM,
Atkinson PW. 1992. Evidence from 12S ribosomal RNA
sequences that onychophorans are modified arthropods.
Science 258:1345–8.
Bate M. 1976. Embryogenesis of an insect nervous system:
I. A map of thoracic and abdominal neuroblasts in Locusta
migratoria. J Embryol Exp Morphol 35:107–23.
Bergström J. 1992. The oldest Arthropoda and the origin of the
Crustacea. Acta Zool 73:287–91.
Bitsch C, Bitsch J. 2004. Phylogenetic relationships of basal
hexapods among the mandibulate arthropods: a cladistic
analysis based on comparative morphological characters.
Zool Scr 33:511–50.
Briggs DEG, Fortey RA. 1989. The early radiation and relationships of the major arthropod groups. Science 246:241–3.
Broadus J, Doe CQ. 1995. Evolution of neuroblast identity:
seven-up and prospero expression reveal homologous and
divergent neuroblast fates in Drosophila and Schistocerca.
Development 121:3989–96.
Brown SJ, Parrish JK, Beeman RW, Denell RE. 1997. Molecular
characterization and embryonic expression of the evenskipped ortholog of Tribolium castaneum. Mech Dev
61:165–73.
Brown SJ, Parrish JK, Denell RE, Beeman RW. 1994. Genetic
control of early embryogenesis in the red flour beetle,
Tribolium castaneum. Am Zool 34:343–52.
Cabrera CV, Martinez-Arias A, Bate M. 1987. The expression of
three members of the achaete-scute gene complex correlates
with neuroblast segregation in Drosophila. Cell 50:425–33.
204
A. Stollewerk and A. D. Chipman
Chipman AD, Arthur W, Akam M. 2004a. A double segment
periodicity underlies segment generation in centipede
development. Curr Biol 14:1250–5.
Eckert C, Aranda M, Wolff C, Tautz D. 2004. Separable stripe
enhancer elements for the pair-rule gene hairy in the beetle
Tribolium. EMBO Rep 5:638–42.
Chipman AD, Arthur W, Akam M. 2004b. Early development
and segment formation in the centipede, Strigamia maritima
(Geophilomorpha). Evol Dev 6:78–89.
Emerson MJ, Schram FR. 1998. Theories, patterns, and reality:
Game plan for arthropod phylogeny. In: Fortey RA,
Thomas RH, editors. Arthropod relationships London:
Chapman & Hall. p 67–86.
Chipman AD, Stollewerk A. 2006. Specification of neural precursor identity in the geophilomorph centipede Strigamia
maritima. Dev Biol 290:337–350.
Copf T, Rabet N, Celniker SE, Averof M. 2003. Posterior patterning genes and the identification of a unique body region
in the brine shrimp Artemia franciscana. Development
130:5915–27.
Damen W. 2002. Parasegmental organization of the spider
embryo implies that the parasegment is an evolutionary
conserved entity in arthropod embryogenesis. Development
129:1239–50.
Damen W, Weller M, Tautz D. 2000. Expression patterns of
hairy, even-skipped, and runt in the spider Cupiennius salei
imply that these genes were segmentation genes in a basal
arthropod. Proc Natl Acad Sci USA 97:4515–19.
Davis GK, Jaramillo CA, Patel NH. 2001. Pax group III genes
and the evolution of insect pair-rule patterning. Development
128:3445–58.
Dawes R, Dawson I, Falciani F, Tear G, Akam M. 1994. Dax,
a locust Hox gene related to fushi-tarazu but showing no
pair-rule expression. Development 120:1561–72.
Dearden PK, Donly C, Grbic M. 2002. Expression of pair-rule
gene homologues in a chelicerate: early expression patterning
of the two-spotted spider mite Tetranychus urticae.
Development 129:5461–72.
Dohle W. 1972. Über die Bildung und Differenzierung des
postnauplialen Keimstreifs von Leptochelia spec. Crustacea,
Tanaidacea). Zool Jb Anat 89:503–66.
