PhD thesis

Comparative neurogenesis, muscle
development, and gene expression
analyses in Brachiopoda
PhD thesis
Andreas Altenburger

THE PHD SCHOOL OF SCIENCE
FACULTY OF SCIENCE
DEPARTMENT OF BIOLOGY
UNIVERSITY OF COPENHAGEN
DENMARK
PhD thesis
Andreas Altenburger
Comparative neurogenesis, muscle
development, and gene expression analyses in
Brachiopoda
Principal supervisor
Associate Prof. Dr. Andreas Wanninger
Co-supervisor
Prof. Dr. Pedro Martinez, University of Barcelona
December , 2010

2 
Principal supervisor
Assoc. Prof. Dr. Andreas Wanninger
Department of Biology
Research Group for Comparative Zoology
University of Copenhagen
Copenhagen, Denmark
Co-supervisor
Prof. Dr. Pedro Martinez
Department of Genetics
University of Barcelona
Barcelona, Spain
Opponents
Prof. Dr. Billie Swalla
Department of Biology
University of Washington
Seattle, USA
Prof. Dr. Bernard Degnan
School of Biological Sciences
The University of Queensland
Brisbane, Australia
Faculty opponent
Assoc. Prof. Dr. Jørgen Olesen
Zoological Museum
Natural History Museum of Denmark
Copenhagen, Denmark
Cover legend
Front: Myoanatomy of Joania (Argyrotheca) cordata. Maximum projection
micrograph of a confocal laserscanning microscope stack. F-actin is labelled
in red, cell nuclei are labelled in blue to indicate the outline of the specimen.
Anterior faces upward and the specimen is approximately 280 µm long.
Back: Schematic illustration of the specimen shown on front. The musculature
comprises pedicle muscles (beige), longitudinal muscles (orange), central
mantle muscles (brown), a U-shaped muscle (green), setae pouch muscles
(red circles), circular mantle muscle (light blue), serial mantle muscles (dark
orange), setae muscles (purple), apical longitudinal muscles (dark blue), and
an apical transversal muscle (yellow).


3
Content
Preface ......................................................................................... 4
Danish abstract ............................................................................... 5
Abstract ......................................................................................... 6
Short abstract ................................................................................. 7
Acknowledgements ......................................................................... 8
Chapter I ....................................................................................... 9
Introduction ................................................................................ 9
Brachiopoda .......................................................................... 9
Nervous system .................................................................... 10
Muscular system ................................................................... 10
Gene expression ....................................................................11
Material and methods ................................................................. 12
Immunocytochemistry and phalloidin labeling .............................. 12
Labeling of Pax3/7 proteins ..................................................... 12
Detection of proliferating cells with BrdU (5-bromo-2-deoxyuridine)
staining ............................................................................... 13
Gene expression analyses ...................................................... 13
Illustrations .......................................................................... 14
Results and discussion ................................................................ 16
Larval development ............................................................... 16
Myogenesis ......................................................................... 20
Neurogenesis with special focus on the apical organ of lophotrochozoan
larvae ................................................................................. 20
Distribution of Pax3/7 proteins in larvae of Terebratalia transversa 22
Growth patterns of Terebratalia transversa ................................. 24
Not and Cdx expression analyses ............................................. 26
References ............................................................................... 28
Chapter II ..................................................................................... 37
Altenburger, A. & Wanninger, A. 2009 Comparative larval myogenesis
and adult myoanatomy of the rhynchonelliform (articulate) brachiopods
Argyrotheca cordata, A. cistellula, and Terebratalia transversa. Frontiers
in Zoology 6: 1-14 ................................................................. 37
Chapter III .................................................................................... 52
Altenburger, A. & Wanninger, A. 2010 Neuromuscular development
in Novocrania anomala: evidence for the presence of serotonin and a
spiralian-like apical organ in lecithotrophic brachiopod larvae. Evolution
& Development 12: 16-24 ....................................................... 52
Chapter IV ................................................................................... 62
Altenburger, A., Martinez, P. & Wanninger, A. First expression study of
homeobox genes in Brachiopoda: the role of Not and Cdx in bodyplan
patterning and germ layer specification. Submitted ...................... 62

4 
Preface
This thesis presents the results of three years of research at the University of
Copenhagen from May 2007 until December 2010, including a research visit of
one year at the University of Barcelona in 2009. The research on neurogenesis,
myogenesis, and gene expression patterns in Brachiopoda was supervised by
Assoc. Prof. Dr. Andreas Wanninger at the Research Group for Comparative
Zoology, Department of Biology, University of Copenhagen, Denmark. The
research on gene expression patterns was mainly carried out in the lab of Prof.
Dr. Pedro Martinez, Department of Genetics, University of Barcelona, Spain.
The PhD project was funded by The Danish Agency for Science, Technology
and Innovation (grant no. 645-06-0294 to Andreas Wanninger).
This project included several research visits of altogether nine weeks at the
Sven Lovén Center for Marine Sciences in Kristineberg, Sweden, three weeks
at the Moreton Bay Research Station on North Stradbroke Island, Australia,
three weeks at the Banyuls-sur-mer Oceanological Observatory, France, and
ten weeks at the Friday Harbor Laboratories, USA. Additional impact on my
thinking about the field of evolution and development had the summer school on
Evolution and Development of the Metazoans by Prof. Dr. Billie Swalla and Prof.
Dr. Ken Halanych at the Friday Harbor Laboratories, University of Washington,
USA, the Summer School on Evolutionary Developmental Biology by Prof. Dr.
Alessandro Minelli and Assist. Prof. Giuseppe Fusco, University of Padua, Italy,
and the EMBO course on Marine Animal Models in Evolution and Development
organized by Prof. Dr. Detlev Arendt at the University of Gothenburg, Sweden.
This thesis is composed of four chapters. Chapter I constitutes a short
introduction to the research field and discusses the presented results in a
broader perspective. Chapters II-IV contain two published papers and one
submitted manuscript, which report the major findings made during this PhD
project.
Copenhagen, December 2010
Andreas Altenburger


5
Danish abstract
Brachiopoda udgør en dyrerække med en unik kropsbygning. Rækken omfatter
ca. 370 nulevende arter opdelt i tre undergrupper, Rhynchonelliformea,
Craniiformea og Linguliformea, men der er over 12.000 beskrevne fossile arter
daterende helt tilbage til tidlig Kambrium. Der er uenighed om brachiopodernes
fylogenetiske position som ofte debateres. Mit projekt har belyst dette problem
gennem ny indsigt i brachipodernes ontogeni. Jeg har beskrevet udviklingen
af nerve- og muskelsystemerne hos de rhynchonelliforme og craniiforme
brachiopod larver af henholdsvis Terebratalia transversa og Novocrania
anomala ved hjælp af immunohistokemiske indfarvninger kombineret med
konfokal laserskanning mikroskopi og 3D-rekonstruktioner. Muskeldannelsen
er beskrevet for både larver og voksne af Joania (Argyrotheca) cordata og
Argyrotheca cistellula og ekspressionsmønstret af transskriptionsfaktorerne
DP311, DP312 (Pax3/7) er beskrevet for larver og voksne af Terebratalia
transversa. Ekspressionsmønstret af homeobox-generne TtrNot og TtrCdx er
beskrevet for larver og juvenile af Terebratalia transversa ved hjælp af whole
mount in situ hybridisering. De væsentligste resultater er: (1) Muskelanatomien
hos rhynchonelliforme brachiopodlarver udviser stor lighed trods store forskelle i
larvernes ydre morfologi. (2) Rhynchonelliforme og craniiforme brachiopodlarver
af henholdsvis Terebratalia transversa og Novocrania anomala udviser et
serotoninholdigt nervesystem, som omfatter fire eller otte flaskeformede celler
i apikalorganet. Et sådant apikalorgan med flaskeformede celler er muligvis
en morfologisk apomorfi for Lophotrochozoa. (3) Ekspressionsmønstret af
TtrNot genet hos larverne af Terebratalia transversa indikerer en oprindelig
funktion af dette gen i forbindelse med gastrulation, ektoderm specifikation
og anlæggelse af nervebaner. For TtrCdx indikerer ekspressionsmønstret en
oprindelig funktion i forbindelse med gastrulation samt dannelsen af den bageste
del af det ektodermale væv hos Brachiopoda. Resultaterne bliver diskuteret
i et fylogenetisk perspektiv gennem sammenligninger med andre rækker
indenfor Lophotrochozoa, og implikationerne for evolutionen af Brachiopoda er
fremhævet.

6 
Abstract
Brachiopods are a small phylum with a unique body plan comprising around
370 living species and over 12.000 described fossil species dating back until the
Lower Cambrian. The phylogenetic position of brachiopods is under controversial
discussion. This project led to new insights into the ontogeny of brachiopods,
which are divided into three clades, Rhynchonelliformea, Craniiformea,
and Linguliformea. By use of immunocytochemistry combined with confocal
laserscanning microscopy and 3D reconstruction software I describe the
development of the nervous and muscular system in the rhynchonelliform and
craniiform brachiopod larvae of Terebratalia transversa and Novocrania anomala.
Myogenesis is described for larvae and adults of Joania (Argyrotheca) cordata
and Argyrotheca cistellula and distribution of the transcription factor proteins
DP311, DP312 (Pax3/7) for larvae and juveniles of Terebratalia transversa. The
expression patterns of the developmental homeobox containing genes TtrNot
and TtrCdx in larvae of Terebratalia transversa are described by use of whole
mount in situ hybridization. The main results are: (1) The larval myoanatomy of
rhynchonelliform brachiopod larvae is very similar, despite gross morphological
differences in their outer morphology. (2) The rhynchonelliform and craniiform
brachiopod larvae of Terebratalia transversa and Novocrania anomala show
a serotonergic nervous system comprising eight or four flask-shaped cells
in the apical organ. Such an apical organ with flask-shaped cells might be a
morphological apomorphy of Lophotrochozoa. (3) The expression pattern of
the TtrNot gene in larvae of Terebratalia transversa suggests an ancestral
role of this gene in gastrulation and ectoderm specification in Brachiopoda.
The expression pattern on TtrCdx suggests an ancestral role of this gene in
gastrulation and the formation of posterior ectodermal tissue in Brachiopoda.
The results are discussed in a phylogenetic framework compared to other
lophotrochozoan phyla and implications of the results for the evolution of
Brachiopoda are pointed out.


7
Short abstract
This thesis deals with selected aspects of brachiopod ontogeny. By use of
immunocytochemistry combined with confocal laserscanning microscopy and
3D reconstruction software the development of the nervous and muscular
system of rhynchonelliform and craniiform brachiopod larvae is described. The
expression patterns of the developmental homeobox containing genes TtrNot
and TtrCdx are described by use of whole mount in situ hybridization. The main
results are: (1) The larval myoanatomy of rhynchonelliform brachiopod larvae is
similar despite gross morphological differences in their outer morphology. (2) The
rhynchonelliform and craniiform brachiopod larvae show a serotonergic nervous
system comprising eight or four flask-shaped cells in the apical organ. An apical
organ comprising flask-shaped cells might be a morphological apomorphy of
Lophotrochozoa. (3) The expression pattern of the TtrNot gene in larvae of
Terebratalia transversa suggests an ancestral role of this gene in gastrulation
and ectoderm specification in Brachiopoda. The expression pattern on TtrCdx
suggests an ancestral role of this gene in gastrulation and the formation of
posterior ectodermal tissue in Brachiopoda.

8 
Acknowledgements
The endeavour of such a thesis is impossible without the help of many people
for whose support I am very grateful. Foremost I want to thank my principle
supervisor Andreas Wanninger whose office door was always open and who
did a great job in motivating and directing me towards the exciting parts of this
study and especially the publication of the results.
I am grateful to Pedro Martinez and his lab, namely Marta Chiodin, Amandine Bery,
Eduardo Moreno, and Alexander Alsen for an inspiring time in Barcelona.
I thank the teachers I had during PhD courses and who had a great influence on
my thinking about the field of evo-devo, especially Billie Swalla, Ken Halanych,
Alessandro Minelli, and Detlev Arendt.
I thank the colleagues with whom I had the pleasure to share the room, lab,
office, or a beer, Henrike Semmler, Nora Brinkmann, Tim Wollesen, Alen Kristof,
Ricardo Neves, Julia Merkel, Birgit Meyer, Lennie Rotvit, Louise Würtz, Jan
Bielecki, Jens Høeg, Lisbeth Haukrogh, Jan Lybeck, and visiting guests at the
lab in Copenhagen.
A special thank you to Anders Garm who translated the abstract into Danish.
Many thanks go to the staff at the marine stations where I collected animals,
in particular the Friday Harbor Laboratories, the Sven Lovén Centre for Marine
Sciences, the Observatoire Océanologique de Banyuls-sur-mer, and the
Moreton Bay Research Station.
A special thank you goes to my wife Ruth who supported my work wherever
she could and who took especially during the time in Barcelona the “burden” of
caring full time almost alone for our son.
This study was financially supported by a grant from the Danish Agency for
Science, Technology and Innovation (grant no. 645-06-0294 to Andreas
Wanninger) and a travel grant from Friday Harbor Labs to the author for
participation in their summer course.

Introduction
9
Chapter I
Introduction
Brachiopoda
The phylogenetic relationship of Brachiopoda is intensely debated among
biologists and paleontologists alike. Brachiopods were already known by Linné,
and 370 extant and more than 12.000 described fossil species are known (Linné
1758; Ax 2003; Logan 2007). Brachiopods were significant members of the early
Cambrian marine fauna and thus are one of the few phyla which are represented
throughout the 550 million years of the Phanerozoic era, which extends from
the first widespread appearance of organisms with mineralized skeletons until
modern times (James et al. 1992). Historically, brachiopods have been assigned
to different invertebrate groups, including molluscs (Lamarck 1801; Cuvier
1805), bryozoans (Huxley 1853; Hancock 1858), bryozoans and phoronids
(Hatschek 1888 ‘Tentaculata’; Hyman 1959 ‘Lophophorata’), or annelids (Morse
1871). The three lophophorate groups or Brachiopoda alone have subsequently
sometimes been regarded as deuterostomes (Brusca and Brusca 1990; Schram
1991; Eernisse et al. 1992; Nielsen 1995). Since the appearance of molecular
research tools, brachiopods have commonly been accepted to be protostomes
(Field et al. 1988; Lake 1990; Halanych 1995; Hejnol et al. 2009). Brachiopod
internal phylogeny distinguishes three clades; the inarticulate Linguliformea
and Craniiformea and the articulate Rhynchonelliformea (Williams et al. 1996).
Members of Linguliformea live buried in mud and have swimming juveniles
instead of a true larval stage (Yatsu 1902). Members of craniiformea live with
their ventral valve attached to stones and have two-lobed lecithotrophic larvae
(Rowell 1960). Members of Rhynchonelliformea have a pedicle with which they
attach themselves to rocks or other hard substrates (Williams et al. 1997). Their
larvae have three lobes and are lecithotrophic (Freeman 2003). Traditionally,
Linguliformea and Craniiformea have been grouped together as Inarticulata,
while Rhynchonelliformea have been named Articulata because their valves
are connected by a hinge (James et al. 1992).
Brachiopods are certainly a comparatively minor phylum when only the number
of recent species is considered. Nevertheless, they are present in all of the
world’s oceans within all depth zones and the approximately 12.000 fossils
species represent a rich source of paleontological information (Logan 2007).

10 Introduction
Nervous system
Microanatomical features related to the nervous system and the musculature of
brachiopod larvae are virtually unknown. The literature on the nervous system
of adult brachiopods boils down to descriptions by two authors on four species,
Gryphus vitreus, Novocrania anomala, Discinisca lamellosa and Lingula anatina
(van Bemmelen 1883; Blochmann 1892a, 1892b). Subsequent reviews of the
same data are available from several authors (Helmcke 1939; Hyman 1959;
Bullock and Horridge 1965a; Williams et al. 1997). In the rhynchonelliform
brachiopod Gryphus vitreus the main body of nervous tissue is found around
the esophagus and nerves emanate laterally from two ganglia, one subenteric
ventral of the esophagus and one supraenteric dorsal of the esophagus
(Rudwick 1970). The nervous system of brachiopod larvae or juveniles is
only known for the linguliform Lingula anatina and Glottidia sp. and consists
of a ventral lophophore system innervating the ciliary bands and a dorsal
lophophore system innervating the body musculature (Hay-Schmidt 1992,
2000). In order to fill the gap of knowledge concerning the brachiopod nervous
system in rhynchonelliform and craniiform brachiopods, this study investigates
the larval and juvenile neuroanatomy of Novocrania anomala (Craniiformea)
and Terebratalia transversa (Rhynchonelliformea).
Muscular system
Adult brachiopods possess two main forms of muscular tissue. These are either
bundles of muscle fibers that control the movement of the valves or myoepithelia
in the lophophore (Williams et al. 1997). The muscles may be smooth, cross
striated, or obliquely striated (Reed and Cloney 1977). Adult rhynchonelliform
brachiopods comprise a pair of adductors, a pair of diductors, and a dorsal
and a ventral pair of adjustor muscles that extend between the pedicle and the
valves, moving the entire shell relative to the pedicle (Richardson and Watson
1975). The adult craniiform Novocrania anomala comprises a pair of posterior
as well as anterior adductors, a pair of oblique internal, and a pair of oblique
lateral muscles (Bulman 1939). The muscular system of brachiopods and their
larvae has been described by several authors (Hancock 1858; Kowalevski 1883;
Blochmann 1892b; Helmcke 1939; Rudwick 1961; Reed and Cloney 1977), but
no studies are available that use the benefit of up-to-date techniques such as
immunocytochemistry in combination with confocal laserscanning microscopy
and 3D reconstruction software in order to visualize in detail the more cryptic
muscle sets of larval and adult brachiopods. Investigation of myogenesis was
carried out in the course of the present PhD study in order to obtain a clearer
picture of the entire brachiopod muscular bauplan as well as the dynamics of

Introduction
11
muscular remodeling during metamorphosis using the following species: Joania
cordata (previously Argyrotheca cordata), Argyrotheca cistellula, Novocrania
anomala, and Terebratalia transversa.
Gene expression
Data on the molecular processes that regulate animal development have
greatly expanded within recent years (Carroll 2005). The investigation of gene
families that encode signaling molecules with roles in the control of cell fate
specification, proliferation, movement, and segment polarity has considerably
improved our understanding of metazoan ontogeny (Davidson and Levine
2008). So far, only few sequences of developmental genes have been
identified in brachiopods, such as members of the Wnt gene family (Holland
et al. 1991) and Hox genes (de Rosa et al. 1999), but nothing has so far been
published on the expression of these genes during ontogeny. This might not
be too surprising, since marine animals as little accessible as brachiopods are
unlikely to be favored as candidate model organisms for this kind of studies
(Sommer 2009). However, since the bauplan of some brachiopods has not
changed significantly since the Early Cambrian, gene expression data from this
phylum are very interesting because they may shed light on gene functions in
the brachiopod ancestor. This information might contribute to understand the
evolution of early bilaterian animals. In this study, the expression patterns of
the developmental homeobox genes Not and Cdx were investigated in larvae
of the rhynchonelliform brachiopod Terebratalia transversa. This was done in
order to reveal the functions of these genes in Brachiopoda and to assess their
ancestral function in animal development.
Not is a homeobox gene and representatives of its family play an important role
during notochord formation in vertebrates (Abdelkhalek et al. 2004). Its role in
invertebrate development is not well known (Martinelli and Spring 2004). Cdx
is a homeobox gene that is expressed in posterior tissues of almost all phyla
investigated so far (Hejnol and Martindale 2008). In addition to the posterior
tissues it was found to be expressed in mesoderm, gut, brain, and the central
nervous system of mice, lancelets, and annelids, as well as in the gut of
Drosophila and the mesoderm of Artemia (Macdonald and Struhl 1986; Duprey
et al. 1988; Brooke et al. 1998; Copf et al. 2004; Fröbius and Seaver 2006). The
gene expression patterns presented in this thesis are the first of their kind for
the phylum Brachiopoda.

