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Advanced Review
The evolution of vision
Walter J. Gehring∗
In this review, the evolution of vision is retraced from its putative origins
in cyanobacteria to humans. Circadian oscillatory clocks, phototropism, and
phototaxis require the capability to detect light. Photosensory proteins allow
us to reconstruct molecular phylogenetic trees. The evolution of animal eyes
leading from an ancestral prototype to highly complex image forming eyes can be
deciphered on the basis of evolutionary developmental genetic experiments and
comparative genomics. As all bilaterian animals share the same master control
gene, Pax6, and the same retinal and pigment cell determination genes, we conclude that the different eye-types originated monophyletically and subsequently
diversified by divergent, parallel, or convergent evolution. © 2012 Wiley Periodicals, Inc.
How to cite this article:
WIREs Dev Biol 2014, 3:1–40. doi: 10.1002/wdev.96
INTRODUCTION
T
he eye is a most fascinating organ, and the
elucidation of eye evolution is one of the most
challenging tasks in evolution biology. In ‘The
Origin of Species’ Charles Darwin1 devoted an entire
chapter to ‘Difficulties of the Theory’ and the eye
as an organ of extreme perfection is discussed
extensively. At first sight, it seems absurd that an
eagle’s eye with all its perfections could have evolved
simply by random variation and selection. However,
Darwin found a possible solution to this enigma
and proposed that the complex, highly perfected
eyes like the eagle’s eyes originated from a relatively
simply prototypic organ consisting of an optic ‘nerve
(photoreceptor cell) surrounded by pigment cells and
covered by translucent skin, but without any lense or
other refractive body’. Such a prototypic eye would
enable its carrier to determine the direction of the
incoming light and provide a substantial selective
advantage over those organisms which can only
distinguish between light and dark. Darwin postulated
this prototype merely on theoretical grounds, but
subsequently it was found in e.g. the planarian
Polycelis auricularia and trochophora larvae. Darwin
also clearly stated that the evolution of the prototype
cannot be explained by selection, because selection
can only act on organs having at least a minimal
function. Therefore, the evolution of the prototypic
∗
Correspondence to: Walter.Gehring@unibas.ch
Growth and Development, Biozentrum, University of Basel, Basel,
Switzerland
Volume 3, January/February 2014
eye must have been a purely stochastic and hence
a very rare event. Once the prototype was formed,
random variation and selection could lead to the
evolution of the various more complex eye-types.
By contrast the neodarwinists like Ernst Mayr2,3
assumed that the various eye-types found in different
animal phyla arose 40–60 times independently.
Their arguments were mainly based on morphology
and physiology, but also on the different modes
of development. However, more recent genetic
experiments indicate that the various metazoan eyetypes are controlled by the same set of transcription
factors, in particular, Pax6 which serves as a master
control gene for eye morphogenesis in taxa as different
as insects and mammals4,5 and is found in all bilaterian
phyla. This demonstrates that the various eye-types
can be ‘built’ with the same tool kit. In 2001, Mayr3
admitted that his former notion was not quite correct.
The evolution of vision will be traced all the way
back to its putative origins in cyanobacteria, the oldest
known fossils on earth. Cyanobacteria have evolved a
circadian clock and phototaxis both of which depend
on vision. They are the first organism with ‘eye spots’,
i.e. organelles for photoreception. The evolution of
circadian clocks and of phototaxis and phototropism
are discussed, followed by the phylogenetic analysis
of the three major classes of photosensory proteins,
phytochromes, cryptochromes, and rhodopsins. The
evolution of the various animal eye-types will be
followed from their prototypes to their adaptive
radiation. Finally, some general genetic mechanisms
governing eye evolution are proposed.
© 2012 Wiley Periodicals, Inc.
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wires.wiley.com/devbio
Advanced Review
THE ORIGIN OF VISION:
CYANOBACTERIA
Cyanobacteria are among the oldest macroscopic
fossils on earth. They are known as stromatolites1–3,6–9 , which are sedimentary rocks formed by
layers of cyanobacteria and represent the earliest evidence for life on earth. Stromatolites from Pilbara in
Western Australia were dated to be approximately
3500 million years old. Living stromatolites can still
be seen today in Western Australia for example at
Shark Bay or in Yellowstone National Park in America. Cyanobacteria are prokaryotic eubacteria capable
of photosynthesis. They use the photosystems I and
II, chlorophyll a and b, the Calvin-cycle for CO2 fixation, H2 O as an electron donor, and they produce O2 .
They contribute substantially to the oxygen content
of the atmosphere. Their cells contain stacks of thylacoid membranes on which photosynthesis takes place.
They can be single-celled, like Gloeobacter or Synechococcus or filamentous, like Anabaena and Nostoc.
They represent a large fraction of the phytoplankton
of the oceans and also occur in fresh water and terrestrial habitats and can grow under extreme conditions.
In the course of evolution, some were taken up as
symbionts by eukaryotes and became chloroplasts.
Light detection confers a considerable selective
advantage to photosynthetic organisms which tend
to be attracted by sunlight and repelled by light of
excessively high intensity surpassing the capacity of
the photosynthetic machinery, and they also avoid UV
light which causes DNA damage (see cryptochromes).
There are two lines of evidence that cyanobacteria can sense light: circadian clocks calibrated by the
light–dark cycle of the sun, and Phototaxis have both
been found in cyanobacteria. The observation that
stromatolites are deposited in a regular centimeterscale spacing similar between modern and ancient
stromatolites suggests that they both were deposited
rhythmically with a period of approximately 20 h
corresponding to their circadian clock.