Dohle W. 1976. Die Bildung und Differenzierung des
postnauplialen Keimstreifs von Diastylis rathkei (Crustacea,
Cumacea) II. Die Differenzierung und Musterbildung des
Ektoderms. Zoomorphologie 84:235–77.
Eriksson BJ, Tait NN, Budd GE. 2003. Head development in the
Onychophoran Euperipatoides kanangrensis with particular
reference to the central nervous system. J Morphol 255:1–23.
Field KG, Olsen GJ, Lane DJ, Giovannoni SJ, Ghiselin MT,
Raff EC, Pace NR, Raff RA. 1988. Molecular phylogeny of
the animal kingdom. Science 239:748–53.
Friedrich M, Tautz D. 1995. Ribosomal DNA phylogeny of
the major extant arthropod classes and the evolution of
myriapods. Nature 376:165–7.
Galant R, Skeath JB, Paddock S, Lewis DL, Carroll SB. 1998.
Expression pattern of a butterfly achaete-scute homolog
reveals the homology of butterfly wing scales and insect
sensory bristles. Curr Biol 8:807–13.
Gerberding M. 1997. Germ band formation and early neurogenesis of Leptodora kindti (Cladocera): first evidence for
neuroblasts in the entomostracan crustaceans. Invertebr
Reprod Dev 32:63–73.
Gerberding M, Scholtz G. 1999. Cell lineage of the midline cells
in the amphipod crustacean Orchestia cavimana (Crustacea,
Malacostraca) during formation and separation of the germ
band. Dev Genes Evol 209:91–102.
Giribet G, Ribera C. 1998. The position of arthropods in the
animal kingdom: a search of a reliable outgroup for internal
arthropod phylogeny. Mol Phylogenet Evol 9:481–8.
Goodman CS, Doe CQ. 1993. Embryonic development of the
Drosophila central nervous system. In: Bate M, MartinezArias A, editors. The development of Drosophila melanogaster.
New York: Cold Spring Harbor Laboratory Press. p 1131–206.
Harzsch S. 2001. Neurogenesis in the crustacean ventral nerve
cord: homology of neuron stem cells in Malacostraca and
Branchiopoda? Evol Dev 3:154–69.
Dohle W. 1998. Myriapod–insect relationships as opposed to
an insect–crustacean sister group relationship. In: Fortey RA,
Thomas RH, editors. Arthropod relationships London:
Chapman & Hall. p 305–15.
Harzsch S. 2003. Ontogeny of the ventral nerve cord in
malacostracan crustaceans: a common plan for neuronal
development in Crustacea and Hexapoda? Arthropod
Struct Dev 32:17–38.
Dohle W. 2001. Are the insects terrestial crustaceans? A discussion of some new facts and arguments and the proposal
of the proper name “Tetraconata” for the monophyletic unit
Crustacea and Hexapoda. Ann Soc Entomol Fr 37:85–103.
Harzsch S, Dawirs RR. 1994. Neurogenesis in larval stages of
the spider crab Hyas araneus (Decapoda, Brachyura): proliferation of neuroblasts in the ventral nerve cord. Roux’s Arch
Dev Biol 204:93–100.
Dohle W, Scholtz G. 1988. Clonal analysis of the crustacean
segment: the discordance between genealogical and segmental
borders. Development 104 (Suppl):147–60.
Harzsch S, Dawirs RR. 1996. Neurogenesis in the developing
crab brain: postembryonic generation of neurons persists
beyond metamorphosis. J Neurobiol 29:384–98.
Dove H, Stollewerk A. 2003. Comparative analysis of neurogenesis in the myriapod Glomeris marginata (Diplopoda)
suggests more similarities to chelicerates than to insects.
Development 130:2161–71.
Harzsch S, Miller J, Benton J, Dawirs RR, Beltz B. 1998.
Neurogenesis in the thoracic neuromeres of two crustaceans
with different types of metamorphic development. J Exp Biol
201:2465–79.
Duman-Scheel M, Patel NH. 1999. Analysis of molecular marker
expression reveals neuronal homology in distantly related
arthropods. Development 126:2327–34.
Heitzler P, Bourouis M, Ruel L, Carteret C, Simpson P. 1996.