12 Material and methods
Material and methods
Immunocytochemistry and phalloidin labeling
A range of morphological and molecular methods were applied to representative
species of two main groups of Brachiopoda: Rhynchonelliformea and
Craniiformea. The musculature was investigated by use of fluorescent
conjugated phalloidin. Phalloidin is a toxin found in the mushroom Amanita
phalloides and it binds irreversibly to F-actin.
The antibodies applied to stain the nervous system bind specifically to neurotransmitters
such as serotonin (5-Hydroxytryptamine [5 HT]), neuropeptides
such as FMRFamide, or tubulins such as α-tubulin.
An overview of the species investigated, the methods, and the antibodies
applied is given in Table 1.
Labeling of Pax3/7 proteins
Arthropods and annelids generate new body segments from a posterior growth
zone (Anderson 1973; Meier 1984; Scholtz and Dohle 1996). It has been
proposed that the situation in Brachiopoda is comparable to the segmented
Annelida (Morse 1871). The larval lobes in rhynchonelliform brachiopods
suggest a segmented body plan and a segmented worm like ancestor of
Brachiopoda (Morse 1873). In order to investigate if the rhynchonelliform
brachiopod larvae of Terebratalia transversa show remnants of segmentation
from a potentially segmented ancestor, the larvae were stained with antibodies
that bind specifically on proteins of the Pax3/7 gene family.
The antibodies DP311 and DP312 detect domains of the Pax 3/7 and non-Pax3/7
proteins in Drosophila and Schistocerca (grasshopper) embryos (Davis et al.
2005). The monoclonal antibodies were raised in mouse and made available
by Michalis Averof (Institute of Molecular Biology & Biotechnology, Greece).
DP311 stains the following proteins in Drosophila: paired (prd), gooseberry
(gsb), gooseberry-neuro (gsbn), aristaless, homeobrain, and repo. DP312
stains prd, gsb, gsbn and Rx.
Larvae and juveniles of Terebratalia transversa were collected and fixed as
described in Chapter II. The primary antibodies were used in a concentration of
1:30 and the staining was applied as described in Chapters II and III. The stained
specimens were analyzed with a Leica DM RXE 6 TL fluorescence microscope
equipped with a TCS SP2 AOBS laserscanning device (Leica Microsystems,
Wetzlar, Germany).

Material and methods
13
Detection of proliferating cells with BrdU (5-bromo-2-deoxyuridine)
staining
BrdU labeling was carried out, in order to identify possible growth zones in
rhynchonelliform brachiopod larvae. BrdU is incorporated into the DNA of
proliferating cells during the S-phase of the cell cycle. Staining of BrdU thus
allows for visualization of dividing cells and their progenies. Larvae of Terebratalia
transversa of the following developmental stages: 6, 11, 24, 35, 48, 60, and 96
hours after fertilization (hpf) were incubated in 0.1mM BrdU (Sigma-Aldrich,
St. Louis, MO, USA) in seawater at 11.5ºC for 6 – 48h. In another experiment
larvae were cultured in 10mM BrdU in seawater for 30 min and subsequently
the larvae were cultured in BrdU free seawater (pulse-chase experiment). After
the treatment with BrdU the larvae were fixed in 4% paraformaldehyde in PBS
for 1 hour at room temperature and then treated for 10 min at 37ºC in 0.01mg/
ml proteinase K in PBS. After that they were kept for 10 min in 0.1N HCl on
ice, 1 hour at 37ºC in 2N HCl, 1 hour in PBS with three changes, and 15 min in
PBT (PBS with Tween 20). Then, the larvae were incubated in 1:500 mouseanti-BrdU
antibody in PBT over night at 4 ºC, washed for 1 hour in PBS with
three changes, 1 hour in 1:200 diluted TRITC, and finally 1 hour in PBS with
three changes. Stained larvae were mounted in glycerol and analyzed with a
Leica DM RXE 6 TL fluorescence microscope equipped with a TCS SP2 AOBS
laserscanning device (Leica Microsystems, Wetzlar, Germany).
Gene expression analyses
The expression of developmental genes was studied by whole mount in
situ hybridization (WMISH). Thereby, target mRNA is visualized with a
complementary RNA probe which contains DIG labelled uridine (Digoxigenin-
11-uridine-5’-triphosphate). The digoxigenin is subsequently stained with a
Anti-DIG-AP, fab fragments antibody that contains alkaline phosphatase (AP)
which in turn is made visible by a reaction with BCIP (5-Bromo-4-chloro-3-
indolyl phosphate) and NBT (nitro blue tetrazolium chloride). In this reaction
BCIP is dephosphorylated by AP and dimerizes to leucoindigo. This dimer is
then oxidized by NBT to an insoluable dark blue 5,5’-dibromo-4,4’ precipitate
(Trinh et al. 2007). The precipitate is visible in daylight conditions and also
reflects laser light which allows the use of this technique in combination with a
confocal laserscanning microscope (Jekely and Arendt 2007).
There are several WMISH protocols available which usually have to be adapted
to the organism they are intended for. Protocols developed for several species
were tested in this study, namely one for the sea urchin Strongylocentrotus

14 Material and methods
purpuratus, the cnidarian Nematostella vectensis, and the polychaete Platynereis
dumerilii, respectively (Arendt et al. 2001; Long and Rebagliati 2002; Martindale
et al. 2004; Venuti et al. 2004). The N. vectensis protocol was found to be the
best of the tested protocols for the brachiopod Terebratalia transversa and was
used accordingly to investigate the expression patterns of TtrNot and TtrCdx
(Chapter IV).
Illustrations
Illustrations were done with Photoshop CS3 and Illustrator CS3 software
(Adobe, San Jose, CA, USA).


15
Table 1. List of species investigated, methods applied, and antibodies used. (+) indicates positive
results, (-) indicates that no clear signal could be obtained, 5HT – stains nervous tissue, ad –
adult, BrdU – 5-bromo-2-deoxyuridine (stains proliferating cells), CLSM – confocal laserscanning
microscopy, DAPI – (stains nucleic acids), engrailed – labels segment boundaries in Drosophila,
Immunostar – producer of antibodies, juv – juvenile, Pax 3/7 – labels segment boundaries in
Drosophila, Phalloidin – stains F-actin, Sigma – Sigma-Aldrich, producer of antibodies, Tubulin
– stains cilia and nervous tissue, WMISH – whole mount in situ hybridization.
Clade
Species
Stages
investigated
larval juv ad
Method
applied
Antibodies
applied
(signal + or -)
Chapter
Rhynchonelliformea
Joania
(Argyrotheca)
cordata
+ - + CLSM
5 HT (Sigma) (-)
DAPI (+)
FMRF (-)
Phalloidin (+)
Tubulin (+)
II
Rhynchonelliformea
5 HT (Sigma) (-)
Argyrotheca
+ - + CLSM
FMRF (-)
II
cistellula
Phalloidin (+)
5 HT
(Immunostar) (+)
BrdU (+)
Cdx (+)
Rhynchonelliformea
Terebratalia
transversa
+ + -
CLSM
WMISH
DAPI (+)
Engrailed (-)
FMRF (-)
I, II, IV
Not (+)
Pax 3/7 (+)
Phalloidin (+)
Tubulin (+)
Phalloidin (+)
Craniiformea
5 HT
Novocrania
+ + - CLSM
(Immunostar) (+)
III
anomala
Tubulin (+)
FMRF (-)

16 Results and discussion
Results and discussion
Larval development
Terebratalia transversa, a representative of Rhynchonelliformea
Larval development of Terebratalia transversa and regional specification during
embryogenesis has been described previously (Freeman 1993). My results
are congruent with these data. The oocyte (Fig. 1A) divides approximately 2
hours after fertilization (hpf) at a water temperature of 11.5 °C and two polar
bodies are formed (Fig. 1B). Cleavage is radial and the first two cleavages are
holoblastic (Fig. 1B, C). The early blastula is composed of rounded cells (Fig.
1D) and gastrulation occurs approximately at 19 hpf (Fig. 1E). In the gastrula,
the wall of the archenteron forms contact with the cells of the ectoderm, i.e.,
the blastocoel virtually disappears (Fig. 1F). Later in development the gastrula
elongates and the blastopore becomes slit-like elongated (Fig. 1G). The three
larval lobes start to form as the embryo elongates further and an apical tuft
appears, which is lost later in development (Fig 1H, I). At this stage the larvae
become positively phototactic and usually swim in the upper part of the water
column. At approximately 75 hpf the larvae are almost fully developed and the
apical, mantle, and pedicle lobe are formed. Only the setae continue to grow
at this point of development. The fully developed larvae eventually become
negatively phototactic. Then, they swim towards the bottom of the culture dish
and repeatedly touch the surface with their apical lobe, probably in order to test
if the substrate is suitable for metamorphosis. Larvae settle and metamorphose
between 120 and 300 hpf. The juveniles still retain the larval setae and the
lophophore starts to form after settlement (Fig. 1J). Metamorphosis appears to
be catastrophic since all tissues seem to be reformed during metamorphosis
(Stricker and Reed 1985a, 1985b).

Results and discussion
17
A B C
D
0 2 3 10
at
E F ec G
H
AL
AL
en
* *
*
18 24
30 36
I
se
AL
ML
PL
J
se
se
se
Lo
75 hpf Pe 360 hpm
se
Figure 1. Developmental stages of Terebratalia transversa at a water temperature of 11.5 °C.
Numbers indicate the age in hours after fertilization (hpf) for all stages except of J where it is
hours after the onset of metamorphosis (hpm). Size of all stages is around 120 µm in diameter,
except for J where it is around 200 µm. Anterior is oriented upwards and cilia are omitted for
clarity. (A) unfertilized oocyte (black) with an egg shell (grey). (B) Lateral view of two cell stage
with two polar bodies and the egg shell (grey). (C) Apical view of a four cell stage. (D) Sagittal
section through an early blastula. (E) Sagittal section through a late blastula at the onset of
gastrulation. (F) Gastrula with ectoderm (ec), endoderm (en), and blastopore (asterisk). The
gastrula starts to swim at this point of development. (G) Elongated late gastrula with slit-like
blastopore (asterisk) and first signs of a distinguished apical lobe (AL). (H) Larva with further
developed lobes, almost closed blastopore (asterisk), and apical tuft (at). (I) Fully established
larva with apical lobe (AL), mantle lobe (ML), and pedicle lobe (PL). Four sets of setae bundles
(se, only two visible) originate from the mantle lobe. (J) Juvenile with lophophore (Lo), and
pedicle (Pe). The remaining larval setae (se) extend beyond the two valves.
se
Novocrania anomala, a representative of Craniiformea
Development of Novocrania anomala and regional specification during
embryogenesis has been described previously (Nielsen 1991; Freeman 2000).
My results are congruent with these data. However, the two authors disagree
about the development of the coelom and the formation of the mesoderm.
According to Nielsen, the sheet of cells that invaginates during gastrulation is
composed of two cell populations, endoderm and mesoderm, whereas Freeman
states that the mesoderm is formed by individual cells which immigrate from the
endodermal cell layer after invagination has been completed (Nielsen 1991;
Freeman 2000). Nielsen describes the coelom as consisting of an anterior
coelomic pouch in the apical lobe and three pairs of coelomic cavities in the

18 Results and discussion
posterior lobe of the larva, whereas Freeman denies the existence of larval
coelomic structures and states that the coelom develops after the larvae have
undergone metamorphosis (Nielsen 1991; Freeman 2000). The methods used
here do not allow a conclusive statement concerning coelom and mesoderm
formation in larvae of N. anomala, there is more work needed to resolve the
controversies on an ultrastructural level.
Cleavage is radial and the first two divisions are holoblastic (Fig. 2B). The gastrula
is first spherical and invagination takes place at the vegetal pole of the larva.
The archenteron cells come to lie opposite of the ectoderm. Subsequently, the
blastocoel disappears completely (Fig. 2C). Later in development the gastrula
elongates and the blastopore comes to lie at the postero-ventral side of the
swimming larva (Fig. 2D). The elongated gastrula subsequently differentiates
into two larval lobes, an apical lobe and a posterior lobe (Fig. 2E, F). Larval
development completes with the growth of three pairs of dorsal setal bundles
on the posterior lobe (Fig. 2G). Prior to settlement, the larva swims along the
bottom of the culture dish, probably in order to test if the substrate is suitable
for settlement. In contrast to the descriptions by Nielsen (1991), the larvae do
not curl before metamorphosis. Although curled larvae are found in the culture
dishes, these seem to be unable to metamorphose. What causes the curling
is unclear, however it can clearly be seen in the musculature of settled larvae
that the remaining larval muscles are elongated and relaxed in contrast to the
contracted musculature of curled larvae (Fig. 3A, B, and Chapter III).
At a water temperature of 14 °C, metamorphosis takes place around six to ten
days after fertilization (dpf). During metamorphosis the larva attaches to the
substrate, secretes the shell, and retains its larval lobes, which are subsequently
transformed and form the lophophore and other adult organs (Figs. 2H, 3B,
C).

Results and discussion
19
A B C D
0 4
25 * 32
E F G H
AL
se
se
se
se
AL
PL
40 72 105
se
AL
PL
se
se
ec
en
*
se
se
se
se
se
s
AL
PL
ec
en
se
se
200
Figure 2. Developmental stages of Novocrania anomala at a water temperature of 14 °C.
Numbers indicate the age in hours after fertilization (hpf) for all stages except for H where it is
hours after the onset of metamorphosis (hpm). Size of all stages is around 130 µm in diameter.
Anterior is oriented upwards. Cilia have been omitted for clarity (A) Unfertilized oocyte (black)
with egg shell (grey). (B) Apical view of a four cell stage with the egg shell at 4hpf. (C) Frontal
view of a gastrula with blastopore (asterisk), ectoderm (ec), and endoderm (en). The gastrula
starts to swim at this point of development. (D) Lateral view of an elongated gastrula with
ectoderm (ec) and endoderm (en). The blastopore (asterisk) is situated on the posterior end
of the gastrula. (E) Dorsal view of an elongated gastrula with almost distinct apical lobe (AL).
(F) Ventral view of an early two-lobed larva with apical lobe (AL) and posterior lobe (PL). The
blastopore is closed and larval setae (se) start to grow on the posterior side. (G) Dorsal view of
a fully developed larva with apical lobe (AL), posterior lobe (PL), and three pairs of dorsal setae
bundles (se). (H) Ventral view of a juvenile after metamorphosis. The larval apical lobe (AL) and
pedicle lobe (PL) are still visible. The juvenile shell (s) is formed on the dorsal side with larval
setae (se) extending from it.
Figure 3. Metamorphosis of Novocrania anomala. Scale bars equal 50 µm, anterior is up. A and
B are overlays of confocal maximum projections of phalloidin stainings and light micrographs.
C is a light micrograph of a live specimen. (A) Ventral view of a curled larva with contracted
musculature (empty arrow), apical lobe (AL), and posterior lobe (PL). (B) Musculature of a
settled juvenile with remaining elongated larval musculature (empty arrowheads), juvenile
anterior adductor muscles (aad), larval setae pouch muscles (arrows), larval anterior lobe (AL),
posterior lobe (PL), and juvenile shell (s). (C) Dorsal view of a settled juvenile with remaining
larval setae (se), shell (s), posterior lobe (PL), and apical lobe (AL) which has started to form
the lophophore (Lo).

20 Results and discussion
Myogenesis
Results of larval myogenesis and adult myoanatomy are presented in Chapters
II and III.
Actin and myosin are molecules present in all metazoans including basal groups
such as sponges and Trichoplax (Thiemann and Ruthmann 1989; Kanzawa et
al. 1995). It has been proposed that the basal pattern of musculature in the
bilaterian ancestor was a grid of outer circular and inner longitudinal musculature,
the Hautmuskelschlauch (HMS), which has in some taxa been modified in
combination with the evolution of hard exoskeletons (Schmidt-Rhaesa 2007a).
Brachiopods have discrete bundles of muscle fibers that control the movement
of the valves and the tentacles. Brachiopods have further myoepithelia which
are found on the inner side of coelomic epithelia, in the parietal bands, in mantle
lobes, and in the lophophore (Williams et al. 1997). Additionally, I could show
that adults of the species Joania cordata, Argyrotheca cistellula, Novocrania
anomala, and Terebratalia transversa contain discrete bundles of mantle
retractor muscles (Chapters II, III), a character that is probably present in all
brachiopods.
The larval musculature is similar among the rhynchonelliform brachiopods
investigated herein (Chapter II). Remnants of a HMS could not be distinguished.
Accordingly, if the ancestor of Brachiopoda had a HMS, it was lost during the
evolution of this phylum. Interestingly, the larval musculature of the craniiform
brachiopod Novocrania anomala is very different from the musculature of
the investigated rhynchonelliform brachiopod larvae (Chapter III). This hints
towards an early split in the evolution of these two groups. This is confirmed by
the fossil record, which estimates the split between the rhynchonelliform and
craniiform clade to have taken place before the Ordovician 485 million years
ago (Freeman and Lundelius 2005).
Neurogenesis with special focus on the apical organ of
lophotrochozoan larvae
Results on neurogenesis in brachiopod larvae and juveniles are presented in
Chapters III and IV.
Adult rhynchonelliform brachiopods have a nervous system which is concentrated
around the esophagus and comprises two ganglia, one dorsal and one ventral of
the esophagus, as well as circumenteric nerves that innervate the lophophore,
ventral mantle nerves, and dorsal mantle nerves (van Bemmelen 1883; Bullock
and Horridge 1965a). The nervous system of adult Novocrania anomala lacks
the dorsal ganglion. The circumenteric nerves emanate laterally from the ventral

Results and discussion
21
ganglion and form a ring around the esophagus. Additional lateral and brachial
nerves emanate from the ventral ganglion (Blochmann 1892b). The nervous
system of the lecithotrophic rhynchonelliform brachiopod larvae of Terebratalia
transversa comprises two sets of four serotonergic flask-shaped cells in the
apical organ that are connected by neurites to a larval neuropil in the apical lobe
(Chapter IV). The nervous system of the lecithotrophic craniiform brachiopod
larvae of Novocrania anomala comprises four centrally positioned serotonergic
flask-shaped cells in the apical organ connected to two ventral nerve cords that
extend ventrolaterally along the body (Chapter III). Linguliform planktotrophic
brachiopod juveniles of Lingula anatina and Glottidia sp. possess a nervous
system comprising an apical ganglion as well as dorsal and ventral lophophore
nerves (Hay-Schmidt 1992). The apical ganglion of Glottidia sp. contains
numerous serotonergic cells that are associated with two serotonergic tracts
which project into the ciliary band (Hay-Schmidt 2000). This system is probably
not homologous to the apical organs found in T. transversa and N. anomala,
since there are numerous serotonergic cells in Glottidia sp. and none of these
cells are flask-shaped.
The evolution of nervous systems has been reviewed by several authors
(Bullock and Horridge 1965b; Holland 2003; Schmidt-Rhaesa 2007b; Arendt
et al. 2008; Benito-Gutiérrez and Arendt 2009; Wanninger 2009; Harzsch and
Wanninger 2010). All eumetazoans are able to transmit information between
cells. Sponges use electric signals albeit lacking neurons (Leys et al. 1999),
cnidarians have a nerve net with electrical and chemical synapses (Anderson
and Trapido-Rosenthal 2009), and bilaterians have a nervous system that often
comprises some sort of “brain” and nerve cords or neurite bundles (Rieger et al.
2010). The last common ancestor of cnidarians and bilaterians most likely had
a nerve net which developed under the control of anteroposterior patterning
genes (Westfall 1996; Westfall and Elliott 2002; Watanabe et al. 2009). The
question whether the ancestor of Protostomia and Deuterostomia had a diffuse
nervous system or a centralized nervous system is still hotly debated and a
final statement can not yet be made (Younossi-Hartenstein et al. 1997; Arendt
and Nübler-Jung 1999; Holland 2003; Lowe et al. 2003; 2006; Telford 2007;
De Robertis 2008; Reichert 2009; Harzsch and Wanninger 2010). Recent
studies showed that larval Entoprocta and adult Mollusca show a tetraneurous
condition consisting of one pair of ventral and on pair of more dorsally positioned
lateral nerve cords. In addition, the creeping-type entoproct larva and the
polyplacophoran larvae exhibit a complex apical organ consisting of around
eight centrally positioned serotonergic flask-shaped cells which are surrounded
by several peripheral cells. (Wanninger et al. 2007; Fuchs and Wanninger 2008;

