Integrative and Comparative Biology
Integrative and Comparative Biology, volume 53, number 1, pp. 103–117
doi:10.1093/icb/ict001
Society for Integrative and Comparative Biology
SYMPOSIUM
Metazoan Circadian Rhythm: Toward an Understanding of a
Light-Based Zeitgeber in Sponges
Werner E. G. Müller,1,* Heinz C. Schröder,* Dario Pisignano,†,‡ Julia S. Markl* and
Xiaohong Wang2,*,§
*ERC Advanced Investigator Grant Research Group at Institute for Physiological Chemistry, University Medical Center of
the Johannes Gutenberg University, Duesbergweg 6, D-55128 Mainz, Germany; †Dipartimento di Matematica e Fisica
‘‘Ennio De Giorgi’’ and National Nanotechnology Laboratory of Istituto Nanoscienze-CNR, Università del Salento, via
Arnesano, I-73100 Lecce, Italy; ‡Center for Biomolecular Nanotechnologies, Istituto Italiano di Tecnologia (I.I.T.), via
Barsanti 1, I-73010 Arnesano-LE, Italy; §National Research Center for Geoanalysis, 26 Baiwanzhuang Dajie, CN-100037
Beijing, China
From the symposium ‘‘Keeping Time During Animal Evolution: Conservation and Innovation of the Circadian Clock’’
presented at the annual meeting of the Society for Integrative and Comparative Biology, January 3–7, 2013 at San
Francisco, California.
1
2
E-mail: [email protected]
E-mail: [email protected]
Synopsis In all eukaryotes, the 24-h periodicity in the environment contributed to the evolution of the molecular
circadian clock. We studied some elements of a postulated circadian clock circuit in the lowest metazoans, the siliceous
sponges. First, we identified in the demosponge Suberites domuncula the enzyme luciferase that generates photons. Then
(most likely), the photons generated by luciferase are transmitted via the biosilica glass skeleton of the sponges and are
finally harvested by cryptochrome in the same individual; hence, cryptochrome is acting as a photosensor. This
information-transduction system, generation of light (luciferase), photon transmission (through the siliceous spicules),
and photon reception (cryptochrome), all occur in the same individual. Therefore, we propose that this photoreception/
phototransduction process might function as a nerve-cell-like signal transmitting system. This was corroborated by the
fact that S. domuncula reacts to different wavelengths of light, originating from the sponge environment, with a differential gene expression of the transcription factor SOX. Recently, we succeeded in demonstrating that in sponges a light/
dark controlled gene is expressed, which encodes for nocturnin, a protein showing poly(A)-specific 30 -exoribonuclease
activity. Quantitative real-time polymerase chain reaction analyses revealed that primmorphs, 3D cell aggregates of sponge
cells, after transfer from light to dark, show a 10-fold increased expression of the nocturnin gene. In contrast, the
expression level of the gene encoding glycogenin decreases in the dark by three- to four-fold. It is concluded that
sponges are provided with the molecular circadian clock protein nocturnin which is highly expressed in the dark.
This finding together with the proposed light-transduction and spicule-based signaling system strongly supports the
view that already the lowest metazoans, the sponges, have elements of a circadian rhythm, characteristic of higher
metazoans.
Introduction
All organisms on earth are under the steady influence
of environmental factors, e.g. daily or seasonal environmental changes, resulting from the planet’s rotation around its own axis and the sun. It is evident
that these changes are driven by physical factors, e.g.
light intensity, airflow, water currents, or food
supply. It was first proposed in 1995, based on
molecular biological data, that sponges evolved first
from a common ancestor of all metazoan phyla
(Müller 1995), from the Urmetazoa, about
600–1000 million years ago (reviewed by Müller
et al. 2007a and Wang et al. 2010). This view was
substantiated multifold (e.g., Harcet et al. 2010).
Advanced Access publication March 8, 2013
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Careful studies of the different metabolic pathways in
sponges revealed that these multicellular animals already comprise all basic structural and functional
systems known from higher metazoan taxa, with
the exception of the nervous and contractile systems
(Müller et al. 2004).
It is well established that in mammals there is a
circadian system that is composed of many individual (tissue-specific) clocks. To generate an integrated
physiological and behavioral response, the phases of
these different clocks are finely tuned by a master
circadian pacemaker residing in the suprachiasmatic
nuclei of the brain. This central circadian clockwork
consists of an interconnected system of positive and
negative feedback loops, with the dictating positive
limb involving two basic helix–loop–helix transcriptional activators CLOCK/BMAL1 that, after heterodimerization, bind to the E-boxes located in the
regulatory region of the period (per) and cryptochrome (cry) genes (reviewed by Albrecht and
Eichele 2003; Ko and Takahashi 2006). The corresponding gene products, cryptochrome and period
proteins, are translocated into the nucleus where
they repress their own transcriptional expression
via inhibition of transcription of CLOCK/BMAL1.
In addition, the interconnected positive and negative
limbs are controlled by the nuclear orphan receptor
REV-ERB, which in turn represses the transcription
of Bmal1 through direct binding to the REV-ERB
response element within the Bmal1 promoter (Huang
et al. 2012). The central clock is synchronized to geophysical time mainly via photic cues perceived by
photosensory organs, e.g. the retina, from where the
electrical signals are transmitted to the central neurons
which in turn influence circadian physiology and behavior. This network emphasizes that neuronal and
humoral cues dictate the synchronization of both central and local oscillators operating in most organs and
tissues (reviewed by Kulczykowska et al. 2010).
Light in the darkness of the aquatic
environment
Luminous planktonic bacteria frequently establish a
symbiosis with fish, squid, and other organisms in
the aquatic/marine environment (Harvey 1921; Belas
et al. 1982) (Fig. 1A). Bacteria (Vibrio sp.) settle on a
fish kept for 1–2 days in an open environment.
Frequently, growing colonies of luminous bacteria
can be photographed by their own light (Fig. 1A).
This visible light is produced by living organisms and
hence is termed bioluminescence, in contrast to
luminescence emitted by a nonliving substance
(Haddock et al. 2010). Bioluminescence and also
W. E. G. Müller et al.
Fig. 1 Bioluminescence (A and B) and biofluorescence in aquatic
organisms (E). (A) Bacteria growing on a fish in the air during a
1- to 2-day period. (A-a) Daylight image. (A-b) Dark image. Right
panel: a mask layered on top of the bacterial colony prevents
bacteria-generated light (logo of the University Mainz). (B)
Generation of light (bioluminescent flashing) by tissue from S.
domuncula; (B-a) A cube of a dark adapted animal exposed to the
detection film. (B-b) The emitted light from a chemiluminescent
detection film. A dark spot on the film was resolved following
development of the X-ray film. (C) Underwater image showing
surface light as seen from the bottom of a tropical sea at a depth of
300 m; the light can be resolved into different qualities of blue. (D)
Diving for the Baikalian sponge L. baicalensis, living under an ice
cover 1–2 m thick. (E) Biofluorescence. (E-a) A photograph of L.
baicalensis taken with a daylight flash and showing sponges with
different color variations from green to whitish, dependent on the
amount of chlorophyll contained in their algal symbionts. (E-b)
Illumination of the same specimens in the dark. Under that condition the biofluorescence residing in the oscules can be monitored; a protruding lobe of the crust is marked (45). The signals
emanating from these areas probably originate from symbiotic
microorganisms. Partially modified from Wang et al. (2012b).
biofluorescence (emission of light by an organic template that has absorbed light from the environment)
are much more frequently seen in the marine
environment than in freshwater. A well-established
biofluorescence is seen in the Lake Baikal sponge
Lubomirskia baicalensis (Wiens et al. 2009) (Fig. 1E).
