Annelida

Abstract

Annelids are a taxon of protostomes comprising more than 1–7,0–00 worldwide–013;distributed species, which can be found in marine, limnic, and terrestrial habitats (Zhang 2–011). Their phylogeny was under discussion for a long time, but recent phylogenomic analyses resulted in a solid backbone of this group (Struck et al. 2–011; Weigert et al. 2–014). According to these analyses, most of the annelid diversity is part of Errantia or Sedentaria, which both form reciprocally monophyletic sister groups (Fig. 9.1) and are now known as Pleistoannelida (Struck 2–011). The Sedentaria also include the Clitellata, Echiura, and Pogonophora (Siboglinidae) as derived from the annelid taxa. Outside Sedentaria and Errantia, several groups can be found in the basal part of the annelid tree, namely, Sipuncula, Amphinomida, Chaetopteridae, Magelonidae, and Oweniidae. The latter two taxa together represent the sister taxon of all other annelids. Given this hypothesis, it has to be assumed that the early diversification of extant annelids took place at least in the Lower Cambrian (5–20 Ma ago) (Weigert et al. 2–014). The phylogenetic position of Myzostomida, a group of commensals or parasites of echinoderms (and, rarely, cnidarians), remains still uncertain. Whereas there is strong support for an annelid ancestry, its exact position awaits to be determined (Bleidorn et al. 2–014). Likewise, the phylogenetic position of several interstitial taxa is still under debate (Westheide 1–987; Worsaae and Kristensen 2–005; Worsaae et al. 2–005; Struck 2–006). A position of Diurodrilidae outside Annelida, as suggested by Worsaae and Rouse (2–008), was rejected by molecular data (Golombek et al. 2–013), and the position of the enigmatic Lobatocerebrum and Jennaria remains unresolved (Rieger 1–980, 1–991). Likewise, the position of Annelida within Protostomia is still uncertain. However, recent phylogenomic analyses recover a clade uniting annelids with Mollusca, Nemertea, Brachiopoda, and Phoronida, but without strong support for any sister group relationship (Edgecombe et al. 2–011).

Full text

193
A. Wanninger (ed.), Evolutionary Developmental Biology of Invertebrates 2: Lophotrochozoa (Spiralia)
DOI 10.1007/978-3-7091-1871-9_9, © Springer-Verlag Wien 2015
C. Bleidorn (*) • C. Helm • A. Weigert
Molecular Evolution and Systematics of Animals ,
Institute of Biology, University of Leipzig ,
Talstraße 33 , Leipzig D-04103 , Germany
e-mail: bleidorn@uni-leipzig.de
M. T. Aguado
Departamento de Biología, Facultad de Ciencias ,
Universidad Autónoma de Madrid ,
Canto Blanco , Madrid 28049 , Spain
9
Chapter vignette artwork by Brigitte Baldrian.
© Brigitte Baldrian and Andreas Wanninger.
Annelida
Christoph Bleidorn , Conrad Helm , Anne Weigert ,
and Maria Teresa Aguado

194
INTRODUCTION
Annelids are a taxon of protostomes compris-
ing more than 17,000 worldwide-distributed
species, which can be found in marine, limnic,
and terrestrial habitats (Zhang 2011 ). Their
phylogeny was under discussion for a long
time, but recent phylogenomic analyses
resulted in a solid backbone of this group
(Struck et al. 2011 ; Weigert et al. 2014 ).
According to these analyses, most of the anne-
lid diversity is part of Errantia or Sedentaria,
which both form reciprocally monophyletic
sister groups (Fig.
9.1 ) and are now known as
Pleistoannelida (Struck 2011 ). The Sedentaria
also include the Clitellata, Echiura, and
Pogonophora (Siboglinidae) as derived annelid
taxa. Outside Sedentaria and Errantia, several
groups can be found in the basal part of the
annelid tree, namely, Sipuncula, Amphinomida,
Chaetopteridae, Magelonidae, and Oweniidae.
The latter two taxa together represent the sister
taxon of all other annelids. Given this hypoth-
esis, it has to be assumed that the early diversi-
fi cation of extant annelids took place at least in
the Lower Cambrian (520 Ma ago) (Weigert
et al. 2014 ). The phylogenetic position of
Myzostomida, a group of commensals or para-
sites of echinoderms (and, rarely, cnidarians),
remains still uncertain. Whereas there is strong
support for an annelid ancestry, its exact posi-
tion awaits to be determined (Bleidorn et al.
2014 ). Likewise, the phylogenetic position of
several interstitial taxa is still under debate
(Westheide 1987 ; Worsaae and Kristensen
2005 ; Worsaae et al. 2005 ; Struck 2006 ). A
position of Diurodrilidae outside Annelida, as
suggested by Worsaae and Rouse ( 2008 ), was
rejected by molecular data (Golombek et al.
2013 ), and the position of the enigmatic
Lobatocerebrum and Jennaria remains unre-
solved (Rieger 1980 , 1991 ). Likewise, the
position of Annelida within Protostomia is still
uncertain. However, recent phylogenomic
analyses recover a clade uniting annelids
with Mollusca, Nemertea, Brachiopoda, and
Phoronida, but without strong support for any
sister group relationship (Edgecombe et al.
2011 ).
Annelids show a huge diversity of body plans,
and it is diffi cult to describe a consistent anatomy
matching most of this variety (Fig. 9.2 ). Most
annelids are coelomate organisms, possessing
multiple segments which occur repetitively along
the anterior-posterior body axis (Purschke 2002 ).
If segmentation is present, the annelid body is
divided into a prostomium, an either homono-
mously (i.e., identical segments) or heterono-
mously (i.e., segments differ from each other)
segmented trunk and a pygidium (Fauchald and
Rouse
1997 ). In many annelid taxa, the prosto-
mium contains the brain; however, in Clitellata
the brain may be found in the following segments
(Bullock 1965 ). The head of the annelids may
bear appendages, as palps or antennae, but these
are lacking in a number of taxa. The mouth can
be found in the fi rst segment which is termed
peristomium. Several members of the Errantia as
well as Amphinomida bear sclerotized mandibu-
lar structures, which may be replaced by a mech-
anism resembling molting (Paxton 2005 ). The
segments of many annelids contain a pair of
nephridia (usually metanephridia), coelomic cav-
ities, ganglia, and ventral and dorsal groups of
chitinous chaetae which might be organized in
parapodia (Purschke 2002 ; Bartolomaeus et al.
2005 ). Segments are generated by a posterior
growth zone which is located in front of the
pygidium (Nielsen 2004 ). The pygidium contains
the anus, which is usually either dorsally or ter-
minally located and is often equipped with pairs
of cirri.
Annelids show a wide variety in the
organization of their nervous system (Bullock
1965 ; Orrhage and Müller 2005 ; Müller 2006 ).
Müller ( 2006 ) proposed a nervous system with
paired circumesophageal connectives, four cere-
bral commissures, fi ve connectives, and numer-
ous commissures in the ventral nerve cord as a
hypothetical ground pattern. However, many
variations of this pattern exist, and many taxa
have not been investigated at all. Accordingly,
alternative hypotheses suggest that a strict
rope-ladder-like nervous system with segmental
C. Bleidorn et al.

195
ganglia interconnected by a pair of connectives
and commissures was not present in the last com-
mon ancestor of annelids (Purschke et al. 2014 ).
Instead, an orthogonal arrangement of the periph-
eral nervous system and the presence of addi-
tional longitudinal nerves might constitute the
ground pattern of Annelida (Lehmacher et al.
2014 ).
Annelids show varying grades of brain com-
plexity which may comprise a number of ganglia.
Fig. 9.1 Phylogeny of Annelida based on Weigert et al. ( 2014 ). Placement of well-investigated model annelids indi-
cated with asterisks
9 Annelida

196
ABC
DEF
G
HI J
KLM
NOP
Fig. 9.2 The diversity of marine Annelida. ( A ) Brada
villosa , Flabelligeridae. ( B ) Glycera capitata , Glyceridae.
( C ) Lepidonotus squamatus , Polynoidae. ( D ) Nereis pela-
gia , Nereididae. ( E ) Chaetopterus sp., Chaetopteridae.
( F ) Syllinae indet., Syllidae. ( G ) Branchiomma arctica ,
Sabellidae. ( H ) Lumbrineris sp., Lumbrineridae. ( I )
Amblyosyllis sp., Syllidae. ( J ) Phyllodoce sp.,
Phyllodocidae. ( K ) Polynoidae indet. ( L ) Sabellidae
indet., Sabellidae. ( M ) Spirobranchus giganteus ,
Serpulidae. ( N ) Lysidice sp., Eunicidae. ( O ) Serpula sp.,
Serpulidae. ( P ) Travisia sp., Travisia (All images pro-
vided by Alexander Semenov (
www.clione.ru ). ©
Alexander Semenov, 2015. All Rights Reserved)
C. Bleidorn et al.

197
Mushroom bodies have been reported for several
taxa of the Phyllodocida (Heuer et al. 2010 ). As
for the nervous system, many different variations
can be found in the muscular system. The pres-
ence of an outer layer of circular muscle fi bers
and an inner layer of four longitudinal bands of
muscle fi bers is often regarded as a possible
ground pattern (Tzetlin and Filippova 2005 ;
Lehmacher et al. 2014 ), but circular muscles are
missing in several annelid taxa and may thus not
constitute a basal annelid feature. Additionally,
other muscular fi ber bundles referred to as
oblique, diagonal, bracing, or dorso-ventral fi bers
might be present and are often compensating
missing circular musculature (Purschke and
Müller
2006 ).
However, many taxa, such as myzostomids,
sipunculids, or echiurids, are clearly deviating
from the pattern described above in various
aspects, and all of them seem to have lost seg-
mentation convergently. Interestingly, all these
examples still show some traces hinting to a sec-
ondary loss of segmentation (Purschke et al.
2000 ; Hessling 2003 ; Kristof et al. 2008 ; Helm
et al. 2014 ). Other taxa such as clitellates and
many other sedentarians lost their parapodia.
Siboglinidae (Pogonophora + Vestimentifera)
show many reductions as adaptation to their life-
style in close association with bacterial endosym-
bionts (Schulze and Halanych 2003 ). Loss of key
characters in Annelida is well-documented and is
regarded as one of the problems to converge to a
well-accepted phylogeny of the whole group
(Purschke et al. 2000 ; Bleidorn 2007 ; Miyamoto
et al. 2013 ).
Several systems of sensory organs are
described for annelids, including a type of che-
mosensory organ called “nuchal organ.” This
type of sensory organ can be found in the pos-
terior part of the prostomium and usually con-
sists of ciliated supporting cells, sensory cells,
and retractor muscles (Purschke
1997 ).
Clitellates as well as several other annelids
such as the basal branching Oweniidae and
Magelonidae lack nuchal organs completely.
Many annelids possess some kind of light
receptive photoreceptors which show great
structural diversity (Purschke et al.
2006 ).
Generally, rhabdomeric, ciliary, and phaoso-
mous photoreceptor cell types are distin-
guished, and they might represent either larval
or adult eyes (Purschke et al. 2006 ; Arendt
et al. 2009 ). Larval eyes are simple organized,
and the eye spots of the trochophore of
Platynereis dumerilii consist of a rhabdomeric
photoreceptor cell and a pigment cell which
provide a direct coupling of light-sensing cili-
ary locomotory control (Jekely et al.
2008 ).
Eyes of adult annelids might be present on the
head, palps, segments (usually laterally), or
even the pygidium (Purschke et al. 2006 ).
Annelids show a variety of reproductive
strategies, and sexual as well as asexual repro-
duction is well-documented for many taxa
(Wilson 1991 ; Bely 2006 ). For sexual reproduc-
tion, different types of free spawning, brooding,
and encapsulation of embryos in cocoons can be
distinguished, and all types involve either plank-
totrophic or lecithotrophic developmental stages
(Thorson 1950 ; Wilson 1991 ). Multiple modes
of development (poecilogeny) are reported for
some annelid species, with the spionid
Streblospio benedicti as the best-investigated
example (Levin 1984 ; Zakas and Wares 2012 ).
By far the most spectacular diversity of repro-
ductive modes can be found across syllids,
including swarming and external fertilization,
internal fertilization, viviparity, and parthogen-
esis, as well as different forms of hermaphrodit-
ism (Franke 1999 ). Several annelid taxa show a
pronounced sexual dimorphism resulting in
dwarf male forms as found, for example, in
some echiurids, siboglinids, or antonbruunids
(Spengel
1879 ; Hartman and Boss 1965 ;
Worsaae and Rouse 2010 ). Not surprisingly,
annelids are also a prime example for the inves-
tigation of heterochrony, and several putative
paedomorphic taxa have been hypothesized
(Westheide 1987 ; Struck 2006 ; Bleidorn 2007 ;
Osborn et al. 2007 ).
9 Annelida