The terrestrial cyanobacterium, Leptolyngbya
sp., forms long, slender trichomes (filaments) and
moves very slowly by parallel movements of its
trichomes toward a light source (positive phototaxis).
The apical cell is characterized by an orange spot
at its distal tip which ressembles the ‘eye spot’ or
‘stigma’ of carotenoid-rich lipid globules which is
almost invariably present in phototactic flagellated
algae.10 Microspectrophotometric analysis showed an
absorption spectrum with a major peak at 456 nm
and a second peak at 504 nm, which was tentatively
identified as rhodopsin. Hydroxylamine treatment
which reacts with all aldehydes including retinal in
aqueous solution to form stable oximes, abolishes
2
this peak at 504 nm and prevents the phototactic
movements of the trichomes. This suggests that these
cyanobacteria have evolved photoreceptor organelles
endowed with rhodopsin as we find them in flagellated
algae, e.g. Chlamydomonas (Figure 2).
THE EVOLUTION OF CIRCADIAN
CLOCKS
Virtually, all organisms from cyanobacteria to humans
have developed cellular oscillations and mechanisms
to synchronize these cellular oscillations to environmental cycles, in particular, to the light–dark cycle
of sunlight. Internal timing mechanisms may facilitate physiological and behavioral adaptations to these
environmental conditions. These timing mechanisms
involve an internal molecular oscillator, a clock, which
is synchronized, entrained, in response to environmental signals, the Zeitgeber (German for time-giver). The
pioneering genetic work of Konopka and Benzer11 in
Drosophila led to the discovery of numerous genes
involved in the control of circadian rhythms. Genetics
in combination with recombinant DNA technology
and biochemistry, led to the isolation of the first clock
genes and to the elucidation of the mechanisms of the
biological clocks (for review, see Ref 12).
Circadian Oscillators in Cyanobacteria
Cyanobacteria such as Synechococcus have an
endogenous timing mechanism that can generate and
maintain a 24 h (= circadian) rhythm controlling
expression of the entire genome (see Ref 13 for
review). This rhythmicity extends to many other
physiological functions. The periodicity of these
processes reflects the periodicity of sunlight, the
primary source of energy for these photoautotrophic
organisms.
The basis of the circadian rhythm is a timing mechanism called circadian clock. The timing
mechanism must keep time and continue to drive oscillatory behaviors even under constant environmental
conditions.14
The best studied model organism is the photoautotrophic cyanobacterium Synechococcus elongatus.
To synchronize and reset the circadian clock, the cultures are kept in the dark for 6–12 h before exposing
them to constant illumination. Timing under these
constant conditions is designated as a ‘free-run’. The
period of any oscillation during the free-run is called
the free-running period. The first half of the freerunning period is the subjective day, the latter half
the subjective night. In addition to free-running selfsustained oscillations, true circadian clocks can be
WIREs Developmental Biology
Evolution of vision
entrained. Entrainment is the process by which the
environmental light–dark cycle determines the period
and phase relationship of an otherwise self-sustained
oscillation. The entrainment requires a mechanism
for perception of the environmental rhythm (light–dark cycle) i.e. some form of light perception. An
authentic circadian clock must also compensate for
temperature changes such that the free-running period
remains constant over the physiological range of
temperatures.
The most advanced studies on the circadian
clock have been carried out in S. elongatus, a
cyanobacterium which is amenable to molecular
genetic analysis. First, a large set of circadian clock
mutants were isolated by using a reporter strain that
carries a luciferase gene attached to a clock-controlled
promoter from a photosystem II gene.15 The respective
genes were isolated and characterized biochemically.
The circadian clock system can be divided into
three components: the oscillator that generates the
basic timing periodicity, the input pathway which
transmits signals from the environment to entrain
(synchronize) the oscillator, and the output pathway
that transmits the temporal information from the
oscillator to control various cellular functions, like
gene transcription. The cyanobacterial oscillator has
been identified16 and reconstituted in vitro.17
The Kai (circadian clock) locus encodes three
contiguous genes, KaiA, KaiB, and KaiC and these
genes are expressed from two promoters: KaiA has
its own promoter, whereas B and C are transcribed as a dicistronic messenger from a promoter
immediately upstream of KaiB. Transcriptional information is not directly required for circadian time
keeping. A sequential program of dual phosphorylation of KaiC forms the basis for the circadian rhythm. The phosphorylation cycle consists of
four steps (1) phosphorylation of threonine (T432),
(2) phosphorylation of serine 431 (S 431). This dual
phosphorylation converts KaiC from an autokinase to
an autophosphatase. (3) Dephosphorylation of T432
and (4) dephosphorylation of S431 and reconversion
of the KaiC autophosphatase to an autokinase. Nakajima et al.17 succeeded in reconstructing a robust
circadian oscillation of KaiC phosphorylation in vitro
by mixing KaiA, KaiB, KaiC, and ATP. So far, a complete reconstruction of the oscillator clock in vitro has
only been accomplished in cyanobacteria.