This endemic sponge lives in a symbiotic relationship with dinoflagellates (Müller et al. 2006a), an interaction that allows the sponges to survive for
105
Universal zeitgebers in sponges
6 months of the year under adverse situations under
the ice cover. Diving expeditions (Fig. 1D) revealed
that most of these sponges are colored a deep green
by the chlorophyll residing in the dinoflagellates.
Illuminating the sponges with daylight shows their
green color (Fig. 1E-a). However, if they are illuminated with blue light, some of the specimens have a
green fluorescence (Fig. 1E-b) surely attributable to
the accumulation of chlorophyll, and perhaps functioning in attracting microorganisms, especially
around the oscule region (Wilson and Hastings 1998).
The phenomenon of bioluminescence occurs across
a broad range of major groups of organisms from bacteria and protists to squid and fishes (Haddock et al.
2010). The luminescence described in many organisms
such as sponges, bryozoans, and salps was considered
doubtful (Herring 1987), until a luciferase-based
system of generating light was recently identified in
sponges (Müller et al. 2009a; also see below). In the
ocean, bioluminescence is a very common phenomenon in prokaryotic and eukaryotic taxa ranging in size
from microns to meters, and including autotrophs,
herbivores, and carnivores (Herring 1987). Bioluminescence in those organisms has been assumed to attract commensal microorganisms or repel attackers.
As outlined below, there are experimental data indicating that the spicules in sponges indeed act as optical
fibers. In the ocean, blue light can penetrate nearly
1000 m; at a depth of 300–400 m the relief of the
seabed can still be seen because of different intensities
of the blue color (Fig. 1C).
The ‘‘nervous’’-like spicule-based signaltransduction system in sponges
While most metazoans have a hard skeleton based on
calcium phosphates or calcium carbonates, most of
the sponges (Hexactinellida and Demospongia), as
well as diatoms, use amorphous silica as an inorganic
polymer to build their skeletal elements. The selection of this inorganic component by sponges, the
oldest still extant metazoans, is not surprising since
in the ancient ocean, in which the sponges emerged,
was a ‘‘soda ocean’’ with a probable pH value of
above 9. Under such conditions, the concentration
of silica, the dioxide form of silicon, in seawater was
presumably higher than it is today. Sponges have the
simplest body plan (Müller et al. 2004), but the formation and structure of their biomineral skeleton are
already highly complex. They form their silica, better
termed bio-silica, enzymatically using silicatein
(Shimizu et al. 1998; Cha et al. 1999; Krasko et al.
2000). Recently, Wang et al. (2012a) summarized the
different hierarchical levels that are involved in the
formation of the siliceous spicules, from the genetic,
via the cellular, to the biological one. As expected,
but not previously proven experimentally, the construction/architecture of the sponge skeleton is genetically controlled and functions as a crucial
element in morphogenetic processes (Müller 2006).
The skeletal framework of sponges is highly ordered,
as seen in the demosponge L. baicalensis (Müller
et al. 2006c) and the hexactinellid Monorhaphis
chuni (Wang et al. 2009). Most siliceous sponges
are composed of larger megascleres (410 mm long)
and smaller microscleres (510 mm).
Spicules as light waveguides
Sponges are devoid of a nervous system. We were the
first to propose that, besides stabilization, the biological function of the siliceous spicules is to act as
optical fibers substituting for a nervous system
(Müller et al. 2006b). It is amazing that during the
long evolutionary history of sponges, their skeletal
elements, the siliceous spicules, which are composed
of quartz glass (Müller et al. 2008), retained the property of operating as light waveguides (Cattaneo-Vietti
et al. 1996; Aizenberg et al. 2004; Müller et al. 2006b;
Wang et al. 2010). This property has been studied in
several hexactinellids and demosponges (CattaneoVietti et al. 1996; Müller et al. 2006b, 2009a) and
it was proposed that light generated in vivo might
be transmitted through the spicules and subsequently
converted into electrical signals via photoelectric
reactions (Müller et al. 2006b).
For our initial studies, we selected the hexactinellid sponge Hyalonema sieboldi that has extremely
long spicules forming its stalk (Müller et al.
2006b). Later, the same function was also proposed
for the hexactinellid M. chuni (Wang et al. 2007).
The first surprising result was that light transmitted
through the spicules is cut off below 600 nm and
above 1310 nm in a similar manner as that known
for combined high-/low-pass filters (Müller et al.
2006b; Wang et al. 2007). From comprehensive studies of luminous genera, of organisms capable of
emitting light through their bioluminescent systems,
it is known that most pelagic deep-sea organisms
(Fig. 1C) emit light with the maximum of emission
spectra about 480 nm, while terrestrial or freshwater
forms produce light with longer wavelengths
(Haddock et al. 2010).
Organization of spicules within the sponge body—the
light-harvesting spicules
The organization of the spicules within the organism
is genetically well fixed. The spicule-based skeleton of
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sponges is architecturally complex and arranged
in a functionally efficient way. As outlined in the
Challenger Report (Ridley and Dendy 1887), two
main types of spicule arrangements occur in demosponges, the ‘‘Reticulate’’ and the ‘‘Radiate.’’ The
arrangement of spicules follows a genetically determined pattern which is either radial or spiral. The
spiral organization is characteristic, for example, of
the Tethyidae (Fig. 2A [top]) and the radial skeleton,
of the Suberitidae (Fig. 2A [bottom]). In our studies,
we focused on specimens of Suberites domuncula
(Fig. 2B). The advantage of using this sponge is
W. E. G. Müller et al.
that most ESTs have been elaborated from this species (SpongeBase 2010); in addition, it can be kept in
aquaria under controlled conditions (LePennec et al.
2003) and its 3D aggregate culture system, the
‘‘primmorphs,’’ has been elaborated (Müller et al.
1999). Moreover, there is (largely) only one spicule
type that exists in this species, the tylostyles (Fig. 2C,
F, and G). The monaxonal tylostyles (150–320 mm in
length) comprise a central siliceous stalk that is terminated at one end with a globular knob with diameters between 6.53 and 7.28 mm (in the longitudinal
direction of the spicule) and between 8.54 and
Fig. 2 Spicules in siliceous sponges. (A) Network of interacting spicules. Schematic representation of the skeletal organization in
siliceous sponges (Demospongiae). ‘‘Radiate’’ pattern of the spiral type (top) and of the radial type (bottom); the spicules are arranged
in bundles termed ‘‘radiating skeletal fibers’’ (rf). Between the fibers, tissue is organized and harbors the aquiferous canal system. At the
surface of the animals, the skeletal fiber bundles are spliced into the individual cortical spicules (cs); from Ridley and Dendy (1887). (B)
A specimen of S. domuncula living in an aquarium. (C) An isolated tylostyle (S. domuncula) having one spheroidal ball-like terminus and a
spine at the other terminus (size: 0.02 cm). The spicule is illuminated by a light source (l) and allows the light to be transmitted through
its central axis. (D) A giant basal spicule (about 300 cm long) acting as a light waveguide in the hexactinellid M. chuni. The light source is
marked (l). (E) Schematic outline of the arrangement of spicules within the surface of G. cydonium, comprising different layers of
spiculae: the ectochrote (ec), sterraster layer (ster) with microscleres (mic), and the fibrous subcortical crypts (fib) contain many large
megascleres (meg). (F and G) The surfaces of the sponges S. domuncula show protruding spicules (sp), tylostyles that expose their
spherical/elliptical knobs to the external environment, while the tips remain inside. Partially modified from Müller et al. (2007a).
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Universal zeitgebers in sponges
9.21 mm
(in
the
perpendicular
direction).