198
EARLY DEVELOPMENT
Egg Structure and Fertilization
Annelids with planktotrophic development usu-
ally have small, non-yolky eggs, whereas species
with lecithotrophic development bear larger and
yolk-rich eggs (Irvine and Seaver 2006 ).
Ultrastructural studies of annelid eggs are scarce
given the immense diversity of this group. One of
the best-investigated examples is the egg of parch-
ment worms of the genus Chaetopterus . Different
regions are distinguished based on staining
properties, divided into a cortical ectoplasm,
endoplasm, and hyaloplasm (Lillie
1906 , 1909 ),
the latter two regions forming the cytoplasm. The
ectoplasm contains large membrane- bound spher-
ules, nuage (a germ line-specifi c organelle con-
taining several proteins), and intracellular
membrane systems. The endoplasm contains yolk
and lipid, interspersed with mitochondria, granular
bodies, and endoplasmatic reticulum. In contrast,
the hyaloplasm (or teloplasm) is characterized by
the absence of granular bodies (Eckberg 1981 ;
Jeffery 1985 ). Eggs of many annelid species show
a clear polarity with an accumulation of develop-
mental factors in the cortex of future polar regions
(Dorresteijn 2005 ). The spatial distribution of
Platynereis dumerilii as a Model for Evolutionary
Developmental Biology
Platynereis dumerilii is a marine annelid
belonging to the errant family Nereididae,
which emerged as a thoroughly investigated
model species. The life cycle of this indirect
developing gonochoric species with plankto-
trophic larvae is well established and control-
lable in the lab. Immature, atokous worms live
in self-constructed tubes. Sexually mature,
epitokous individuals, which appear mor-
phologically different to atokous individuals,
leave the tube and begin swimming to fi nd
partners for spawning during the night. The
day of swarming is controlled by an endog-
enous lunar cycle which can be triggered
artifi cially in the lab. Cultures were estab-
lished in the lab in 1953 and are bred since
then without interruption. Experimental tech-
niques such as cell ablation, whole-mount
in situ hybridization, RNA interference, and
Morpholino knockdowns are routinely appli-
cable. First transgenic lineages have been cre-
ated, and a project sequencing the genome is
underway for this species (
http://4dx.embl.
de/platy/ ). Comparative genomic studies sug-
gest that the genome of P. dumerilii retains
a more ancestral organization compared to
other protostomian model organisms such as
Drosophila melanogaster or Caenorhabditis
elegans . Important insights into the evolution
of segmentation, vision, and the nervous sys-
tem in Bilateria were provided by evolution-
ary developmental studies on P. dumerilii , and
since a number of labs now use this animal as
a model, important results with considerable
relevance for our understanding of animal
evolution and development are likely to keep
emerging in the near future.
Atokous juvenile of Platynereis dumerilii (After
Fischer and Dorresteijn (
2004 ) )
C. Bleidorn et al.

199
maternal mRNA in the ectoplasm has been
described for Chaetopterus (Jeffery and Wilson
1983 ; Jeffery 1985 ). After fertilization and before
initiation of the fi rst cleavage, a reorganization of
the yolk-free hyaloplasm (teloplasm) has been
observed in several annelids (Dorresteijn 1990 ;
Weisblat and Huang 2001 ). It has been shown for
Platynereis dumerilii that after attachment of the
sperm to the egg surface, cortical granules released
by exocytosis from the ectoplasm start forming an
egg jelly on the outside, which removes supernu-
merary sperm from its surface (Dorresteijn 1990 ).
Ultrastructural investigations of the ectoplasm of
eggs of P. dumerilii and the clitellate Theromyzon
rude reveal an extensive framework of actin fi la-
ments that are involved in remodeling the egg sur-
face after fertilization (Fernandez et al.
1987 ;
Kluge et al. 1995 ).
Following fertilization a reorganization of
the endoplasm can be observed. The distribu-
tion of the two cytoplasmic domains can be cat-
egorized into different types, which seem to be
restricted to certain annelid taxa (Shimizu
1999 ). Most investigated non-clitellate annelids
(e.g., chaetopterids, nereidids, and onuphids)
show a stratifi cation of the endoplasm into two
domains, and, in most cases, the clear hyalo-
plasm (teloplasm) is localized at the animal
pole (Wilson 1892 ; Huebner and Anderson
1976 ; Jeffery and Wilson 1983 ). In contrast,
three domains can be distinguished in the clitel-
late endoplasm, with teloplasm localized at
both the animal and vegetal poles of the egg
(Shimizu 1999 ; Weisblat and Huang 2001 ).
These cytoplasmatic movements are coordi-
nated by complex cytoskeletal mechanisms
which even seem to vary among taxa. Whereas
in the leech Helobdella triserialis , microtu-
bules are shown to play an important role,
movement of the teloplasm in the oligochaete
Tubifex is orchestrated by an actin network
(Astrow et al.
1989 ; Shimizu 1995 ).
Cleavage
Annelids develop by spiral cleavage which is
characterized by cleavage furrows which are
oblique to the egg axis due to an inclination of the
mitotic spindle (see Chapter
7 ). This cleavage
starts with two orthogonal cell divisions which
generate four blastomeres, called A, B, C, and D
(Costello and Henley 1976 ). Correlating with the
differing developmental modes in annelids, blas-
tomeres usually exhibit the same size (equal
cleavage) in species with indirect development
and planktotrophic larvae, whereas direct devel-
opers show pronounced differences in blastomere
size (unequal cleavage) (Anderson 1966 ; Arenas-
Mena 2007 ). However, exceptions to this trend
exist. For example, Platynereis dumerilii and
Platynereis massiliensis both show unequal spi-
ral cleavage patterns, even though the former spe-
cies develops indirectly and the latter directly
(Schneider et al.
1992 ). In most cases unequal
cleavage is achieved due to positioning of the
mitotic spindle. However, in some annelids this
cleavage pattern is facilitated due to the presence
of membrane- bound polar lobes (Freeman and
Lundelius 1992 ). Such a polar lobe is also
reported for the myzostomid Myzostoma cirrif-
erum (Eeckhaut and Jangoux 1993 ). The differ-
ent modes of spiral cleavage across annelids have
been thoroughly reviewed in Dorresteijn ( 2005 ).
The future axis of the developing embryo is
already determined, with A and C corresponding
respectively to the left and right side of the
embryo and blastomeres B to D defi ning the
antero-ventral to postero- dorsal axis (Nielsen
2004 ). Due to uneven cleavage, starting from the
third cell division, a shifting in the angle of the
mitotic spindles, which alternates during subse-
quent divisions, becomes obvious in almost all
annelids. These shifts are either dextral (clock-
wise) or sinistral (anti-clockwise) and lead to the
name-giving spiral arrangement pattern of blas-
tomeres. By oblique divisions animal and vegetal
daughter cells are generated, referred to as micro-
mere quartets and macromere quartets. Some
annelids such as the opheliid Armandia brevis
show equal cleavage also in the third cleavage,
generating micro- and macromeres of equal size
(Hermans
1964 ). In the oweniid Owenia collaris
and some leeches, micromeres are larger than
macromeres in the eight-cell stage, a pattern
which is also known from several nemerteans
9 Annelida

200
(Dohle 1999 ; Smart and Von Dassow 2009 ).
Several authors introduced the idea that a spe-
cifi c, phylogenetically conserved pattern of blas-
tomeres can be seen at this stage, termed the
“annelid cross.” This pattern is regarded as typi-
cal for most annelids (including echiurans) but
cannot be found in sipunculids, which show the
so-called molluscan cross. However, a continuum
of different variants between these patterns is
demonstrated, and consequently, these concepts
have been neglected for phylogenetic purposes
(see also Chapter 6 ; Maslakova et al. 2004a ;
Nielsen
2004 ). The cell fate of individual blasto-
meres is conserved across annelids, and a specifi c
nomenclature is used to trace the fate of
blastomeres throughout development, using
capital letters for macromeres and small letters
for micromeres (Conklin 1897 ; Costello and
Henley 1976 ; Nielsen 2004 ). A number is used as
prefi x to designate the quartet of which the
macromeres or micromeres originated from. The
micromeres continue to divide, and daughter
cells inherit the name of their mother cell, with
modifi cation, to trace its origin (animal vs. vege-
tal) (Fig. 9.3 ). An alternative cell nomenclatural
system is in use for leeches (Dohle 1999 ). Many
clitellates show a cleavage pattern that nearly
obscures the original spiral mode of cleavage. In
these cases where yolk content and egg size are
reduced, the embryo is nourished by the sur-
rounding fl uid within the cocoon (Dohle
1999 ).
Several siboglinids show elongated eggs with a
high yolk content leading to an aberrant pattern
of spiral cleavage (Southward 1999 ). In most
annelids, the cleavage pattern shifts from spiral to
bilaterally symmetric after the formation of the
fourth quartet of micromeres (Meyer and Seaver
2010 ). In P. dumerilii cell fates of sister blasto-
meres along the animal-vegetal axis are specifi ed
by levels of beta-catenin. High levels specify
vegetal sister cell fates, while lower levels spec-
ify animal sister cell fates. Interestingly, no beta-
catenin asymmetry is observed after the fi rst
bilaterally symmetrical and transverse cell divi-
sions (Schneider and Bowerman
2007 ).
Descendants of the fi rst micromere quartet
(1a–1d) form larval head structures including the
apical organ, larval eyes, and the head ectoderm
as well as the primary trochoblasts (Nielsen
2004 ). The trochoblasts received their name as
they will give rise to the prototroch, and this
has been demonstrated for several annelids,
A - quadrant
C - quadrant
D - quadrant
B - quadrant
A - quadrant
D - quadrant
B - quadrant
4D
1b 1c
1c
1c
1c 11
11
1b 21
12
11
12
21
22
11
12
1b
1b
1a
1a
1a
1a
1d 1d
1d
21
22 12
21
22
22
1d
3A
3C
3b 2b 2b 2
2
2
3a
2a 1
1
1
2a
3d
4d
2d
2d
3c
2c
2c 2
1
3B
Fig. 9.3 Diagram illustrating the spiral cleavage cell nomenclature in the 33-cell stage of an unequally cleaving
embryo of Arenicola cristata (Child
1900 ). The four quadrants ( A – D ) are indicated by colors
C. Bleidorn et al.