Although the mechanism of oscillation has
been elucidated, our understanding of the input and
output pathways is still limited. In particular, the
photoreceptor molecule remains to be identified. On
the input side, CikA protein has been identified
as a main component to provide environmental
input. It is a divergent bacteriophytochrome with
characteristic histidine protein kinase domains and
a cryptic response regulator motif. However, rather
than sensing the light directly, it seems to sense
the redox state of the cell, which is dependent on
illumination, but also on redox metabolism. So far,
no photoreceptor that directly relays light information
to the central oscillator has been identified.18
On the output side, SasA, a histidine protein
kinase, interacts directly with the oscillator protein
KaiC and activates the transcription factor RapA,
which can activate all promoters in the Synechocystis
genome. RapA in turn is negatively controlled by LabA
in order to control KaiBC in a circadian fashion.18,19
The question of whether the circadian clock
provides a selective advantage has been approached
experimentally and it has been clearly demonstrated
by direct competition experiments that resonating
circadian clocks enhance fitness in cyanobacteria.20
The Circadian Clock in Plants
The circadian clock in plants regulates a large number of biological processes, such as rhythmic leaf
movements, elongation rates of stems, hypocotyls and
roots, circumnutations of stems, primary and secondary metabolite biosynthesis, hormone biosynthesis
and responses, water-stress responses, water uptake,
opening of the stomata, seed dormancy, and defense
against pathogens (see Ref 21 for review). In some
plants, the clock is used to measure the environmental
photoperiod to induce inflorescence meristems, so that
flowering occurs during the correct season (photoperiodic flowering).22 These phenomena co-ordinately
contribute to increased fitness and therefore, confer a
selective advantage.
The powerful genetics of Arabidopsis led to
the isolation of a substantial number of clock
genes (see Ref 21 for review). However, these are
quite different from those of cyanobacteria. They
generate a cyclic negative feedback loop composed
of transcriptional repressor proteins: morning-phase
proteins, evening-phase proteins, and midday-phase
proteins. The sustainable oscillation is dependent on
similar decay rates for mRNAs and proteins, and large
amounts of proteins at their peak level. Whether a
posttranslational oscillator as found in cyanobacteria
functions also in plants is an open question.
A genetic approach also led to the isolation
of a cryptochrome blue light receptor.23 Arabidopsis
seedlings grown under light have a shorter hypocotyl
than seedlings grown in the dark; this response
is mediated by blue (420–500 nm), red, or farred (700–750 nm) light. Certain hy mutants have
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Advanced Review
selectively lost the capacity to respond to certain
portions of the spectrum. The hy4 (= cry1) mutant
has selectively lost its response to blue light. The
isolation and sequencing of the cry1 gene showed
that it encodes a protein with sequence similarity to
DNA photolyases. The CRY1 protein was shown to
be a flavoprotein, but it lacks photolyase activity
and it is the long-sought blue light receptor and
was named cryptochrome 1 (CRY1). Subsequent
experiments by Somers et al.24 showed that it
functions in the entrainment of the circadian clock
of Arabidopsis. Since then cryptochromes have been
found in many flowering plants and also in ferns and
Chlamydomonas algae. They also play a major role in
animals.
Circadian Oscillators in Fungi
Our knowledge about the circadian clock in fungi
is again limited to model organisms which are
amenable to molecular genetic analysis, in this case,
mostly Neurospora (see Ref 25 for review). Like
in cyanobacteria the biological clock consists of
interconnected positive and negative feedback loops:
The GATA-type transcription factors WC-1 and
WC-2 assemble to form the White-Collar complex
(WCC), which activates the clock gene frequency (frq).
FRQ protein then inhibits the activity of WCC and
thereby feeds back on its own synthesis. Owing to
this feedback, regulation frq RNA and FRQ protein
display high-amplitude circadian abundance rhythms
constituting the biological clock. In an interconnected
positive feedback loop, FRQ supports accumulation
of high levels of WCC at the posttranslational
level. The negative feedback loop takes place in the
nucleus, whereas the positive loop is confined to the
cytosol.26 The maturation of FRQ from a nuclear
repressor toward an inactive cytoplasmic repressor
is due to phosphorylation by casein kinase 1 a. In
the cytoplasm, WCC becomes dephosphorylated and
reactivated by the protein phosphatase PP2A.
On the input side, the WCC is directly activated
by light, which resets the clock. WC-1 protein binds
the flavin (FAD) as a chromophore and serves as
a circadian blue light receptor.27 In addition, a
hierarchical cascade of transcription factors activates
light-induced transcription of hundreds of target
genes. To prevent disturbances of the clock by changes
in light intensity during the day and to ensure
proper synchronization, the cells are desensitized to
ambient light by photoadaptation. In Neurospora,
photoadaptation depends on the blue light receptor
vivid (VVD) which accumulates immediately after
light activation and rapidly silences the expression
of WCC-dependent genes.28
4
Although some of these mechanisms are
reminiscent of the cyanobacterial clock, the clock
genes are completely different suggesting that the clock
may have evolved several times separately.
The Biological Clocks in Animals
We shall limit the discussion of animal clocks to
Drosophila and the mouse, the two model organisms
which are best studied (see Ref 29 for review).
The isolation of mutants in the period (per) gene
of Drosophila11 provided the first evidence that
daily rhythms in the sleep–wake cycle of animals
are genetically controlled. One mutation (pershort )
reduced the period of activity rhythm, whereas perlong
prolonged the period of activity, and a third mutation
per01 abolished the locomotor rhythm altogether. It
took another 15 years until the per gene was cloned
and available for molecular analysis.30 Hardin et al.31
showed that the per mRNA levels are cycling in
circadian rhythm, and that the cycling was advanced
in the pershort mutant, whereas it was delayed in perlong
flies indicating that the PER protein must feed-back
to regulate its own expression.