Illumination of the spicules toward the globular
end reveals that these knobs act as converging
lenses (Fig. 2C) perhaps focusing the light beam. It
might be stressed that the tylostyles from S. domuncula protrude to the external space and in that position can channel light waves to the interior of the
animal. It is striking that these spicules are localized
in a highly ordered pattern, immediately below the
surface cell layers, in palisade-like arrays (Müller
et al. 2010). The orientation of tylostyles is such
that the pointed rods are directed toward the
center of the animal, the medulla, whereas the
knobs are directed toward the surface (Fig. 2F and
G). This directionality might imply light guidance
from the outside into the sponge’s body. This relates
to the light conditions of the habitat; the animals are
found exclusively in shallow water (20–30 m). In
coastal waters, light within a short range of wavelengths (about 500 nm) is preferentially transmitted,
compared with offshore oceanic water. Interestingly,
this range of wavelengths corresponds to the bioluminescence emission spectrum, emitted by the
luciferase of S. domuncula. Hence, it can be deduced
that spicules, exposed at the surface of the sponge,
absorb/harvest and transmit light to a cryptochromecontaining photon-harvesting system.
Other species of sponge, e.g. Geodia cydonium,
have a more complex arrangement of their spicules
(Müller et al. 2007b) (Fig. 2E). Different types of
spicules are arranged in a highly organized pattern
within the body. In cross-section (Fig. 2E), the globular specimens of G. cydonium can reach diameters
of 20 cm, forming a cortex (ectosome) 4–8 mm thick
and which surrounds the central body. The cortex
comprises three regions: (1) the thin ectochrote
with numerous oxyasters and fine ectosomal spicules,
(2) the sterrastral layer that is almost totally filled
with microscleres, and (3) the fibrous layer containing only a few sterrasters. The choanosome, which
follows the subcortical crypts, contains masses of
large oxeas and triaenes that are oriented concentrically in bundles toward the center of the animal. The
scheme in Fig. 2E shows the intimate contacts between the spicules, an arrangement suggesting that
the surface spicules, besides being a mechanical defense against predators, might act as light collectors
that transmit light into the more central regions of
the sponge. Subsequently, this suggestion was proven
for the demosponge Tethya aurantium (Brümmer
et al. 2008). In hexactinellids, e.g. Euplectella aspergillum, the interaction between the spicules is even
more obvious. This species’ spicules measure larger
average sizes (Leys et al. 2007) and often even fuse
into a continuous super-scaffold (Müller et al. 2009b).
The most striking examples of sponge spicules functioning as a light waveguide are the giant basal spicules
of M. chuni (Wang et al. 2009) (Fig. 2D).
Experimental data show that sponges can react to
light originating from their environment. The following paragraph indicates that these animals also can
generate endogenously light via luciferase. Based on
this property, we propose that the sponge can—
within the same individual—generate light through
the luciferase, which is transmitted through the spicules and finally harvested at the end of the spicule by
a cryptochrome. It is proposed that this signaling
pathway adjusts the endogenous rhythm.
Light-generating machinery: luciferase–luciferin
Bioluminescence, a chemiluminescence reaction, is a
widespread phenomenon in nature, a process during
which two molecules react within an organism with
the emission of light. Intensive research started with
the elucidation of the luciferin–luciferase reaction in
the firefly Photinus pyralis (Strehler and McElroy
1949). Later, molecular biological studies revealed
that luciferase is a generic name and includes a
series of different classes of luciferase molecules.
Luciferins have been identified in bacteria (a reduced
riboflavin phosphate, FMNH2, that undergoes oxidation by luciferase in association with long chain aldehyde and oxygen molecules), in dinoflagellates
(it is conformationally shielded from luciferase at
the basic pH of 8 but is set free and accessible to
oxidation at the more acidic pH of 6), in the marine
ostracod Vargula (it is acquired by ingestion of bacteria), in coelenterates (its activity is controlled by
the concentration of Ca2þ and the molecule shares
homology with the calcium-binding protein calmodulin), and in insects (fireflies, glowworms, and click
beetles; in fireflies [Photinus or Luciola] luciferin, a
benzothiazole, has the unique property of requiring
ATP as a co-factor to convert into an active molecule; reviewed in Meighen [1991] and Greer and
Szalay [2002]).
The first evidence that S. domuncula contains a
light-generating system was obtained by exposing
tissue slices from a dark-adapted animal to light.
When a tissue sample was applied to a chemiluminescent detection film for 24 h, a clear signal could
be detected after development with the chemiluminescent substrate (Fig. 1B) (Müller et al. 2009a). As a
control, tissue from an animal pretreated with rotenone was exposed to the film. Under otherwise
identical conditions, this tissue caused no signal on
the film. In order to identify the luciferase in
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S. domuncula, our EST database was screened with
the main focus on the polypeptides that comprise
the characteristic domains for metazoan luciferase
(Müller et al. 2009a), e.g., the acetyl-coenzyme A
synthetase region and the luciferin-binding site residues. The database that comprises 30,000 ESTs can
be considered to cover most of the genes in these
animals. Indeed, one fragment was found that was
similar in sequence to firefly luciferase. Firefly luciferase converts the substrate luciferin, a heterocyclic
carboxylic acid, and ATP into the corresponding
luciferyl adenylate and (with the consumption of
molecular oxygen) into a product with an electronically excited state. That product emits a photon of
visible light (Branchini et al. 2007). The binding site
for luciferin on luciferase has been studied in detail
in P. pyralis (Branchini et al. 2007). In Luciola cruciata, the dehydroluciferin moiety interacts in the
hydrophobic pocket of the luciferase with the regions
consisting of -8 (amino acid residues), b-12, b-13,
b-14, b-15, and an additional loop. During the flashing reaction, luciferin is converted by monooxygenase luciferase to oxyluciferin (Gomi and Kajiyama
2001). In turn, oxyluciferin displays a strong inhibitory effect on the firefly luciferase reaction in a competitive manner. Hence, intensive screening for
luciferin-regenerating system(s) was initiated and resulted in the identification and molecular cloning of
a luciferin-regenerating enzyme from P. pyralis and
related insects. This enzyme converts oxyluciferin in
the presence of D-cysteine to intermediate 2-cyano6-hydroxybenzothiazole.
After completion of the molecular biological studies, we unequivocally could demonstrate the presence
of the gene encoding luciferase (Müller et al. 2009a).
We also reported on the existence of a gene for the
luciferin-regenerating enzyme. Furthermore, after
having demonstrated for the first time both that
demosponge spicules act as optical waveguides and
that tissue samples from the animals have the capacity to produce light, we showed that the recombinant
luciferase was biocatalytically active and able to cause
bioluminescence in vitro. Finally, we showed that the
level of expression of the luciferase gene is strongly
downregulated if S. domuncula cells are kept in the
light. In these initial studies, primmorphs, a special
form of 3D cell culture (Müller et al. 1999), were
applied. Recently, tissue slices through the cortex/
surface regions of S. domuncula were hybridized in
situ with a labeled probe of luciferase. When tissue
slices from a light-adapted animal were analyzed,
hardly any hybridization signals could be detected
(Fig. 3A). When a slice from a dark-adapted
animal was incubated with the luciferase probe,
W. E. G. Müller et al.
however, a strong signal was seen in the surface
layer of the sponge (Fig. 3B). This finding is in accordance with a previous analysis showing that total
RNAs from light-adapted primmorphs have a significantly lower level of luciferase transcripts, compared
with the level measured in dark-adapted primmorphs
(Müller et al. 2009a).
The emission spectrum of the S. domuncula
enzyme revealed a maximum at 548 nm and a
minor peak at 590 nm (pH 8.0), in accordance
with the bioluminescence spectra of the firefly luciferase/luciferin system that generally vary between 520
and 620 nm. In turn, the bioluminescence emission
spectrum, with peaks at 548 and 590 nm, overlaps
with the light-transmission spectra through hexactinellid spicules (Müller et al. 2009a). These results
suggest that spicules act as waveguides for both biogenous light and environmental light (Müller et al.