201
e.g., capitellids, dinophilids, and nereidids
(Wilson 1892 ; Eisig 1898 ; Nelson 1904 ).
Trochophore larvae of Myzostomida lack or
show a reduced prototroch (Rouse 1999 ), but as it
has been shown for the nemertean Carinoma
tremaphorus , this need not be refl ected in the for-
mation of trochoblasts (Maslakova et al. 2004b ).
Three sets of trochoblast cells are involved in
prototroch formation, a pattern which is highly
conserved across Spiralia (Henry et al. 2007 ).
Besides the primary and accessory trochoblasts,
which are derived from the fi rst micromere quar-
tet, this includes secondary trochoblasts formed
by some descendants of the second micromere
quartet (2a–2c). Some annelids deviate from this
pattern, e.g., the terebellid Amphitrite ornata ,
who lacks the accessory trochoblasts (Damen
and Dictus
1994 ). Other descendants of the sec-
ond micromere quartet generally develop into the
foregut (stomodaeum) as well as part of the ecto-
derm (Nielsen 2004 ). The 2d cell is the somato-
blast, developing into the major part of the body
ectoderm posterior of the prototroch (Meyer and
Seaver 2010 ). Using cell ablation studies, it has
been shown for Capitella teleta that the 2d cell is
responsible for organizing activity during early
embryonic development, as well as bilateral sym-
metry and dorso- ventral axis organization of the
head, and formation of neural, foregut, and meso-
derm tissue (Amiel et al. 2013 ). In clitellates,
four pairs of ectoteloblasts (called N, O, P, Q) are
descendants of the 2d cell and give rise to four
germbands including smaller cells (Dohle 1999 ;
Goto et al. 1999 ). Cells derived from the third
micromere quartet (3a–3d) form the foregut and
ectomesoderm and might be the origin of proto-
nephridia (Nielsen
2004 ; Ackermann et al. 2005 ).
Interestingly, in C. teleta , mesodermal bands are
generated by 3c and 3d (Meyer et al. 2010 ).
Usually the mesoderm and endoderm are
formed by cells of the fourth micromere quartet
(4a–4d), as characteristic for spiral cleavage in
Lophotrochozoa in general (Chapter 7 ; Gline
et al. 2011 ). Of special interest is the fate of the 4d
cell, which has been called “mesenteloblast” or
“primary mesoblast” (Wilson
1898 ). This cell
gives rise to the adult mesoderm in most spira-
lians including mollusks or entoprocts (Chapters
6 and 7 ). In Clitellata, a fi fth teloblast (M) is
derived from the 4d cell, specifying a mesodermal
germband (Goto et al. 1999 ). Progenitors of 4d
form bilaterally symmetrical mesodermal anlagen
in Platynereis dumerilii (Ackermann et al. 2005 ;
Fischer and Arendt 2013 ). Prior to gastrulation,
four secondary mesoblast cells bud from descen-
dants of the 4d cell and show the morphology and
gene expression signature of primary germ cells
(Rebscher et al. 2012 ). These primary germ cells
stay in mitotic arrest until individuals enter game-
togenesis (Lidke et al. 2014 ). In C. teleta the 4d
cell generates few muscle cells, primordial germ
cells, and the anus (Meyer et al.
2010 ). It has been
suggested that mesoteloblast‐like mesodermal
stem cells forming continuous mesodermal bands
are part of the Pleistoannelida ground pattern
(Fischer and Arendt 2013 ).
Gastrulation
The process of gastrulation in annelids has been
reviewed in detail by several authors (Okada
1957 ; Anderson 1973 ; Weisblat and Huang 2001 ;
Irvine and Seaver 2006 ), and the following
descriptions provide a generalized pattern found
in clitellate and non-clitellate annelids.
Gastrulation of embryos with less yolk starts
with the invagination (embolic gastrulation) of
putative midgut cells, and epithelia derived from
the micromere cap grow toward the ventral side.
Mesoteloblasts can be found in a posterior posi-
tion in the blastocoel, whereas ectoteloblasts are
located adjacent to them, below the larval ecto-
derm. The fate of the blastopore differs across
annelid taxa, and protostomy, where the blasto-
pore becomes the mouth, is found in most anne-
lid taxa. Notably, deuterostomy, where the
blastopore becomes the anus, has been demon-
strated for eunicids (Åkesson
1967 ). The concept
of amphistomy, in which both the mouth and the
anus are derived from the corresponding ends of
the blastopore, which was claimed to be present
in Polygordius (Arendt and Nübler-Jung 1997 ),
might not occur in any organism at all (Hejnol
and Martindale 2009 ). Some differences apply
for the gastrulation of yolk-rich annelid embryos.
9 Annelida

202
Here, the process is rather described as epiboly,
where the micromere cap grows over putative
midgut cells and teloblasts (Irvine and Seaver
2006 ). In clitellates, the blastopore is found in the
point where the germinal bands coalesce to form
the germinal plate.
LATE DEVELOPMENT
Larval Ciliary Bands
Larval morphological characters vary across dif-
ferent annelid families (Fig. 9.4 ). A trochophore
has distinct larval ciliary regions forming promi-
nent bands or tufts, and the presence of a pro-
totroch is regarded as defi ning (Bhaud and
Cazaux 1987 ; Rouse 1999 ). However, detailed
investigations concerning the homology of the
respective ciliated regions within the different
annelid families are lacking. At the anterior end
of the episphere, the apical tuft marks the posi-
tion of the larval apical organ, a feature well
known for most invertebrate taxa with ciliated
larvae (Marlow et al. 2014 ). Appearing early in
development, the apical tuft forms a sensory
region that is located in the direction of larval
movement but often disappears in early larval
stages (see Chapter 7 for details on apical tuft
morphology). Although an apical tuft is wide-
spread within annelids, larvae without an apical
tuft are known for most cirratulids, histriobdel-
lids, lopadorhynchids, orbiniids, sabellids, and
tomopterids (Rouse 1999 ).
The prominent prototroch is represented by an
equatorial ring consisting of usually compound
cilia formed by a group of specifi c trochoblasts
(Damen and Dictus
1994 ). Situated anterior to
the mouth opening, a prototroch is known for
most annelids, mollusks, and entoprocts (Nielsen
2012 ). Dividing the larval body in an anterior
episphere and a posterior hyposphere (see
Chapter 7 ), the prototroch is present mainly in
planktotrophic annelid larvae and some leci-
thotrophic stages but absent in direct developing
taxa such as clitellates, aelosomatids, and histri-
obdellids (Rouse 1999 ). In some annelid taxa, the
cilia of the prototroch may cover almost the
whole episphere in early developmental stages
(e.g., in Chaetopterus ; see Fig.
9.4L ). The pro-
totroch may be formed by equatorially arranged
ciliary tufts ( Myzostoma cirriferum ; see
Fig. 9.4G ), or the whole larva may be covered by
cilia, and a defi ned prototroch is hardly distin-
guishable, e.g., in early larvae of the eunicid
Marphysa (Fig. 9.4E ). An epispheral ciliated
band is represented by the meniscotroch, which
is only known for Phyllodocida (Bhaud and
Cazaux 1982 ; Rouse 1999 ). Forming a tuft of
short cilia, the meniscotroch is located in a
ventral position within the episphere. Posterior to
the latter structure, some annelids possess a
ciliated band situated anterior to the prototroch –
the akrotroch (Häcker
1896 ). Forming a com-
plete ring separated from the apical tuft and the
prototroch, an akrotroch can be found in syl-
lids, orbiniids (e.g., in Scoloplos armiger , see
Fig. 9.5 ), onuphids, cirratulids, and several
Eunicida (Rouse 1999 ).
Situated posteriorly to the prototroch, the
metatroch is represented by a ciliated ring that
often beats opposed to the latter one and lies
in a pre-segmental (= peristomial) position
(Strathmann 1993 ; Nielsen 2012 ). Being present
in most annelid families, a metatroch seems to
be absent in Echiura (Fig. 9.4J ) and Opheliidae.
For Capitellidae, Siboglinidae, and Syllidae, the
presence of a metatroch is still discussed (Rouse
2000a ). Planktotrophic larvae of several polynoid
scale worms possess another bundle of long cilia,
the oral brush, which seems also to be involved in
feeding mechanisms (Phillips and Pernet 1996 ).
A prominent ventral ciliary band is represented
by the neurotroch, which is known at least in some
annelid families including many sedentarian taxa,
e.g., Orbiniidae (Fig.
9.5 ), Sabellidae (Fig. 9.6 ),
and Maldanidae. In both planktotrophic and leci-
thotrophic developmental stages, the neurotroch
forms a distinct ventral ciliated area interconnect-
ing proto- and telotroch, which often appears later
in larval development (Rouse 1999 ). The telo-
troch, defi ned as a posterior ring of cilia used for
locomotion (Strathmann 1993 ), also appears later
in development and is known for both planktotro-
phic and lecithotrophic developmental stages
(Strathmann 1993 ). The telotroch marks the
C. Bleidorn et al.

203
ABC
D
EF
GHI
JK
L
Fig. 9.4 Diversity of annelid trochophore larvae. Anterior
(apical) is up in all aspects. ( A ) Polygordius sp.
(Polygordiidae) after Woltereck (
1904 ). ( B ) Magelona fi li-
formis (Magelonidae) after Wilson (
1982 ). ( C ) Eurythoe
complanata (Amphinomidae) after Kudenov (
1974 ). ( D )
Owenia collaris (Oweniidae) after Smart and Von Dassow
(
2009 ). ( E ) Marphysa sanguinea (Eunicidae) after
Prevedelli et al. (
2007 ). ( F ) Phyllodoce maculata
(Phyllodocidae) after Voronezhskaya et al. (
2003 ). ( G )
Myzostoma cirriferum (Myzostomida) after Eeckhaut and
Jangoux (
1993 ). ( H ) Platynereis dumerilii (Nereididae)
after Fischer and Dorresteijn (
2004 ). ( I ) Phascolosoma per-
lucens (Sipuncula) after Jaeckle and Rice (
2002 ). ( J )
Urechis caupo (Echiura) after Pilger (
2002 ). ( K ) Osedax sp.
(Siboglinidae) after Rouse et al. (
2009 ). ( L ) Chaetopterus
variopedatus (Chaetopteridae) after Henry (
1986 ).
Abbreviations: at apical tuft, ch chaetae, lc lateral cilia, mt
metatroch, nt neurotroch, pt prototroch, tt telotroch
9 Annelida

204
pt
eye
ak
pt
ch1
tt
nt
cilia
ak
g1
g2
g3
tt
cilia
eye mouth
nt
ak
eye
pt
mt
g1
ch2 g2
cilia
ch5
tt
nt
pt
mt
mt
mt
A
B
CD
EF G H
Fig. 9.5 Development of Scoloplos armiger (intertidal
clade) after Anderson (
1959 ). Anterior (apical) is up in all
aspects. ( A ) Unfertilized egg. ( B ) Early 4-day embryo,
dorsal view. ( C ) Late 5-day embryo, dorsal view. ( D ) Late
5-day embryo, ventral view. ( E ) Late 6-day embryo, dor-
sal view. ( F ) Late 6-day embryo, lateral view. ( G ) Early
7-day embryo, dorsal view. ( H ) early 7-day embryo, ven-
tral view. Abbreviations: ak akrotroch; ch 1, ch 2, ch 5
chaetiger 1, 2, 5; g 1, g 2, g 3 gastrotrochs of chaetigers;
mt metatroch; nt neurotroch; pt prototroch; tt telotroch
position of the posterior growth zone (Nielsen
2012 ). Further ciliated bands that may occur in
several families are the gastro- and nototroch
(= segmentally arranged ventral and dorsal ciliary
bands), which are sometimes referred to as
paratrochs (Bhaud and Cazaux 1982 ).
Post-trochophore Development
and Larval Forms
Within annelid ontogeny, the metatrochophoral
stage usually follows the prototrochophore/
trochophore (Fig. 9.7 ). In this developmental
stage, the fi rst signs of segmentation are visible,
e.g., the formation of the fi rst parapodia and
chaetae. In accordance with individual develop-
ment, several subdivisions of the metatrochopho-
ral stage are possible (Fischer et al. 2010 ). In
some terebellids and pectinariids, the
metatrochophore builds a tube and is called aulo-
phore (Bhaud and Cazaux 1982 ).
In Oweniidae a special type of trochophore
occurs, the so-called mitraria (Fig. 9.4D ). The
mitraria larva exhibits prominent proto- and
metatrochal bands, as well as an apical tuft.
C. Bleidorn et al.