During the past 20 years research on the clock in
Drosophila and in parallel in the mouse has revealed
that the clock mechanism is remarkably conserved
between insects and mammals, including humans.
The regulatory scheme of the molecular clock
is outlined in Figure 1. It is basically a bimodal
switch of two negative feedback loops. The cycle
is driven by a heterodimer protein CLOCK-CYCLE
(CLK-CYC). This core heterodimer coordinates the
antiphasic expression of two sets of genes, the
morning and the evening genes that generate a bimodal
switch mechanism which is self-sustaining and takes
approximately 24 h (circadian) to complete. During
daytime the heterodimer CLK-CYC is active and the
E-box genes (including per, tim, vri, pdp-1ε, and
cwo) are switched on, whereas the V/P-box genes
(including Clk and Cry) are turned off due to rapid
accumulation of the VRI-repressor (see Figure 1).
At night, when CLK-CYC is inactive, E-box genes
are turned off, because the PER-TIM heterodimer
blocks CLK-CYC and direct repression of E-boxes
by CWO, whereas V/P-box genes are switched on by
PDP1ε which replaces VRI repression. The activity
of clock proteins is modulated by phosphorylation
by protein kinases leading to protein degradation,
whereas protein phosphatases stabilize the respective
proteins. The phosphorylation status also guides the
subcellular localization of the clock proteins which
accumulate either in the nucleus or the cytoplasm.
Light input to the clock is mediated by
the blue-light receptor protein called Cryptochrome
Evolution of vision
Morning transcripts
V/P-box = TTATGTAA
per, tim
vri, pdple,
cwo
Clk, Cry
ZT
0
3
Morning transcripts
V/P-box = TTATGTAA
Evening transcripts
E-box = CACGTG
6
9
12
PER-TIM
VRI
15
PDP1ε
18
CWO?
21
bHLH-orange
24/0
Clk, Cry
3
CRY
bHLH-PAS
PAS
bZIP
Cryptochrome
CWO
cwo
Clk
CLK
Cry
CYC
per
PER
tim
TIM
CRY
pdple
vri
PDP1ε
VRI
FIGURE 1 | The molecular circadian clock of Drosophila. Upper panel: Abundance profiles of head mRNA (dashed lines) and protein (solid lines)
for rhythmic components of the core oscillator mechanism. Numbers indicate Zeitgeber time (ZT), where 0–12 = lights on (white bar) and 12–24/0 =
lights off (black bar). mRNA transcripts that peak in the morning (blue) and evening (orange) are shown with the main cis -regulatory DNA sequence
that is targeted to generate rhythmic expression indicated above each peak. CRY protein is not shown, although it is degraded by light and, therefore,
accumulates during the night. The CYC protein levels are constitutive due to constant cyc expression. Lower panel: Flow diagram showing
transcriptional regulation from left to right and feedback from right to left in the core circadian mechanism. The panels are aligned and color coded so
that the interactions can be correlated with the abundance profiles, which they generate. Key: solid arrows = up-regulation; blunt-ended lines =
down-regulation; dashed arrows = transcription/translation; dotted line = probable interaction between CRY and PER in the dark; lightning bolt
depicts light-induced activation of CRY; inset depicts the major transcription factor families which are involved.29 (Reprinted with permission from Ref
29. Copyright 2011 the Biochemical Society)
(CRY).32 The central clock cells are neuronal cells,
i.e. photoreceptor cells and nine groups of specific
interneurons in the brain.29 However, there are
also non-neuronal clock cells (glial cells, cells of
the Malpighian tubules, and the rectum). There
are also clock cells in the abdomen, thorax, legs,
wings, antennae, proboscis, ocelli, retina, and brain.
These peripheral clock cells also synchronize with the
external light–dark cycle, which suggests that they
all express cryptochrome.33 However, those neurons
which do not express cryptochrome (CRY? neurons)
and cannot synchronize with the external light-dark
cycle, receive light input from photoreceptor cells in
the eyes and ocelli.34
The biological clock mechanism in mammals is
remarkably similar to that of Drosophila. Many of
the core factors of the clock are conserved between
Drosophila and mouse. The mammalian clock also
consists of a bimodal switch (interlocked-feedbackloop). However, during vertebrate evolution two
genome duplications have occurred, so that several
clock genes have been duplicated. For example, there
are two mammalian Clock (Clk) genes, and two cyc
genes, whose protein products form heterodimers and
activate E-box genes. There are also two cryptochrome
genes (Cry1 and Cry2) whose protein products no
longer respond to light. The only clock cells that
receive light input are located in the suprachiasmatic
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Advanced Review
nuclei (SCN) of the brain. Light input to the SCN
comes from a specialized group of Melanopsinexpressing retinal-ganglion cells in the eye.35,36 In
response to light, these retinal ganglion cells switch
the Per genes on in SCN neurons.37 Increased levels
of Per-Cry reset the circadian clock. The most striking
difference between Drosophila and mouse involves
feedback that generates the interlocked-feedbackloop, which involves a different class of genes than
those of Drosophila. However, the conservation of
the clock mechanisms between insects and mammals
is quite remarkable.