2010).
Flashing sponge: light emission within individuals of
S. domuncula
As a more direct proof that sponges can indeed
rhythmically flash light, a specimen was connected
with a light-recording device (Wiens et al. 2010).
The animal was placed in a 1.0-l beaker under
optimal conditions. Prior to the experiment, the specimen remained in complete darkness in a vibrationfree cabinet for 3 days. Then, a tissue slice, 2–3 mm
thick, was ablated from the surface (cortex) during a
30-s period of illumination. Using a forceps, the
samples were then lightly pressed against the glass
wall. Subsequently, the animals remained for 3 days
in complete darkness. A photomultiplier consisting
of a photocathode to detect photons of 300–
890 nm wavelength was coupled to a glass beaker
1.5 mm thick, and the signals were recorded through
a customized script on a PC with an IEEE-488 interface card. In a typical experiment, the specimens
started to emit light after a lag phase of 1400–
2000 min (23–33 h). A characteristic recording profile
is shown in Fig. 4A. In this experiment, the specimen
started to flash after 1688 min of complete inactivity.
Almost at the onset of flashing, the highest number
of counts (2250) was recorded in a rhythmic way.
The activity lasted approximately 4000 min (66 h)
(Fig. 4A). Flashing was abruptly terminated, as
indicated by a sudden decrease in the number of
counts (Fig. 4A). At present, it is only possible to
speculate about the function of the proposed light
flashing/signaling process. Earlier reports proposed
that in corals, the light-responsive cryptochrome
system is involved in the synchronization of, for
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Universal zeitgebers in sponges
Fig. 3 Luciferase of S. domuncula. (A and B) The expression of the luciferase was assessed by in situ hybridization. The antisense cDNA
was labeled with digoxigenin and the hybridization signals were detected with nitrotetrazolium blue chloride/5-bromo-4-chloro-3indolyl phosphate (Müller et al. 2010). (A) Only weak luciferase signals are seen in tissue slices from animals exposed to light, while (B)
strong signals are seen in samples from dark-adapted animals. (C) The proposed photoreception/phototransduction system of sponges.
Biogeneous light is generated by luciferase, which is postulated to be transmitted through spicules that act as light waveguides. The
schematic outline shows that the luciferase-mediated oxidation of luciferin results in the generation of bioluminescence. Photons are
transmitted through the spicule and then are detected by a chromophore/redox system associated with cryptochrome. Concomitantly,
oxyluciferin is the substrate for the luciferin-regenerating enzyme. Partially modified from Müller et al. (2010).
example, mass spawning (Levy et al. 2003), concurrently suggesting that the coral cryptochrome system
controls an ancient rhythm resembling a circadian
clock.
In a further attempt to elucidate involvement of a
downstream signal-transduction molecule in the
transmission of light signals, a G-protein was identified (published data). This S. domuncula G-protein
shares highest similarity of sequence with vertebrate
transducin. Future studies must clarify if there is any
functional relationship between sponge G-protein
and vertebrate transducin (Hargrave et al. 1993)
during the proposed light transduction signalling
(Fig. 4B).
Light detection system: cryptochrome
In Fig. 5A and B, it is shown that the spicules in the
subsurface region organize to bundles that often
combine and run toward the center of the sponge
specimens. This morphological arrangement of the
spicules corroborates again that the spicules have besides of the proposed defense role, and the function
to stabilize the sponge body relevance for light-harvesting and transduction/transmission.
The discovery of cryptochrome(s), which are
crucially involved in light-harvesting reactions in
corals (Levy et al. 2007), stimulated a systematic
screening for homologous poriferan proteins and resulted in the identification of candidate molecules in
110
W. E. G. Müller et al.
Fig. 4 Flashing light in the sponge S. domuncula. (A) A specimen was exposed to a light-detecting photomultiplier. Typically, flashing
started after a lag phase of 1400–20,00 min. In the representative experiment shown here the first signals (counts, reflecting intensities
of the photon) were recorded after a period of 1688 min. The strongest signals were detected approximately 1900 min after the start
of the experiment. Magnification of the peaks revealed a characteristic shape; one multi-peak sequence of flashes is shown between
1702 and 1707 min. Such a typical flashing can be divided into an initial strong burst (1), followed by an exponential decrease (2), which
then enters a short plateau (3); one flash unit ends abruptly (4). (B) Recently, a G-protein has been identified that shares high sequence
similarity with vertebrate transducin, a key molecule in the light-signal transmission in vertebrates, in which G-protein is coupled to
receptors, called opsin, that contain the chromophore 11-cis retinal. The metabolic pathway of carotene/retinal/retinoic acid was
elucidated by Müller et al. (2011, 2012b). The initial product, b-carotene, is synthesized by sponge-associated bacteria, while the
processing enzymes are sponge-coded. Future studies are needed to show whether retinal interacts with cryptochrome in a
light-sensory system in sponges; the building blocks are there.
demosponges (S. domuncula) and in hexactinellids
(Crateromorpha meyeri) (Müller et al. 2010). As in
the coral model (Levy et al. 2007), the expression of
poriferan cryptochrome is controlled by light (Müller
et al. 2010). More specifically, the expression of the
S. domuncula cryptochrome gene after exposure to
light (330–900 and 700–1100 nm) (Müller et al.
2010) is primarily restricted to the surface zone of
the animal, suggesting that the photoactivated protein is compartmentalized in the cortex (the outer
tissue layer). Similarly, the expression of poriferan
luciferase is light-adapted (Müller et al. 2009a).
The gene expression of cryptochrome is dependent
on light. Samples of cell aggregates or tissue were
exposed to light for 1–8 h, using a short-pass filter
(spectral range 330–900 nm) or a long-pass filter
(spectral range 700–1100 nm). Afterward, RNA was
extracted. Subsequent Northern blot analyses revealed that after dark adaptation of primmorphs
and tissue, no expression of cryptochrome was detectable. After 2 h of light exposure (330–900 nm),
however, an increased expression of cryptochrome
could be seen. There was further increase after 4 or
8 h of exposure to light, both in tissue and primmorphs. In a crucial set of RNA experiments, a
quantitative real-time polymerase chain reaction
(qPCR) was applied to determine cryptochrome
transcription over 24 h, including light–dark transition. Subsequently, cryptochrome expression was
correlated to the expression of the housekeeping
gene tubulin. Thus, during 12-h exposure to light,
cryptochrome expression increased to 0.53 and
then decreased, until after 12 h of darkness a ratio
of 0.15 was reached. Accordingly, cryptochrome expression during exposure to light was up to 3.5-fold
higher than in darkness (Müller et al. 2010).
These data demonstrate that exposure of
S. domuncula (tissue and primmorphs) to light stimulates cryptochrome expression. In contrast to studies performed with corals (Levy et al. 2007), the
light-induced response in sponges occurs regardless
of the spectral range. Because cryptochrome is
Universal zeitgebers in sponges
111
Fig. 5 Proposed photo-transduction mechanism in demosponges. (A and B) Arrangements of the spicules in the surface (su) region of
S. domuncula. The outer surface zone (su), about 2 mm thick, is densely packed with tylostyles spicules. In the region near the surface, the
spicules are packed into bundles (bu) that run toward the center of the specimen. These bundles are formed from parallel spicules that
often combine. (C) The proposed light-flashing signaling process in S. domuncula. During development and during exposure to light binding
of the morphogen retinoic acid to its receptor RXR induces an expression of a transcription factor, the SOX-related protein (SOX). In
vertebrates, SOX operates during differentiation in general and neuronal development in particular. Flanked by Pax-6, a transcription
factor without apparent poriferan homolog, SOX-2 causes coordinated development of cells during embryogenesis and during the
proposed neurogenesis. In sponges, the SOX-related protein causes consecutive expression of luciferase (effector) and cryptochrome
(receiver). Both are components of the poriferan light generating/harvesting system, with the siliceous spicules as light waveguides. In this
manner, coordinated transmitted signals might regulate, for example, pulsation of pinacocytes and contraction of myocytes.
particularly localized around the spicules, these
skeletal elements might operate as waveguides for
environmental light that penetrates with significant
intensity to water depths of approximately 20 m in
the northern Adriatic Sea, a natural habitat of
S. domuncula. Thus, spicules at the surface of the
animal might function as collectors that transmit
light to cryptochrome, the putative photosensory
receptor of sponges (Figs. 3C and 5C). In addition,
light might be produced within the body of
the sponges via luciferase that might again be collected and transmitted via the same photo-optical
system.