22 Results and discussion
Wanninger 2008; 2009). In Nemertea, the lecithotrophic, non-pilidium like larva
of Quasitetrastemma stimpsoni shows a pair of serotonergic flask-shaped cells
in the apical organ plus a pair of subapical cells and two posterior neurons
that are located ventrolaterally (Chernyshev and Magarlamov 2010). Annelid
larvae show a serotonergic apical organ comprising up to four cells. The apical
organ is associated with the prototrochal nerve ring which in turn is connected
to two ventral nerve cords (Voronezhskaya et al. 2003; McDougall et al. 2006;
Brinkmann and Wanninger 2008). The apical organ of ectoproct cyphonautes
larvae comprises two pairs of serotonergic cell bodies from which lateral nerves
project towards the corona (Hay-Schmidt 2000; Gruhl 2009). One of the two cell
clusters in the apical organ contains flask-shaped cells (Nielsen and Worsaae
2010). In the apical organ of the ectoproct coronate larva of Bugula neritina
two flask-shaped serotonergic cells are present (Pires and Woollacott 1997;
Shimizu et al. 2000). In the actinotroch larva of Phoronida, the apical organ
contains numerous serotonergic cells, but these are probably not flask-shaped
(Santagata 2002; Santagata and Zimmer 2002; Wanninger 2008).
Taken together, the data that have recently become available on lophotrochozoan
larval neuroanatomy suggest that an apical organ comprising serotonergic
flask-shaped cells was present in larvae of the last common lophotrochozoan
ancestor (Wanninger 2008). Accordingly, an apical organ containing such cells
might be a morphological apomorphy of Lophotrochozoa.
Distribution of Pax3/7 proteins in larvae of Terebratalia transversa
A sister group relationship of Brachiopoda with Annelida has been hypothesized
based on molecular data as well as on paleontological data and is supported by
the notion that annelids and brachiopods share similarities in the ultrastructure
of their setae (Gustus and Cloney 1972; Orrhage 1973; Field et al. 1988; Lake
1990; Conway Morris and Peel 1995; Lüter 2000b). Several developmental
genes that are involved in the establishment of segments and segmentation in
animals have been characterized, some of which belong to the Pax3/7 group.
Pax3 and Pax7 genes probably arose by duplication from unique ancestral Pax3/7
genes and have similarities in their protein sequence and expression (Hayashi et
al. 2010). Pax3/7 genes are also known as Pax group III genes and include the
pair-rule gene paired (prd), the segment polarity genes gooseberry (gsb), and
gooseberry-neuro (gsbn), a gene that is expressed in the developing nervous
system and, together with engrailed, establishes the posterior commissures in
the fruit fly Drosophila melanogaster (Noll 1993; Colomb et al. 2008). Together
with their vertebrate homologs (Pax-3 and Pax-7) the Pax3/7 group forms one

Results and discussion
23
of four classically defined subgroups of the Pax family transcription factors
(Balczarek et al. 1997). Pax3/7 shares its expression among distantly related
insects and shows several patterns including pair-rule, segment polarity,
and neural patterning (Davis et al. 2005). In crustaceans Pax3/7 genes are
expressed in iterated stripes (Davis et al. 2005). In myriapods and chelicerates
Pax3/7 gene expression exhibits iterated stripes that form early in the posteriormost
part of the germ band (Davis et al. 2005). In the tardigrade Hypsibius
dujardini, the Pax3/7 proteins localize in a segmentally iterated pattern in the
ectoderm, after establishment of endomesoderm segmentation, but before the
visible segmentation of the ectoderm (Gabriel and Goldstein 2007). Pax3/7 is
also localized within the developing head region of the tardigrade embryo, but
no pair-rule pattern is visible during any stage of embryogenesis (Gabriel and
Goldstein 2007). Tardigrades, together with arthropods and onychophorans
belong to Panarthropoda (Halanych 2004).The expression pattern of Pax3/7 in
H. dujardini suggests that the pair-rule function of Pax3/7 may have arisen near
the base of Arthropoda.
In the annelid Platynereis dumerilii Pax3/7 proteins are found in the peripheral
nervous system (Kerner et al. 2009). In larvae of the brachiopod Terebratalia
transversa DP311 and DP312 show identical staining patterns. Pax 3/7 starts
to be present in four cells of the apical lobe in the late elongated gastrula (Fig.
4B). The cells containing Pax3/7 products are later distributed in a ring on the
apical lobe of early three-lobed larvae without setae (Fig. 4C). Fully established
larvae show a loose distribution of cells that contain Pax3/7 products in their
apical lobe (Fig. 4D, E). In juveniles Pax3/7 containing cells are mainly found
in the growing lophophore (Fig. 4F). The presence of Pax3/7 gene products
in the apical lobe indicates a function of those genes during neurogenesis in
T. transversa. However, further experiments are necessary in order to assess
whether the staining specifically shows Pax3/7 protein products, since the
antibodies used were developed against the Pax3/7 sequences of Drosophila
melanogaster. Ideally, cloning of the sequences of the Pax3/7 homologs of
Terebratalia transversa should be carried out, followed by mapping of the
epitopes of DP311 and DP312 on peptide arrays with the known peptide
sequences of T. transversa and other metazoans (Harlow and Lane 1999). The
final proof would then be in situ hybridizations with the specific corresponding
probes. In addition, a double staining with serotonin would be necessary in
order to prove that the cells containing Pax3/7 gene products are co-localized
with the nervous system.

24 Results and discussion
Growth patterns of Terebratalia transversa
Figure 4. Staining of
Pax3/7 proteins with
DP311. Overlay of confocal
maximum projections on
light micrographs. Anterior
is up and scale bars equal
50 µm. (A) Gastrula with
blastopore (asterisk) and
no signal. (B) Late gastrula
with slit-like blastopore
(asterisk). Pax3/7 proteins
are stained in four cells
in the future apical lobe
(al). (C) Early three-lobed
larva with almost closed
blastopore (asterisk).
Pax3/7 proteins are present
in several cells of the apical
lobe (al) and distributed in
a ring around it. No signal
is found in the mantle lobe
(ml) and in the pedicle lobe
(pl) (D) Lateral view of a
larva with apical lobe (al),
mantle lobe (ml), pedicle
lobe (pl), and setae (se).
Pax3/7 protein containing
cells are concentrated in
the dorsal part of the apical
lobe. (E) Fully established
larva with apical lobe (al),
mantle lobe (ml), pedicle
lobe (pl), and setae (se).
Cells with Pax3/7 proteins
are loosely distributed in
the apical lobe. (F) Juvenile
after metamorphosis.
Pax3/7 proteins are loosely
expressed in the developing
lophophore (Lo) of the
juvenile. The dorsal shell (s)
of this specimen is slightly
shifted upwards relative
to its natural position, and
larval setae (se) extend out
of the valves
In order to identify possible growth zones in brachiopod larvae, proliferating
cells in Terebratalia transversa were labeled with 5-bromo-2-deoxyuridine
(BrdU). Dividing cells are equally distributed in the blastula stage (Fig. 5A),
the gastrula (Fig. 5B), and the elongated gastrula (Fig. 5C). In the elongated
gastrula, cells divide mostly in the center of the larva and form the mantle lobe,
which is marked by a ring of dividing cells (Fig. 5D). Thereafter, dividing cells
are again equally distributed throughout the larva (Fig. 5E). Larvae competent
for metamorphosis also show an equal distribution of proliferating cells after a
pulse-chase experiment, which once again indicates that there are no distinct
growth zones that form most parts of the larval body, but that dividing cells are
found throughout the developing specimen (Fig. 5F). The BrdU data suggest
that from the viewpoint of proliferation zones, there are no similarities between

Results and discussion
25
Figure 5. Pattern of
BrdU staining in larvae of
Terebratalia transversa.
Overlay of confocal
maximum projections and
light micrographs. Scale
bars equal 50 µm. All stages
show an equal distribution
of proliferating cells,
there are thus no distinct
growth zones identifiable.
(A) Blastula. (B) Early
gastrula with blastopore
(asterisk). (C) Late slightly
elongated gastrula with
blastopore (asterisk). (D)
Early three lobed stage
with the developing apical
lobe (al), mantle lobe (ml),
and pedicle lobe (pe). (E)
Three lobed stage with
apical lobe (al), mantle
lobe (ml), and pedicle lobe
(pl). This stage is at the
onset of setae formation.
(F) Fully developed threelobed
stage with apical
lobe (al), mantle lobe (ml),
and pedicle lobe (pl).
the development of Annelida and Brachiopoda. For annelids, it has been shown
that, although the post-metamorphic segments originate from a posterior growth
zone, the precise location of the growth zone can vary (Seaver et al. 2005;
Brinkmann and Wanninger 2010). However, the rhynchonelliform brachiopods
are regarded derived amongst brachiopod subgroups (Carlson 1995). The
distribution of proliferating cells in Terebratalia transversa can therefore not
completely rule out the possibility that the brachiopod ancestor had a growth
zone. Similar experiments in linguliform and craniiform brachiopods are needed
in order to further assess this issue.
The Annelida-Brachiopoda sister group hypothesis based on the ultrastructure
of the setae has been questioned by Lüter who showed that there is a difference
in the ultrastructure of larval and adult setae in the brachiopods Lingula anatina,
Notosaria nigricans, and Calloria inconspicua, suggesting a convergent
evolution of setae in Annelida and Brachiopoda (Lüter 2000b). An additional

26 Results and discussion
argument against segmentation in brachiopod larvae is that the segmented
appearance with three larval lobes is not recognizable by the inner bauplan
on the ultrastructural level (Lüter 2000a). This has been shown for Notosaria
nigricans and Calloria inconspicua. In these species, a single coelomic anlage
forms one compartment with all mesodermally derived cells separated only by
cellular membranes. Thus, there is only one mesoderm compartment in these
larvae, which encloses one coelomic cavity (Lüter 2000a). In the segmented
Annelida the coelom forms one pair of coelomic cavities in each segment
(Anderson 1973).
Not and Cdx expression analyses
Results of gene expression patterns of the homeobox genes TtrNot and TtrCdx
are presented in Chapter IV.
In Terebratalia transversa, the ortholog of the homeobox gene Not, TtrNot, is
expressed in the ectoderm from the beginning of gastrulation until completion
of larval development, which is marked by a three-lobed body with larval setae.
Expression starts at gastrulation in two areas lateral to the blastopore and
subsequently extends over the animal pole of the gastrula. With elongation of
the gastrula, expression at the animal pole narrows to a small band, whereas
the areas lateral to the blastopore shift slightly towards the future anterior region
of the larva. Upon formation of the three larval body lobes, TtrNot expressing
cells are present only in the posterior part of the apical lobe. Expression ceases
entirely at the onset of larval setae formation. TtrNot expression is absent in
unfertilized eggs, in embryos prior to gastrulation, and in settled individuals
during and after metamorphosis. Comparison to the expression patterns of Not
genes in other metazoan phyla suggests an ancestral role in gastrulation, germ
layer (ectoderm) specification, and neural patterning, with co-opted functions in
notochord formation in chordates and left/right determination in ambulacrarians
and vertebrates (Chapter IV).
In Terebratalia transversa the ParaHox gene TtrCdx is expressed on the
posterior side of the blastopore and its expression stays in this region until the
three-lobed larva is fully formed. The expression of TtrCdx suggests a function
of this gene during gastrulation and ectoderm patterning in Brachiopoda. The
pattern of Cdx in other metazoans ranges from expression in the mesoderm,
gut, brain, central nervous system to posterior tissues (Fröbius and Seaver
2006). The basal function of Cdx is probably in patterning of posterior tissues.

General conclusions and perspectives for future research
27
General conclusions and perspectives for future research
The results presented herein are the first developmental gene expression
studies in Brachiopoda, as well as the first detailed comparative description of
myogenesis and neurogenesis in brachiopod larvae based on antibody staining,
confocal laserscanning microscopy, and 3D reconstruction software. This study
shows that microanatomical data can yield new insights into the evolution and
development of lesser known metazoan phyla such as Brachiopoda. It provides
the first evidence of an apical organ in brachiopod larvae that comprises
serotonergic flask-shaped cells, similar to those found in ectoprocts and
spiralians. This result strongly suggests that such an apical organ constitutes a
morphological apomorphy of Lophotrochozoa.
Gene expression analyses of TtrNot imply an ancestral role of this gene in
gastrulation and ectoderm specification in Brachiopoda. The function of Not in
notochord formation in chordates and left/right determination in ambulacrarians
and vertebrates might thus be co-opted in these deuterostome clades. Analysis
of the TtrCdx gene expression suggests an ancestral role in gastrulation and the
formation of posterior tissues in Brachiopoda as well as in Bilateria in general.
Further studies should extend the database of brachiopod morphogenesis
and gene expression patterns to more organ systems as well as to the third
brachiopod subtaxon, Linguliformea. This would allow for a full representation
of the phylum Brachiopoda with its three clades Craniiformea, Linguliformea,
and Rhynchonelliformea and should allow significant inferences concerning
gene function and organ system evolution within this lophophorate phylum.
Such data would allow insights into the evolution of organ systems, and body
plans in Brachiopoda. Additionally, investigation of gene expression patterns in
Brachiopoda is needed in order to compare the function of genes, co-option,
and ancestral gene functions among Brachiopoda and other animal phyla. An
expressed sequence tags or genome-based approach would be the best choice
in order to obtain the sequences of the whole range of developmental genes.
Preferably, this should be done for one representative of each brachiopod clade.
Morphological and molecular data together would facilitate the reconstruction of
the evolution of organ systems in Brachiopoda once the phylogenetic position
of Brachiopoda and its sister groups has been settled.

28 References
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Chapter II
37
Chapter II
Altenburger, A. & Wanninger, A. 2009 Comparative larval
myogenesis and adult myoanatomy of the rhynchonelliform
(articulate) brachiopods Argyrotheca cordata, A. cistellula, and
Terebratalia transversa. Frontiers in Zoology 6: 1-14

38 Chapter II
Frontiers in Zoology
BioMed Central
Research
Comparative larval myogenesis and adult myoanatomy of the
rhynchonelliform (articulate) brachiopods Argyrotheca cordata, A.
cistellula, and Terebratalia transversa
Andreas Altenburger and Andreas Wanninger*
Open Access
Address: University of Copenhagen, Department of Biology, Research Group for Comparative Zoology, Universitetsparken 15, DK-2100
Copenhagen Ø, Denmark
Email: Andreas Altenburger - aaltenburger@bio.ku.dk; Andreas Wanninger* - awanninger@bio.ku.dk
* Corresponding author
Published: 3 February 2009
Frontiers in Zoology 2009, 6:3
doi:10.1186/1742-9994-6-3
This article is available from: http://www.frontiersinzoology.com/content/6/1/3
Received: 5 November 2008
Accepted: 3 February 2009
© 2009 Altenburger and Wanninger; licensee BioMed Central Ltd.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0),
which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Abstract
Background: Despite significant methodological progress, Brachiopoda remains one of the
lophotrochozoan phyla for which no recent ontogenetic data employing modern methodologies
such as fluorescence labelling and confocal microscopy are available. This is particularly astonishing
given the ongoing controversy concerning its phylogenetic position. In order to contribute new
morphogenetic data for phylogenetic and evolutionary inferences, we describe herein the ontogeny
and myoanatomy of larvae and adults of the rhynchonelliform brachiopods Argyrotheca cordata, A.
cistellula, and Terebratalia transversa using fluorescence F-actin labelling combined with confocal
laserscanning microscopy.
Results: Fully grown larvae of A. cordata and T. transversa consist of three distinct body regions,
namely an apical lobe, a mantle lobe with four bundles of setae, and a pedicle lobe. Myogenesis is
very similar in these two species. The first anlagen of the musculature develop in the pedicle lobe,
followed by setae muscles and the mantle lobe musculature. Late-stage larvae show a network of
strong pedicle muscles, central mantle muscles, longitudinal muscles running from the mantle to
the pedicle lobe, setae pouch muscles, setae muscles, a U-shaped muscle, serial mantle muscles,
and apical longitudinal as well as apical transversal muscles. Fully developed A. cistellula larvae differ
from the former species in that they have only two visible body lobes and lack setae. Nevertheless,
we found corresponding muscle systems to all muscles present in the former two species, except
for the musculature associated with the setae, in larvae of A. cistellula. With our survey of the adult
myoanatomy of A. cordata and A. cistellula and the juvenile muscular architecture of T. transversa we
confirm the presence of adductors, diductors, dorsal and ventral pedicle adjustors, mantle margin
muscles, a distinct musculature of the intestine, and striated muscle fibres in the tentacles for all
three species.
Conclusion: Our data indicate that larvae of rhynchonelliform brachiopods share a common
muscular bodyplan and are thus derived from a common ancestral larval type. Comparison of the
muscular phenotype of rhynchonelliform larvae to that of the other two lophophorate phyla,
Phoronida and Ectoprocta, does not indicate homology of individual larval muscles. This may be
due to an early evolutionary split of the ontogenetic pathways of Brachiopoda, Phoronida, and
Ectoprocta that gave rise to the morphological diversity of these phyla.
Page 1 of 14
(page number not for citation purposes)