Comparing the clock mechanisms of cyanobacteria and plants, fungi, and animals, we have to assume
that there have been multiple circadian clock origins
during evolution.12 However, there are also highly
conserved elements, particularly between insects and
mammals. Also, the use of cryptochrome for resetting
the clock is very ancient and has been substituted
by melanopsin only in mammals. We shall trace
cryptochrome evolution further in the discussion of
visual pigments.
THE EVOLUTION OF PHOTOTAXIS
AND PHOTOTROPISM
The second line of evidence that cyanobacteria can
detect light comes from studies of phototaxis and
comparative genomics.
Phototaxis38 is a locomotory movement of a
whole organism in response to a light stimulus. Movement toward the light is called positive phototaxis,
whereas avoidance of illumination is designated as
negative phototaxis. In sessile organisms like plants
and fungi, the term phototropism is used for the
corresponding light response, meaning that growth
is determined by the direction of the light source.
In both cases, a mechanism for sensing light is
required.
Some prokayotes are phototactic. They often
use a random walk strategy, using a type I sensory
rhodopsin for light sensing and a two-component
signaling system to induce a reversal of the flagellar
beat. This strategy only allows phototaxis along a
steep light gradient, as found in microbial mats or
sediments. Some filamentous bacteria have evolved the
ability to steer toward a light vector. This allows them
to navigate in two dimensions, gliding on a surface.
In contrast, eukaryotes have evolved the capacity
to follow a light vector in three dimensions in open
water. This capability requires a polarized organism
with a defined shape, helical swimming driven by cilia,
and a shading device for the light sensor which allows
the organism to sense the direction of light. Such
6
an eye spot or prototypic eye confers a considerable
selective advantage, and preceeds the evolution of
more sophisticated eye-types.
Phototaxis in Cyanobacteria
Cyanobacteria, like Synechococcus and Anabaena can
slowly orient along a light vector.39 This phototactic
orientation occurs in filaments or colonies. It is a slow
gliding motility, a uniform motion on a solid surface
in a direction parallel to the cell’s long axis and occasionally interrupted by reversals.40 The unicellular
cyanobacterium Synechocystis sp. PCC 6803 shows
gliding motility on agar plates or glass slides, a small
and intermittent translocation on a solid surface, in
which the direction of movement often changes. This
species can be transformed genetically and a number
of genes involved in phototactic motility have been
identified.41,42 Wildtype Synechocystis cells form flat
sheet-like irregularly shaped colonies which can move
on solid surfaces. Mutant cells with disrupted pil
genes involved in the formation of type IV pili form
round colonies and have lost motility. Type IV pili
are 5–7 nm in diameter and several micrometers in
length and are involved in several functions, including
gliding movement, phototaxis, and DNA transfer to
other cells.
The action spectra for phototaxis are quite
complex, depending on the species of cyanobacteria used. However, in most species they overlap
with the absorbtion spectra of phycobilins, which
serve as chromophores in phytochromes. In Thermosynechococcus, the action spectrum shows several
peaks at 530, 570, 640, and 680 nm, but at higher
fluence rates the red action peak (at 640–680 nm)
disappeared and far-red peaks at 720 and 740 nm
emerged. When cells are illuminated simultaneously
with red and far-red light, phototaxis is drastically reduced, which also suggests that phytochromes
are involved.43 Two classes of cyanobacterial phytochromes have been found and were designated as
cyanobacteriochromes classes 1 and 2. In Synechococcus, mutations in the PixJ1 mutations in the PixJ1
(= Tax1) gene which abolish positive phototaxis,
but show instead negative phototaxis, have identified a photoreceptor for phototaxis.44 It contains
GAF and PAS domains as they are typically found in
phytochromes, which are the major red/far-red light
receptors in plants (see below). It has been shown that
the Tax1 homolog of Thermosynechococcus is localized at the poles of this rod-shaped cyanobacterium,
which suggests that it may be involved in sensing the
light direction.43 Negative phototaxis is controlled by
another cyanobacteriochrome locus42 and two adjacent response regulator loci which together encode
Evolution of vision
a UV-A-activated signaling system. The cyanobacteriochrome receptor gene encodes a protein with
three transmembrane domains two PAS, a GAF, and
a histidinekinase domain, which is characteristic for
phytochromes. Illumination by UV-A light (360 nm)
induces strong negative phototaxis which represents
a UV-avoidance reaction (see also cryptochromes).
In cyanobacteria, cyanobacteriochromes have undergone adaptive radiation and at least 41 different genes
have been identified encoding these phytochromes
involved in light perception. Whole genome sequencing of other cyanobacteria has identified several other
photoreceptors: In Anabaena, one sensory rhodopsin
(a green light receptor) has been found and two Cryptochromes/Photolyases (putative blue light receptors).
Gloeobacter has a proteorhodopsin gene closely similar to that found in dinoflagellates (see Russian Doll
Hypothesis). Gloeobacter also has a cryptochromelike gene, similar to animal cryptochromes which are
related to (6-4) photolyases (see cryptochromes).
Phototaxis in Halophylic Archaea
Archaea like Halobacterium salinarum have sensory rhodopsins for phototaxis.45,46 Rhodopsins are
seven transmembrane proteins which bind retinal as
a chromophore.47 Light triggers the all-trans/13-cis
isomerization of retinal,48 which leads to phototransductory signaling through a two-component phosphotransfer relay system. In H. salinarum two types of
rhodopsins, sensory rhodopsin I (SRI) and SRII have
been found, which signal via the halobacterial transducer proteins Htrl and Htrll.49,50 Signaling in phototactic archaea involves CheA, a histidine kinase, which
phosphorylates the response regulator CheY.51 Phosphorylated CheY induces swimming reversals. SRI
functions as an attractant receptor for orange light and
a repellent receptor for near-UV light, whereas SRII
is repellent receptor for blue light. This mechanism of
phototaxis only works with a steep light gradient.