Light-dependent alterations of induction of genes
involved in the light-response system
The findings that in response to light both luciferase
and cryptochrome are expressed to a higher level
prompted us to search for upstream localized transcription factors (TFs) controlling the expression of
cryptochrome and luciferase, the key molecules
involved in the proposed photoreception/photogeneration system in sponges (Wiens et al. 2010). No
poriferan homolog of Pax-6, a molecule required for
morphogenesis of the visual system in triploblasts
(Gehring and Ikeo 2009), could be detected.
Therefore, we screened for SOX genes, encoding a
112
family of metazoan-specific TF (Koopman et al.
2004) that have been found to play important roles
in a variety of developmental processes, particularly
during organogenesis (Wegner 1999). All SOX
proteins comprise a highly conserved high mobility
group box. Among the 430 family members known
so far, in particular one protein, SOX-2 (Hagstrom
et al. 2005), has been implicated in early development of the human eye and brain. In our studies
(Wiens et al. 2010), we analyzed the spatio-temporal
expression of S. domuncula genes, coding for a
SOX-related protein, a luciferase, and a cryptochrome, following ablation of the surface cell layers
and after exposure to light (to be published). After
the increased expression of the SOX-related protein,
a higher steady-state level of luciferase and
cryptochrome is seen, suggesting that the expression
of the latter two molecules is not only under the
control of regeneration/morphogenetic events but
also under the control of exposure to light (Wiens
et al. 2010) (Fig. 5C).
We postulate that the resulting upregulation of
the proposed flashing-collection-guidance-detection
pathway is followed by a physiological effect on pinacocytes (expanding and contracting cells) and myocytes (contractile cells). In turn, the data gathered
suggest that, similar to corals, sponges are provided
with a photosystem that controls distinct behavioral
patterns and physiological reactions (e.g., rhythmic
pulsations of aquiferous canals in the sponge’s body).
Potential zeitgebers and their
controlling molecules in sponges
Sponges are provided with a zeitgeber ‘‘light’’ that
controls their (proposed) endogenous clock. As outlined above, environmental light can be perceived or
light can be generated endogenously by the sponges.
This endogenous clock might be required for tuning
the physiological and molecular rhythms to use the
environmentally determined conditions in an economically optimal way. Like any other organism,
sponges living in the shallow illuminated/dimmed
zone or in the deep sea are exposed to about-daily
or seasonal changes that occur in their environment
(e.g., light, temperature, and feeding cycles).
Until recently, no studies of the endogenous oscillators or clocks in sponges have been published that
led to an understanding of the mechanism whereby
sponges are able to maintain a sustained oscillation,
or rhythm, perhaps even in the absence of external
environmental cues. Since sponges lack a nervous
system or even a central nervous system, we were
not surprised at failure to find (until now) a
W. E. G. Müller et al.
homology of the vertebrate CLOCK and BMAL1
molecules. We were asking whether sponges also
have the abundant downstream molecules acting as
secondary zeitgebers in vertebrates’ cryptochrome
and nocturnin. As outlined above, manifold evidence
has been presented that cryptochrome also acts as a
functional light receiver in lower animals, e.g.
sponges and hydrozoans. From studies with mammals it is known that cryptochrome is crucially involved in the general day–night rhythm (van der
Horst et al. 1999; Kume et al. 1999).
One major link between circadian biology and metabolism in vertebrates is nocturnin (Gilbert et al.
2011). It has been established that the expression
of nocturnin peaks in the early night in liver,
kidney, spleen, and also retina (Wang et al. 2001).
Cryptochrome
The existence and function of this protein in S.
domuncula are outlined above.
Nocturnin
Recently, we successfully demonstrated the existence
of nocturnin in S. domuncula and in turn outlined the
function of this protein in the demosponge (Müller
et al. 2012a). Sponges are filter feeders and are decisively dependent on the nutritional status of the environment. Considering that in vertebrates, it is
known that one of the circadian-expressed proteins,
nocturnin, is induced during fasting in the white adipose tissue of mice (Gilbert et al. 2011) and in turn
plays a role in sensing the circadian clock regulating
lipid mobilization in fat cells. Nocturnin is a
deadenylase, a 30 -exoribonuclease, with specificity for
poly(A) tracts in messenger RNA (mRNA), whose
steady-state expression is under both circadian and
acute regulation (Green and Besharse 1996; Green
et al. 2007). We initially discovered an poly(A)specific 30 -exoribonuclease and purified it from calf
thymus (Müller et al. 1977, 1980; Schröder et al.
1980). It has been assigned to an EC number (EC
3.1.13.4—poly(A)-specific ribonuclease [http://www.
brenda-enzymes.org/php/result_flat.php4?ecno¼3.1.
13.4]). Subsequently, the 30 -exoribonuclease was
found to control the half-life of mRNAs through hydrolysis of their poly(A) tracts (Müller et al. 1978).
The complete S. domuncula nocturnin sequence
was obtained (Müller et al. 2012a). The deduced
sponge protein shows high sequence similarity with
the corresponding human sequence and with the
cnidarian sequence from Nematostella vectensis;
the sponge nocturnin comprises the domain for
the exonuclease–endonuclease–phosphatase domain
Universal zeitgebers in sponges
113
Fig. 6 Scheme presenting the major molecules contributing to the core machinery of the molecular clock. (A) The transcription factors
BMAL1 and CLOCK constitute the positive limbs of the core circadian network by acting positively on the expression of the
downstream gene targets, primarily the period, cryptochrome, Ror, and Rev-Erb genes. It is assumed that the light/dark cycle in general and
the luciferase light-generating system, here in sponges, adjust the fidelity of the clock’s oscillation. The gene products PER (period) and
CRY (cryptochrome), the major negative limb, inhibit the activity of BMAL1 and CLOCK. A further clock controlled gene product, the
nuclear receptor Rev-Erb likewise negatively affects the expression of Bmal1 via interaction with the Rev-Erb/ROR responsive elements (RORE). In addition, the expression of the clock controlled gene Noc is positively controlled by BMAL1 and CLOCK. The protein
(continued)
114
superfamily. Functional studies on the S. domuncula
primmorph system found that these 3D cell aggregates change their morphology depending on light
supply. Upon exposure to light, primmorphs show
a faster and stronger increase in DNA, protein, and
glycogen compared with primmorphs that remain in
the dark. qPCR analyses revealed that after transfer
from light to dark, primmorphs show a 10-fold increased expression of the nocturnin gene. In contrast,
the transcript level of glycogenin, the starter enzyme
for glycogen synthesis (Shearer et al. 2005; Moslemi
et al. 2010), decreases in the dark by three- to
four-fold. Exposure of primmorphs to light causes
a decrease in nocturnin transcripts and an opposed
increase in glycogenin transcripts.