Chapter II
39
Frontiers in Zoology 2009, 6:3
http://www.frontiersinzoology.com/content/6/1/3
Background
Brachiopoda is a small lophophorate phylum with a
prominent fossil record since the Lower Cambrium [1].
More than 12.000 fossil and approximately 380 recent
species are known to date [2,3]. The phylum is commonly
divided into three taxa, the articulate Rhynchonelliformea
and the two inarticulate clades Craniiformea and Linguliformea
[4], and has traditionally been grouped together
with Phoronida and Ectoprocta into the superphylum
Lophophorata. However, this classification has recently
been challenged by paleontological and molecular datasets.
While some analyses employing morphological data
assign Brachiopoda to Deuterostomia [e.g., [5,6]], recent
molecular data either propose sistergroup relationships to
various spiralian phyla including Mollusca, Annelida, and
Nemertea [7-11], or support the notion that Phoronida
are an ingroup of Brachiopoda [12,13].
Apart from some mainly gross morphological studies [14-
21], detailed data using modern techniques such as fluorescence
labelling and confocal laserscanning microscopy
are not yet available. This is especially true with respect to
the development of the musculature, despite the fact that
myo-anatomical features may provide useful characters
for reconstructing phylogenetic relationships [22,23].
Recently, some data on larval muscle development for the
proposed brachiopod sister groups Phoronida and Ectoprocta
have become available [24-28]. Accordingly, larval
myogenesis in Brachiopoda constitutes an important gap
of knowledge in comparative developmental studies on
Lophophorata. With the first thorough, comparative
account of brachiopod larval myogenesis provided herein
for the rhynchonelliform species Argyrotheca cordata
(Risso, 1826), Argyrotheca cistellula (Searles-Wood, 1841),
and Terebratalia transversa (Sowerby, 1846), we aim at
stimulating the discussion concerning lophophorate bodyplan
evolution, phylogeny, and development. Furthermore,
we contribute to questions concerning the
muscular ground pattern of rhynchonelliform brachiopod
larvae. We supplement our ontogenetic data with a
detailed description of the adult muscle systems of all
three species.
Results
Embryonic and larval development of Argyrotheca
cordata
Embryos and larvae of Argyrotheca cordata are brooded by
the mother animal and are released as late-stage larvae
competent to undergo metamorphosis. Accordingly, larval
development is entirely lecithotrophic. After cleavage
and gastrulation (Fig. 1A), a three-lobed larva is established,
which comprises an anterior apical lobe, a mantle
lobe in the mid-body region, and a posterior pedicle lobe
(Fig. 1B–F). In very early three-lobed stages, the blast-
Scanning development Figure 1electron of Argyrotheca micrographs cordata of the embryonic and larval
Scanning electron micrographs of the embryonic and
larval development of Argyrotheca cordata. Anterior
faces upward and scale bars equal 50 μm. (A) Early gastrula
with blastopore (arrow). (B) Ventral view of an embryo at
the onset of differentiation of the three-lobed larval bodyplan
comprising apical lobe (AL), mantle lobe (ML), and pedicle
lobe (PL). The arrowhead points to the region of the larval
apical ciliary tuft. The arrow points to the larval mouth which
corresponds to the blastopore. (C) Dorsal view of a larva
with distinct anlagen of the three body lobes. (D) Ventral
view of a specimen of the same ontogenetic stage as the one
in C with reduced larval apical ciliary tuft (arrowhead) and
with the almost closed blastopore (arrow). (E) Three-lobed
larva at the onset of setae formation (double arrowheads),
dorso-lateral view. (F) Lateral view of a fully differentiated
larva showing two of the four pairs of larval setae (double
arrowheads) and a distinct primordial hump (asterisk).
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opore is visible at the base of the apical lobe (Fig. 1B). This
larval mouth closes during subsequent larval development
(Fig. 1D).
The apical lobe is ciliated and bears, in early three lobed
stages, an apical tuft which is lost in later stages (Fig. 1B,
D). When the three lobes are fully established, four bundles
of larval setae are formed at the posterior margin of
the mantle lobe (Fig. 1E). Finally, in larvae competent to
undergo metamorphosis, the anlage of the pedicle
becomes visible as a distinct primordial hump at the posterior
pole of the pedicle lobe (Fig. 1F).
Myogenesis and adult myoanatomy of Argyrotheca
cordata
The larvae investigated were about 230–270 μm long and
210–240 μm wide. The first F-actin-positive signal is visible
as distinct spots in the area that later forms the mantle
lobe (Fig. 2A). These distinct spots are F-actin-positive
microvilli which are situated in the lower part of the setal
sacs where the setae are formed [cf. [29]]. The strong fluorescence
signal of the microvilli disappears once setae formation
is completed, due to the increasing predominance
of the larval musculature (Fig. 2D–F).
The pedicle muscles start to form in three-lobed larvae
that still lack setae (Fig. 2B). In older larvae with short
setae (corresponding to the stage shown in Fig. 1E), setae
muscles start to develop. These run from the setal pouches
in anterior direction and connect to the apical longitudinal
muscles at the border between apical and mantle lobe
(lateral setae muscles) or to the central mantle muscles
(dorsal setae muscles), respectively (Fig. 2C). The apical
longitudinal muscles extend laterally within the apical
lobe and terminate anteriorly at an apical transversal muscle
(Fig. 2C). At this stage, longitudinal muscles are also
found within the pedicle lobe. From there, they run into
the mantle lobe, where they connect to longitudinal muscles
which originate at the muscle interconnection point
at the border between apical and mantle lobe. The larval
gut rudiment is visible as a tube in the centre of the larvae
(Fig. 2C).
In fully developed larvae, setae pouch muscles are established
and interconnected by a circular mantle muscle
(Fig. 2D). From this circular mantle muscle emerge serial
mantle muscles, which are dorsolaterally closed by the
central mantle muscles. The central mantle muscles are
connected to the dorsal setae muscles and to the apical
longitudinal muscles at the border of the apical and the
mantle lobe (Fig. 2E–F). Anteroventrally, the serial mantle
muscles are enclosed by a U-shaped muscle which extends
ventrally from the pedicle muscles towards the circular
mantle muscle (Fig. 2D–F; see also additional file 1). The
primordial hump is devoid of any musculature (Fig. 2E–
F).
Adult A. cordata studied were 0.8–1.3 mm wide and 0.9–
1.4 mm long. We can confirm four pairs of muscles which
have been described previously [30]. These are one pair of
adductors and one pair of diductors, which attach to both
the dorsal and to the ventral valve. In addition, there are
two pairs of pedicle adjustors, one of which being
attached to the ventral valve and the pedicle, and one
being attached to the dorsal valve and the pedicle (Fig.
3A–B). In addition, we found a distinct musculature in the
tentacles of the lophophore and in the digestive system.
Each tentacle contains several bands of striated muscle
fibers (Fig. 3D–E), while the stomach and intestine are
each lined by numerous delicate ring muscles (Fig. 3C).
Moreover, minute muscles are distributed along the dorsal
and ventral mantle margin, which probably function
as mantle retractor muscles. These mantle retractors are
abundant and are oriented perpendicularly to the mantle
margin that lines the shell (Fig. 3A–B).
Myogenesis and adult myoanatomy of Argyrotheca
cistellula
Similar to Argyrotheca cordata, larvae of A. cistellula are lecithotrophic
and are brooded by the mother animal. A. cistellula
larvae lack setae and the mantle lobe encloses the
pedicle lobe during development. Thus, the fully developed
larvae have only two visible lobes, namely the apical
and the mantle lobe. The investigated larvae were around
117–139 μm long and 78–104 μm wide. The first muscles
appear in larvae with all lobes fully differentiated. These
are two dorsal mantle muscles which extend dorsally from
anterior to posterior in the mantle lobe (Fig. 4A). Parallel
and further lateral to these dorsal mantle muscles run the
early lateral mantle muscles, and the first rudiments of the
serial mantle muscles arise at this stage in the mantle lobe.
These develop subsequently into a network of muscles
that extends dorsally and ventrally from the two lateral
mantle muscles (Fig. 4A–F). These lateral mantle muscles
connect to the apical longitudinal muscles at the anterior
pole and to the posterior muscle ring at the posterior pole
of the larvae (Fig 4B–F). During subsequent development,
the ventral mantle muscles and the pedicle muscles
emerge (Fig. 4C). The pedicle muscles, situated in the centre
of the mantle lobe, are the most prominent muscles in
fully grown larvae (Fig. 4D). They connect to the apical
longitudinal muscles, which in turn are in contact with
the apical transversal muscles. The latter form a muscle
ring in the apical lobe (Fig. 4E–F). The musculature of
fully developed larvae includes the pedicle muscles, which
are connected to the apical longitudinal muscles, the ventral
mantle muscles, and the dorsal mantle muscles that
connect to the pedicle muscles. Furthermore, serial mantle
muscles, which extend dorsally and ventrally from the
lateral mantle muscles, are present. Ventrally, the serial
mantle muscles terminate at the ventral mantle muscles
(Fig. 4F).
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Myogenesis in Argyrotheca cordata
Figure 2
Myogenesis in Argyrotheca cordata. CLSM maximum projection micrographs, anterior faces upward. F-actin is labelled in
red and cell nuclei are labelled in blue to indicate the outline of the specimens. Scale bars equal 50 μm. (A) Early larva in dorsal
view with the first F-actin signals from microvilli (mi) within the setal canals. (B) Early three-lobed larval stage, postero-dorsal
view, showing apical lobe (AL), mantle lobe (ML), pedicle lobe (PL), first rudiments of the pedicle musculature (pm), and microvilli
(mi) in the setae pouches. (C) Larval stage with fully differentiated lobes and short setae in ventral view (corresponding to
the larval stage shown in Fig. 1E). Visible are the apical transversal muscle (atm), the apical longitudinal muscles (alm), the interconnecting
apical muscles (iam), the interconnecting mantle muscles (imm), the longitudinal muscles (lm), the foregut rudiment
(fg), the hindgut rudiment (hg), the pedicle muscles (pm), microvilli (mi), the setae pouch musculature (arrowheads), and the
setae muscles (sm). (D) Lateral right view of a fully developed three-lobed larva with the U-shaped muscle (empty arrows) on
the ventral side. At this stage, the setae pouches are interconnected by a circular mantle muscle (arrow). New at this stage are
the central mantle muscles (empty arrowhead). Further indicated are the setae pouch musculature (arrowheads), the setae
muscles (sm), the serial mantle muscles (double arrowheads), the pedicle musculature (pm), and the apical longitudinal muscles
(alm). (E) Same stage as in D, ventro-lateral view. The U-shaped muscle (empty arrows) is directly connected to the pedicle
muscles (pm). In addition, the apical transversal muscle (atm), the apical longitudinal muscles (alm), the serial mantle muscles
(double arrowhead), the central mantle muscles (empty arrowheads), the setae pouch muscles (arrowheads), the setae muscles
(sm), the circular mantle muscle (arrow), and the primordial hump (asterisk) are indicated. (F) Fully grown larva in ventral
view with circular mantle muscle (arrows), serial mantle muscles (double arrowheads), setae pouch muscles (arrowheads),
setae muscles (sm), pedicle muscles (pm), longitudinal muscles (lm), apical longitudinal muscles (alm), apical transversal muscle
(atm), interconnecting apical muscles (iam), primordial hump (asterisk), and central mantle muscles (empty arrowheads).
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Similar to the condition found in Argyrotheca cordata, four
pairs of shell muscles are found in adult A. cistellula (Fig.
5). One pair of shell adductors attaches medially to the
dorsal and to the ventral valve (Fig. 5). Two pairs of pedicle
adjustors extend posterior into the pedicle, whereby
one attaches to the dorsal and one to the ventral valve.
Finally, one pair of diductors attaches at the posterior end
of the ventral valve and runs to the dorsal valve.
Each tentacle of the lophophore contains a number of striated
muscle fibres. Mantle margin muscles are arranged
perpendicularly to the shell periphery along the edge of
the dorsal and the ventral valve (Fig. 5A–B).
Myogenesis, metamorphosis, and juvenile myoanatomy of
Terebratalia transversa
Larvae of Terebratalia transversa are lecithotrophic and
develop for approximately four days at 11°C in the water
column until they are competent to undergo metamorphosis.
The investigated larvae were three-lobed, 120–178
μm long and 94–141 μm wide, whereby the pedicle lobe
was partly overgrown by the mantle lobe. The first developing
muscles are the pedicle muscles and early rudiments
of the serial mantle muscles (Fig. 6A). Thereafter,
the musculature of the four setae pouches forms (Fig. 6B).
In later stages, the setae pouch muscles interconnect with
the circular mantle muscle (Fig. 6C). A U-shaped muscle
extends on the ventral side of the larvae from the pedicle
muscles towards the circular mantle muscle. The serial
mantle muscles and the setae muscles span between the
circular mantle muscle and the U-shaped muscle strand.
The latter run from the setae pouches to the central mantle
muscles (Fig. 6D). The central mantle muscles extend
from the dorsal setae muscles, which run from the dorsal
setae pouches towards the apical lobe. They connect to the
apical longitudinal muscles at the border of the apical and
the mantle lobe (Fig. 6D). Subsequently, the apical musculature
develops, which consists of an apical transversal
muscle and two lateral apical longitudinal muscles that
are connected to the serial mantle muscles (Fig. 6E). In
late three-lobed larvae, the pedicle muscles are, together
with the central mantle muscles, the most prominent
muscular structures. The central mantle muscles connect
to the serial mantle muscles, the setae pouch muscles, the
setae muscles, and the apical musculature (Fig. 6F).
During metamorphosis, parts of the larval musculature
appear to get resorbed and juvenile muscles develop (Fig.
7A). We were, however, unable to clarify whether or not
certain components of the larval musculature are incorporated
into the juvenile muscular bodyplan.
The juvenile musculature comprises early rudiments of
the tentacle muscles, early rudiments of the mantle margin
musculature, the musculature of the intestine, adductors,
ventral pedicle adjustors which are connected to the
diductors, and dorsal pedicle adjustors (Fig. 7B–D).
Discussion
Comparison of larval and adult rhynchonelliform
myoanatomy
The gross morphology of Argyrotheca cistellula differs considerably
from that of A. cordata and Terebratalia transversa
in that the pedicle lobe gets enclosed by the mantle lobe
during development [19]. Thus, A. cistellula appears twolobed
and lacks setae, while the other two species express
three distinct body lobes and setae. Despite these differences,
myogenesis follows a similar pattern in all three
species (Table 1). When fully developed, prominent pedicle
muscles, apical longitudinal as well as apical transversal
muscles, and serial mantle muscles are present in all
three species. In addition, A. cordata and T. transversa
show a circular mantle muscle which we consider homologous
to the posterior muscle ring in A. cistellula. This
homology is based on the similar position of this muscle
in the mantle lobe and the fact that the U-shaped muscle
of A. cordata and T. transversa and the ventral mantle muscles
of A. cistellula all insert at this muscle. The central
mantle muscles of A. cordata and T. transversa are in our
opinion homologous to the dorsal mantle muscles of A.
cistellula due to the similar position of these muscles and
their connection to the apical and the serial mantle muscles
in all three species. The U-shaped muscle of A. cordata
and T. transversa corresponds to the ventral mantle muscles
in A. cistellula due to their similar position and the fact
that these muscles enclose the serial mantle muscles
antero-ventrally.
Despite these similarities, we found distinct differences in
the myoanatomy of the three species investigated. As
such, the setae pouch muscles, the setae muscles, and the
longitudinal muscles, which run from the mantle lobe to
the pedicle lobe, are only present in A. cordata and T. transversa,
while the lateral mantle muscles are only present in
larvae of A. cistellula. These differences between A. cistellula
on the one hand and A. cordata and T. transversa on
the other correspond to the gross morphological observation
that A. cistellula lacks setae.
Larval setae in brachiopods have been proposed to function
as a defence device and to control buoyancy [31]. The
setae of A. cistellula larvae have probably been secondarily
lost, as these larvae are brooded and may settle shortly
after release from the mother animal. However, A. cordata
larvae have retained their setae despite being brooded,
which may hint towards an extended planktonic period of
these larvae.
The muscles in the pedicle lobe have been proposed earlier
to be of functional use during metamorphosis
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Adult myoanatomy of Argyrotheca cordata
Figure 3
Adult myoanatomy of Argyrotheca cordata. F-actin is labelled in red and cell nuclei are labelled in blue. Scale bars equal
100 μm in all aspects except in E, where it equals 25 μm. (A) Overlay of CLSM maximum projection micrograph and light
micrograph, anterior faces upward, dorsal view. Indicated are the tentacle muscles (tm), the mantle margin muscles (mm), the
tentacles of the lophophore (te), the mantle cavity (mc), the intestine (in), the shell (s), the adductors (ad), the ventral pedicle
adjustors (vpa), which extend from the ventral valve into the pedicle, the dorsal pedicle adjustors (dpa), which extend from the
dorsal valve into the pedicle, and the diductors (di). One diductor is lacking as a result of the removal of the animal from the
substrate. (B) Overlay of a CLSM maximum projection micrograph and a light micrograph, anterior faces upward, ventral view.
Indicated are the same structures as in A. (C) Enlarged view of the ring musculature lining the intestine (in). In addition, one
adductor (ad), the diductors (di), and the ventral pedicle adjustors (vpa) are visible. (D) Enlarged view of the tentacles of the
lophophore and the corresponding tentacle musculature (tm). (E) Detail of a tentacle muscle fibre showing typical striation pattern
(double arrows).
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[32,33]. When larvae settle, a glandular region at the tip of
the primordial hump functions as site of attachment to
the substrate [34]. Subsequently, the primordial hump
forms the first rudiment of the juvenile pedicle. After larval
settlement, the mantle lobe is inverted over the apical
lobe and eventually forms the juvenile mantle. The apical
lobe gets enclosed by the valves and forms the lophophore
and all anterior adult structures [32,35]. At the
onset of metamorphosis, the U-shaped muscle may, due
to its connection to the pedicle muscles and the circular
mantle muscle, aid in inverting the mantle lobe. During
metamorphosis, the larval pedicle muscles are still present
at the time of ventral pedicle adjustor and diductor formation.
However, whether the larval pedicle muscles are
resorbed or are (partly) incorporated into the juvenile
diductor and/or pedal adjustor muscles could not be clarified
by the present study.
Argyrotheca cordata is the sole species from this study for
which data on the larval myoanatomy had previously
been available. In the first descriptions from 1873 and
1883, "muscles abdominaux", that run from the pedicle
lobe into the mantle lobe, had been identified [14,30]. A
different description was given slightly later, when a network
of muscles in the fully developed larva was
described. The muscles were denoted "Muskel des lateralen
Borstenbündels", "Muskel des medialen Borstenbündels",
"musculus contractor", "musculus rotator
dorsalis", and "musculus abductor" [15]. Our findings
confirm the results of the first papers with respect to the
pedicle muscles and the setae muscles. However, in our
specimens, the pedicle muscles were not directly connected
to the setae muscles as depicted in the first descriptions,
but were instead connected to the U-shaped muscle.
In adult Argyrotheca cordata, four pairs of muscles had
been identified previously [30]. The pair of adductor muscles
has two insertion sites, one anterior to the other at the
dorsal valve, and an additional one at the ventral valve.
The pair of diductor muscles inserts at the posterior part
of both the ventral and the dorsal valve. One of the two
pairs of adjustors inserts at the ventral valve and the pedicle,
while the other pair inserts at the dorsal valve and the
pedicle [30].
The muscular systems of adult A. cordata and A. cistellula
are similar to each other and comprise one pair of adductors,
two pairs of pedicle adjustors and one pair of diductors.
The tentacles contain several fibres of striated
musculature which have previously been described as
"rows of striated fusiform myoepithelial cells" in the
lophophore of T. transversa [36].
For the juvenile musculature of Terebratalia transversa we
followed the nomenclature used by Eshleman and
Wilkens [37]. The juvenile musculature, five days after
metamorphosis, comprises rudiments of the tentacle
muscles, rudiments of the mantle margin musculature,
one pair of adductors, one pair of diductors, one pair of
dorsal, and one pair of ventral pedicle adjustors. The ventral
pedicle adjustors are connected to the diductors in the
juvenile.
Comparative myogenesis of Lophophorata
For the Phoronida, data on muscle development are currently
available for three species, namely Phoronis pallida,
P. harmeri, and P. architecta [24,26,27]. The larvae of these
species are of the actinotroch-type and differ considerably
from brachiopod larvae in both their gross anatomy and
in their lifestyle, because these phoronid larvae are planktotrophic,
while the brachiopod larvae investigated herein
are of the typical three-lobed, lecithotrophic type. Accordingly,
a considerable part of the larval phoronid musculature
is linked to the digestive system (e.g., the oesophageal
ring muscles) and to the maintenance of a cylindrical
body shape (e.g., a meshwork of circular and longitudinal
muscles in the bodywall). In addition, trunk retractor
muscles, that originate from the posterior collar ring muscles
and insert in the telotrochal region, are present in
phoronid larvae [27]. The collar region contains mainly
ring muscles and few longitudinal muscles. The subumbrellar
and exumbrellar layers of the hood contain circular
muscles and a series of longitudinal muscles, which, in
the exumbrellar layer, function as hood elevators [27].
Furthermore, the tentacles of phoronid actinotroch larvae
contain elevator and depressor muscles which consist of
two loops in the elevators and a single loop in the depressors.
These tentacle muscles are interconnected by the ring
muscle of the collar [27]. We did not identify any muscles
in the larvae of the three brachiopod species described
herein that could potentially correspond to the actinotroch
muscle systems known so far.
The muscular architecture in ectoproct larvae is very
diverse, thus following the high plasticity of larval gross
morphology and the notion that lecithotrophic larvae
might have evolved up to six times within Ectoprocta [38].
To date, the larval muscular systems have been described
for Membranipora membranacea (cyphonautes larva), Flustrellidra
hispida (pseudocyphonautes larva), Celleporaria
sherryae and Schizoporella floridana (both coronate larva),
Bowerbankia gracilis (vesiculariform larva), Bugula stolonium
and B. fulva (both buguliform larva), Sundanella
sibogae, Nolella stipata, Amathia vidovici, Aeverrillia setigera,
and Alcyonidium gelatinosum (all ctenostome larva), and
Crisia elongata (cyclostome larva) [25,28]. Recently, a
number of homologies have been proposed for various
larval ectoproct muscle systems [25]. These are the coronal
ring muscle, which underlies the ciliated, ring-shaped
swimming organ of most larval types, the anterior median
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Myogenesis in Argyrotheca cistellula
Figure 4
Myogenesis in Argyrotheca cistellula. Overlay of CLSM maximum projection micrograph and light micrograph, anterior
faces upward. Scale bars equal 50 μm. Note that only two lobes are visible: the apical lobe (AL) and the mantle lobe (ML),
which encloses the pedicle lobe. (A) Early larva in dorsal view with the dorsal mantle muscles (empty arrowheads), the early
lateral mantle muscles (lmm), and early rudiments of the serial mantle muscles (double arrowheads). (B) Dorsal view of a later
larval stage with the lateral mantle muscle strand (lmm), rudiments of the posterior muscle ring (arrow), dorsal mantle muscles
(empty arrowheads), and the serial mantle muscles (double arrowhead). (C) Later larva in ventro-lateral left view with pedicle
muscles (pm) that are connected to the ventral mantle muscles (empty arrows). The serial mantle muscles (double arrowhead)
are connected to the lateral mantle muscles (lmm), the apical longitudinal muscles (alm) start to develop, and the early posterior
muscle ring is visible (arrow). (D) Same stage as in C, dorsal view. The pedicle muscles (pm) are prominent and connect to
the dorsal mantle muscles (empty arrowheads). In addition, the lateral mantle muscles (lmm), the serial mantle muscles (double
arrowheads), a part of the posterior muscle ring (arrow), and the apical longitudinal muscles (alm) are visible. (E) Fully developed
larva, ventral view. The apical transversal (atm) and the apical longitudinal muscles (alm) are fully developed and connect
to the pedicle muscles (pm). The connection between pedicle muscles and dorsal mantle muscles (empty arrowheads) is visible
in the anterior region of the pedicle muscles. Further indicated are the ventral mantle muscles (empty arrows), the serial mantle
muscles (double arrowheads), the lateral mantle muscles (lmm), and the area of the posterior muscle ring (arrow). (F) Same
larval stage as in E, ventro-lateral left view. The pedicle muscles (pm) are the most prominent muscles in the centre of the mantle
lobe. They are connected to the apical longitudinal muscles (alm), which terminate at the apical transversal muscle (atm),
which in turn forms a muscle ring in the apical lobe. The ventral mantle muscles (empty arrows) and dorsal mantle muscles
(empty arrowhead) are also connected to the pedicle muscles. The serial mantle muscles (double arrowhead) extend dorsally
and ventrally from the lateral mantle muscles (lmm). The latter terminate at the posterior muscle ring (arrow).
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muscle, which runs anteriorly from ventral to dorsal in
most species, lateral muscles, which project laterally in
dorso-ventral direction in most larvae, longitudinal muscles
along the posterior body axis, and transversal muscles,
which are situated transversally in the central body
region of F. hispida, M. membranacea, and A. gelatinosum.
Besides these proposed homologous muscles, each larval
type shows unique muscles in the body wall and/or inside
the larval body, reflecting at least partly the functional
adaptations to a planktotrophic versus a lecithotrophic
lifestyle. No muscles corresponding to any of the ectoproct
muscle types were found in the brachiopod species
investigated in this study (and noticeably no homologous
muscles between the lecithotrophic ectoproct and brachiopod
larval types could be identified), again demonstrating
the high plasticity of lophophorate larval anatomy.
Conclusion
All rhynchonelliform brachiopod larvae studied to date
are three-lobed with four bundles of setae [39], except for
the larva of Argyrotheca cistellula, which is externally
bilobed and lacks setae, and the three-lobed thecideid larvae,
which likewise lack setae [40]. Despite these gross
morphological differences, myogenesis in the three brachiopod
species investigated is very similar. Thus, we propose
a larval muscular groundpattern for
rhynchonelliform brachiopods comprising apical longitudinal
muscles, apical transversal muscles, circular mantle
muscles, central mantle muscles, longitudinal muscles,
serial mantle muscles, pedicle muscles, setae pouch muscles,
setae muscles, and a U-shaped muscle. However, a
final statement can only be made once data on the musculature
of theceid and rhynchonellid larvae become
available.
Comparing this proposed larval muscular groundpattern
to the hitherto investigated phoronids, ectoprocts, and
spiralian taxa such as polychaetes, molluscs,
plathelminths or entoprocts does not reveal any homologies
of larval brachiopod muscles and the muscles of other
lophotrochozoan larvae, regardless of whether the respective
larvae are lecithotrophic or planktotrophic [23,41-
47]. From these data we conclude that the ontogenetic
pathways of the individual lophophorate phyla have split
early in evolution from that of other Lophotrochozoa,
which then resulted in the wide morphological diversity
of larval and adult lophophorate bodyplans.
Methods
Animal collection and fixation
Argyrotheca cordata and A. cistellula
Adults were obtained from encrusting coralline red algae
(coralligène), which was collected in the vicinity of the
Observatoire Océanologique de Banyuls-sur-mer, France
(42°29'27.51" N; 3°08'07.67" E), by SCUBA from 30–40
m depth in July 2002 and June 2007. All developmental
stages from unfertilized eggs to fully differentiated larvae
were obtained by dissection from the adults. The specimens
were relaxed at room temperature in 7.14% MgCl 2 ,
fixed in 4% paraformaldehyde (PFA) in 0.1 M phosphate
Adult myoanatomy of Argyrotheca cistellula
Figure 5
Adult myoanatomy of Argyrotheca cistellula. Overlay of CLSM maximum projection micrograph and light micrograph,
anterior faces upward. Scale bars equal 300 μm. (A) Dorsal view. (B) Ventral view. Indicated are the mantle margin muscles
(mm), the shell (s), the adductors (ad), the diductors (di), the dorsal pedicle adjustor (dpa), the ventral pedicle adjustor (vpa),
the intestine (in), the mantle cavity (mc), and the tentacle muscles (tm).
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Myogenesis in Terebratalia transversa
Figure 6
Myogenesis in Terebratalia transversa. Overlay of CLSM maximum projection micrograph and light micrograph, anterior
faces upward. Scale bars equal 50 μm. (A) Ventral view of an early three-lobed stage with apical lobe (AL), mantle lobe (ML),
and pedicle lobe (PL). Discernable are the pedicle musculature (pm), the first anlagen of the serial mantle muscles (double
arrowhead), and the setae (se). (B) Ventral view of a slightly older larva with prominent pedicle musculature (pm), anlagen of
the setae pouch musculature (arrowheads), and setae (se). (C) Later larval stage, ventral view with pedicle musculature (pm),
setae pouch muscles (arrowhead), serial mantle muscles (double arrowhead), and central mantle muscles (empty arrowheads),
which are extensions of the dorsal setae muscles. The serial mantle muscles are posteriorly connected to the circular mantle
muscle (arrows) and antero-ventrally connected to the U-shaped muscle (empty arrows), which extends from the pedicle muscles
to the circular mantle muscle. (D) Lateral view of a later larva with the muscle systems described in C. In addition, the first
anlagen of the apical longitudinal musculature (alm), the setae muscles (sm), and the setae (se) are visible. (E) Same stage as in
D with prominent pedicle muscles (pm) that are connected to the apical longitudinal muscles (alm). The latter connect to the
apical transversal muscle (atm). In addition, the setae pouch muscles (arrowheads), the setae muscles (sm), and the setae (se)
are indicated. (F) Fully developed larva, ventral view, with central mantle muscles (empty arrowheads), pedicle muscles (pm),
circular mantle muscle (arrows), U-shaped muscle (empty arrows), serial mantle muscles (double arrowheads), setae pouch
musculature (arrowheads), setae muscles (sm), apical longitudinal muscles (alm), apical transversal muscle (atm), and setae (se).
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Metamorphosis and adult myoanatomy of Terebratalia transversa
Figure 7
Metamorphosis and adult myoanatomy of Terebratalia transversa. (A-C) Overlay of CLSM maximum projection
micrograph and light micrograph, anterior faces upward. F-actin is labelled in red and cell nuclei are labelled in blue. Scale bars
equal 50 μm. (A) Larva during metamorphosis. A mosaic of larval and juvenile features are present including the pedicle (pe),
the larval pedicle muscles (pm), the first rudiments of the juvenile tentacle musculature (tm), one diductor (di), and the ventral
pedicle adjustors (vpa). (B) Juvenile 5 days after metamorphosis, dorsal view with the remaining larval setae (se), the mantle
margin muscles (mm), the tentacle muscles (tm), the adductors (ad), the musculature of the intestine (in), the diductors (di),
the ventral pedicle adjustors (vpa), the dorsal pedicle adjustors (dpa), and the pedicle (pe). (C) Juvenile 5 days after metamorphosis,
ventral view with the remaining larval setae (se), rudiments of the mantle margin muscles (mm), rudiments of the tentacle
muscles (tm), the adductors (ad), the ventral pedicle adjustors (vpa), the diductors (di), the dorsal pedicle adjustors (dpa),
and the pedicle (pe). (D) Reconstruction of the 3D arrangement of the juvenile musculature based on the CLSM dataset used
in C showing the dorsal pedicle adjustors (red), the adductors (dark blue), the mantle margin muscles (light blue), and the tentacle
muscles (magenta). The ventral pedicle adjustors (yellow) are ventrally connected to the diductors (green).
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buffer (PB) for 2 hours or for 3–5 hours, and subsequently
washed thrice with 0.1 M PB for 15 min each. The samples
were stored in 0.1 M PB with 0.1% NaN 3 at 4°C. Material
fixed for 2 hours was used for immunocytochemistry
(ICC) and material fixed for 3–5 hours was used for scanning
electron microscopy (SEM).
Terebratalia transversa
Adults were collected in the San Juan Archipelago, USA, in
the vicinity of the Friday Harbor Laboratories, and were
kept in running seawater tables. To obtain larvae, females
were dissected and their eggs transferred into beaker
glasses with filtered seawater. The seawater was changed
several times in order to wash off follicle cells, and the
eggs were left overnight for germinal vesicle breakdown.
Males were opened and left in filtered seawater overnight.
Thereafter, their testes were scraped out, macerated, and
diluted with filtered seawater to obtain a sperm suspension.
Prior to fertilization, sperm cells were activated by
adding three drops of a 1 M Tris buffer solution (Sigma-
Aldrich, St. Louis, MO, USA) to approximately 50 ml of
sperm suspension. Larvae were maintained in embryo
dishes at around 11°C and the filtered seawater was
changed twice daily. Free swimming larvae, metamorphic
stages, and juveniles five days after metamorphosis were
relaxed in 7.14% MgCl 2 and fixed in 4% PFA in 0.1 M PB
for 30 min at room temperature. Larvae were washed
thrice for 15 min in 0.1 M PB and stored in 0.1 M PB with
0.1% NaN 3 at 4°C.
Scanning electron microscopy
For scanning electron microscopy (SEM), the specimens
were postfixed in 1% OsO 4 , dehydrated in a graded acetone
series, critical point dried, and sputter coated with
gold. Digital images were acquired using a LEO 1430 VP
SEM (Zeiss, Jena, Germany).
Table 1: Comparative larval myoanatomy of the rhynchonelliform brachiopods Argyrotheca cordata, Terebratalia transversa, and A.
cistellula
Species
Muscle Argyrotheca cordata Terebratalia
transversa
Argyrotheca
cistellula
Location
Symbol in figures
apical longitudinal
muscles
apical transversal
muscle
+ + + apical lobe alm
+ + + (apical muscle ring) apical lobe atm
central mantle muscles + + +
(dorsal mantle
muscles)
mantle lobe
empty arrowheads
circular mantle muscle + + +
(posterior muscle ring)
mantle lobe
arrows
lateral mantle muscle - - + mantle lobe lmm
longitudinal muscles + + - mantle and pedicle
lobe
lm
pedicle muscles + + + pedicle lobe pm
serial mantle muscles + + + mantle lobe double arrowsheads
setae muscles + + - mantle lobe sm
setae pouch
musculature
+ + - mantle lobe arrowheads
U-shaped muscle + + +
(ventral mantle
muscle)
mantle lobe
empty arrows
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50 Chapter II
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F-actin labelling, confocal laserscanning microscopy
(CLSM), and 3D reconstruction
Prior to staining, larvae were washed thrice for 15 min in
PB and incubated for 1 h in PB containing 0.1% Triton X-
100 (Sigma-Aldrich) to permeabilize the tissue. Then, the
specimens were incubated in 1:40 diluted Alexa Fluor 488
phalloidin (Invitrogen, Molecular Probes, Eugene, OR,
USA) and 3 μg/ml DAPI (Invitrogen) in the permeabilization
solution overnight at 4°C. Subsequently, specimens
were washed thrice for 15 min in 0.1 M PB and embedded
in Fluoromount G (Southern Biotech, Birmingham, AL,
USA) on glass slides. The same procedure was used for
juveniles and adults, with the addition of a decalcifying
step using 0.05 M EGTA (Sigma-Aldrich) at room temperature
overnight prior to permeabilization and staining.
Negative controls omitting the phalloidin dye were performed
on all species in order to avoid potential misinterpretations
caused by autofluorescence.
The samples were analysed with a Leica DM RXE 6 TL fluorescence
microscope equipped with a TCS SP2 AOBS
laserscanning device (Leica Microsystems, Wetzlar, Germany).
Animals were scanned at intervals of 0.49 μm or
0.64 μm, respectively, and the resulting image stacks were
merged into maximum projection images. Photoshop
CS3 (Adobe, San Jose, CA, USA) was used to create overlay
images of CLSM and light micrographs and for assembling
the figure plates. 3D reconstruction was performed
on CLSM datasets using volume rendering algorithms of
the graphics software Imaris 5.7.2 (Bitplane, Zurich, Switzerland).
Competing interests
The authors declare that they have no competing interests.
Authors' contributions
AA performed research and drafted the manuscript. AW
designed and coordinated research, performed the SEM
analysis, and contributed significantly to the writing of
the manuscript. Both authors read and approved the final
version of the manuscript.
Additional material
Additional file 1
Larval musculature of Argyrotheca cordata. Movie of a confocal scan
through a fully developed larva of Argyrotheca cordata to illustrate the
three-dimensional arrangement of the larval musculature.
Click here for file
[http://www.biomedcentral.com/content/supplementary/1742-
9994-6-3-S1.mpg]
Acknowledgements
We are grateful to Henrike Semmler (Copenhagen) for rearing and fixing
Terebratalia larvae during the Comparative Invertebrate Embryology class
2006 at the Friday Harbor Laboratories and for comments on an early draft
of the manuscript. We further thank the divers and the staff of the Marine
Biological Station Banyuls-sur-mer for collecting the coralligène and for
providing laboratory space. Scott Santagata (Brookville, New York) is
thanked for comments on the manuscript and Jana Hoffmann (Berlin, Germany)
for providing access to some of the classic literature. The valuable
comments of an anonymous reviewer helped to improve the manuscript.
This study was funded by the Danish Agency for Science, Technology and
Innovation (grant no. 645-06-0294 to AW) and the Danish Research
Agency (grant no. 21-04-0356 to AW). Research in the lab of A. Wanninger
is further supported by the EU-funded Marie Curie Network MOLMORPH
(contract grant number MEST-CT-2005-020542).
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52 Chapter III
Chapter III
Altenburger, A. & Wanninger, A. 2010 Neuromuscular development
in Novocrania anomala: evidence for the presence of serotonin
and a spiralian-like apical organ in lecithotrophic brachiopod
larvae. Evolution & Development 12: 16-24