Genes of the Enhancer of split and achaete-scute complexes
are required for a regulatory loop between Notch and Delta
Neurogenesis in myriapods and chelicerates
205
during lateral signalling in Drosophila. Development
122:161–71.
Mathew AP. 1956. Embryology of Heterometrus scaber (Thorell),
Arachnida, Scorpionidae. Zool Mem Univ Travancore 1:1–96.
Heitzler P, Simpson P. 1993. Altered epidermal growth factorlike sequences provide evidence for a role of Notch as a
receptor in cell fate decisions. Development 117:1113–23.
Mittmann B. 2002. Early neurogenesis in the horseshoe crab
Limulus polyphemus and its implication for arthropod
relationships. Biol Bull 203:221–2.
Hejnol A, Schnabel R. 2005. The eutardigrade Thulinia
stephaniae has an indeterminate development and the potential to regulate early blastomere ablations. Development
1332:1349–61.
Mouchel-Vielh E, Blin M, Rigolot C, Deutsch JS. 2002.
Expression of a homologue of the fushi tarazu (ftz) gene in
a cirripede crustacean. Evol Dev 4:76–85.
Hughes CL, Kaufman TC. 2002. Exploring the myriapod body
plan: expression patterns of the ten Hox genes in a centipede.
Development 129:1225–38.
Hwang UW, Friedrich M, Tautz D, Park CJ, Kim W. 2001.
Mitochondrial protein phylogeny joins myriapods with
chelicerates. Nature 413:154–7.
Janssen R, Prpic N-M, Damen W. 2004. Gene expression
suggests decoupled dorsal and ventral segmentation in the
millipede Glomeris marginata (Myriapoda: Diplopoda).
Dev Biol 268:89–104.
Jiménez F, Campos-Ortega JA. 1979. A region of the Drosophila
genome necessary for CNS development. Nature 282:310–12.
Jiménez F, Campos-Ortega JA. 1990. Defective neuroblast
commitment in mutants of the achaete-scute complex and
adjacent genes of Drosophila melanogaster. Neuron 5:81–9.
Kadner D, Stollewerk A. 2004. Neurogenesis in the chilopod
Lithobius forficatus suggests more similarities to chelicerates
than to insects. Dev Genes Evol 214(8):367–79.
Kettle C, Johnstone J, Jowett T, Arthur H, Arthur W. 2003.
The pattern of segment formation, as revealed by engrailed
expression, in a centipede with a variable number segments.
Evol Dev 5:198–207.
Kraus O, Kraus M. 1994. Phylogenetic system of the Tracheata
(Mandibulata): on “Myriapoda”–Insecta interrelationships,
phylogenetic age and primary ecological niches. Verh
Naturwiss Ver Hamburg 34:5–31.
Nakao K, Campos-Ortega JA. 1996. Persistent expression of
genes of the Enhancer of split complex suppresses neural
development in Drosophila. Neuron 16:275–86.
Nardi F, Spinsanti G, Boore JL, Carapelli A, Dallai R, Frati F.
2003. Hexapod origins: monophyletic or paraphyletic?
Science 299:1887–9.
Nilsson DE, Osorio D. 1998. Homology and parallelism in
arthropod sensory processing. In: Fortey RA, Thomas RH,
editors. Arthropod relationships London: Chapman & Hall.
p 333–47..
Osorio D, Averof M, Bacon JP. 1995. Arthropod evolution: great
brains, beautiful bodies. Trends Ecol Evol 10:449–54.
Patel NH. 1994. Developmental evolution: insights from studies
of insect segmentation. Science 266:581–90.
Patel NH, Ball EE, Goodman CS. 1992. Changing role of evenskipped during the evolution of insect pattern formation.
Nature 357:339–42.
Patel NH, Kornberg TB, Goodman CS. 1989. Expression of
engrailed during segmentation in grasshopper and crayfish.
Development 107:201–12.
Peel A. 2004. The evolution of arthropod segmentation
mechanisms. BioEssays 26:1108–16.