While the clock-controlled genes are acting back
on the expression of the central core clock molecules
Bmal1/BMAL1 and Clock/CLOCK either negatively
(Per/period and Cry/cryptochrome) or positively
(orphan receptor- related to Ror/retinoic acid),
Noc/nocturnin remains in the cytoplasm and
controls the half-life and stability of key mRNA species involved in the adjustment of the homeostasis of
intermediary metabolism, especially of fatty acids
and glycogen (Fig. 6A). Nocturnin is a circadian
deadenylase that was originally described as a 30 -exonuclease from calf thymus (see above). A function
for this enzyme was proposed to be a selective hydrolysis of the poly(A) tail of mRNAs, allowing a control
of the post-transcriptional net-polyadenylation of
heterogeneous nuclear RNA, especially during the
phase of DNA synthesis. In our studies in the
S. domuncula primmorph system, we reported a
light/dark-correlated change of the steady level of
the transcripts for glycogenin. The expressed glycogenin protein/enzyme is a crucial regulator of glycogen synthesis, acting both as a glycosyltransferase and
primer. Based on these data, we concluded that
W. E. G. Müller et al.
sponges are provided with the molecular circadian
clock protein nocturnin that is highly expressed in
the dark, thereby controlling/downregulating the
function of this key metabolic enzyme of glycogenin
(Müller et al. 2012a).
The day/night rhythm of expression of Per and
Cry genes that is activated by the CLOCK-BMAL1
heterodimer in a negative feedback loop has been
implicated in regulation of gluconeogenesis in
mouse liver through CRY-caused inhibition of
glucagon-stimulated cAMP production (Hatori and
Panda 2010). It is likely that the circadian clock
protein CRY may affect the rate of gluconeogenesis
and glycolysis in sponge tissue in a similar way
(Fig. 6B). It is known that the allosteric regulator
fructose-2,6-bisphosphate plays a crucial role in controlling gluconeogenesis and glycolysis at the level of
the common metabolite fructose-1,6-bisphosphate.
The concentration of this allosteric activator
depends on the cAMP-stimulated phosphorylation
of phosphofructokinase-2. At high cAMP concentrations, fructose 1,6-bisphosphatase (gluconeogenesis)
is activated and phosphofructokinase (glycolysis) is
inhibited, while at low cAMP concentrations
phosphofructokinase (glycolysis) is stimulated and
fructose-1,6-bisphosphatase (gluconeogenesis) is inhibited. Consequently, it can be expected that at
the beginning of the day, glycolysis will be activated
and gluconeogenesis will be inhibited, while at the
beginning of night gluconeogenesis will be upregulated and glycolysis will be downregulated (Fig. 6B).
Conclusion—does a universal
zeitgeber-effector process exist from
sponges to mammals?
Many biological processes within unicellular and
multicellular organisms oscillate with distinct
Fig. 6 Continued
product, nocturnin, is a deadenylase that we originally described as a 30 -exoribonuclease. This enzyme controls the decay kinetics of
mRNAs via specific removal of their poly(A) tail. It is assumed that nocturnin removes (especially in the dark/night) key regulatory
proteins of the metabolism of fatty acids and glycogen. (B) Hypothetical regulation of the glycolysis and gluconeogenesis pathways in
sponges via the circadian clock protein CRY during the diel cycle. It is assumed that CRY differentially affects the rate of gluconeogenesis and glycolysis via modulation of cAMP production, in analogy to the model in mouse liver (Hatori and Panda 2010). At the
beginning of the day (left box), the levels of CRY are high, resulting in a strong reduction of cAMP production, while at the beginning of
the night (right box), CRY levels are low with no, or only a small, effect on cAMP production. The level of cAMP affects gluconeogenesis and glycolysis via modulation of the level of fructose-2,6-bisphosphate. This allosteric regulator activates the formation of
fructose-1,6-bisphosphate, a common metabolite of glycolysis and gluconeogenesis (shown in the middle of the figure) from
fructose-6-phosphate via phosphofructokinase (glycolysis), while the conversion of fructose-1,6-bisphosphate to fructose-6-phosphate
via fructose-1,6-bisphosphatase (gluconeogenesis) is inhibited by fructose-2,6-bisphosphate. Hence, at low concentrations of cAMP,
when the fructose-2,6-bisphosphate level is high (beginning of the day), phosphofructokinase (glycolysis) is stimulated (green arrows)
and fructose-1,6-bisphosphatase (gluconeogenesis) is inhibited (red arrows); at high cAMP concentrations (beginning of the night), when
the fructose-2,6-bisphosphate level is low, fructose 1,6-bisphosphatase (gluconeogenesis) is activated (green arrows) and phosphofructokinase (glycolysis) is inhibited (red arrows). Partially modified from Müller et al. (2012a) and Hatori and Panda (2010).
Universal zeitgebers in sponges
peak-to-peak intervals, or periods, of activity (BellPedersen et al. 2005). Most of the circadian oscillators can be entrained to a given phase and period
through zeitgebers (signals from the environment)
(Oda and Friesen 2011). The most dominant zeitgebers are the light–dark cycle and alteration in nutritional supply. The present data show that light plays
an important role in the overall regulation of metabolism and perhaps, as suggested herein, also as a
zeitgeber for the postulated circadian rhythm.
The present experimental data show that sponges
already have some elements of a circadian rhythm,
involving the clock elements cryptochrome and nocturnin. In S. domuncula that lives in the dawn zone
(about 20 m deep), the endogenous clock, involving
cryptochrome and nocturnin, is synchronized by the
well-established cue ‘‘light’’ (Liu and Green 2002).
Studies are now in progress to evaluate whether
the nutritional status of the environment, as well as
‘‘light,’’ can act as a cue to synchronize the metabolism of sponges. This direction of study is important
for an understanding of any potential rhythm in
deep-sea animals, where surely exogenous light will
have only a subordinate role; it has been postulated
that surface bacteria might produce light that could
be channeled through the spicules to a
light-responsive harvesting system in sponge cells
(Epstein et al. 2011).
Funding
W.E.G.M. is a holder of an ERC Individual
Advanced Grant (no. 268476 BIOSILICA). This
work was supported by grants from the German
Bundesministerium für Bildung und Forschung
(project ‘‘Center of Excellence BIOTECmarin’’), the
Deutsche Forschungsgemeinschaft (Schr 277/10-2),
the International Human Frontier Science Program,
the European Commission (project no. 311848—
BlueGenics; project no. 244967—Mem-S Project;
and project no. 268476—MarBioTec*EU-CN*), the
Public Welfare Project of Ministry of Land and
Resources of the People’s Republic of China (grant
no. 201011005-06), and the International S & T
Cooperation Program of China (grant no.
2008DFA00980).
References
Aizenberg J, Sundar V, Yablon AD, Weaver JC, Chen G.
2004. Biological glass fibers: correlation between optical
and structural properties. Proc Natl Acad Sci USA
101:3358–63.
Albrecht U, Eichele G. 2003. The mammalian circadian clock.
Curr Opin Genet Dev 13:271–7.
115
Belas R, Mileham A, Cohn D, Hilman M, Simon M,
Silverman M. 1982. Bacterial bioluminescence: isolation
and expression of the luciferase genes from Vibrio harveyi.
Science 218:791–3.
Bell-Pedersen D, Cassone VM, Earnest DJ, Golden SS,
Hardin PE, Thomas TL, Zoran MJ. 2005. Circadian
rhythms from multiple oscillators: lessons from diverse
organisms. Nat Rev Genet 6:544–56.
Branchini BR, Ablamsky DM, Murtiashaw MH, Uzasci L,
Fraga H, Southworth TL. 2007. Thermostable red and
green light-producing firefly luciferase mutants for bioluminescent reporter applications. Anal Biochem 361:253–62.
Brümmer F, Pfannkuchen M, Baltz A, Hauser T, Thiel V.