205
AB
CD
EF
Fig. 9.6 Development of the lecithotrophic developmen-
tal stages of Megalomma vesiculosum revealed by anti-
tubulin staining. All images are in ventral view except of
( D) which is in dorsal view. Anterior is up. Confocal
maximum projections. ( A ) The early, nonfeeding trocho-
phore exhibits a prominent prototroch ( pt ). An apical tuft
is lacking. ( B ) The later trochophore gains a well-devel-
oped prototroch ( pt ) and cilia ( ci ) at the anterior pole.
Furthermore, the ventral neurotroch ( nt ) and the posterior
telotroch ( tt ) develop at this stage. ( C ) The early
metatrochophore exhibits three pairs of parapodia ( pa )
and a metatroch ( mt ). Neurotroch ( nt ), prototroch ( pt ), and
telotroch ( tt ) are still present in this free- swimming but
nonfeeding stage. ( D ) Shortly before metamorphosis, the
late nectochaete has lost the main ciliary bands and starts
development of the adult tentacles ( te ). ( E ) After meta-
morphosis the juveniles settle within a tube and start feed-
ing. The parapodia ( pa ), the tentacles ( te ), and remnants
of the metatroch ( mt ) are exhibited. ( F ) Late juvenile
worms show an adult-like morphology. The tentacles ( te )
are well-developed, and the animals start to elongate by
posterior segment addition. an anus, ci cilia, mo mouth
opening, mt metatroch, ne nephridia, nt neurotroch, pa
parapodia, pt prototroch, tt telotroch. Scale bars = 100 μm
(© Conrad Helm, 2015. All Rights Reserved)
9 Annelida

206
AB
CD
EF
Fig. 9.7 Development of Platynereis dumerilii . ( A )
Cleaving embryo, where the third cleavage forms micro-
meres and macromeres. ( B ) Early trochophore, with pro-
totroch and apical tuft. ( C ) Late trochophore, with
simultaneous appearance of the fi rst three larval segments.
( D ) Mid metatrochophore, with developing chaetae reach-
ing over the body wall. ( E ) Early nectochaete, with forma-
tion of the metatroch and elongation of the trunk. ( F )
Juvenile, with rapidly growing jaws and further addition
of body segments. Combined after Fischer et al. (
2010 ).
Abbreviations: ac anal cirrus, ae adult eyes, adc anterior
dorsal cirrus, ant antenna, apt apical tuft, avc anterior ven-
tral cirrus, ch chaetae, chs chaetal sac, j jaws, ld lipid
droplet, le larval eye, mg , midgut, mm macromere, mt
metatroch, pat 1 , fi rst paratroch, pat2 second paratroch, pl
palps, pp parapodia, pro proctodeum, pt prototroch, sf sto-
modeal fi eld, sto stomodeum, tt telotroch, 4CS 4th chae-
tigerous segment, 5CS 5th chaetigerous segment
C. Bleidorn et al.

207
Notably, all ciliary bands are monociliated, an
unusual feature for annelid larvae. The hypo-
sphere of the mitraria is strongly reduced, and the
juvenile segmental body develops within the lar-
val body (Wilson 1932 ; Smart and Von Dassow
2009 ). Another unusual larval type is repre-
sented by the rostraria in Amphinomidae and
Euphrosinidae (Mileikovsky 1960 , 1961 ). After
a trochophore stage with a proto- and metatroch
and an apical tuft (Fig. 9.4C ), the episphere of
the metatrochophore elongates, and tentacles are
formed for feeding (Jägersten 1972 ). Remarkably
elongated metatrochophore stages can be found
within siboglonids (Southward
1999 ). The
metatrochophore of investigated vestimentiferan
siboglinids is sessile and bears a prostomium, a
peristomium, and two chaetigers. A prototroch,
a neurotroch, and an apical organ are present as
well as juvenile/adult organs such as tentacles
and pyriform glands (Bright et al. 2013 ).
The end of the metatrochophore stage is usu-
ally marked by the point when the parapodia are
fully developed. If present, the next larval stage is
represented by the nectochaete (Fig. 9.7E ), which
is characterized by the presence of functional para-
podia which are used mainly for swimming. In
Poecilochaetus (Spionidae) this stage is some-
times called nectosoma; in other spionids, it refers
to the chaetosphaera stage (Bhaud and Cazaux
1982 ). In this stage or the previous one, most lar-
vae start body elongation and segment formation
through the posterior growth zone (Irvine and
Seaver 2006 ). A unique swimming larval form
called pelagosphaera is known for Sipuncula (Rice
1976 ). The body of this larval type can be divided
into three body regions: head, mid region includ-
ing metatroch, and a large trunk. These larvae can
be either lecithotrophic or planktotrophic forms,
whereas the latter can live up to 6 months in the
plankton (Jaeckle and Rice
2002 ).
After the nectochaetal (or pelagosphaera)
stage, metamorphosis occurs. During metamor-
phosis, the free-swimming larvae change behav-
ioral characteristics and start an adult-like
lifestyle as juveniles. This metamorphic step can
differ drastically between various species and is
markedly pronounced in larvae of Polygordius ,
which rupture the larval body that is later either
eaten or discarded (Rouse
2006 ). Such a transi-
tion of lifestyles seems to be less distinct or miss-
ing in annelids with lecithotrophic developing
stages, where ciliary bands are absorbed or the
chaetal morphology changes (Figs. 9.5 and 9.6 ).
Although late development and subsequent
metamorphosis may differ between several taxa,
in almost all annelid larvae, the larval episphere
becomes the adult prostomium, and the poste-
rior hyposphere becomes the pygidium and the
posterior growth zone (Fig. 9.8 ). The remaining
hyposphere forms the peristomium, which lacks
chaetae in adult annelids (Nielsen
2004 ). The
segmented body between the peristomium and
the pygidium develops by segment formation
from the posterior growth zone (Irvine and
Seaver 2006 ).
Larval Feeding Modes
Several types of larval feeding behaviors and
developmental modes occur in different annelid
families, mostly divided into either feeding and
free-swimming planktotrophic larvae with “indi-
rect” development or nonfeeding and mostly less
motile embryonic and juvenile forms with
“direct” development. The latter rely on maternal
sources of nutrition in the form of yolk stored in
the egg during oogenesis, feeding on yolk-rich
nurse eggs, or translocation of nutrition directly
from the parent (Qian and Dahms 2006 ). An
overview of larval feeding in annelids is summa-
rized by Rouse ( 2006 ). As direct development
occurs without a metamorphosis, developing
stages are usually referred to as “embryonic” or
“juveniles,” avoiding the term “larvae” (Nielsen
2009 ; Winchell et al. 2010 ). However, debates
remain about the defi nition of the term “larva,”
and alternative terminologies exist (McEdward
and Janies 1993 ; Pechenik 1999 ).
Many authors regard a biphasic life cycle
including a planktotrophic trochophore larva as
representing the plesiomorphic condition of
annelid development (Heimler 1988 ; Nielsen
2012 ). Nevertheless, this idea is doubted by some
authors (Haszprunar et al.
1995 ; Rouse 2000a , b ).
Based on cladistic analyses and ancestral state
9 Annelida

208
Fig. 9.8 Development of annelids indicating the contri-
bution from the four quadrants (A–D), after Nielsen
(
2004 ). Note that the whole segmented body and the
pygidium develop from descendants of the (D) quadrant.
Abbreviations: at apical tuft, pt prototroch, mt metatroch,
tt telotroch, nt neurotroch, gt gastrotroch
reconstruction, multiple events in the evolution
of feeding larvae from lecithotrophic ancestors
seem more parsimonious to assume. According
to this hypothesis, the prototroch had a primary
function for locomotion and became indepen-
dently associated with feeding in several lineages
(Rouse
2000a ). By studying larval forms of
sabellids, Pernet ( 2003 ) was able to demonstrate
the persistence of reduced ciliary structures for
food uptake in nonfeeding larvae. Consequently,
it is suggested that the direction of evolution goes
from planktotrophy to lecithotrophy in this case.
Further on, a functional relation between egg size
and gut development has been hypothesized.
Non-feeding stages usually bear larger eggs, and
due to the increased cell size, gut development
might be hindered, resulting in nonfunctional
guts in nonfeeding stages (Pernet 2003 ). As this
scenario may be generalized for all annelids,
planktotrophy seems to be the likely ancestral
state in annelids.
Most planktotrophic larvae exhibit a prominent
proto- and metatroch, used for locomotion
and food uptake by “downstream feeding”
(Rouse
2000a ). This is the case for larvae of
Amphinomidae, Chrysopetalidae, Glyceridae,
Nephtyidae, Oweniidae, Pectinariidae, Polynoidae,
and Sabellaridae. However, other annelid families
with planktotrophic larval development use differ-
ent modes of feeding behavior. Taxa with nonfeed-
ing larvae possessing maternally derived nutrition
can be found all over the annelid tree within vari-
ous families, and lecithotrophy may have evolved
secondarily (Rouse
2000a ). A special case of leci-
thotrophic development is represented by adel-
phophagy, where larvae develop by uptake of
nutrients from nurse eggs, which are only pro-
duced as nutrition reserve (Fig. 9.9 ), as in some
C. Bleidorn et al.