Pisani D, Poling LL, Lyons-Weiler M, Hedges SB. 2004. The
colonization of land by animals: molecular phylogeny and
divergence times among arthropods. BMC Biology 2:1.
Kraus O, Kraus M. 1996. On myriapod/insect interrelationships.
Mém Mus Natl Hin Nat 169:283–90.
Pistillo D, Skaer N, Simpson P. 2002. scute expression in
Calliphora vicina reveals an ancestral pattern of longitudinal
stripes on the thorax of higher Diptera. Development
129:563–72.
Kusche K, Burmester T. 2001. Diplopod hemocyanin sequence
and the phylogenetic position of the Myriapoda. Mol Biol
Evol 18:1566–73.
Regier JC, Shultz JW, Kambic RF. 2005. Pancrustacean phylogeny: hexapods are terrestrial crustaceans and maxilopods
are not monophyletic. Proc Biol Sci 272:395–401.
Ligoxygakis P, Yu SY, Delidakis C, Baker NE. 1998. A subset of
Notch functions during Drosophila eye development require
Su(H) and E(spl) gene complex. Development 125:2893–900.
Romani S, Campuzano S, Modolell J. 1987. The achaete-scute
complex is expressed in neurogenic regions of Drosophila
embryos. EMBO J. 6:2085–92.
Mallatt JM, Garey JR, Shultz JW. 2004. Ecdysozoan phylogeny
and Bayesian inference: first use of nearly complete 28S and
18S rRNA gene sequences to classify the arthropopds and
their kin. Mol Phylogenet Evol 31:178–91.
Scholtz G. 1984. Untersuchungen zur Bildung und
Differenzierung des postnaupliaren Keimstreifs von
Neomysis integer Leach (Crustacea, Malacostraca,
Peracarida). Zool Jb Anat 112:295–349.
Martin-Bermudo MD, Carmena A, Jimenez F. 1995. Neurogenic
genes control gene expression at the transcriptional level
in early neurogenesis and in mesectoderm specification.
Development 121:219–24.
Scholtz G. 1990. The formation, differentiation and segmentation of the post-naupliar germ band of the amphipod
Gammarus pulex L. (Crustacea, Malacostraca, Peracarida).
Proc R Soc Lond B Biol Sci 239:163–211.
Martin-Bermudo MD, Martinez C, Rodriguez A, Jiménez F.
1991. Distribution and function of the lethal of scute
gene product during early neurogenesis in Drosophila.
Development 113:445–54.
Scholtz G. 1992. Cell lineage studies in the crayfish Cherax
destructor (Crustacea, Decapoda): germ band formation,
segmentation and early neurogenesis. Roux’s Arch Dev Biol
202:36–48.
206
Scholtz G, Gerberding M. 2002. Cell lineage of crustacean
neuroblasts. In: Wiese K, editor. The Crustacean nervous
system. Berlin, Heidelberg, New York: Springer. p 406–416.
Schram FR, Emerson MJ. 1991. Arthropod pattern theory: a new
approach to arthropod phylogeny. Mem Qld Mus 31:1–18.
Seugnet L, Simpson P, Haenlin M. 1997. Transcriptional
regulation of Notch and Delta: requirement for neuroblast
segregation in Drosophila. Development 124:2015–25.
Shultz JW, Regier JC. 2000. Phylogenetic analysis of arthropods
using two nuclear protein-encoding genes supports a
crustacean þ hexapod clade. Proc Biol Sci 267:1011–19.
Simpson P. 1990. Lateral inhibition and the development of the
adult sensory bristles of the peripheral nervous system of
Drosophila. Development 109:509–19.
Skaer N, Pistillo D, Simpson P. 2002. Transcriptional heterochrony of scute and changes in bristle pattern between two
closely related species of blowfly. Dev Biol 252:31–45.
Skeath JB. 1999. At the nexus between pattern formation and
cell-type specification: the generation of individual neuroblast
fates in the Drosophila embryonic central nervous system.
BioEssays 21:922–31.
Skeath JB, Panganiban G, Selegue J, Carroll SB. 1992. Gene
regulation in two dimensions: the proneural achaete and
scute genes are controlled by combinations of axis patterning
genes through a common intergenic control region. Genes
Dev 6:2606–19.