2008. Light inside sponges. J Exp Mar Biol Ecol 367:61–4.
Cattaneo-Vietti R, Bavestrello G, Cerrano C, Sarà A,
Benatti U, Giovine M, Gaino E. 1996. Optical fibres in
an Antarctic sponge. Nature 383:397–8.
Cha JN, Shimizu K, Zhou Y, Christianssen SC, Chmelka BF,
Stucky GD, Morse DE. 1999. Silicatein filaments and subunits from a marine sponge direct the polymerization
of silica and silicones in vitro. Proc Natl Acad Sci USA
96:361–5.
Epstein AK, Pokroy B, Seminara A, Aizenberg J. 2011.
Bacterial biofilm shows persistent resistance to liquid
wetting and gas penetration. Proc Natl Acad Sci USA
108:995–1000.
Gehring WJ, Ikeo K. 2009. Pax 6: mastering eye morphogenesis and eye evolution. Trends Genet 15:371–7.
Gilbert MR, Douris N, Tongjai S, Green CB. 2011. Nocturnin
expression is induced by fasting in the white adipose tissue
of restricted fed mice. PLoS One 6:e17051.
Gomi K, Kajiyama N. 2001. Oxyluciferin, a luminescence
product of firefly luciferase, is enzymatically regenerated
into luciferin. J Biol Chem 276:36508–13.
Green CB, Besharse JC. 1996. Use of a high stringency differential display screen for identification of retinal mRNAs
that are regulated by a circadian clock. Brain Res Mol
Brain Res 37:157–65.
Green CB, Douris N, Kojima S, Strayer CA, Fogerty J,
Lourim D, Keller SR, Besharse JC. 2007. Loss of
Nocturnin, a circadian deadenylase, confers resistance to
hepatic steatosis and diet-induced obesity. Proc Natl Acad
Sci USA 104:9888–93.
Greer LF, Szalay AA. 2002. Imaging of light emission from the
expression of luciferase in living cells and organisms: a
review. Luminescence 17:43–74.
Haddock SHD, Moline MA, Case JF. 2010. Bioluminescence
in the sea. Ann Rev Marine Sci 2:443–93.
Hagstrom SA, Pauer GJ, Reid J, Simpson E, Crowe S,
Maumenee IH, Traboulsi EI. 2005. SOX2 mutation causes
anophthalmia, hearing loss, and brain anomalies. Am J
Med Genet 138 A:95–8.
Harcet M, Roller M, Cetkovic H, Perina D, Wiens M,
Müller WEG, Vlahovicek K. 2010. Demosponge EST sequencing reveals a complex genetic toolkit of the simplest
metazoans. Mol Biol Evol 27:2747–56.
Hargrave PA, Hamm HE, Hofmann KP. 1993. Interaction of
rhodopsin with the G-protein, transducin. Bioessays
15:43–50.
Harvey EN. 1921. Studies on bioluminescence. XIII.
Luminescence in the coelenterates. Biol Bull 41:280–4.
116
Hatori M, Panda S. 2010. CRY links the circadian clock and
CREB-mediated gluconeogenesis. Cell Res 20:1285–8.
Herring PJ. 1987. Systematic distribution of bioluminescence
in living organisms. J Biolum Chemilum 1:147–63.
Huang N, Chelliah Y, Shan Y, Taylor CA, Yoo SH, Partch C,
Green CB, Zhang H, Takahashi JS. 2012. Crystal structure
of the heterodimeric CLOCK:BMAL1 transcriptional
activator complex. Science 337:189–94.
Ko CH, Takahashi JS. 2006. Molecular components of
the mammalian circadian clock. Human Mol Genet
15:R271–7.
Koopman P, Schepers G, Brenner S, Venkatesh B. 2004.
Origin and diversity of the Sox transcription factor gene
family: genome-wide analysis in Fugu rubripes. Gene
328:177–86.
Krasko A, Batel R, Schröder HC, Müller IM, Müller WEG.
2000. Expression of silicatein and collagen genes in the
marine sponge Suberites domuncula is controlled by silicate
and myotrophin. Eur J Biochem 267:4878–87.
Kulczykowska E, Popek W, Kapoor BG. 2010. Biological clock
in fish. Enfield (NH): Science Publishers.
Kume K, Zylka MJ, Sriram S, Shearman LP, Weaver DR,
Jin X, Maywood ES, Hastings MH, Reppert SM. 1999.
mCRY1 and mCRY2 are essential components of the negative limb of the circadian clock feedback loop. Cell
98:193–205.
LePennec G, Perović S, Ammar MSA, Grebenjuk VA,
Steffen R, Müller WEG. 2003. Cultivation of primmorphs
from the marine sponge Suberites domuncula: morphogenetic potential of silicon and iron. J Biotechnol 100:93–108.
Levy O, Dubinsky Z, Achituv Y. 2003. Photobehavior of stony
corals: responses to light spectra and intensity. J Exp Biol
206:4041–9.
Levy O, Appelbaum L, Leggat W, Gothlif Y, Hayward DC,
Miller DJ, Hoegh-Guldberg O. 2007. Light-responsive cryptochromes from a simple multicellular animal, the coral
Acropora millepora. Science 318:467–70.
Leys SP, Mackie GO, Reiswig HM. 2007. The biology of glass
sponges. Adv Mar Biol 52:1–145.
Liu X, Green CB. 2002. Circadian regulation of nocturnin
transcription by phosphorylated CREB in Xenopus retinal
photoreceptor cells. Mol Cell Biol 22:7501–11.
Meighen EA. 1991. Molecular biology of bacterial bioluminescence. Microbiol Rev 55:123–42.
Moslemi AR, Lindberg C, Nilsson J, Tajsharghi H,
Andersson B, Oldfors A. 2010. Glycogenin-1 deficiency
and inactivated priming of glycogen synthesis. N Engl J
Med 362:1203–10.
Müller WEG. 1995. Molecular phylogeny of metazoa
[Animals] (1995) Monophyletic origin. Naturwiss 82:321–9.
Müller WEG. 2006. The stem cell concept in sponges
(Porifera): metazoan traits. Semin Cell Develop Biol
17:481–91.
Müller WEG, Schröder HC, Arendes J, Steffen R, Zahn RK,
Dose K. 1977. Alterations of activities of nucleases and
polyadenylate polymerase in synchronized mouse L-cells.
Eur J Biochem 76:531–40.
Müller WEG, Arendes J, Zahn RK, Schröder HC. 1978.
Control of enzymatic hydrolysis of polyadenylate segment
of messenger RNA: role of polyadenylate-associated proteins. Eur J Biochem 86:283–90.
W. E. G. Müller et al.
Müller WEG, Schröder HC, Zahn RK, Dose K. 1980.
Degradation of 20 –50 -linked oligoriboadenylates by
30 -exoribonuclease and 50 -nucleotidase from calf thymus.
Hoppe-Seyler’s Z Physiol Chem 361:469–72.
Müller WEG, Wiens M, Batel R, Steffen R, Schröder HC,
Borojevic R, Custodio MR. 1999. Establishment of a primary cell culture from a sponge: Primmorphs from
Suberites domuncula. Mar Ecol Prog Ser 178:205–19.
Müller WEG, Wiens M, Adell T, Gamulin V, Schröder HC,
Müller IM. 2004. The Bauplan of the Urmetazoa: The
basis of the genetic complexity of Metazoa using the siliceous sponges [Porifera] as living fossils. Int Rev Cytol
235:53–92.
Müller WEG, Belikov SI, Kaluzhnaya OV, Perović-Ottstadt S,
Fattorusso E, Ushijima H, Krasko A, Schröder HC. 2006a.