209
AB
CD
EF
Fig. 9.9 Development of ciliation in larvae of the adel-
phophagous spionid Boccardia cf. polybranchia revealed
by anti-tubulin staining. All images are in ventral view;
anterior is up. Confocal maximum projections. ( A ) Early
larvae lack ciliary regions and are full of yolk. ( B ) The
trochophoral stage attaches to a nurse egg ( ne ) and starts
to digest its nutrients. Ciliation is only exhibited within
the mouth opening ( mo ) and in the region of the ventral
ciliated patches ( vp ), which are used for attachment. ( C )
The late trochophore develops a distinct prototroch ( pt )
arranged of ciliated patches and a less-prominent
metatroch ( mt ). ( D ) In the metatrochophoral stage, the
prototroch ( pt ), metatroch ( mt ), the ventral ciliated patches
( vp ), and a telotroch ( tt ) are present. First signs of segmen-
tation are visible in this stage. ( E ) In the late metatrocho-
phore the ventral ciliated patches ( vp ) are reduced.
Instead, a distinct neurotroch ( nt ) and several gastrotrochs
( gt ) form. ( F ) In the nectochaetal stage, shortly before
leaving the egg capsule, three bands of gastrotrochal cili-
ary bands ( gt ) are developed, and notopodial cilia ( no ) are
exhibited. The remaining ciliated bands are still present.
gt gastrotroch, mo mouth opening, mt metatroch, ne nurse
egg, no nototroch, nt neurotroch, pt prototroch, tt telo-
troch, vp ventral ciliated patch. Scale bars = 50 μm (©
Conrad Helm, 2015. All Rights Reserved)
9 Annelida

210
spionid taxa (Gibson and Carver 2013 ). Moreover,
poecilogeny, showing different types of develop-
ment in one species, seems to be common in some
spionid genera (Blake and Kudenov 1981 ; Levin
1984 ; Chia et al. 1996 ).
Segmentation
Segmented annelids generate their fi rst seg-
ments, usually simultaneously, as larvae, and
later segments are sequentially added from a
posterior growth zone (Irvine and Seaver
2006 ).
Consequently, some authors differentiate
between primary and secondary segments, an
idea that goes back to Iwanoff ( 1928 ) who postu-
lated a distinct ontogenetic origin for both sets of
segments. Segment formation on the cellular
level is well understood for clitellate embryos,
which are all direct developers and often show
huge and therefore experimentally manipulable
eggs. Segmentation in leeches is strongly corre-
lated with cell division patterns of teloblast cells
and their descending blast cells (Fig. 9.10 ). All
teloblasts originate from descendants of the D
quadrant, with a pair of mesoteloblasts (M)
being derived from 4d and four pairs of ectoder-
mal teloblasts (N, O, P, Q) from 2d descendants.
Mechanisms of the specifi cation of the ectotelo-
blast lineage are different between the hirudin-
ean Helobdella and the oligochaete Tubifex . In
Helobdella the O/P teloblasts constitute an
equivalence group, as they are both pluripotent
and may subsequently follow either the O or the
P fate (Weisblat and Blair 1984 ). On the con-
trary, in Tubifex , the fate of the P teloblast is
determined by birth, whereas the O teloblast is
initially pluripotent and is restricted to the fate of
the O lineage due to signaling from the P lineage
(Arai et al.
2001 ). Each teloblast undergoes
repeated series of unequal division, producing
bandlets of the so-called blast cells. The N and Q
lineages produce two different types of blast
cells, which appear in alternation. The four ecto-
dermal bandlets of each side of the bilateral
embryo join and form together with the meso-
dermal band, the germinal band, which lies at the
surface of the embryo. During gastrulation, the
germinal bands from both sides of the embryo
coalesce into the germinal plate, the origin of
segments. The stereotyped cell divisions of all
blast cells contribute to the forming segments
(Shimizu and Nakamoto
2001 ; Weisblat and
Huang 2001 ; Irvine and Seaver 2006 ). Each
ectodermal band contributes neural and epider-
mal progeny, but two thirds of the neurons are
derived from the N teloblast (Weisblat and
Huang 2001 ). Ectodermal segmentation can be
divided into two steps. In the fi rst step, distinct
cell clusters are generated autonomously by each
bandlet. These separate clusters are aligned with
the cell cluster derived from the mesodermal
bandlet in a second step (Shimizu and Nakamoto
2001 ). It has been shown experimentally for
Helobdella that hemilateral ablation of mesoder-
mal precursor cells results in the loss of ectoder-
mal segmental organization. In contrast,
mesodermal boundaries are determined autono-
mously, without positional cues from ectodermal
tissue (Blair 1982 ).
The cellular basis of segment generation in
non-clitellate annelids is less well studied. One
reason seems to be that they are more diffi cult to
handle experimentally due to the smaller size of
their embryos and larvae (Irvine and Seaver
2006 ). An obvious difference is the absence of
large visible teloblasts in non-clitellate annelids
(Seaver et al. 2005 ). However, using semiauto-
mated cell tracking, mesoteloblast-like stem cells
were revealed for Platynereis dumerilii which
also form mesodermal bands of daughter cells
(Fischer and Arendt 2013 ). For this species, two
distinct sets of stem cells could be described for
the posterior growth zone, where many rounds of
division of small populations of teloblast-like
stem cells generate new segments (Gazave et al.
2013 ). In contrast, Seaver et al. ( 2005 ) could not
fi nd evidence of a teloblastic growth zone in
Capitella teleta and the serpulid Hydroides
elegans using incorporation of BrdU, therewith
confi rming older studies (Wilson 1890 ; Shearer
1911 ). Instead, segments in larvae arise from a
fi eld of mitotically active cells located in lateral
body regions. However, the authors could not
C. Bleidorn et al.

211
rule out if inconspicuous teloblast-like cells
might be present. For Chaetopterus , it has been
suggested that at least the fi rst 15 segments are
formed by subdivision of existing anlagen and
not by a posterior growth zone (Irvine et al.
1999 ). Similarly, formation of repetitive struc-
tures in myzostomids differs from an addition
governed by a posterior growth zone. As such,
during development the third pair of parapodial
structures appears fi rst, followed by the fourth
pair, second and fi fth pair (simultaneously), and
fi rst pair (Jägersten 1940 ). Future studies of espe-
cially non-clitellate annelids are necessary to fur-
ther assess the existing variability in the process
of segment formation throughout Annelida.
Muscular System
Annelids show a huge variety of muscular orga-
nization, with longitudinal musculature orga-
nized in separate bands or massive plates and
circular musculature that can be fully developed,
incomplete, or even completely missing (Tzetlin
and Filippova 2005 ). Therefore, it comes without
surprise that differences in the development of
the muscular system have been found across the
investigated taxa. Phalloidin labeling coupled
with confocal microscopy revealed an origin of
muscular development posterior of the apical
organ in the phyllodocid Phyllodoce groenland-
ica , with distinct transversal muscles starting to
140
90
80
29
26
M
N
P
Q
10
0
Fig. 9.10 Schematic
representation of the stem
cell-mediated, lineage-
dependent segmentation in
leeches and other clitellate
annelids. Anterior is to the
top . Pairs of diagonal lines
indicate discontinuities in
the depicted structures.
One bilateral pair of
mesodermal stem cells ( M
teloblasts) and four
bilateral pairs of ectoder-
mal stem cells ( N , O/P ,
O/P , and Q teloblasts)
constitute the posterior
growth zone. Two types of
blast cells are contributed
by the N lineage,
designated ns ( red ) and nf
( blue ), which arise in
alternation. The numbers
on the left side indicate the
progressing time during
segment formation given in
hours of clonal age.
Arrows indicate the
delimitation of two
ganglionic primordia
(From Rivera and Weisblat
2009 , with permission
from the publisher)
9 Annelida

212
grow posteriorly. Subsequently, several longitu-
dinal muscle fi bers start to develop and grow in
posterior direction. Simultaneously, outer circu-
lar muscle fi bers begin to appear in a progression
from anterior to posterior. The longitudinal mus-
cle fi bers reach the anal region approximately
7 days after hatching, and additional circular
muscle fi bers forming distinct rings develop from
anterior to posterior. Additional longitudinal
muscle fi bers develop in the dorsal region, form-
ing a continuous layer. Musculature of the diges-
tive system is hardly recognizable in early stages.
Notably, the organization of the body wall mus-
culature starts before the formation of the fi rst
segments (Helm et al.
2013 ). Similarities to this
kind of musculature development have been
found in several other annelids. As such, an ante-
rior origin in either lecithotrophic embryos or
planktotrophic larvae is also reported for, e.g.,
capitellids, clitellates, nereidids, and sabellariids
(Hill 2001 ; Bergter and Paululat 2007 ; Hunnekuhl
et al. 2009 ; Brinkmann and Wanninger 2010a ;
Fischer et al. 2010 ). No circular musculature
could be detected in developing stages of the
maldanid Axiothella rubrocincta , even though
they are present in adults (Brinkmann and
Wanninger 2010b ).
In contrast to the description for Phyllodoce ,
musculature of the digestive system develops
before the body wall musculature in planktotro-
phic larvae of serpulids and sabellariids
(McDougall et al. 2006 ; Brinkmann and
Wanninger 2010a ). Temporal shifts in the devel-
opmental onset of several muscle groups, a phe-
nomenon described as heterochrony, are a
common theme when comparing myogenesis
between different and even closely related anne-
lid species and are even more pronounced in the
comparison of planktotrophic with lecithotro-
phic developing species (McDougall et al.
2006 ;
Brinkmann and Wanninger 2010a ; Helm et al.
2013 ). As in Phyllodoce , many annelid species
show a successive appearance of circular mus-
culature from anterior to posterior, which has
been also described, e.g., for the tubifi cid
Limnodrilus and the serpulid Filograna implexa
(Bergter et al. 2007 ; Wanninger 2009 ). In con-
trast, in sipunculids, Platynereis massiliensis
and the leech Erpobdella octoculata , anterior
circular muscles are formed synchronously
(Wanninger et al.
2005 ; Bergter et al. 2007 ;
Kristof et al. 2011 ; Helm et al. in press ).
Muscular development in the non-segmented
sipunculids as analyzed for Phascolion strom-
bus , Phascolosoma agassizii , Thysanocardia
nigra , and Themiste pyroides shows that the fi rst
anlagen of circular body wall musculature
appear simultaneously. Rudiments of four lon-
gitudinal retractor muscles appear at the same
time, with longitudinal muscle fi bers forming a
pattern of densely arranged fi bers around the
retractor muscles (Kristof et al.
2011 ).
Neurogenesis
Annelids show a huge variety of adult nervous
system organization, and until today the ancestral
ground pattern remains under discussion (Bullock
1965 ; Orrhage and Müller 2005 ). The develop-
ment of the nervous system, however, has been
investigated in a surprisingly small number of
taxa. Several transmission electron microscopy
(TEM)-based studies on the larval nervous sys-
tem of phyllodocids and serpulids were published
by Lacalli ( 1981 , 1984 , 1986 ). Some detailed
comparative studies were conducted concerning
the anatomy of the larval apical organ.
Immunocytochemical studies revealed an almost
universal occurrence of an apical organ with
fl ask-shaped cells in larvae of Annelida,
Mollusca, Entoprocta, and Platyhelminthes,
exhibiting FMRFamide- and serotonin-like
immunoreactivity (e.g., Hay-Schmidt 2000 ;
Wanninger
2009 ). Usually, the apical organ in
annelid trochophores is simple, containing a few
fl ask-shaped cells which have slender necks,
dense cytoplasm, and a single projecting cilium
(Lacalli 1981 ). Whereas these cells are missing
in echiurans and many other annelids, sipuncu-
lans show a more complex apical organ with up
to ten fl ask-shaped cells (Wanninger 2008 ).
Marlow et al. ( 2014 ) analyzed the molecular fi n-
gerprint of apical organ cells in Platynereis
dumerilii . They found that orthologs of six3 and
foxq2 are involved in the formation of the apical
C. Bleidorn et al.