Phototaxis in Algae and Protists
Red algae lack cilia in all stages of their life cycle and
therefore, the ability of helical swimming. Also, they
lack eye spots (stigmata) and as a consequence are
incapable to perform three-dimensional phototaxis.
However, they find their optimal light conditions by
surface gliding and two-dimensional phototaxis as
described for Porphyridium.52
Green algae (Chlamydomonas) have a photoreceptor organelle called eye spot or stigma
located in the outermost portion of the chloroplast,
directly underneath the two chloroplast membranes
indicating that it is derived from cyanobacteria
(see unicellular photoreceptor organelles). Indeed,
stigmata-like structures have been found in certain
cyanobacteria (see: The origin of vision: cyanobacteria). The stigma consists of tens to hundreds of
lipid globules containing carotenoid pigments which
provide a screening function, shielding the light from
one side and serving as a light reflector (Figure 2). The
stigma is not to be mistaken for the photoreceptor. The
stigma only provides directional shading for the adjacent membrane-bound photoreceptors. Stigmata can
also reflect and focus light onto the photoreceptors
like a concave mirror, thereby increasing sensitivity
(Figure 2). Carotenoids can also serve as light antennae: Xanthorhodopsin of the eubacterium Salinibacter
ruber contains a single energy-donor carotenoid (salinixanthin) noncovalently bound to a small 25 kDa
membrane protein with a single retinal acceptor.53
In the best studied green alga Chlamydomonas
reinhardtii phototaxis is steered by a rhodopsin pigment. This was first shown by restoring phototaxis in
blind mutants by retinal analogs.55 Genomic sequencing revealed two Channelrhodopsins-1 and 2.56–58
These proteins have an N-terminal 7-transmembrane
domain similar to archaebacterial proteorhodopsin
and a C-terminal membrane-associated domain. They
act as light-gated cation channels and trigger depolarizing photocurrents.56,57,59 Channelopsin-1 was
shown to localize in the stigma region by immunofluorescence studies.60
Dinoflagellates have evolved the most sophisticated photoreceptor organelles starting out from
relatively simple stigmata with carotene-containing
vesicles61 to some species like Cryptoperidinium
foliaceum with lamellar bodies consisting of regularly stacked membranes strikingly resembling mammalian rod and cone cells. Some dinoflagellates
like Warnowia and Erythropsis even have lenses
for focusing the incoming light to a retina-like
structure, the retinoid (see Unicellular Photoreceptor Organelles).62–64 Dinoflagellates possess two cilia,
a transverse cilium in the groove around the equator
of the cell and a longitudinal one inserting directly
behind the eye organelle, serving as a steering cilium.
Genomic studies have identified a proteorhodopsin
(type I rhodopsin) in Pyrocystis lunula,65 in Oxyrrhis
(Ernst Bamberg, personal communication) and in Erythropsis (Suga et al., unpublished data) which are
closely related to cyanobacterial proteorhodopsins
from Gloeobacter indicating that the eye organelle
is indeed derived from their chloroplasts.
Euglena belongs to a diverse group of biciliate protists which have taken up a secondary
plastid by endosymbiosis from an algal prey.
They are autotrophic, also phototactic and have
7
Advanced Review
wild-type cells the stigma blocks one of the preferred
directions.66 The paraflagellar body contains a flavinbinding photoreceptor, which in Euglena has been
identified as photoactivated adenylcyclase (PAC). PAC
mediates both positive and negative phototaxis.67
(a)
Phototropism in Flowering Plants
Phototropism in plants was already studied by Charles
Darwin who showed that the growing coleoptiles of
oat seedlings exposed to lateral illumination bend
toward the light, which is called positive phototropism. The curvature is caused by differential cell
elongation. If the tip of the coleoptile is covered by
an opaque cover, this bending is inhibited, indicating
that some factor is induced by light at the tip and
transmitted further down the coleoptile to induce the
bending. The respective substance was later shown by
Went68 to be a plant hormone, called auxin and its
structure determined as indole acetic acid.
Phototropism in flouring plants such as Arabidopsis is directed by blue light receptors called
phototropins (see below). Other photoreceptors found
in plants include phytochromes that sense red light
and cryptochromes which are also blue light receptors. The evolutionary history of these photopigments
is discussed below.
2 Hz
5 μm
Plasma
membrane
(b)
Retinal
switch
H+
ChR1
Ca2+
VGCC
H+
Na+
Ca2+
ChR2
Light
ΔΨ
Phototropism in Fungi
125 nm
= λ/4
FIGURE 2 | The eyespot of Chlamydomonas. (a) A Chlamydomonas
cell with two flagella, a large green chloroplast and a yellow-orange
eyespot (eye organelle). (b) Function of the eyespot, Channelrhodopsin
1 (ChR1) is a light-gated proton channel located in the plasma
membrane above the pigment spot. The entering protons diffuse
laterally and activate a voltage or proton-gated Ca++ channel (NGCC).