Chapter III
53
EVOLUTION & DEVELOPMENT 12:1, 16–24 (2010)
DOI: 10.1111/j.1525-142X.2009.00387.x
Neuromuscular development in Novocrania anomala: evidence for the
presence of serotonin and a spiralian-like apical organ in lecithotrophic
brachiopod larvae
Andreas Altenburger and Andreas Wanninger
Department of Biology, Research Group for Comparative Zoology, University of Copenhagen, Universitetsparken 15,
DK-2100 Copenhagen Ø, Denmark
Author for correspondence (email: awanninger@bio.ku.dk)
SUMMARY The phylogenetic position of Brachiopoda remains
unsettled, and only few recent data on brachiopod
organogenesis are currently available. In order to contribute
data to questions concerning brachiopod ontogeny and evolution
we investigated nervous and muscle system development
in the craniiform (inarticulate) brachiopod Novocrania
anomala. Larvae of this species are lecithotrophic and have
a bilobed body with three pairs of dorsal setal bundles that
emerge from the posterior lobe. Fully developed larvae exhibit
a network of setae pouch muscles as well as medioventral
longitudinal and transversal muscles. After settlement, the
anterior and posterior adductor muscles and delicate mantle
retractor muscles begin to form. Comparison of the larval
muscular system of Novocrania anomala with that of
rhynchonelliform (articulate) brachiopod larvae shows that
the former has a much simpler muscular organization. The
first signal of serotonin-like immunoreactivity appears in fully
developed Novocrania anomala larvae, which have an apical
organ that consists of four flask-shaped cells and two ventral
neurites. These ventral neurites do not stain positively for the
axonal marker a-tubulin in the larval stages. In the juveniles,
the nervous system stained by a-tubulin is characterized by
two ventral neurite bundles with three commissures. Our data
are the first direct proof for the presence of an immunoreactive
neurotransmitter in lecithotrophic brachiopod larvae and
demonstrate the existence of flask-shaped serotonergic cells
in the brachiopod larval apical organ, thus significantly
increasing the probability that this cell type was part of the
bauplan of the larvae of the last common lophotrochozoan
ancestor.
INTRODUCTION
The phylogenetic position of Brachiopoda remains unresolved,
although most molecular analyses agree on their inclusion
within Lophotrochozoa (Hejnol et al. 2009; Paps et al.
2009). Alternatively, some recent works support the more
traditional view that Brachiopoda clusters with Ectoprocta
and Phoronida to form the Lophophorata, the direct sistergroup
of Spiralia (Trochozoa) (Gee 1995; Nielsen 2002; Halanych
2004). Current brachiopod internal phylogeny suggests
division of the phylum into the three clades Linguliformea,
Craniiformea, and Rhynchonelliformea (Williams et al. 1996).
Craniiform brachiopods share morphological traits with both
linguliforms and rhynchonelliforms. For example, craniiforms
and linguliforms possess a circumferential mantle cavity, a
muscle system with oblique muscles, and two pairs of shell
adductors, a transitional median tentacle during lophophore
development and a median division of the brachial canals into
two separate cavities within the lophophore. Craniiforms and
rhynchonelliformes exhibit a proteinaceous calcitic shell, a
16
single row of tentacles on a trocholophous lophophore,
gonads suspended in the mantle sinus, and lecithotrophic
larvae (Rowell 1960; Atkins and Rudwick 1962; Williams
et al. 1996).
Experimental embryology has shown that the animal half
of the egg forms the ectodermal epithelium of the apical lobe,
whereas the vegetal half forms endoderm, mesoderm, and the
ectoderm of the mantle lobe in Novocrania anomala (Mu¨ ller
1776) (previously assigned to various genera and thus also
referred to in the literature as Crania anomala or Neocrania
anomala, respectively) (Lee and Brunton 1986, 2001; Freeman
and Lundelius 1999; Freeman 2000; Holmer 2001; Cohen et
al. 2008). During metamorphosis, both the ventral and the
dorsal valve are formed from the dorsal epithelium of the
larva (Nielsen 1991).
Recent immunocytochemical studies have revealed the almost
universal occurrence of an apical organ that contains
flask-shaped cells in larvae of Annelida, Mollusca, Sipuncula,
Entoprocta, and Platyhelminthes (see Wanninger 2009 for
review). These flask-shaped cells express serotonin-like
& 2010 Wiley Periodicals, Inc.

54 Chapter III
Altenburger and Wanninger
immunoreactivity and may also show FMRFamidergic
immunoreactivity. The wide occurrence of serotonin indicates
that this neurotransmitter was part of the ancestral metazoan
nervous system (Hay-Schmidt 2000). Surprisingly, neither
serotonin-like immunoreactivity nor the existence of flaskshaped
cells have hitherto been proven for lecithotrophic larvae
of any brachiopod clade, thus leaving a significant gap in
our understanding of the evolution of the brachiopod nervous
system and the origin of this cell type within the lophophorates.
Accordingly, we provide herein the first thorough
immunocytochemical study on neurogenesis in a brachiopod
with a lecithotrophic larva, the craniiform Novocrania anomala,
and compare our findings with data on other lophotrochozoan
phyla. In our general quest to shed light on
brachiopod organogenesis, we also present data on Novocrania
anomala myogenesis, which for the first time allows conclusive
comparisons between the muscular systems of
craniiform and rhynchonelliform brachiopod larvae and thus
contributes to answering questions concerning the ancestral
muscular bodyplan of brachiopod larvae.
MATERIALS AND METHODS
Animal collection, breeding, and fixation
Rocks with attached adults of Novocrania anomala where obtained
by dredging in the vicinity of the Sven Love´ n Centre for Marine
Sciences, Gullmarsfjord, Sweden (58115 0 921 00 N, 11125 0 103 00 E) in
October 2007 and September 2008. The rocks were maintained in
the laboratory in running seawater and adults were removed and
dissected for gametes. For artificial fertilization, eggs and sperm
were removed from the gonads with pulled glass pipettes and
placed in separate glass beakers with filtered seawater at ambient
seawater temperature (141C). The water containing the eggs was
changed at least four times to wash off follicle cells and superfluous
gonad tissue. Eggs were regularly checked for germinal vesicle
breakdown and sperm cells were checked for motility under a
compound microscope. After approximately 12 h, 2 ml of a highly
diluted sperm suspension (testes of three to five adults in approximately
100 ml filtered sea water) were added to the beakers containing
eggs. Developing larvae were fixed at various stages after
fertilization (from 34 h post-fertilization [hpf] to 17 days post-settlement)
in 4% paraformaldehyde in 0.1 M phosphate buffer (PB)
for 90 min. Thereafter, larvae were washed three times for 15 min
each in 0.1 M PB and finally stored in 0.1 M PB containing 0.1%
NaN 3 at 41C.
Immunocytochemistry, confocal laserscanning
microscopy (CLSM), and three-dimensional (3D)
reconstruction
Before staining, larvae were washed thrice for 15 min each in PB
and incubated for 1 h in PB containing 0.2% Triton X-100
(Sigma-Aldrich, St. Louis, MO, USA) at room temperature to
permeabilize the tissue. For F-actin staining, specimens were left
overnight at 41C in0.1M PB containing 0.2% Triton X-100 and
1:40 diluted Alexa Fluor 488 phalloidin (Invitrogen, Molecular
Probes, Eugene, OR, USA). For serotonin and a-tubulin staining,
specimens were first incubated overnight at 41C in 6% normal goat
serum in 0.1 M PB and 0.2% Triton X-100 (blocking solution).
Second, specimens were incubated for 24 h at 41C in blocking solution
containing either a 1:800 diluted polyclonal primary serotonin
antibody (Zymed, Carlton Court, CA, USA), or a 1:500
diluted monoclonal primary acetylated a-tubulin antibody (Sigma-
Aldrich). Third, specimens were washed in the permeabilization
solution overnight at 41C with four changes. Then, the secondary
antibodies (either Alexa Fluor 633-conjugated goat anti-rabbit,
Invitrogen or TRITC-conjugated goat anti-rabbit, Sigma-Aldrich)
were added in a 1:300 dilution to the blocking solution and the
samples were incubated for 24 h. Subsequently, the specimens were
washed three times for 15 min each in 0.1 M PB and embedded in
Fluoromount G (Southern Biotech, Birmingham, AL, USA) on
glass slides. Negative controls omitting either the phalloidin dye or
the respective secondary antibody were performed in order to test
for signal specificity and rendered no signal. The samples were
analyzed with a Leica DM RXE 6 TL fluorescence microscope
equipped with a TCS SP2 AOBS laserscanning device (Leica Microsystems,
Wetzlar, Germany). Animals were scanned with 0.16–
0.49 mm step size, and the resulting image stacks were merged into
maximum projection images. In addition, light micrographs were
recorded to allow overlay with the CLSM images for exact orientation
and localization of the muscle and nervous systems within
the animals. Adobe Photoshop CS3 software (Adobe, San Jose,
CA, USA) was used to create overlay images and for assembling
the figure plates. The sketch drawings were generated with Adobe
Illustrator CS3 (Adobe), and the 3D reconstructions were created
with the Imaris imaging software version 5.7.2 (Bitplane, Zu¨ rich,
Switzerland) based on the CLSM image stacks.
RESULTS
Brachiopod neuromuscular development 17
Myogenesis
The first signals of F-actin were found in the setae pouches of
bilobed larvae at the onset of setae formation. The six setae
pouches are distributed in pairs along the dorsal ridge of the
posterior lobe (Fig. 1A). As the setae grow, the setae pouch
muscles develop further into spherical systems (Figs. 1, B and
G–I and 2A). Later in development, the setae pouch muscles
get interconnected by two bundles of medioventral longitudinal
muscles, which run ventrally from anterior to posterior
(Figs. 1, C and D and G–I and 2A). The medioventral longitudinal
muscle strands get interconnected by transversal
muscles (Fig. 1, B–D and G–I) that are distributed homogenously
in early stages (Fig. 1B) and concentrate into three
bundles in later stages (Fig. 1D). Accordingly, the metamorphic
competent larva has setae pouch muscles, medioventral
longitudinal muscles, and transversal muscles. During metamorphosis,
the larval musculature is replaced by the juvenile
musculature, which most likely develops entirely de novo, that
is, independent of the larval muscle systems (Fig. 1E). The