Cold stress defense in the freshwater sponge Lubomirskia
baicalensis: role of okadaic acid produced by symbiotic dinoflagellates. FEBS J 274:23–36.
Müller WEG, Wendt K, Geppert C, Wiens M, Reiber A,
Schröder HC. 2006b. Novel photoreception system in
sponges? Unique transmission properties of the stalk spicules from the hexactinellid Hyalonema sieboldi. Biosens
Bioelectron 21:1149–55.
Müller WEG, Kaluzhnaya OV, Belikov SI, Rothenberger M,
Schröder HC, Reiber A, Kaandorp JA., Manz B,
Mietchen D, Volke F. 2006c. Magnetic resonance imaging
of the siliceous skeleton of the demosponge Lubomirskia
baicalensis. J Struct Biol 153:31–41.
Müller WEG, Li J, Schröder HC, Qiao L, Wang XH. 2007a.
The unique skeleton of siliceous sponges (Porifera;
Hexactinellida and Demospongiae) that evolved first from
the Urmetazoa during the Proterozoic: a review.
Biogeosciences 4:219–32.
Müller WEG, Schloßmacher U, Eckert C, Krasko A,
Boreiko A, Ushijima H, Wolf SE, Tremel W,
Schröder HC. 2007b. Analysis of the axial filament in spicules of the demosponge Geodia cydonium: different silicatein composition in microscleres [asters] and megascleres
[oxeas and triaenes]. Eur J Cell Biol 86:473–87.
Müller WEG, Jochum K, Stoll B, Wang XH. 2008. Formation
of giant spicule from quartz glass by the deep sea sponge
Monorhaphis. Chem Mater 20:4703–11.
Müller WEG, Kasueske M, Wang XH, Schröder HC, Wang Y,
Pisignano D, Wiens M. 2009a. Luciferase a light source
for the silica-based optical waveguides (spicules) in the
demosponge Suberites domuncula. Cell Mol Life Sci
66:537–52.
Müller WEG, Wang XH, Burghard Z, Bill J, Krasko A,
Boreiko A, Schloßmacher U, Schröder HC, Wiens M.
2009b. Bio-sintering processes in hexactinellid sponges:
fusion of bio-silica in giant basal spicules from
Monorhaphis chuni. J Struct Biol 168:548–61.
Müller WEG, Wang X, Schröder HC, Korzhev M,
Grebenjuk VA, Markl JS, Jochum KP, Pisignano D,
Wiens M. 2010. A cryptochrome-based photosensory
system in the siliceous sponge Suberites domuncula
(Demospongiae). FEBS J 277:1182–201.
Müller WEG, Binder M, von Lintig J, Guo YW, Wang XH,
Kaandorp JA, Wiens M, Schröder HC. 2011. Interaction of
the retinoic acid signaling pathway with spicule formation
in the marine sponge Suberites domuncula through
Universal zeitgebers in sponges
activation of bone morphogenetic protein-1. Biochim
Biophys Acta 1810:1178–94.
Müller WEG, Wang XH, Grebenjuk VA, Korzhev M,
Wiens M, Schloßmacher U, Schröder HC. 2012a.
Nocturnin in the demosponge Suberites domuncula: a potential circadian clock protein controlling glycogenin synthesis in sponges. Biochem J 448:233–42.
Müller WEG, Wang XH, Binder M, von Lintig J, Wiens M,
Schröder HC. 2012b. Differential expression of the demosponge (Suberites domuncula) carotenoid oxygenases in response to light: protection mechanism against the
self-produced toxic protein (Suberitine). Mar Drugs
10:177–99.
Oda GA, Friesen WO. 2011. Modeling two-oscillator circadian systems entrained by two environmental cycles. PLoS
One 6(8):e23895.
Ridley SO, Dendy A. 1887. Report on the Monaxonida
collected by H.M.S. ‘challenger’. Report on the scientific
results of the voyage of the H.M.S. challenger during the years 1873–76. Vol. 20. London: Eyre &
Spottiswoode.
Schröder HC, Zahn RK, Dose K, Müller WEG. 1980.
Purification and characterization of a poly(A)-specific exoribonuclease from calf thymus. J Biol Chem 255:4535–8.
Shearer J, Graham TE, Battram DS, Robinson DL, Richter EA,
Wilson RJ, Bakovic M. 2005. Glycogenin activity and
mRNA expression in response to volitional exhaustion in
human skeletal muscle. J Appl Physiol 99:957–62.
Shimizu K, Cha J, Stucky GD, Morse DE. 1998. Silicatein
alpha: cathepsin L-like protein in sponge biosilica. Proc
Natl Acad Sci USA 95:6234–8.
SpongeBase. 2010. Available online: https://octavia.vk.medizin.
uni-mainz.de/login.cgi (accessed September 28, 2012).
Strehler BL, McElroy WD. 1949. Purification of firefly luciferin. J Cell Physiol 34:457–66.
van der Horst GT, Muijtjens M, Kobayashi K, Takano R,
Kanno S, Takao M, de Wit J, Verkerk A, Eker AP, van
Leenen D, et al. 1999. Mammalian Cry1 and Cry2 are essential for maintenance of circadian rhythms. Nature
398:627–30.
Wang Y, Osterbur DL, Megaw PL, Tosini G, Fukuhara C,
Green CB, Besharse JC. 2001. Rhythmic expression of
117
Nocturnin mRNA in multiple tissues of the mouse. BMC
Dev Biol 1:9.
Wang XH, Li J, Qiao L, Schröder HC, Eckert C, Kropf K,
Wang YM, Feng QL, Müller WEG. 2007. Structure and
characteristics of giant spicules of the deep sea hexactinellid
sponges of the genus Monorhaphis (Hexactinellida:
Amphidiscosida: Monorhaphididae). Acta Zoologica Sinica
53:557–69.
Wang XH, Schröder HC, Müller WEG. 2009. Giant siliceous
spicules from the deep-sea glass sponge Monorhaphis chuni:
morphology, biochemistry and molecular biology. Int Rev
Cell Mol Biol 273:69–115.
Wang XH, Hu S, Gan L, Wiens M, Müller WEG. 2010.
Sponges (Porifera) as living metazoan witnesses from the
Neoproterozoic: biomineralization and the concept of their
evolutionary success. Terra Nova 22:1–11.
Wang XH, Schröder HC, Wang K, Kaandorp JA,
Müller WEG. 2012a. Genetic, biological and structural hierarchies during sponge spicule formation: from soft
sol-gels to solid 3D silica composite structures. Soft
Matter 8:9501–18.
Wang XH, Fan XT, Schröder HC, Müller WEG. 2012b.
Flashing light in sponges through their siliceous fiber network: a new strategy of ‘‘neuronal transmission’’ in animals. Chinese Sci Bull 57:3300–11.
Wegner M. 1999. From head to toes: the multiple facets of
SOX proteins. Nucleic Acids Res 27:1409–20.
Wiens M, Wrede P, Grebenjuk VA, Kaluzhnaya OV,
Belikov SI, Schröder HC, Müller WEG. 2009. Towards a
molecular systematics of the Lake Baikal/Lake Tuva
Sponges. In: Müller WEG, Grachev MA, editors. Biosilica
in evolution, morphogenesis, and nanobiotechnology. Prog
Mol Subcell Biol [Marine Molecular Biotechnology]. Vol.
47. Berlin: Springer-Verlag. p. 111–44.
Wiens M, Wang X, Unger A, Schröder HC, Grebenjuk VA,
Pisignano D, Jochum KP, Müller WEG. 2010. Flashing light
signaling circuit in sponges: endogenous light generation
after tissue ablation in Suberites domuncula. J Cell
Biochem 111:1377–89.
Wilson T, Hastings JW. 1998. Bioluminescence. Annu Rev
Cell Dev Biol 14:197–230.