Channelrhodopsin 2 (ChR2) is conductive for H+ , Na+ and Ca++ . The
voltage change is transmitted along the membrane and sensed by
the VGCC channels in the flagellar membrane. A sudden Ca influx
induces a switch of flagellar motion. The carotene reflects the light to
activate the ChRs.54 (Reprinted with permission from Ref 54. Copyright
2004 American Physiological Society)
a red carotenoid-containing shading stigma and a
paraflagellar body, a photosensory structure, at the
base of the long cilium. In contrast to green algae, the
stigma in Euglena is not part of the chloroplast, which
is presumably the result of a secondary endosymbiosis. The presence of both the stigma and the adjacent
paraflagellar body are required for phototaxis. The
direction finding is brought about by a dichroic
mechanism, based on the paracrystalline structure of
the paraflagellar body. Stigma-less mutants are still
phototactic but swim in two directions, whereas in the
8
Fungi respond to light, particularly blue light to regulate development and behavior, presumably for the
optimization of spore production and dispersal, and
for growth. In the 1950s the use of Phycomyces in
sensory transduction research was promoted by the
Nobel laureate Max Delbrück.69 Blue light regulates
various aspects of the Phycomyces biology: it regulates
the development of fruiting bodies (sporangiophores),
stimulates β-carotene biosynthesis and modifies the
growth of sporangiophores in response to light, designated as phototropism. In organisms that do not move,
phototropism corresponds to phototaxis in motile
organisms. In addition, the Phycomyces sporangiophore can change the direction of growth after sensing
other environmental signals, like gravity, wind, touch,
and the presence of nearby objects, making this unicellular structure a unique experimental system.70
Phycomyces responds to a wide range of light
intensities, extending over 10 orders of magnitude, approximating that of the human eye. This
is achieved through the action of two photosystems operating at different light intensities. A genetic
screen for phototropic mutants in Delbrück’s laboratory led to the isolation of mad mutants and a
first outline of the sensory transduction pathway.71
Evolution of vision
The discovery of additional mad mutants and their
detailed characterization led to the identification of 10
unlinked genes, mad A through mad I.72,73 Mutants of
mad A and mad B are defective in phototropism and
other light responses suggesting that their gene products play key roles in Phycomyces photobiology.70
Most of our understanding of fungal photobiology comes from Neurospora (see above). Mutations in
wc-1 and wc-2 disrupt all the responses of Neurospora
to blue light. The WC-1 protein contains a zinc-finger
motif, a chromophore-binding domain (called LOV)
and PAS domains for protein–protein interactions.27
The LOV domain binds the flavin chromophore (FAD)
allowing WC-1 to function as a circadian blue light
receptor.
In Phycomyces madA encodes a protein similar
to the Neurospora blue light receptor WC-1, and
madB corresponds to WC-2. The proteins MAD A
and MAD B also contain a zinc-finger motif and can
interact to form a complex. These findings indicate
that phototropism and other responses to light
in Phycomyces are mediated by a photoresponsive
transcription complex as in Neurospora.74
Phototaxis in Metazoa
Phototaxis in Sponges
In the demosponge Reneira75 the embryos develop in
brooding chambers inside the sponge. The chambers
contain 20–150 embryos which develop into larvae.
The larvae have an outer layer of monociliated cells
possessing 20 μm long cilia, but there are two protruding bare patches of cells at both poles of the larva.
At the posterior pole there is a ring of pigmented cells
bearing much longer (120–150 μm) cilia, which are
thought to be the photoreceptor cells with steering
cilia. The larvae are released from the brooding chamber and in the presence of light, swim continuously
forward rotating clockwise (as observed from the posterior end of the larva). Upon release, the larvae swim
first upward from the adult sponge, but they are negatively phototactic until at least for 12 h after release. A
sudden increase in light intensity causes the long cilia
to freeze which presumably allows the larvae to steer
away from areas of bright light toward darker areas,
to settle down and form a new sponge underneath the
corals in the reef. Although all other metazoan phyla
have rhodopsin as photoreceptor pigment, the genome
of the sponge Amphimedon queenslandica76 does not
contain a rhodopsin gene, and searches for rhodopsin
genes in the marine sponge Microciona as well as
the freshwater sponge Ephydatia were negative.77 The
action spectrum of photosensitivity of Reneira75 has
a maximum in blue light (440 nm) and a smaller,
secondary response peak (at 600 nm) in orange-red
light. This suggests that the photoreceptor pigment in
sponges may be a flavin or carotenoid.
This is in line with findings in adult Demospongiae (Suberites domuncula)78–80 which have identified
cryptochrome as photoreceptor pigment based on the
light-reactive behavior of these siliceous sponges. They
have identified an intriguing quartz glass-based system of spicules, which can generate light enzymatically
with a luciferase; the spicules serving as optical waveguide and cryptochrome as photosensor. This suggests
that also the larvae of demosponges use cryptochrome
as a photoreceptor. Cryptochrome does have a flavin
chromophor as suggested by the spectral analysis.
Whether sponges never had a rhodopsin gene or
whether they have lost it, remains an open question.
Phototaxis in Cnidarians
In the planula larva of the box jellyfish Tripedalia the
most primitive kind of ‘eye’ has been found in the form
of single pigmented photoreceptor cells which are
interspersed between the ciliated epithelial cells covering the larva81 (Figure 2). These photoreceptor cells
are also carrying a flagellum (cilium) and microvilli
on their surface and they contain pigment granules.