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18 EVOLUTION & DEVELOPMENT Vol. 12, No. 1, January--February 2010
Fig. 1. Muscle development in Novocrania anomala. Overlay of maximum projection micrographs from phalloidin staining and light
micrographs. Anterior faces upwards and scale bars equal 50 mm. (A) Larva with anterior lobe (AL), posterior lobe (PL), and early signs of
F-actin in the three pairs of setae pouches (arrows) along the dorsal ridge of the posterior lobe. (B) Larva with setae (se), anterior lobe (AL),
posterior lobe (PL), setae pouch muscles (arrows), homogenously distributed transversal muscles (asterisks), and a distinct F-actin-rich area
(arrowheads), which might be involved in cementing the larva to the substrate during settlement. (C) Later larval stage with setae pouch
muscles (arrows), medioventral longitudinal muscles (empty arrows), and F-actin-rich area (arrowhead) on the dorsal side. (D) Metamorphic
competent larva in ventral view with setae (se) and setae pouch muscles (arrows), which are ventrally interconnected by two strands
of medioventral longitudinal muscles (empty arrows). The medioventral longitudinal muscles are interconnected by transversal muscles,
which at this stage are concentrated into three bundles (asterisks). (E) Specimen during metamorphosis with remnants of larval setae pouch
muscles (arrows) and larval medioventral longitudinal muscles (empty arrows), which are most probably undergoing resorption. The adult
anterior adductor muscles (aad) start to develop. (F) Juvenile with mantle margin muscles (mm), anterior adductor muscle (aad), oblique
muscle (ob), and posterior adductor muscles (pad). (G–I) Three-dimensional reconstruction of the dataset shown in (D). (G) Ventral view of
the musculature of a fully developed larva with medioventral longitudinal muscles (red), setae pouch muscles (yellow), and transversal
muscle (asterisk). (H) Same specimen as in (G), anterior view. (I) Same specimen as in (G), dorsal view.

56 Chapter III
Altenburger and Wanninger
Brachiopod neuromuscular development 19
Fig. 2. Semischematic representation of
the larval musculature of craniiform and
rhynchonelliform brachiopods. (A) Musculature
of Novocrania anomala with
setae pouch muscles (red circles), medioventral
longitudinal muscles (white), and
transversal muscles (yellow-grey). Size of
the specimen is approximately 150 mm.
(B) Musculature of Argyrotheca cordata
based on Altenburger and Wanninger
(2009) with pedicle muscles (beige), longitudinal
muscles (orange), central mantle
muscles (brown), U-shaped muscle
(green), setae pouch muscles (red circles),
circular mantle muscle (light blue), serial
mantle muscles (dark orange), setae muscles
(purple), apical longitudinal muscles
(dark blue), and apical transversal muscle
(yellow). Size of the specimen is approximately
280 mm.
juvenile musculature comprises mantle margin muscles,
oblique muscles, as well as anterior and posterior adductor
muscles (Fig. 1F).
Neurogenesis
The first signals of serotonin-like immunoreactivity appear in
fully developed, metamorphic competent, bilobed larvae at
approximately 86 hpf (Table 1). At this stage, four flaskshaped
cells are present in the anterior-most part of the apical
lobe (Fig. 3, A–D). They are oriented in different directions
with only one pointing toward the apical pole of the larva.
The flask-shaped cells are connected to two ventral neurites
that extend posteriorly (Fig. 3, A–D). The flask-shaped cells
are lost during metamorphosis, and early juveniles have two
ventral neurites that project from the anterior lobe into the
posterior lobe (Fig. 3E). During subsequent development, the
ventral neurites become interconnected by a median commissure
in the mid-part of the juvenile (Fig. 3F).
The axonal marker a-tubulin is first expressed in juveniles 5
days after metamorphosis (Fig. 4A). Two solid neurite bundles
develop ventrolaterally in the anterior lobe of the juvenile and
subsequently grow in posterior direction into the posterior lobe
(Fig. 4B). Later in development, these neurite bundles close by
an anterior and a posterior commissure, and the median commissure
is established (Fig. 4, C–F). Serially arranged mantle
neurites extend from the anterior part of the ventral neurite
bundles in a lateral direction toward the mantle margin of
the juvenile (Fig. 4, B–F). Comparison of the position of the
a-tubulin signal in the juvenile and the serotonin-like signal in
the larva suggests that the larval ventral neurites are the earliest
neurites of the future ventral neurite bundles of the juvenile.
Table 1. Landmarks of Novocrania anomala development at 141C
Age (hours post
fertilization)
Gross morphology
Myoanatomy as inferred by
F-actin staining
3–4 First cleavage No signal No signal
26–30 Swimming, spherical gastrula No signal No signal
42–49 Swimming, elongated gastrula No signal No signal
65–73 Swimming, bilobed larva with
setae starting to develop
First signals of actin in setae
pouches (Fig. 1A)
No signal
86–96 Fully established, swimming,
bilobed larva with long setae
168 Settled juvenile after metamorphosis
Fully developed larval musculature
with setae pouch muscles, longitudinal
muscles, and transversal muscles
(Fig. 1D)
Juvenile with mantle margin muscles,
anterior adductors, and posterior
adductors (Fig. 1F)
Neuroanatomy as inferred by
antibody staining
Larval nervous system with four
flask-shaped cells in the apical organ
and two ventral neurites (Fig.
3, B–D)
Juvenile with two ventral neurite
bundles, commissures, and serially
arranged neurites (Figs. 3, E and
F and 4, A–F)

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20 EVOLUTION & DEVELOPMENT Vol. 12, No. 1, January--February 2010
Fig. 3. Development of the serotonergic
nervous system in Novocrania anomala.
(A, B, E, and F) Overlay of maximum
projection micrographs of serotonin
staining and light micrographs. (C and
D) Three-dimensional reconstruction of
the dataset shown in B. Anterior faces
upwards and scale bars equal 50 mm. (A
and B) Metamorphic competent larva
with three pairs of setae bundles (se) and
four flask-shaped serotonergic cells (asterisks)
in the anterior part of the apical
lobe (AL), as well as two ventral
serotonergic neurites (arrows) running
from the apical lobe toward the posterior
lobe (PL). The stage in (A) is slightly
younger than that depicted in (B). (C and
D) Same dataset as in (B) with four flaskshaped
serotonergic cells (red) and two
ventral neurites, which are interconnected
anteriorly (yellow). (C) Ventral view. (D)
Lateral view. The flask-shape is visible
only in one cell due to the different position
of the cells. (E) Juvenile during
metamorphosis with two ventral neurites
(arrows), which run from the region of
the former anterior lobe (AL) into the
region of the former posterior lobe (PL).
Larval setae (se) and juvenile shell (s) are
present. (F) Later stage of a juvenile with
two ventral neurites (arrows) which are
interconnected by a median commissure
(mco).
DISCUSSION
Comparative brachiopod myoanatomy
The musculature of fully developed Novocrania anomala larvae
consists of setae pouch muscles, the medioventral longitudinal
muscles that interconnect these setae pouch muscles,
and transversal muscles that interconnect the medioventral
longitudinal muscles (Table 1 and Fig. 2A). This relatively
simple muscular organization differs significantly from that of
articulate brachiopod larvae, which comprises pedicle muscles,
longitudinal muscles, a circular mantle muscle, central
mantle muscles, a U-shaped muscle, serially arranged mantle
muscles, setae muscles, setae pouch muscles, apical longitudinal
muscles, and an apical transversal muscle (Fig. 2B; see
also Altenburger and Wanninger 2009). Unfortunately, very
little is known about brachiopod larval ecology and behavior

58 Chapter III
Altenburger and Wanninger
Brachiopod neuromuscular development 21
Fig. 4. Development of the nervous system
in Novocrania anomala as revealed by
acetylated a-tubulin staining. (A–D)
Overlay of maximum projection micrograph
of a-tubulin staining and light micrograph.
(E and F) Three-dimensional
reconstructions of the dataset shown in
(D). Anterior faces upwards and scale
bars equal 50 mm. (A) First a-tubulin signal
in a juvenile 5 days after metamorphosis.
The former larval apical lobe
(AL) and posterior lobe (PL) are still
visible under the shell (s) of the juvenile.
Two ventral neurite bundles develop in
the anterior lobe (arrows). The juvenile
body is still covered by larval cilia (ci).
Some serially arranged neurites (sn) extend
inwards from the ventral neurite
bundles. (B) The ventral neurite bundles
(arrows) elongate further in posterior direction.
A median commissure (mco)
starts to form. From the anterior portion
of the ventral neurite bundles, serially arranged
mantle neurites (smn) extend distally
outwards, and serially arranged
neurites (sn) extend inwards. The cilia of
the juvenile gut (gu) are visible in the
median region of the juvenile. (C) Juvenile
with the same structures as in (B).
The median commissure (mco) is closed
and the ventral neurite bundles (arrows)
have fused anteriorly to form the anterior
commissure (aco). (D) Neural anatomy
of a juvenile 17 days after metamorphosis
with an anterior commissure (aco), a median
commissure (mco), and a posterior
commissure (pco) that interconnect the
ventral neural bundles (arrows). In addition,
the serially arranged mantle neurites
(smn), which extend toward the edge of
the juvenile mantle, are visible. (E) Threedimensional
reconstruction of the dataset
shown in (D), dorsal view. (F) Three-dimensional
reconstruction of the dataset
shown in (D). Postero-dorsal view demonstrating
that the ventral neurite bundles
(yellow) and the serially arranged
mantle neurites (green) bend ventrally.
(James et al. 1992). Rhynchonelliform larvae show a change
from positive to negative phototactism when reaching metamorphic
competence. In laboratory cultures, they swim in the
culture dish with the anterior lobe or the ventral side of the
body repeatedly forming contact with the bottom of the dish,
probably probing for a suitable place for settlement (Chuang
1996). We observed a similar behavior in Novocrania anomala
larvae before metamorphosis.
At the current state of knowledge, it remains difficult to
relate the differences in larval myoanatomy to aspects concerning
the ecology of the respective brachiopod larvae, because
the latter remains virtually unknown (James et al. 1992).
An earlier study showed that larvae of Novocrania anomala
are able to settle 4 days after fertilization (Nielsen 1991), although
we observed this behavior only in 7-day-old larvae.
Rhynchonelliform brachiopods are known to settle after 3

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22 EVOLUTION & DEVELOPMENT Vol. 12, No. 1, January--February 2010
(Terebratulina retusa), 5–14 (Terebratalia transversa) (own
observations), or up to 160 days (Liothyrella uva) (see Peck
and Robinson 1994 for listed overview). However, whether or
not these differences in the planktonic lifespan of brachiopod
larvae accounts for the differences in their myoanatomy remains
speculative. Instead, we consider the dissimilarities in
how metamorphosis is achieved in craniiform (inarticulate)
and rhynchonelliform (articulate) larvae as a possible reason
for this morphological variation. During metamorphosis, larvae
of Novocrania anomala curl ventrally by contraction of
the paired medioventral muscles and attach to the substrate
via the epithelium at the posterior end of the larva. The brachial
valve is then secreted by the median part of the dorsal
epithelium and the pedicle valve is secreted by the attachment
epithelium (Nielsen 1991). Larvae of the rhynchonelliform
brachiopod T. transversa attach via a secretory product produced
by the distal tip of the pedicle lobe at the posterior end
of the larva. After attachment, the mantle lobe flips over the
apical lobe and secretes a protegulum containing calcium
carbonate (Stricker and Reed 1985; Freeman 1993).
The phylogenetic relationship of craniiforms to the other
brachiopod subtaxa is still controversial. Based on their lack of
a valve-to-valve articulation they have traditionally been
grouped together with other inarticulated groups (Williams
and Rowell 1965a, b). This view is supported by molecular
analyses based on 18S rDNA sequences, which either place the
craniiforms within the linguliforms (Cohen 2000) or as the
direct sister-group to the linguliforms (Cohen and Weydmann
2005). Other morphological characters such as the presence of
an anus and a lophophore without internal mineralized support
underpins a close relationship of craniiform and linguliform
brachiopods (Carlson 1995). However, based on the
lecithotrophy of the larvae and the presence of a calcareous
shell in the adults, craniiform brachiopods have been proposed
to be closer related to the rhynchonelliforms rather than to
the linguliforms, which have a free-swimming planktotrophic
life cycle stage that closely resembles the morphology of juvenile
brachiopods (Nielsen 1991). An alternative scenario proposes
that lecithotrophic larvae equipped with larval setae are
basal for Brachiopoda and that the swimming ‘‘paralarvae’’ of
lingulids constitute a planktonic juvenile stage, thereby implying
that the linguliforms have secondarily lost the lecithotrophic
larva (Lu¨ ter 2001). Our data corroborates this view.
The musculature of postmetamorphosic Novocrania anomala
comprises anterior adductors, posterior adductors, and
oblique lateral muscles. This corresponds to the musculature
found in adults, which in addition have brachial protractor
muscles at the base of the lophophore, an unpaired median
muscle, and oblique internal muscles (Bulman 1939; Helmcke
1939; Williams and Rowell 1965a, b). In the present study we
found mantle retractor muscles, which had previously been
undescribed for Novocrania anomala and which correspond to
the respective muscles found in the rhynchonelliform brachiopods
Argyrotheca cordata, Argyrotheca cistellula, and
Terebratalia transversa (Altenburger and Wanninger 2009).
Given the distinct differences in the larval musculature of
craniiforms and rhynchonelliforms, it is difficult to infer a
muscular ground pattern for brachiopod larvae. However, it
appears likely that a hypothetical ancestral brachiopod larva
had at least setae pouch muscles and a musculature that interconnect
these setae pouch muscles.
Neurogenesis
The serotonergic nervous system of Novocrania anomala starts
to develop in fully established larvae (see Table 1), and shows
an apical organ consisting of four flask-shaped cells and two
lateroventral neurites, which grow from the anterior lobe into
the posterior lobe. These results constitute the first unambiguous
account of the presence of an apical organ with
serotonergic flask-shaped cells in a lecithotrophic brachiopod
larva. Similar apical organs containing flask-shaped cells have
been found in a wide range of lophotrochozoans including
entoprocts (Wanninger et al. 2007), mollusks (Voronezhskaya
et al. 2002; Wanninger and Haszprunar 2003), annelids
(Voronezhskaya et al. 2003), and ectoprocts (Pires and Woollacott
1997; Shimizu et al. 2000). The finding of an apical
organ with serotonergic flask-shaped cells in a lecithotrophic
brachiopod larva suggests that such an apical organ was also
present in the larva of the last common lophotrochozoan ancestor
(Wanninger 2009). Interestingly, such flask cells are
also present in larvae of the demosponge Amphimedon queenslandica,
but whether or not they express serotonin-like
immunoreactivity in this species remains unknown (Sakarya
et al. 2007). Accordingly, it appears that the evolution of
flask-shaped cells in metazoan larvae predated the poriferan–
eumetazoan split, whereby it remains possible that these cells
only acquired serotonin-like immunoreactivity in the lophotrochozoan
lineage. In case of such a scenario, serotoninexpressing
flask cells would be a distinct apomorphy for the
entire Lophotrochozoa.
Similar to the vast majority of lophotrochozoan larvae,
but significantly different to the situation found in the entoproct
creeping-type larva and the larva of polyplacophoran
mollusks, the apical organ of Novocrania anomala is comparatively
simple, thus supporting the notion that a simple apical
organ was present in the ‘‘ur-lophotrochozoan’’ larva,
whereas a complex apical organ is likely to be a synapomorphy
of a monophyletic Entoprocta1Mollusca (Tetraneuralia
concept; see Wanninger 2009).
A serotonergic nervous system has been described previously
for planktotrophic linguliform brachiopod ‘‘paralarvae.’’
There, the apical organ is located at the base of the
median tentacle and comprises numerous serotonergic cells
(Hay-Schmidt 1992). Although it is tempting to speculate that
this neural structure might correspond to the spiralian-type

60 Chapter III
Altenburger and Wanninger
apical organ described herein for Novocrania anomala, it is
important to note (i) that a flask-shaped character could not
be assigned to the apical organ cells of these linguliform paralarvae
and (ii) that the number of cells in their apical organ
is considerably higher than that of the other spiralian larvae.
Overall, the ‘‘apical organ’’ of linguliform larvae resembles
more closely the one found in phoronid and deuterostome
larvae (Santagata 2002), the homology of which remains to be
proven. The suggested derived character of the nervous system
of linguliform brachiopod paralarvae is consistent with
the view that linguliforms have lost the lecithotrophic larva
and have secondarily acquired a planktotrophic life cycle
stage via a stage that resembles a swimming juvenile rather
than a ‘‘true’’ brachiopod larva (Lüter 2001).
We found a-tubulin-positive neural tissue solely in postmetamorphic
specimens of Novocrania anomala. The a-tubulin
signal is located in the same region as the serotonin-like
signal and shows two ventral neurite bundles that are interconnected
by one commissure at the anterior end, one at the
posterior end, and by a median commissure. The fact that we
did not find a-tubulin in the Novocrania anomala larvae that
exhibit serotonergic neurites demonstrates that tubulin
alone is not a reliable marker for nervous structures in
lophotrochozoan larvae. The tubulinergic nervous system in
juvenile Novocrania anomala outlines the adult nervous
system, which consists of two ventral neurite bundles, a subesophageal
and a supraesophageal commissure, and mediodorsal
mantle neurites (Blochmann 1892; Bullock and
Horridge 1965). The anterior ventral neurite bundles form
the arm neurites of the lophophore. Perpendicular from these
arm neurites extend accessory brachial neurites (Williams and
Rowell 1965a, b).
Despite some classical studies, the adult neural anatomy of
brachiopods is only poorly known (James et al. 1992). In the
articulate Gryphus vitreus the nervous system comprises a
transverse supraenteric ganglion and a subenteric ganglion
lying above and below the esophagus, as does the subesophageal
and supraesophageal commissure in Novocrania
anomala (Bullock and Horridge 1965). Our study provides a
first step toward an understanding of the larval anatomy,
neurotransmitter distribution, and development of the nervous
system in brachiopod taxa with lecithotrophic larvae.
Although additional data are needed to assess the brachiopod
neural ground pattern, the finding that serotonergic flaskshaped
cells similar to those found in spiralian larvae do occur
in the apical organ of Novocrania anomala larvae strengthens
the hypothesis that this cell type was also present in the last
common ancestor of Lophotrochozoa (see Wanninger 2009).
Acknowledgments
We are grateful to Matthias Obst and the staff of the Sven Lovén
Centre for Marine Science, Kristineberg, Sweden for help with collection
of adult animals and for providing laboratory space. This
study was funded by the Danish Agency for Science, Technology and
Innovation (grant no. 645-06-0294 to A. W.). Research in the laboratory
of A. W. is further supported by the EU-funded Marie Curie
Network MOLMORPH (contract grant no. MEST-CT-2005-020542).
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L. 1996. A supra-ordinal classification of the Brachiopoda. Proc. R. Soc.
B 351: 1171–1193.
Williams, A., and Rowell, A. J. 1965a. Brachiopod anatomy. In R. C.
Moore (ed.). Treatise on Invertebrate Paleontology, Part H, Brachiopoda.
Geological Society of America and University of Kansas Press, Lawrence,
KS, pp. 6–57.
Williams, A., and Rowell, A. J. 1965b. Evolution and phylogeny. In R. C.
Moore (ed.). Treatise on Invertebrate Paleontology, Part H, Brachiopoda.
The Geological Society of America and The University of Kansas Press,
Lawrence, KS, pp. 164–199.