213
plate, whereas the apical tuft is formed in a cen-
tral six3 -free area of the apical plate.
Besides this, only few comprehensive studies
for developmental sequences of annelids com-
bining immunocytochemical staining coupled
with confocal laser scanning microscopy exist
(Hessling 2002 ; Hessling and Westheide 2002 ;
Voronezhskaya et al. 2003 ; McDougall et al.
2006 ; Brinkmann and Wanninger 2008 ; Kristof
et al. 2008 ; Fischer et al. 2010 ; Winchell et al.
2010 ; Helm et al. 2013 , in press ). Main targets
for these studies were serotonin, a biogenic
amine involved in neuronal signaling, and the
neuropeptide FMRFamide. Labeling of tubulin
is additionally used to stain neurotubules. In
summary, these studies show that neurogenesis
in annelids is variable, following different devel-
opmental pathways. Planktotrophic larvae typi-
cally bear a serotonergic nerve ring underlying
the prototroch and an apical organ that bears
serotonergic and FMRFamidergic cells. The
development of the larval nervous system usu-
ally starts from two subsystems (Fig.
9.11 ).
FMRFamidergic immunoreactivity increases
from anterior toward posterior during nervous
system development. In Phyllodoce and some
other annelids, a single serotonergic neuron
located at the posterior pole of the larva is pres-
ent (Fig. 9.11A ). From here, anteriorly project-
ing nerve fi bers start to grow, outlining the future
ventral nerve cords (Voronezhskaya et al. 2003 ).
Such a posterior origin of serotonin-like immu-
noreactivity was also detected in another phyl-
lodocid, in syllids, nereidids, and orbiniids
(Orrhage and Müller 2005 ; McDougall et al.
2006 ; Fischer et al. 2010 ; Helm et al. 2013 ). In
contrast, the investigated sabellariids, spirorbids,
and sipunculans show no evidence for a posterior
serotonergic cell (Brinkmann and Wanninger
2008 ; Kristof et al. 2008 ; Brinkmann and
Wanninger 2009 ). Later, the adult nervous sys-
tem starts to develop along pathways established
by the earliest peripheral neurons of the larva.
However, other authors propose a separate devel-
opment of the larval and adult nervous system
(Lacalli
1984 ) or fi nd that the larval nervous sys-
tem is integrated in the adult one (Hay-Schmidt
1995 ). The relative timing of events during
neurogenesis shows major shifts between com-
pared species and is regarded as cases of heter-
ochrony (McDougall et al.
2006 ; Brinkmann and
Wanninger 2008 ; Helm et al. 2013 ).
A different picture is found in direct develop-
ing lecithotrophic species as investigated for
nereidids. In Nereis arenaceodentata , the ner-
vous system has developed already much of the
complexity of the adult at hatching. This includes
a large brain and the presence of circumesopha-
geal connectives, nerve cords, and segmental
nerves. Within 1 week after hatching, cephalic
sensory structures and brain substructures are dif-
ferentiated, and the nervous system architecture
resembles that of adults (Winchell et al.
2010 ). A
similar developmental pattern of the nervous sys-
tem has been described for Platynereis massilien-
sis (Helm et al. in press ).
Analyses of the development of the nervous
system of the non-segmented Echiura and
Sipuncula gained major interest, as they pro-
vided direct ontogenetic evidence for the indi-
rectly inferred loss of segmentation in these taxa
as suggested by molecular phylogenies. Using
immunocytochemistry, a metameric organiza-
tion of the nervous system has been demon-
strated for two echiuran species: Urechis caupo ,
which has planktotrophic larvae, and Bonellia
viridis with directly developing lecithotrophic
stages (Hessling 2002 ; Hessling and Westheide
2002 ). The development of the nervous system
in Bonellia viridis proceeds from anterior to
posterior. This is obvious in early larvae, which
show a full set of serotonergic perikarya in the
anterior region, while this pattern is incomplete
in the posterior area. This pattern suggests the
presence of a posterior growth zone (Hessling
and Westheide
2002 ). Similarly, in larval stages
of Urechis caupo , a serial repetitive distribution
of serotonin-containing neurons and their cor-
responding pairs of peripheral nerves, both
formed in an anterior-posterior gradient, imply
a segmental pattern. Moreover, larvae show a
paired origin of the ventral nerve cord (Hessling
2002 ). Another case of “ontogeny recapitulating
phylogeny” has been demonstrated for sipuncu-
lans, where neurogenesis of Phascolosoma
agassizii follows a segmental pattern. During
9 Annelida

214
AB
CD
EF
Serotonin Serotonin + FMRFFMRF
Fig. 9.11 Schematic
representation of neuronal
development in larval
stages of Phyllodoce
groenlandica exhibiting
formation from two
subsystems (Modifi ed from
Helm et al. (
2013 ))
Diagrams are in ventral
view; anterior is up. Major
types of neuronal
structures are color coded.
( A ) 0.5 days past hatching
( dph ). First serotonergic
immunoreactivity appears
at the posterior pole. ( B ) 2
dph. Nervous system
originates from posterior
(serotonin) and anterior
(FMRFamide). ( C ) 7 dph.
Serotonergic and
FMRFamidergic immuno-
reactivities start to overlap
to the greatest extent. ( D )
11 dph. Numerous
serotonergic cells are
detectable within the
epi- and the hyposphere.
( E ) 20 dph. Serotonergic
and FMRFamidergic nerve
cells are limited mainly to
anterior regions. ( F ) 34
dph. The larval nervous
system is fully developed.
at apical tuft, pt prototroch
C. Bleidorn et al.

215
development of this species, a pair of
FMRFamidergic and serotonergic axons gains
four pairs of associated serotonergic perikarya
and interconnecting commissures in an anterior-
posterior progression. During later larval stages,
the commissures disappear and the two seroto-
nergic axons fuse, forming a single ventral
nerve cord, and after cell migration, a nonmeta-
meric central nervous system can be found in
adults (Kristof et al. 2008 ). Interestingly, neuro-
genesis of the sipunculan Phascolion strombus
lacks any signs of a segmented origin, and sero-
tonergic structures are missing completely in
their larvae, which may be the result of the
abbreviated larval phase in this species
(Wanninger et al.
2005 ).
Nephridia and Coelomogenesis
Most adult annelid taxa possess metanephridia as
excretion organs; however, in the development
they are usually preceded by protonephridia,
which can be found in larvae and sometimes also
in developing juveniles (Bartolomaeus and Quast
2005 ). These larval excretory organs were termed
“head kidneys” by Hatschek ( 1886 ) and are
located anteriorly in trochophore stages, closely
behind the larval eyes (Bartolomaeus and Quast
2005 ). Later, homologous organs were also
described from lecithotrophic developmental
stages, as shown for the direct developing species
Scoloplos armiger (intertidal clade), which has
no free-swimming larval stage (Bartolomaeus
1998 ). Head kidneys are present in some echiu-
rans but are missing in sipunculid developmental
stages. It is hypothesized that the ancestral state
of larval annelid protonephridia was an organ
composed of three cells: a terminal cell, a nephro-
pore cell, and a duct cell (Bartolomaeus and
Quast
2005 ). This simple construction has been
modifi ed in adaptation to different developmental
modes in several annelid lineages and especially
in planktotrophic larvae, where the nephropore
cell is often missing (Kato et al. 2011 , 2012 ).
Adult segmental nephridia differentiate from a
single anlage, consisting of few cells which line a
small lumen fi lled with microvilli. This duct
becomes ciliated, and the most proximal cells are
separated during coelomogenesis. During coelo-
mic cavity growth, the proximal part of the anlage
is passively opened, forming the metanephridial
funnel. A truncation of this process due to sup-
pression of the separation of duct cells leads to a
differentiation into a protonephridium, as, for
example, observed in several taxa of the
Phyllodocida (Bartolomaeus 1999 ). The develop-
ment of metanephridia in the non-segmented
sipunculids as investigated for Golfi ngia minuta
seems to be similar as in segmented annelids. An
overview of annelid nephridial organs is given by
Bartolomaeus and Quast (
2005 ).
Segmented annelids show a heteronomous
coelomogenesis, and the coelomic lining is of
mesodermal origin. Prior to metamorphosis (if
present), a pair of unsegmented coelomic cavities
stretches out over the fi rst larval segments. This
process has been studied in detail for the serpulid
Spirorbis spirorbis , where two caudally located
mesodermal cell clusters proliferate cells, which
merge to surround the gut ventrally close to the
anus. Fluid starts to accumulate between spaces
of desmosomes, and at the same time myofi brils
appear, and due to growth and separation pro-
cesses, this myoepithelium develops into the coe-
lomic lining. After migration, the two coelomic
cavities meet dorsally, completely surrounding
the gut. Postmetamorphic stages develop strictly
segmental coelomic cavities during segment for-
mation. Coelomic cavities are highly reduced or
completely missing in leeches and several meio-
fauna annelids with a presumed progenetic origin
(Koch et al. 2014 ).
GENE EXPRESSION
Only few model annelids are well characterized
concerning gene expression patterns. Prime can-
didate taxa are leeches of the genera Helobdella
and Hirudo , the sedentary annelid Capitella
teleta , and the errant annelid Platynereis dumeri-
lii . Besides this, some studies exist for the
chaetopterid Chaetopterus , some sipunculid
worms, and a few serpulids as well as for addi-
tional nereidid and clitellate species. These
9 Annelida

216
studies mainly used candidate gene approaches,
where orthologs were chosen based on studies in
arthropods and vertebrates. Main points of
interest are the genomic basis of segmentation,
Hox gene expression, and nervous system
development as well as gastrulation and gut
development.
Segmentation
Metameric segmentation can be found in verte-
brates, arthropods, and annelids, and the distant
phylogenetic position of these taxa gave rise to
the question of how often this feature evolved in
animals (Seaver
2003 ). Traditionally well inves-
tigated is the molecular background of segmenta-
tion in vertebrates and arthropods, which show
profound differences (Tautz 2004 ). Given the fact
that a close relationship between arthropods and
annelids was suspected as formulated in the
Articulata hypothesis (Scholtz 2002 ), a possible
common ancestry of segmentation in these taxa
became a focus of many evolutionary develop-
mental studies of annelids. Genes or gene fami-
lies identifi ed to play a vital role in segment
formation in arthropods were used as candidates
in several studies (Wedeen and Weisblat 1991 ;
Prud’homme et al. 2003 ; Seaver and Kaneshige
2006 ; Saudemont et al. 2008 ; Dray et al. 2010 ;
Steinmetz et al. 2011 ).
Best investigated is the molecular background
of segmentation in Platynereis dumerilii and
Capitella teleta . For Drosophila it has been dem-
onstrated that para-segmental borders are gener-
ated by an interaction between the segment
polarity genes wingless and engrailed (Tautz
2004 ). The gene wingless (or Wnt1 ) is part of the
Wnt gene family, and engrailed is a
homeodomain- bearing transcription factor. As in
arthropods, a role in segment formation is sug-
gested for this pair of genes in P. dumerilii
(Prud’homme et al. 2003 ), where an expression
of continuous ectodermal stripes is observed for
these genes at the border of the segments during
their formation. However, investigation of other
annelid taxa questions the conserved nature for
engrailed as a “segmentation gene” in annelids.
In Chaetopterus , engrailed is expressed during
all larval stages in different structures or organs,
and no signs of a putative segment polarity pat-
tern of expression are obvious (Seaver et al.
2001 ). Congruently, no conserved segment polar-
ity pattern was found investigating the expression
of this gene in developing C. teleta or Hydroides
elegans individuals (Seaver and Kaneshige
2006 ). Finally, ablation of individual cells
expressing engrailed in the leech Helobdella did
not hinder remaining segmental clones in their
normal development (Seaver and Shankland
2001 ). Consequently, the establishment of seg-
ment polarity in the leech (and possibly many
other annelid taxa) seems to be independent of
cell interactions across the anterior- posterior axis
as known for arthropods (Seaver and Shankland
2001 ).
A set of pair rule genes is expressed in
Drosophila and many other arthropods to pattern
the embryo across the anterior-posterior axis,
including eve , hairy , and runt (Damen 2007 ). All
or some of these genes were investigated in detail
for C. teleta and Helobdella robusta . In contrast
to Drosophila , where it is expressed in stripes in
the growth zone, the Capitella ortholog of hairy
( Cap - hes1 ) shows an expression limited to a
small band of cells in each larval segment. In
juveniles its expression is limited to a small
mesodermal domain of the posterior growth zone
(Thamm and Seaver 2008 ). In vertebrates and
some arthropods, the expression of hairy is con-
trolled by the Notch pathway (Stollewerk et al.
2003 ). In Capitella , Notch and hairy do not show
a broadly overlapping expression, with a Notch
localization in already formed segments, anterior
to the hairy signal (Thamm and Seaver
2008 ). In
Helobdella , hairy is expressed in teloblasts and
primary blast cells. The expression peak corre-
lates with the production of blast cells by the
teloblasts. However, no striped pattern suggest-
ing a pair rule function was found (Rivera et al.
2005 ). Similarly, Notch is also expressed in telo-
blasts and blast cells, and functional studies
revealed that the disruption of the Notch / hairy
signaling results in a disruption of segmentation
(Song et al. 2004 ; Rivera and Weisblat 2009 ). For
Platynereis dumerilii , 15 hairy paralogs could be
C. Bleidorn et al.