Snodgrass RE. 1938. Evolution of the Annelida, Onychophora
and Arthropoda. Smithson Misc Collect 97:1–159.
Snodgrass RE. 1950. Comparative studies on the jaws of
mandibulate arthropods. Smithson Misc Collect 116:1–85.
Snodgrass RE. 1951. Comparative studies on the head of
mandibulate arthropods. Ithaca, NY: Comstock.
Stollewerk A. 2002. Recruitment of cell groups through Delta/
Notch signalling during spider neurogenesis. Development
129:5339–48.
Stollewerk A. 2004. Secondary neurons are arrested in an
immature state by formation of epithelial vesicles during
neurogenesis of the spider Cupiennius salei. Front Zool 1:3.
Stollewerk A, Tautz D, Weller M. 2003. Neurogenesis in the
spider: new insights from comparative analysis of morphological processes and gene expression patterns. Arthrop Struct
Dev 32:5–16.
Stollewerk A, Weller M, Tautz D. 2001. Neurogenesis in the
spider Cupiennius salei. Development 128:2673–88.
Telford MJ. 2000. Evidence for the derivation of the Drosophila
fushi tarazu gene from a Hox gene orthologous to lophotrochozoan Lox5. Curr Biol 10:349–52.
A. Stollewerk and A. D. Chipman
Turberville JM, Pfeiffer DM, Field KG, Raff RA. 1991. The
phylogenetic status of arthropods as infered from 18S
rRNA sequences. Mol Biol Evol 8:669–86.
Wheeler SR, Carrico ML, Wilson BA, Brown SJ, Skeath JB. 2003.
The expression and function of the achaete-scute genes in
Tribolium castaneum reveals conservation and variation
in neural pattern formation and cell fate specification.
Development 130:4373–81.
Wheeler SR, Skeath JB. 2005. The identification and expression
of achaete-scute genes in the branchiopod crustacean Triops
longicaudatus. Gene Expr Patterns 5:695–700.
Wheeler WC. 1998. Arthropod fossils and phylogeny. In:
Edgecombe GD, editor. Arthropod fossils and phylogeny.
New York: Columbia University Press. p 9–32.
Wheeler WC, Cartwright P, Hayashi CY. 1993. Arthropod
phylogeny: a combined approach. Cladistics 9:1–39.
Whitington PM. 1996. Evolution and neural development in the
arthropods. Semin Cell Dev Biol 7:605–14.
Whitington PM, Leach D, Sandeman R. 1993. Evolutionary
change in neural development within the arthropods: axonogenesis in the embryo of two crustaceans. Development
118:449–61.
Wills MA, Briggs DEG, Fortey RA, Wilkinson M, Sneath PHA.
1998. Arthropod fossils and phylogeny. In: Edgecombe GD,
editor. An arthropod phylogeny based on fossil and recent
taxa. New York: Columbia University Press. p 33–105.
Winter G. 1980. Beiträge zur Morphologie and Embryologie
des vorderen Körperabschnitts (Cephalosoma) der
Pantopoda Gerstaecker, 1863. I. Zur Entstehung des
Zentralnervensystems. Z Zool Syst Evol Forsch 18:27–61.
Wülbeck C, Simpson P. 2000. Expression of achaete-scute
homologues in discrete proneural clusters on the developing
notum of the medfly Ceratitis capitata, suggests a common
origin for the stereotyped bristle patterns of higher Diptera.
Development 127:1411–20.
Wülbeck C, Simpson P. 2002. The expression of pannier and
achaete-scute homologues in a mosquito suggests an ancient
role of pannier as a selector gene in the regulation of the dorsal
body pattern. Development 129:3861–71.
Yoshikura M. 1955. Embryological studies on the liphistiid spider, Heptathela kimurai, part II. Kumamoto J Sci B 2:1–86.
Zrzavý J, Štys P. 1997. The basic body plan of arthropods:
insights from evolutionary morphology and developmental
biology. J Evol Biol 10:353–67.