62 Chapter IV
Chapter IV
Altenburger, A., Martinez, P. & Wanninger, A. First expression
study of homeobox genes in Brachiopoda: the role of Not and Cdx
in bodyplan patterning and germ layer specification. Submitted

Chapter IV
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63
First expression study of homeobox genes
in Brachiopoda: the role of Not and Cdx in
bodyplan patterning, neurogenesis, and germ
layer specification
Andreas Altenburger 1 , Pedro Martinez 2, 3, Andreas Wanninger 1*
1
University of Copenhagen, Department of Biology, Research Group for Comparative Zoology,
Universitetsparken 15, DK-2100 Copenhagen Ø, Denmark
2
Universitat de Barcelona, Facultat de Biología, Departament de Genètica, Av. Diagonal 645,
ES-08028 Barcelona, Spain
3
Institució Catalana de Recerca i Estudis Avançats (ICREA)
*corresponding author. E-mail: awanninger@bio.ku.dk
ABSTRACT
Not is a homeobox containing gene
that regulates the formation of the
notochord in chordates, while Caudal
(Cdx) is a ParaHox gene involved in
the formation of posterior tissues of
various animal phyla. Here, we present
the first expression data of a Not
and a Cdx homolog in the articulate
brachiopod Terebratalia transversa.
The T. transversa homolog, TtrNot,
is expressed in the ectoderm from
the beginning of gastrulation until
completion of larval development,
which is marked by a three-lobed body
with larval setae. Expression starts at
gastrulation in two areas lateral to the
blastopore and subsequently extends
over the animal pole of the gastrula.
With elongation of the gastrula,
expression at the animal pole narrows
to a small band, whereas the areas
lateral to the blastopore shift slightly
towards the future anterior region of
the larva. Upon formation of the three
larval body lobes, TtrNot expressing
cells are present only in the posterior
part of the apical lobe. Expression
ceases entirely at the onset of larval
setae formation. TtrNot expression
is absent in unfertilized eggs, in
INTRODUCTION
Homeobox genes are characterized
by the presence of a short, wellconserved
DNA fragment, which
encodes for the homeodomain. The
latter is a protein motif of 60 to 63
amino acids, which was first described
embryos prior to gastrulation, and in
settled individuals during and after
metamorphosis. Comparison with
the expression patterns of Not genes
in other metazoan phyla suggests
an ancestral role in gastrulation and
germ layer (ectoderm) specification
with co-opted functions in notochord
formation in chordates and left/right
determination in ambulacrarians and
vertebrates. TtrCdx is first expressed
after gastrulation in the ectoderm of the
gastrula in the posterior region of the
blastopore. Its expression stays stable
in the ectoderm at the posterior pole
of the blastopore until the blastopore
is closed. Thereafter, the expression
remains in the ventral portion of the
mantle lobe of the fully developed larva.
No TtrCdx expression is detectable in
the juvenile after metamorphosis. The
expression of TtrCdx is congruent with
findings in other metazoans, were
genes belonging to the Cdx/caudal
family are predominantly localized
posteriorly during gastrulation and
subsequently play a role in the
formation of posterior tissues.
for Drosophila melanogaster homeotic
genes, and subsequently was found
in all animal phyla studied to date
(McGinnis et al. 1984, Scott and
Weiner 1984, Lanfear and Bromham
2008). Homeobox genes function as
developmental control genes that

64 Submitted manuscript
Chapter IV
encode transcription factors which
activate gene cascades (Hueber and
Lohmann 2008). In the case of the
Not gene, which plays an important
role during notochord formation in
vertebrates (Stein and Kessel 1995,
Talbot et al. 1995, Gont et al. 1996, Stein
et al. 1996, Abdelkhalek et al. 2004),
the downstream genes are known to
regulate mesoderm formation in sea
urchins as well as left/right patterning,
notochord, mesoderm, and somite
formation in vertebrates (Peterson et
al. 1999, Yasuo and Lemaire 2001,
Beckers et al. 2007). Homologs of
the homeobox gene Not have been,
among others, identified in Xenopus
(Xnot), chick (Gnot1, Gnot2), zebrafish
(flh), mouse (noto), Hydra (HvuNot),
Drosophila (90Bre), and the basal
eumetazoan Trichoplax adhaerens
(TadNot), but the developmental
role of Not in invertebrates without a
notochord is largely unknown (Dessain
and McGinnis 1993, von Dassow et al.
1993, Knezevic et al. 1995, Odenthal
et al. 1996, Gauchat et al. 2000,
Martinelli and Spring 2004, Hoskins
et al. 2007). A Not homolog seems
to be lacking in the model sponge
Amphimedon queenslandica (Bernard
Degnan, personal communication).
However, the presence of a Not gene
in cnidarians and Trichoplax indicates
that it was present prior to the evolution
of the mesoderm, and thus long
before the evolution of the notochord.
Accordingly, the ancestral role of Not
remains elusive. Given its confirmed
absence in the poriferan genome
would make it a good candidate for a
eumetazoan apomorphy.
Cdx is a member of the ParaHox gene
cluster which probably originated by
duplication from an ancestral ProtoHox
gene cluster which led to the Hox and
ParaHox clusters, respectively (Brooke
et al. 1998). Cdx has been found to be
involved in the development of posterior
tissues of almost all animal phyla in
which it has been investigated and is
thus often termed “caudal” (Epstein et
al. 1997, Copf et al. 2004). In addition
to the posterior tissues, it was found
to be expressed in the mesoderm of
taxa as diverse as Artemia, Capitella,
Patella, Branchiostoma, and Mus;
in the gut of Drosophila, Capitella,
Branchiostoma, and Mus; and in the
central nervous system of Capitella,
Branchiostoma, and Mus (Macdonald
and Struhl 1986, Duprey et al. 1988,
Le Gouar et al. 2003, Copf et al.
2004, Fröbius and Seaver 2006). Cdx
is absent in the recently sequenced
poriferan Amphimedon queenslandica
(Larroux et al. 2008, Srivastava et
al. 2010), but a gene related to Cdx
is present in Nematostella vectensis,
a representative of Cnidaria, the
proposed sister group to Bilateria
(Chourrout et al. 2006, Quiquand et
al. 2009).
The phylogenetic position of
Brachiopoda within Bilateria is still
controversial (Williams and Carlson
2007, Hejnol et al. 2009, Paps et al.
2009). Most authors include them within
Lophotrochozoa, but their position
within this clade remains unresolved,
and some authors consider them a
sister group to Deuterostomia (Nielsen
2002). Within the phylum, Brachiopoda
comprises three clades: Linguliformea,
Craniiformea, and Rhynchonelliformea.
Linguliformea and Craniiformea are
often considered sister groups and
were traditionally termed “inarticulate”,

Chapter IV
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65
because their valves are not connected
to each other by a hinge (Cohen and
Weydmann 2005). We investigated the
brachiopod Terebratalia transversa, a
representative of the rhynchonelliform
(articulate) brachiopods, the largest
group within recent Brachiopoda. So
far, several Hox gene sequences have
been characterized for the linguliform
brachiopod Lingula anatina (de Rosa et
al. 1999). However, no expression data
for any Hox or homeobox containing
genes are currently available for
Brachiopoda. With the investigation of
Not and Cdx expression in Terebratalia
transversa we aim to shed light on the
function of these genes in invertebrate
body patterning and thereby contribute
to the discussion concerning their
ancestral roles in eumetazoan (i.e.,
placozoan, diploblast, and triploblast)
development and evolution.
MATERIAL AND METHODS
Animal collection, rearing, and
fixation
Adult animals were dredged in the vicinity
of the Friday Harbor Laboratories,
Washington, USA, at 48º32’869 N;
122º58’452 W during summer 2008
and spring 2009. The animals were
placed in running seawater tables
at ambient seawater temperature
(approx. 11.5ºC). Embryos were
obtained by artificial fertilization. To
this end, gonads were dissected from
the specimens and stored individually
in beaker glasses. The eggs were
washed several times with seawater
and left in 100ml seawater until
germinal vesicle breakdown, which
usually occurred within 10-16 hours
after dissection. Sperm cells were left
until they had acquired a high degree
of motility, which usually occurred after
4-14 hours. Sperm remained active
until up to 48 hours after dissection.
For fertilization, a few drops of the
sperm suspension were added to the
beaker glasses containing the eggs.
Development of embryos and larvae
was monitored closely and the beaker
glasses were cleaned daily from debris
with help of a glass pipette driven by a
peristaltic pump. Larvae were fixed at
various developmental stages in 4%
paraformaldehyde in 0.5M NaCl, 0.1M
MOPS (pH 7.5) for 8-10 hours at 4ºC,
washed in 50% EtOH for 30 min, and
finally stored in 80% EtOH at -20ºC.
Cloning and in situ hybridization
RNA was extracted from larvae
at various developmental stages
with a miRCURY RNA Isolation Kit
(Exiqon, Vedbaek, Denmark). It was
reversely transcribed into cDNA with
a RETROscript Kit using oligo(dT)
primers (Applied Biosystems/Ambion,
Austin, TX, USA). In order to screen
for homeobox containing genes, the
cDNA was used as template for PCR
reactions with the following degenerate
primers: HoxF 5’-GCT CTA GAR YTN
GAR AAR GAR TT-3’, which recognizes
the peptide sequence ELEKEF, and
HoxR 5’-GGA ATT CRT TYT GRA
ACC ADA TYT T-3’, which recognizes
the peptide sequence KIWFQN
(Murtha et al. 1991; Balavoine and
Telford 1995). PCR was carried out
under the following conditions: 3 min
94 °C, followed by 40 cycles of 45s at
94 °C, 45s at 50 °C, and 60s at 72 °C,
followed by a final extension step of
10 min at 72 °C. PCR products were
purified over column with a QIAquick
Gel Extraction Kit (Qiagen, Venlo, The

66 Submitted manuscript
Chapter IV
Netherlands) and subsequently ligated
into a pGEM-T Easy vector (Promega,
Madison, WI, USA). Ligation products
were transformed into One Shot
TOP10 E. coli competent cells
(Invitrogen, Carlsbad, CA, USA). Cells
were allowed to grow over night; clone
DNA was isolated using a QIAprep
Spin Miniprep Kit (Qiagen). Insert
sequences were sequenced at the
sequencing facility of the University
of Barcelona and identified using
the tBLASTx algorithm. Two specific
forward primers were subsequently
designed from the TtrNot and the
TtrCdx PCR sequences: TtrNotF1 5’-
GGA GAA GGA GTT CGA AAG GCA
ACA A-3’, TtrNotF2 5’-CCG AAT CCC
AAG TGA AGA TCT GGT-3’, TtrCdxF1
5’-CCT GGA GCT GGA GAA GGA
GTT CTG T-3’, and TtrCdxF2 5’-AAC
AAC CTT GTA CTT TCA GAG AGA
CAG G-3’. The specific primers were
used nested in a 3’RACE-PCR using
a SMART RACE kit following the
manufacturer’s protocol (Clontech,
Mountain View, CA, USA). The
sequences of the RACE-PCR products
were again checked by BLAST and the
positive clones were used for in situ
probe production using the DIG RNA
Labeling Kit (SP6/T7, Roche, Basel,
Switzerland).
In situs were done following a standard
protocol with a 5 min proteinase K step
and at least 48 hours of hybridization
time at 40ºC or 45ºC (Martindale
et al. 2004; Hejnol and Martindale
2008). For cohorts aged 0-64 hours
after fertilization (hpf), in situs were
performed on developmental stages
that were 2-4 hours apart, for the age
group of 64-154 hpf, in situs were
done every 5-10 hours, while for later
stages longer intervals were chosen.
The latest stages investigated were
540 hpf old, which corresponded to
420 hours after settlement/onset of
metamorphosis (hps). Sense probes
were generated as controls for in situ
hybridization. Since they didn’t give
any signal they are omitted in the
figures.
Stained specimens were photographed
with a Leica ProgRes C3 digital
camera mounted on a Leica MZ
16F stereomicroscope. Schematic
illustrations were generated using
Adobe Illustrator CS3 and CS4
graphics software (Adobe, San Jose,
CA, USA). Analysis of gene sequences
and primer design was done with CLC
Main Workbench 5 (CLC bio, Aarhus,
Denmark).
Immunostaining and confocal
laserscanning microscopy
Larvae were stained with antibodies
against serotonin (ImmunoStar,
Hudson, WI, USA) and acetylated
α-tubulin (Sigma-Aldrich, St. Louis,
MO, USA). In addition, cell nuclei
were labeled using DAPI (Invitrogen,
Eugene, OR, USA). Prior to staining,
larvae were washed thrice for 15min
each in phosphate buffer (PB) and
incubated for 1h in PB containing
0.2% Triton X-100 (Sigma-Aldrich)
at room temperature. Thereafter,
the larvae were incubated over night
at 4ºC in 6% normal goat serum
in 0.1M PB and 0.2% Triton X-100
(blocking solution). Then, the larvae
were incubated for 24 hours at 4ºC in
blocking solution containing a 1:800
dilution of the polyclonal serotonin
antibody, 3µg/ml DAPI, and a 1:800
dilution of the monoclonal acetylated

Chapter IV
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67
Fig. 1 Characterization of the Not sequence of Terebratalia transversa. (A) Not
homeodomain sequence alignment. The accession numbers for the EMBL/GenBank databases
are given in brackets: Terebratalia transversa (Ttr, brachiopod, XXXXXXX), Nematostella
vectensis (Nve, cnidarian, XP_001641364.1), Hydra vulgaris (Hvu, cnidarian, CAB88387.1),
Trichoplax adhaerens (Tad, placozoan, AAQ82694.1), Drosophila melanogaster (Dme, fruit fly,
NP_650701.1), Strongylocentrotus purpuratus (Spu, sea urchin, AAD20328.1), Hemicentrotus
pulcherrimus (Hpu, sea urchin, BAD91047.1), Branchiostoma floridae (Bfl, Florida lancelet,
XP_002601133.1), Danio rerio (Dre, zebrafish, NP_571130.1), Xenopus laevis (X, frog,
NP_001081625.1). The following alternative species and Hox protein sequences were chosen
as outgroups: Drosophila virilis, Antennapedia (DviAnt, fruit fly, AAQ67266.1), Drosophila
melanogaster, Proboscipedia (DmePb, fruit fly, CAA45272), Neanthes virens, Hox7 and
Engrailed (NviHox7 and NviEng, annelid, DQ366682 and DQ366680). Dots represent amino
acid identity with the amino acid sequence of the T. transversa Not protein shown at the top of
the alignment. (B) Alignment tree based on the 52 amino acid sequences shaded in Fig. 1A.
Algorithm = UPGMA; Bootstrap = 10.000 replicates. Bootstrap values are given for each node.
Due to the small number of residues for the analysis, the phylogenetic signal of the tree is
limited. The tree shows, however, that TtrNot clusters with all other Not protein sequences and
thus is a true Not protein.
α-tubulin antibody. Subsequently,
the larvae were washed four times
over a period of 12h in PB containing
0.2% Triton X-100 and an Alexa Fluor
633-conjugated goat anti-rabbit as
well as an Alexa Fluor 488-conjugated
goat anti-mouse secondary antibody
(Invitrogen) in a dilution of 1:400
for 24h at 4ºC. Finally, the larvae
were washed tree times for 15 min
each in 0.1M PB and embedded in
Flouromount G (Southern Biotech,
Birmingham, AL, USA) on glass
slides. The samples were analyzed
with a Leica TCS SP5 II confocal
system (Leica Microsystems, Wetzlar,
Germany). The resulting image stacks
were merged into maximum projection
images and assembled using Adobe
Photoshop CS3 software (Adobe, San
Jose, CA, USA).

68 Submitted manuscript
Chapter IV
Fig. 2 Characterisation of the Cdx sequence of Terebratalia transversa. (A) Cdx
homeodomain sequence alignment. The accession numbers for the EMBL/GenBank databases
are given in brackets: Terebratalia transversa (Ttr, brachiopod, XXXXXXX), Nematostella
vectensis (Nve, cnidarian, DQ500749), Patella vulgata (Pvu, gastropod, AJ518062.1), Capitella
teleta (Cte, annelid, AAZ95508.1), Drosophila melanogaster (Dme, fruit fly, NM_057606.4),
Tribolium castaneum (Tca, flour beetle, NM_001039409.1), Ciona intestinalis (Cin, tunicate,
NP_001071669), Saccoglossus kowalevskii (Sko, hemichordate, NP_001158415), and Mus
musculus (Mmu, mouse, NM_009880.3). The following alternative species and Hox protein
sequences were chosen as outgroups: Drosophila simulans, Abdominal B (DsiAbdB, fruit
fly, XP_002103136), Drosophila melanogaster, Proboscipedia (DmePb, fruit fly, CAA45272),
Drosophila virilis, Antennapedia (DviAnt, fruit fly, AAQ67266.1), Mus musculus, HoxA9
(MmuHoxA9, mouse, NP_034586.1), Neanthes virens, Engrailed (NviEng, annelid, DQ366680).
Dots represent amino acid identity with the amino acid sequence of the T. transversa Cdx
protein shown at the top of the alignment. (B) Alignment tree based on the 49 amino acid
sequences is shaded in Fig. 2A. Algorithm = UPGMA; Bootstrap = 10.000 replicates. Bootstrap
values are given for each node. Due to the small number of residues used in the analysis, the
phylogenetic signal of the tree is limited. The tree shows, however, that TtrCdx clusters with all
other Cdx/caudal protein sequences and thus is a true Cdx protein.
RESULTS
Characterization of the Terebratalia
Not and Cdx genes
The PCR-amplified region of the TtrNot
homeobox encodes for a 45 amino
acids long peptide which is largely
similar to the NveNot sequence of the
cnidarian Nematostella vectensis (82%
sequence identity). This peptide has
clear affinities to the HvuNot protein
of the cnidarian Hydra vulgaris (69%
sequence identity, Fig. 1). The TtrNot
segment cloned by RACE-PCR was
667 base pairs long and is deposited
in GenBank under the accession
number XXXXXX. The transcribed
region downstream of the homeobox
ends with a poly-A stretch and shows
no similarity to other known gene
sequences.
The conserved protein sequence of the
TtrCdx homeobox identified in this study

Chapter IV
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Fig. 3 Not expression in Terebratalia transversa. Scale bars equal 50 µm, age of specimens
is given in hours after fertilization (hpf) or hours after settlement (hps), respectively. Blue
represents areas of TtrNot expression. (A) Fertilized egg lacking TtrNot expression. (B) 16
cell stage. (C) 32-64 cell stage. (D) Blastula at the onset of gastrulation. TtrNot is expressed
in two fields of cells (arrows) lateral to the future blastopore. (E) Early gastrula in vegetal view
showing the blastopore (asterisk) and two fields of TtrNot expressing cells. (F) Same stage
as in E, lateral view, the blastopore is on the lower side (asterisk). The lateral fields of TtrNot
extend into the animal pole of the gastrula. The ectoderm (ec) and endoderm (en) have started
to form and are demarcated by a dashed line for clarity. (G) The TtrNot expressing cells extend
in a horseshoe-like pattern over the entire gastrula. (H) Lateral view of a late gastrula with
widened archenteron. The TtrNot expressing cells extend over the entire ectoderm (ec) lateral
to the blastopore (asterisk). The lateral fields of TtrNot expressing cells are interconnected via
a small band of TtrNot expressing cells (arrowhead) on the animal pole of the gastrula. TtrNot is
not expressed in the endoderm (en). (I) Slightly elongated gastrula stage, vegetal view showing
the blastopore (asterisk). The position of TtrNot expressing cells has slightly shifted towards
the animal pole, i.e., the future anterior region of the larva. (J) Further elongated gastrula with
blastopore (asterisk) and two fields of TtrNot expressing cells, which are interconnected by a
narrow band of TtrNot expressing cells (arrowhead) in the animal region of the gastrula. (K)