217
identifi ed, which are expressed in mesodermal
tissue, forming segments, and during neurogene-
sis, where it may be involved in the patterning of
the nervous system (Gazave et al. 2014 ).
However, these authors also found no overlap
with the expression of Notch .
The expression patterns of the arthropod pair
rule genes eve and runt do not suggest a similar
role in Capitella . Instead, the expression of runt
can be found in the brain and ventral nerve cord,
as well as the fore- and hindgut. Two eve paralogs
were characterized for Capitella , both showing a
complex expression pattern, which does not cor-
respond to segmental stripes as expected by
results from Drosophila (Seaver et al.
2012 ).
Likewise, de Rosa et al. ( 2005 ) did fi nd such a
pattern of eve expression in developing
Platynereis dumerilii . However, these authors
speculate about a role of this gene in the posterior
addition of segments. A detailed functional study
for eve has been conducted for Helobdella (Song
et al. 2002 ). Segments arise sequentially from
fi ve pairs of teloblasts in leeches (see above), and
eve is expressed in these teloblasts and their pri-
mary blast cells in Helobdella . Later embryos
express eve in cells of the ventral nerve cord
which stem from the N teloblast. Morpholino
knockdowns suggest a role of eve in early cell
division through early segmentation in
Helobdella . However, no pair rule pattern is
found for this gene in the leech.
The zinc fi nger transcription factor hunchback
plays the role of a gap gene in Drosophila , which
defi nes expression domains of pair rule and Hox
genes (see Vol. 5, Chapter 1 ). Moreover, hunch-
back is involved in mesoderm development and
neurogenesis. In Platynereis dumerilii , hunch-
back expression is detected in mesodermal cells
belonging to the posterior growth zone of juve-
nile worms. Additionally, expression in the pre-
cursors of the somatic segmented mesoderm,
formed during larval development, could also be
confi rmed, a striking similarity with arthropods
(Kerner et al.
2006 ). However, an expression of
hunchback could not be detected in segmental
precursor cells of the posterior growth zone in
Capitella and Helobdella , and a role in the pat-
terning of the anterior-posterior axis was rejected
for these species (Iwasa et al.
2000 ; Werbrock
et al. 2001 ).
NKL genes are a family of homeodomain
transcription regulators that are involved in the
patterning of mesodermal derivatives in
Drosophila (Holland 2001 ; Jagla et al. 2001 ).
The expression of seven genes of this cluster has
been investigated in developing Platynereis
dumerilii , and all are involved in the specifi cation
of mesodermal derivatives including muscular
precursors (Saudemont et al. 2008 ). Notably, fi ve
of the investigated genes ( NK4 , Lbx , Msx , Tlx ,
and NK1 ) show an expression in complementary
stripes in the mesoderm and/or ectoderm of
developing segments. Moreover, genes of the
Hedgehog signaling pathway show a similar
striped pattern of expression, and segment forma-
tion in P. dumerilii is disrupted when treated with
molecules antagonistic to this signaling (Dray
et al.
2010 ).
Wnt genes regulate a wide range of develop-
mental processes, including axis elongation and
segmentation (Cadigan and Nusse 1997 ). This
gene family ancestrally includes 13 paralog
groups, of which several metazoan lineages lost
some of the genes (Janssen et al. 2010 ). In
Platynereis dumerilii and Capitella teleta , all
paralog groups besides Wnt3 could be discov-
ered. In the leech Helobdella robusta , only nine
paralog groups are present, with additionally
Wnta , Wnt8 , and Wnt9 missing. Most Wnt genes
in P. dumerilii are expressed in ectodermal seg-
mental stripes and/or in the area around the
pygidium (Janssen et al. 2010 ). Expression anal-
yses in H. robusta and C. teleta led to comparable
results (Cho et al. 2010 ). Due to similarities with
arthropods, a role of Wnt genes in segment for-
mation in both annelids and arthropods is sug-
gested by some authors (Janssen et al.
2010 ).
In summary, the candidate gene approach led
to the discovery of many similarities as well as
differences between annelids and arthropods in
gene expression patterns during the formation of
segments. The expression of some genes at seg-
mental boundaries in Platynereis dumerilii
shows a remarkable similarity to arthropods.
However, for other annelid taxa investigated for
these candidate genes (as, e.g., for engrailed or
9 Annelida

218
hunchback ), the picture becomes less clear, and
future studies covering segment formation in
more annelid taxa are clearly wanted. Moreover,
fewer similarities are found compared with
arthropods when investigating pair rule genes. In
the discussion of a putative common ancestry of
segmentation in annelids and arthropods, differ-
ent authors come to different conclusions using
basically the same set of evidence (de Rosa et al.
2005 ; Thamm and Seaver 2008 ). However,
homology of genes expressed during segment
formation must not imply a homology of a seg-
mented body plan itself. At present, available
developmental, paleontological, and phyloge-
netic evidence supports a convergent evolution
of segmentation in arthropods and annelids
(Couso
2009 ; Chipman 2010 ). Given this
hypothesis, co-option of the same set of genes
into the process of segment formation leading to
a convergent pattern of gene expression can
explain the similarities found between annelids
and arthropods (Chipman 2010 ; Ferrier 2012 ).
Hox and ParaHox Genes
Hox genes comprise a family of transcription
factors bearing a DNA-binding homeodomain
(Gellon and McGinnis 1998 ). Hox genes are usu-
ally found as linked chromosomal clusters and
show spatial and temporal collinearity (Garcia-
Fernandez 2004 ). This means that genes from
the 5′-end of the cluster are usually expressed
more anteriorly than the ones from the 3′-end. In
similar fashion we also see a temporarily earlier
onset of genes from the 5′-end compared to those
from the 3′-end. In bilaterian animals Hox genes
are mainly involved in the patterning of body
regions; however, several examples of co-option
into other areas of expression are described
(Wagner et al.
2003 ). All these characteristics
made this set of genes a prime target for evolu-
tionary developmental biologists to understand
major transitions in animal body plan evolution
(Akam
1998 ).
For annelids, the genomic organization of the
Hox cluster is only fully described for Capitella
teleta and Helobdella robusta (Fröbius et al.
2008 ; Simakov et al. 2013 ). In Capitella , assem-
bled whole genome shotgun data found Hox
genes distributed on three scaffolds, with one
scaffold containing the Post1 genes clearly sepa-
rated from the others. In contrast, the leech
Helobdella shows an extensive fragmentation of
the Hox cluster (Simakov et al. 2013 ). For
Capitella , 11 Hox genes ( lab , pb , Hox3 , Dfd , Scr ,
lox5 , Antp , lox4 , lox2 , Post2 , post1 ) correspond-
ing to 11 different paralog groups were detected,
and the presence of these genes are regarded as
ancestral for lophotrochozoans in general
(Fröbius et al.
2008 ; Simakov et al. 2013 ).
Interestingly, Helobdella also shows a derived
pattern here, with the duplication of two paralog
groups (fi ve copies of Scr and two copies of
Post2 ) and the loss of orthologs of pb and Post1
(Simakov et al. 2013 ). For many other annelids,
information about the Hox gene complement are
available through PCR and cloning studies; how-
ever, genomic organization and absence of genes
cannot be derived from this approach (Dick and
Buss 1994 ; Snow and Buss 1994 ; Irvine et al.
1997 ; Cho et al. 2003 , 2006 ; Kulakova et al.
2007 ; Bleidorn et al. 2009 ).
The expression of Hox genes during develop-
ment has been only investigated for a few annelid
taxa, Capitella teleta , Chaetopterus variopeda-
tus , Alitta ( Nereis ) virens , Platynereis dumerilii ,
Hirudo medicinalis , and two Helobdella species
(Irvine and Martindale 2000 ; Peterson et al.
2000 ; Kulakova et al. 2007 ; Fröbius et al. 2008 ;
Gharbaran and Aisemberg 2013 ). The most
inclusive study deals with C. teleta , where for the
fi rst time spatial and temporal collinearity for
Hox genes could be demonstrated for a lophotro-
chozoan taxon (Fröbius et al.
2008 ). Capitella
Hox genes, except for Post1 , are all expressed in
ectodermal domains during larval development,
with a spatial correlation of anterior expression
borders and location of genes in the Hox cluster.
Anterior class Hox genes ( lab , pb , Hox3 ) are the
fi rst genes expressed, occurring before the
appearance of segments. The expression of Dfd
and Scr can be detected after appearance of the
fi rst segments, followed by the expression of
lox5 , Antp , and lox4 . The expression of lox2 and
Post2 appears last. Interestingly, all Hox genes in
C. Bleidorn et al.