70 Submitted manuscript
Chapter IV
Same stage as in J with animal view onto the future dorso-anterior part of the larva. The two
fields of TtrNot expressing cells which are interconnected by a narrow band of TtrNot expressing
cells (arrowhead) are visible. (L) Elongated gastrula with small blastopore (asterisk). The TtrNot
expressing cells are distributed equally along the posterior part of the future larval apical lobe.
(M) Ventral view of an early three-lobed larva with apical tuft (at), anterior lobe (al), mantle lobe
(ml), and pedicle lobe (pl). The blastopore (asterisk) is closed. The border between ectoderm
(ec) and mesoderm (ms) is visible in the mantle and pedicle lobe. TtrNot expressing cells are
distributed in the posterior part of the apical lobe (indicated by the dashed line). (N) Ventroanterior
view of a larva with all larval lobes fully established: apical lobe (al), mantle lobe (ml),
pedicle lobe (pl). The blastopore (asterisk) is closed and TtrNot expressing cells are distributed
in a ring along the posterior part of the apical lobe. (O) Dorsal view of a fully differentiated
larva with apical lobe (al), mantle lobe (ml), pedicle lobe (pl), and setae (s). TtrNot is no longer
expressed. (P) Posterior view of a juvenile at 360 hours after settlement (hps) with pedicle (p),
anlage of both valves (v) ,and larval setae (s), which extend beyond the valves. TtrNot is not
expressed.
Fig. 4 Cdx expression in
Terebratalia transversa. Scale bars
equal 50 µm, age is given in hours
after fertilization (hpf). (A) Gastrula
stage, TtrCdx (purple) is expressed in
the ectoderm of the posterior pole of
the blastopore (asterisk). (B) Gastrula
stage slightly older than the one in A,
TtrCdx expression on the posterior
side of the blastopore (asterisk) is
more intense than in A. (C) Maximum
projection image of a confocal
reflection scan of a specimen with
similar expression pattern as in B,
showing that TtrCdx is still limited to
the ectoderm. (D) Early larva with
modest expression of TtrCdx (arrow)
in the ventral part of the future mantle
lobe (ml), immediately posterior to
the blastopore (asterisk). The future
apical lobe (al) and posterior lobe (pl)
can already be distinguished. (E) Early
three-lobed larva with apical lobe (al),
mantle lobe (ml), and pedicle lobe
(pl). TtrCdx (arrow) is expressed in
the posterior part of the mantle lobe,
further posterior from the almost closed blastopore (asterisk) than in previous stages. The
larval apical lobe (al), mantle lobe (ml), and pedicle lobe (pl) are further developed. (F) Fully
established larva with expression of TtrCdx (arrow) in the center of the ventral part of the mantle
lobe (ml). Apical lobe (al), mantle lobe, and pedicle lobe (pl) are fully developed and four sets
of larval setae (se) extend from the mantle lobe.

Chapter IV
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shows 73% -80% sequence similarity
with Cdx/caudal protein sequences
known from other metazoans and 61%
sequence similarity with the Cdx/Xlox
sequence of Nematostella vectensis
(Fig. 2). The TtrCdx segment cloned
by RACE PCR was 1037 base pairs
long and is deposited in GeneBank
under the accession number XXXX.
The transcribed region downstream of
the homeobox ends with a poly-A tail
and shows no similarity to other known
gene sequences.
Not and Cdx expression during
Terebratalia development
Cleavage in Terebratalia transversa
is radial. Within eight hours after
fertilization (hpf), the embryo develops
into a blastula. Gastrulation starts at 18
hpf. The spherical gastrula has a central
blastopore until approximately 26 hpf.
Thereafter, the gastrula elongates and
the blastopore narrows to a slit, moves
anteriorly, and closes at around 42 hpf.
At this stage, the elongated larva starts
to develop its characteristic three–
lobed body, consisting of an anterior
lobe, a mantle lobe, and a pedicle
lobe. At around 65 hpf, the larval setae
start to form and larval development is
complete by approximately 80-96 hpf.
Under our experimental conditions,
larvae settled between 120 and 240
hpf. The expression patterns of TtrNot
and TtrCdx are shown in Figs. 3–5.
Fertilized eggs and early cleavage
stages did not reveal the presence of
a TtrNot RNA transcript (Fig. 3A-C).
Expression of TtrNot starts laterally
on both sides of the blastopore in
the early gastrula stage at 18 hpf
(Fig. 3D). At 22 hpf, the expression
of TtrNot increases in the ectoderm
on both sides of the blastopore and
extends towards the animal pole of the
gastrula (Figs. 3E, F, 5A). These lateral
bands of TtrNot expressing cells fuse
in slightly older stages (i.e., at 24 hpf;
Fig. 3G). Shortly after that, at 26 hpf,
when the archenteron is enlarged, the
band of TtrNot expressing cells on the
animal pole narrows (Figs. 3H, 5B). At
28 hpf the blastopore starts to become
slit-like, the gastrula elongates, and
TtrNot expressing cells are present in
two fields lateral of the blastopore and
close to the future anterior pole of the
larva. The fields of TtrNot expressing
cells are interconnected by a narrow
band of TtrNot expressing cells on the
animal side (Figs. 3I-K, 5C). At around
36 hpf the blastopore starts to close,
the larva elongates further, and TtrNot
expressing cells are only detected
in the animal region of the gastrula,
which later forms the larval apical lobe
(Fig. 3L). Subsequently, the blastopore
closes completely and the larva
differentiates the three characteristic
body lobes. TtrNot is continuously
expressed in a ring of cells in the
apical lobe until the beginning of setae
formation. The area of the apical lobe
where TtrNot is expressed corresponds
to the region that bears the cilia used
for swimming of the larva (Figs. 3M,
N, 5D). Once the larval lobes are fully
established and setae formation has
started (i.e., at around 75 hpf), TtrNot
is no longer detectable (Figs. 3O, 5E).
Likewise, specimens that have settled
and started to metamorphose do not
show any TtrNot expression (Figs. 3P,
5F).
TtrCdx starts to be expressed in
the ectoderm of the gastrula at the
posterior pole of the blastopore (Figs.

72 Submitted manuscript
Chapter IV
Fig. 5 Schematic representation of Not and Cdx expression in Terebratalia transversa.
Dark grey – ectoderm, light grey – endoderm. All specimens are approximately 120 µm in
diameter. (A) Almost spherical gastrula at 22 hours after fertilization (hpf). TtrNot is expressed
in two lateral fields of the ectoderm. TtrCdx is expressed on the posterior side of the blastopore.
(B) Gastrula at 26 hpf. The two lateral fields of TtrNot expressing cells are interconnected by
a narrow band of TtrNot expressing cells on the animal side of the gastrula. TtrCdx is more
intensely expressed posterior to the blastopore. (C) Early larva at 30 hpf. The fields of TtrNot
expressing cells are positioned further towards the animal pole of the gastrula than in previous
stages. The blastopore is still open. TtrCdx is expressed posterior to the blastopore. (D) Larva
at the onset of lobe formation at 42 hpf. TtrNot is solely expressed in the posterior part of the
apical lobe. Locomotory cilia and an apical tuft are already present. TtrCdx is expressed in the
postero-ventral part of the mantle lobe. (E) Three-lobed larva at 75 hpf. TtrNot is no longer
expressed but TtrCdx is still present in the posterior part of the mantle lobe. (F) Juvenile at
360 hours after settlement. The lophophore has started to develop between the valves. Neither
TtrNot nor TtrCdx are expressed.
4A, 5A). TtrCdx expression remains
in this position in the larval ectoderm
throughout development. Expression
intensifies as the gastrula gets older
(Figs. 4B, C, 5B). In early larvae prior
to the onset of lobe formation (Figs.
4D, 5C), early three-lobed larvae (Figs.
4E, 5D), and fully developed larvae
(Figs. 4F, 5E) TtrCdx is expressed in
the ventral ectoderm posterior to the
closed blastopore.
Neither the expression of TtrNot nor
the expression of TtrCdx is co-located
with the larval serotonergic nervous
system of Terebratalia transversa,
which comprises an apical organ with
eight flask-shaped serotonergic cells
that lie antero-dorsally in the apical
lobe (Fig. 5). The flask-shaped cells
are connected via individual neurites
to a larval neuropil that is situated in
the center of the apical lobe. (Fig. 5A,

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73
Fig. 6 Larval
serotonergic nervous
system of Terebratalia
transversa.
Maximum projections of
a confocal microscopy
image stack. Serotonin
is labeled red, tubulin
is green, cell nuclei are
blue. (A) Lateral view
of a fully established
larva with apical lobe
(al), mantle lobe (ml), and pedicle lobe (pl). The apical organ comprises eight flask-shaped
cells (arrows) which are connected by neurites (empty arrowheads) to a larval anterior neuropil
(asterisk). The apical organ is situated towards the dorsal side of the apical lobe. Note that
only cilia but no neural structures are labeled by the α-tubulin antibody. (B) Detailed view of
the apical organ of the specimen shown in A. The apical organ comprises two sets of four
flask-shaped cells each (arrowheads). The flask-shaped cells are connected to the neuropil
(asterisk) by individual neurites (empty arrowheads).
B). Staining with the pan-neural marker
anti-acetylated tubulin did not reveal
any additional neural structures.
DISCUSSION
The role of the Not gene in metazoan
neurogenesis
The designation of the gene “Not”
refers to the site where its expression
was detected for the first time, namely
in the notochord of the African Clawed
Frog, Xenopus laevis (Gont et al.
1993, von Dassow et al. 1993). The
notochord is a cartilaginous, rod
shaped structure, apomorphic to the
Chordata. It is at least present in some
developmental stages of all chordates,
including the ascidian tadpole larva,
and functions as axial skeleton,
induces the development of the
neural tube, and is thus a key player
in chordate neurogenesis (Stemple
2005). Moreover, Not is expressed in
the neural tube of the mouse, chick,
frog, and zebrafish (Stein and Kessel
1995, Talbot et al. 1995, Yasuo and
Lemaire 2001, Beckers et al. 2007).
Where it has been analyzed in detail,
Not seems to act primarily as a
transcriptional repressor (Yasuo and
Lemaire 2001). In zebrafish, loss-offunction
mutants of floating head (flh;
the zebrafish Not homolog) lack the
notochord altogether, and the somites
fuse below the neural tube (Talbot et al.
1995). Expression studies suggest that
cells lacking flh expression differentiate
into muscle rather than notochordal
tissue (Halpern et al. 1995). Noto,
the Not homolog in the mouse, and
flh repress paraxial mesoderm fate
while maintaining axial mesoderm fate
(Amacher and Kimmel 1998).
Only little is known about the
expression patterns and functions of
Not in non-chordate metazoans. In
Trichoplax adhaerens, the Not gene is
expressed at the bottom of body folds
of intact animals as well as during
wound healing (Martinelli and Spring

74 Submitted manuscript
Chapter IV
2004). In addition, Not expressing cells
in this species, which lacks a nervous
system and even neurons, overlaps
with the site of expression of the
neurotransmitter RFamide (Schuchert
1993). In Drosophila, the Not homolog
90Bre is present in the nervous
system, where it is expressed in the
ventral nerve cord and the posterior
brain anlage of germ band retracted
embryos, as well as in the lateral/
posterior region of the eye/antennal
disc (Dessain and McGinnis 1993).
In the ascidians Halocynthia roretzi
and Ciona intestinalis, Hr-Not and Ci-
Not, respectively, are expressed in the
posterior end of the tail, as well as in
the notochord and a small part of the
anterior neural tube in the larval tailbud
stage (Utsumi et al. 2004). These data
hint towards an ancestral role of Not in
metazoan neurogenesis.
The ring-like expression of TtrNot in
the ciliated region of the apical lobe
in Terebratalia larvae is intriguing.
Noto, the Not ortholog in the mouse,
is known to function in ciliogenesis,
and TtrNot might thus serve a similar
role in our study species. In this
context,it should also be considered
that the putative spiralian homolog
of this larval swimming device, the
prototroch, is underlain, and probably
innervated, by a ring nerve (Wanninger
2009). Accordingly, it is tempting
to speculate that Not may also be
expressed in the prototroch ring nerve
of spiralian trochophore larvae and/
or the prototroch itself. In Terebratalia
transversa, however, we did not find
any corresponding neural structure in
the region of Not expression (Fig. 6).
The larval nervous system of T.
transversa differs in several details
from that of the craniiform brachiopod
Novocrania anomola. In N. anomala,
the apical organ comprises four,
centrally positioned serotonergic flaskshaped
cells that are connected to two
ventral neurites which elongate laterally
along the larval body (Altenburger
and Wanninger 2010). By contrast,
the apical organ of T. transversa has
two sets of flask-shaped cells. Each
set contains four cells and each cell is
connected to the larval anterior neuropil
by a single serotonergic neurite. Due to
these differences, the morphology of
the ancestral brachiopod larval apical
organ remains elusive. However, the
data currently available indicate that an
apical organ comprising serotonergic
flask-shaped cells was part of the
brachiopod ground pattern and most
likely constitutes a morphological
apomorphy of Lophotrochzoa, since
such cells are also found in larval
Entoprocta, Mollusca, Nemertea,
Annelida, and Ectoprocta (Pires and
Woollacott 1997, Shimizu et al. 2000,
Friedrich et al. 2002, Voronezhskaya
et al. 2002, 2003, McDougall et al.
2006, Wanninger et al. 2007, Fuchs
and Wanninger 2008, Chernyshev
and Magarlamov 2010, Nielsen and
Worsaae 2010).
Not expression during gastrulation
and germ layer formation
In all species studied so far, Not
expression starts prior to or at the onset
of gastrulation. This is also the case
in the brachiopod investigated herein,
Terebratalia transversa. In the sea
urchin Strongylocentrotus purpuratus,
Not is expressed in the vegetal plate at
the mesenchyme-blastula stage and
in the secondary mesenchyme, with

Chapter IV
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75
expression ceasing after gastrulation
(Peterson et al. 1999). In ascidians,
Not expression starts at the eight
cell stage in all blastomeres and is
thereafter expressed in the posterior
part of the larval tail, the notochord,
and a small part of the anterior neural
tube at the tailbud stage (Utsumi
et al. 2004). Interestingly, we found
TtrNot being solely expressed in the
ectoderm of T. transversa, while their
homologs are expressed in all three
germ layers during Xenopus and
ascidian embryogenesis (von Dassow
et al. 1993, Utsumi et al. 2004).
Apart from the development of the
nervous system, the notochord,
and various germ layers, Not is also
responsible for left/right patterning
in the mouse, where it is expressed
in the “node”, i.e., the organizer of
gastrulation (Beckers et al. 2007).
In the sea urchins Hemicentrotus
pulcherimus and Strongylocentrotus
purpuratus, Not is expressed in the
archenteron of the gastrula and in
the mesoderm of the right coelomic
pouch of two-armed pluteus larvae,
were it is likewise involved in left/right
determination (Peterson et al. 1999,
Hibino et al. 2006).
The current data suggest an overall role
of Not in gastrulation as well as germ
layer and nervous system patterning.
Whether Not was used in specification
of all three germ layers in Urbilateria
(as exemplified in the ascidians and
Xenopus) or whether its ancestral role
was in ectoderm patterning alone (as
in Terebratalia) remains to be revealed
by future comparative studies. In
any case, it appears that the Not
gene has been co-opted into several
other functions during evolution of
respective metazoan (deuterostome)
lineages, such as notochord formation
in chordates and left/right patterning
in ambulacrarians (sea urchin) and
vertebrates (mouse).
The role of Cdx in metazoan
development
Cdx is a member of the ParaHox cluster
in which three genes are linked in a
manner reminiscent of the Hox genes,
with the gene order 3’-Gsx-Xlox-Cdx-5’
(Brooke et al. 1998). Compared to Hox
genes, ParaHox genes seem to be
much more evolutionary labile, since
they do not appear together in all
species investigated and sometimes
they are not clustered (Ferrier and
Holland 2002).
Cdx expression patterns are known
from several animal phyla and there is
a wide range of tissues in which Cdx is
expressed (Fröbius and Seaver 2006).
A gene related to Cdx is present in
the proposed bilaterian sister group,
the cnidarian Nematostella vectensis
(Chourrout et al. 2006, Ryan et al. 2006,
2007). Cdx was first characterized as a
posterior patterning gene in Drosophila
melanogaster (Mlodzik et al. 1985)
and it appears to serve a similar role
in a number of other taxa including
various arthropods, the nematode
Caenorhabditis elegans, and the basal
gastropod mollusk Patella vulgata
(Waring and Kenyon 1991, Xu et al.
1994, Schulz et al. 1998, Abzhanov
and Kaufman 2000, Dearden and
Akam 2001, Rabet et al. 2001, Copf et
al. 2003, Le Gouar et al. 2003, 2004,
Shinmyo et al. 2005, Olesnicky et al.
2006). In the annelids Platynereis
dumerilii, Nereis virens, Tubifex
tubifex, and Capitella sp., Cdx has an

76 Submitted manuscript
Chapter IV
anterior and a posterior expression
domain (Fröbius and Seaver 2006,
Matsuo and Shimizu 2006, Kulakova
et al. 2008, Hui et al. 2009).
The expression pattern of Cdx in
Terebratalia transversa shows some
similarity to that of Platynereis dumerilii.
In both species Cdx is expressed in the
ectoderm at the onset of gastrulation. In
P. dumerilii, the ectodermal expression
of PduCdx encircles the posterior
portion of the slit-like blastopore and
extends from there anteriorly along its
edges (de Rosa et al. 2005). PduCdx
continues to be expressed in the
posterior part of the trochophore larva
in the posterior midgut and hindgut
(Hui et al. 2009). Expression of Cdx in
the gut is also found in the sea urchin
Strongylocentrotus purpuratus, the
lancelet Brachiostoma floridae, and
the mouse Mus musculus (Duprey
et al. 1988, Brooke et al. 1998,
Arnone et al. 2006). We did not find
expression of TtrCdx in the larval gut
of Terebratalia transversa, which might
be due to the fact that those larvae are
lecithotrophic and that metamorphosis
is catastrophic, i.e., that all major larval
tissues degenerate after settlement
(Stricker and Reed 1985a, 1985b).
Since we did not investigate feeding
juveniles with a functional gut, the role
of TtrCdx in gut formation remains
elusive.
Concerning the role of Cdx in the
protostome-deuterostome ancestor
(PDA), two major hypotheses are
currently discussed. Either, expression
in the PDA might have been in an
anterior and a posterior domain of the
nervous system, as in recent acoels
as well as the lophotrochozoans
Platynereis dumerilii, Capitella sp.,
Tubifex tubifex, and Patella vulgata
(Le Gouar et al. 2003, de Rosa et al.
2005, Matsuo et al. 2005, Fröbius and
Seaver 2006, Hejnol and Martindale
2008). Expression in the hindgut and
posterior tissues of recent animals
would thus have been co-opted. Or,
Cdx expression in the PDA was in
posterior tissues and a dissociation
of Cdx from the ParaHox cluster in
Lophotrochozoa allowed for its cooption
into the anterior domain of the
nervous system, as is the case in
acoels and some lophotrochozoans
(Hui et al. 2009). In this respect it
would be interesting to focus future
investigations on the genomic
arrangement and the expression of
the respective Gsx and Xlox genes of
the ParaHox cluster in brachiopods.
ACKNOWLEDGEMENTS
We thank the Friday Harbor
Laboratories and especially Billie
Swalla (University of Washington) for
providing lab space and assistance in
rearing Terebratalia transversa. Olga
Lévai (Leica Microsystems, Mannheim,
Germany) is thanked for providing the
SP5 confocal system that was used for
the scans upon which Fig. 6 is based.
We are grateful to Marta Chiodin
(University of Barcelona) for guidance
in lab procedures. Bernard M. Degnan
(University of Queensland) is thanked
for sharing previously unpublished
sequence data of the demosponge
Amphimedon queenslandica. This
study was funded by the Danish
Agency for Science, Technology and
Innovation (grant no. 645-06-0294
to AW). Research in the lab of AW
and PM was further supported by
the EU-funded Marie Curie Network

Chapter IV
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77
MOLMORPH (contract grant number
MEST-CT-2005-020542). PM is
grateful to the Spanish Ministerio
de Ciencia e Innovación and the
Generalitat de Catalunya for financial
support.
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