219
C. teleta show their highest expression level at a
unique stage during the course of development,
refl ecting the order of activation for each gene. A
unique Hox gene expression boundary can be
detected for all nine thoracic segments, and the
posterior-most located Hox genes ( lox2 and
Post2 ) are only expressed in the abdomen
(Fig. 9.12 ). Whereas no expression of Hox genes
could be detected in the pygidium of C. teleta ,
expression of Post2 was detected in the pygidium
of nereidids (Kulakova et al.
2007 ). Expression
of Post1 , the gene which seems to be separated
from the rest of the Hox cluster, could not be
detected in any investigated stages of C. teleta ,
besides some signals in chaetal sacs (Fig. 9.12 ;
Fröbius et al. 2008 ). This result is congruent with
analyses of expression of this gene in nereidids
(Kulakova et al. 2007 ).
Fig. 9.12 Hox gene expression profi le in larvae and juve-
niles of Capitella teleta after Fröbius et al. (
2008 ). Solid
bars indicate strong expression; dashed bars indicated
weaker expression. Abbreviations: A1–4 abdominal seg-
ments 1–4, Gz growth zone, Pe peristomium, Pr prosto-
mium, Py pygidium, T1–T9 thoracic segments 1–9
9 Annelida

220
A staggered expression of fi ve Hox genes gen-
erally in line with spatial and temporal collinear-
ity can also be found in Chaetopterus , even
though the genomic organization of the Hox clus-
ter remains unknown in this species (Irvine and
Martindale 2000 ). No strict temporal collinearity
was found in expression studies for nereidid
worms and Helobdella (Kourakis et al. 1997 ;
Kulakova et al. 2007 ). However, all studies sug-
gest an involvement of Hox genes in body pat-
terning along the anterior-posterior axis, a
function that seems to be the ancestral role of
Hox genes in bilaterian animals (Kulakova et al.
2007 ; Butts et al. 2008 ). Notably, annelids show
a predominant expression of Hox genes in neuro-
genic structures such as the ganglia and the ven-
tral nerve cord. This is especially obvious in
Chaetopterus and leeches (Shankland et al. 1991 ;
Aisemberg and Macagno 1994 ; Wong et al. 1995 ;
Kourakis et al. 1997 ; Irvine and Martindale 2000 ;
Gharbaran and Aisemberg 2013 ).
The ParaHox cluster is a paralog of the Hox
gene cluster, containing three genes ( Gsx , Xlox ,
and Cdx ) (Brooke et al. 1998 ). As for Hox genes,
temporal collinearity has been likewise demon-
strated for ParaHox genes in many instances.
However, the ParaHox cluster seems to be lost in
several investigated ecdysozoan taxa which show
a breakup of the cluster and missing genes
(Ferrier and Minguillon 2003 ). All three ParaHox
genes seem to be present in annelids (Ferrier and
Holland 2001 ; Fröbius and Seaver 2006 ; Park
et al. 2006 ). The genomic organization of
ParaHox genes has been studied in detail for
Platynereis dumerilii (Hui et al. 2009 ). In this
species, a head-to-head location of Gsx and Xlox
could be demonstrated, with Cdx located in a
separate position on the same chromosome.
Expression analyses of these genes in Alitta
( Nereis ) virens suggest a role in anterior- posterior
patterning of the digestive system and in the
specifi cation of neuroectodermal cell domains
(Kulakova et al.
2008 ). Especially Gsx seems to
be involved in the development of the brain in all
investigated annelids ( P. dumerilii , A . ( N .) virens ,
C. teleta ), a function which is regarded as ances-
tral for these genes for bilaterians in general
(Fröbius and Seaver 2006 ; Kulakova et al. 2008 ).
Genes Involved in Neurogenesis
The development of the central nervous system
has been deeply studied for Platynereis dumeri-
lii . Neural progenitor cells are located close to
the ventral midline and express axin , a negative
regulator of the Wnt/β-catenin pathway, which
controls the transition between these proliferat-
ing cells and differentiating neurons (Demilly
et al. 2013 ). Wnt-controlled proliferation of neu-
ral progenitors is also well-documented for verte-
brates and arthropods, especially Drosophila
(Bielen and Houart
2014 ). Using a candidate
gene approach, genes with a conserved expres-
sion in developing vertebrate and arthropod
brains were chosen as major targets in studies on
annelids. The developing head of annelid larvae
is demarcated by the expression of six3 and otx
homeobox genes (Fig. 9.13 ), a patterning system
that might be universal to bilaterian animals
(Steinmetz et al. 2010 ). MicroRNAs are short
noncoding RNAs that posttranscriptionally regu-
late gene expression (Ambros 2004 ). The expres-
sion of several microRNAs is highly tissue
specifi c and conserved across animals
(Christodoulou et al. 2010 ). In P. dumerilii and
Capitella teleta , the microRNAs mir - 7 , mir - 137 ,
and mir - 153 show a localized expression in dis-
tinct neurosecretory brain tissue, a pattern which
was also found in zebra fi sh (Tessmar-Raible
et al. 2007 ; Christodoulou et al. 2010 ). The
expression of the complementary pair mir - 9 and
mir - 9 */ mir - 131 is restricted to two sets of differ-
entiated neurons in the developing annelid brain,
with the most apical cells located at the base of
the antennae (Christodoulou et al.
2010 ). The
three transcription factors rx , otp , and nk2.1 are
all expressed in the developing forebrain of P.
dumerilii . All cells expressing these genes, as
well as mir - 7 , are vasotocinergic extraocular pho-
toreceptors. The expression pattern matches
those known from the same cell type in zebra fi sh
(Tessmar-Raible et al. 2007 ). Gene networks
controlling the pattern along the anterior- posterior
axis of the central nervous system are conserved
across bilaterians, and the involved genes are
mainly Hox genes (Ferrier 2012 ). The patterning
of the dorso-ventral axis in P. dumerilii is
C. Bleidorn et al.

221
controlled by a gene network including nk2.2 ,
nk6 , Pax2 / 5 / 8 , Pax6 , Pax3 / 7 , dlx , msx , gsx , sim ,
and dbx (Denes et al. 2007 ; Ferrier 2012 ). A sim-
ilar patterning of the neuroepithelium is obvious
in vertebrates (Denes et al.
2007 ).
Besides the detailed investigations summa-
rized for Platynereis dumerilii , some gene
expression studies dealing with the development
of the nervous system in leeches and Capitella
teleta are published. As shown for P. dumerilii ,
otx shows a largely head-specifi c expression in
Helobdella (Bruce and Shankland 1998 ). The
expression of Lox10 , a putative nk2.1 ortholog,
was detected in the developing brain of
Helobdella , congruent with the results for P.
dumerilii (Nardelli-Haefl iger and Shankland
1993 ). In Hirudo medicinalis , the central class
Hox gene Lox1 controls the differentiation of the
so-called “rostral penile evertor neurons” that
innervate the male penis (Gharbaran and
Aisemberg
2013 ). For the same species, expres-
sion of the axon migration guiding protein netrin
is shown to be involved in forming intergangli-
onic neuronal tracts and in defi ning ventrodorsal
boundaries of peripheral innervation (Gan et al.
1999 ). Another protein family investigated in H.
medicinalis is the innexins, where several cloned
members show a restricted expression in neurons
(Dykes and Macagno 2006 ). In C. teleta , Delta
and Notch expression was detected during brain
development in larvae, as well as in the forming
ganglia of the ventral nerve cord of juveniles,
suggesting a role of Notch signaling in neurogen-
esis (Thamm and Seaver 2008 ).
In summary, development of the central ner-
vous system and expression patterns of genes
localized in the brain show strong similarities
between vertebrates and annelids. Based on
Head
Trunk
Pro-
stomium
Larval eye
Six3
otx
gbx
hox1
hox4
lox5
Peri-
stomium
Meta-
stomium
Pygidium
Fig. 9.13 Expression of six3 , otx , gbx , and Hox genes in
neuroectodermal regions of Platynereis dumerilii larvae.
The six3 and otx expressing regions cover the developing
prostomium and the peristomium, from which the cere-
bral ganglia and eyes develop. Dark gray region marks the
mouth (From Steinmetz et al. (
2010 ) )
9 Annelida

222
these results (and further studies involving
other taxa), the presence of a centralized ner-
vous system in the last common ancestor of
protostomes and deuterostomes seems plausi-
ble for several authors (Arendt et al. 2008 ;
Holland et al. 2013 ).
Genes Acting in the Development
of the Digestive Tract
The gut of annelids consists of a foregut (stomo-
deum) and a hindgut (proctodeum), both origi-
nating from ectoderm, as well as of the midgut
which is of endodermal origin. All three parts
can usually be subdivided into different func-
tional regions (Tzetlin and Purschke
2005 ).
Several genes involved in bilaterian foregut and
hindgut patterning have been investigated for
Platynereis dumerilii (Arendt et al. 2001 ). As
such, the T-box transcription factor brachyury is
expressed in the ventral part of the developing
foregut as well as in the hindgut of late trocho-
phore larvae, resembling the pattern known from
larvae of basal branching deuterostomes. The
homeobox gene goosecoid is fi rst expressed in a
small number of cells at the anterior blastopore
margin which develops into the foregut.
Expression can be additionally detected in adja-
cent cells which will contribute to the develop-
ment of the foregut nervous system. As the
expression patterns of the investigated genes
seem to be conserved in protostomes and deu-
terostomes, a single origin of the tripartite bilat-
erian through gut has been hypothesized (Arendt
et al. 2001 ). This idea has been later challenged
based on expression studies of the same set of
genes in acoels, which are lacking a through gut
(Hejnol and Martindale
2008 ).
Genes involved in the patterning of ectoder-
mal and endodermal parts of the gut have been
studied for Capitella teleta , Chaetopterus vari-
opedatus , and the sipunculid Themiste lagenifor-
mis . In C. teleta , the transcription factor FoxA
and genes of the GATA family are expressed
across the entire developing gut (Boyle and
Seaver 2008 ). Different genes of the GATA fam-
ily are exclusively expressed in the developing
midgut, with a prominent expression of gataB1 at
its boundaries. In contrast, FoxA expression can
be detected surrounding the blastopore during
development as well as in the foregut and hindgut
during organogenesis. Partly similar expression
patterns are reported for Themiste and
Chaetopterus and might refl ect the differences in
the gut architecture of these species with differ-
ent feeding mechanisms (Boyle and Seaver
2010 ). Moreover, expression of the ParaHox
gene Cdx is also reported for anterior and poste-
rior regions of the gut in C. teleta (Fröbius and
Seaver
2006 ). In nereidids, the ParaHox gene
Gsx is expressed during the formation of the fore-
gut and the midgut. The expression of the
ParaHox gene Xlox has been detected in all inves-
tigated annelids, including Helobdella , H. medic-
inalis , C. teleta , and nereidids (Wysocka-Diller
et al. 1995 ; de Rosa et al. 2005 ; Fröbius and
Seaver 2006 ; Kulakova et al. 2008 ; Hui et al.
2009 ).
Bilaterian animals are divided into deutero-
stomes, ecdysozoans, and lophotrochozoans
(Edgecombe et al. 2011 ). Whereas research on
several well-established model organisms in the
former two groups (e.g., Drosophila ,
Caenorhabditis , Danio , Mus ) has provided
detailed insights into molecular mechanisms of
the development, lophotrochozoans have tradi-
tionally been chronically understudied in this
regard (Tessmar-Raible and Arendt 2003 ). The
rise of Platynereis dumerilii (and other annelids
like Capitella and Helobdella ) as EvoDevo mod-
els has provided major insights into the evolution
of the nervous system and segment formation in
annelids, a key lophotrochozoan phylum (see
above). Interestingly, the genomic architecture of
Platynereis seems to be little derived from a
hypothetical bilaterian ground pattern, enabling
many insights into comparative developmental
genomics (Raible et al.
2005 ; Ferrier 2012 ).
Future studies focusing on additional annelid lin-
eages, such as the basal branching oweniids or
the non-segmented sipunculans, will certainly
improve our understanding of the evolution of
bilateria in general.
C. Bleidorn et al.

223
OPEN QUESTIONS
• Segment formation in non-clitellate annelids
• Myogenesis in Echiurida
• Development of the nervous system in basal
branching annelids
• Homology of ciliary bands including the vari-
ous trochi in different annelid and lophotro-
chozoan larvae
• Genetic background of annelids showing
putative deuterostomy
• Comparative expression studies of Hox,
ParaHox, and other key developmental genes
across the various annelid subtaxa, especially
lesser-known groups such as Myzostomida,
Sipuncula, and Echiurida
• Gene expression studies of “segmentation
genes” in non-segmented annelids
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