The microvilli presumably contain rhodopsin and the
flagellum is involved in steering the larva which is
phototactic. The structure of these cells suggests that
they are the most primitive rhabdomeric photoreceptors, whereas the adult box jellyfish has highly evolved
eyes with multiple photoreceptor and pigment cells,
as well as lenses. The photoreceptors of the adult are
of the ciliary-type (see below). In adult Tripedalia, the
eyes are integrated together with a statocyst (gravity sensory organ) into so-called rhopalia, batteries of
sensory organs. Several lens and slit eyes are combined
with a statocyst in the four rhopalia, one on each side
of the box jellyfish. The sensory input is integrated by
nerve connections,82 which represent the first stages
of brain evolution. Therefore, we can assume that the
eye as a sensory organ has evolved before the brain,
providing sensory information which subsequently is
processed by the brain.4,83
Phototaxis in Bilateria
As in cnidarians the larvae of bilateria show the
most primitive optical apparatus. Trochophora larvae
of the marine annelid Platynereis possess a pair
of prototypic eyes consisting of two cells only, a
photoreceptor cell and a shielding pigment cell.84
These simple eyes mediate phototaxis. Selective
illumination of one eye changes the beating of adjacent
cilia resulting in a locally reduced water flow. Similar
to Chlamydomonas, the Platynereis larvae swim in
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Advanced Review
a right handed helix, i.e. in a clockwise forward
movement, rotating around their antero-posterior
axis. The shielding by the pigment cells results in
an on–off illumination of the photoreceptor cell, if
the larvae swims at 90◦ to the incident light, and
a constant illumination, if it swims toward the light
source. This allows the larva to determine the direction
of the light source, which confers a considerable
selective advantage. For negative phototaxis, shielding
by the pigment cell seems not to be required.
The eye spots of unicellular organisms have to
be considered as cellular organelles, whereas these
larval photoreceptors have to be designated as eye
organs because they consist of at least two celltypes a photoreceptor cell and a pigment cell. These
prototypic eyes mediate phototaxis as shown by laser
ablation experiments. Ablation of both eyes leads to a
complete loss of phototaxis, whereas larvae in which
only one eye was ablated are still capable of swimming
toward the light in most cases.
A first step of cellular differentiation leads to the
formation of a pigment cell and a photoreceptor cell.
The photoreceptor is a sensory-motor neuron which
regulates phototactic steering by acetylcholine and
is connected via its axon to two ciliated cells which
contain acetylcholine receptors. Therefore, phototaxis
in Platynereis larvae is due to direct sensory-motor
coupling between the photoreceptor cell and the
locomotor ciliary cell. In Tripedalia jellyfish larvae,
light detection and ciliary locomotor output are
still combined in a single cell (see above). In adult
Platynereis, the eyes are much more elaborate and the
sensory-motor neurons differentiate into sensory and
motor neurons and are connected by interneurons
which eventually become integrated in the brain.
Subsequently, the brain plays an increasingly
important role in processing the visual information.
PHOTOSENSORY PROTEINS
Phytochromes
The red/far-red regulators of photomorphogenesis in
higher plants are called phytochromes (see Ref 85 for
review). Phytochromes share a conserved photosensory protein core which comprises a PAS domain,
a GAF domain, and a Phy domain to which a
linear tetrapyrrole chromophore bilin is covalently
linked. There are three major bilins: phycocyanobilin
(PCB), phytochromobilin (PB), and biliverdin (BV).
Phytochrome-related photosensory proteins have not
only been found in higher plants, but also in cyanobacteria, nonoxygenic photosynthetic bacteria, nonphotosynthetic bacteria, unicellular algae, diatoms, and
fungi. The bilin chromophore is covalently attached
10
to a conserved cysteine residue in the phytochrome
apoprotein. Organisms using PCB or PB have a conserved cysteine in the GAF domain, whereas those
using BV have a conserved Cys N-terminal to the
PAS domain. Plant and some algal phytochromes
lack a specific histidine residue which serves as a
phosphorylation site for bona fide histidine protein kinases which are involved in signaling. The
photoswitching arises via photoisomerization of the
bilin chromophore about its 15/16 double bond, followed by a protein–chromophore relaxation process
that shifts the absorbance spectrum further. In most
phytochromes, the photoproduct is metastable and
slowly reverts to the stable dark state. This process is known as dark reversion. The conformational
changes occurring upon photoisomerization alter the
chromophore–protein interactions, which trigger signaling. Phytochromes typically have a C-terminal
histidine protein kinase domain.
Light-induced protein kinase activity was first
reported in cyanobacteria.86 In Synechocystis, it has
been shown that the pixJ1 gene which encodes a
phytochrome-like photosensory protein is essential
for phototaxis.87 Site-directed mutagenesis revealed
that a Cys-His motif in the second GAF domain
binds a linear tetrapyrrole chromophore characteristic
for phytochromes. PixJ1 protein showed a photoreversible conversion between a blue light-absorbing to a
green light-absorbing form, which in the dark reverts
to the blue light-absorbing form. In contrast to the
plant phytochromes, red or far-red light irradiation
produced no change in the spectra of PixJ1. Therefore, this and other cyanobacterial phytochromes have
been named cyanobacteriochromes.
Cryptochromes
The power of Arabidopsis genetics led to the first
isolation of a cryptochrome blue light photoreceptor protein.23,88 Arabidopsis seedlings grown under
light have a shorter hypocotyl than seedlings grown
in the dark. This response can be mediated by
blue (420–500 nm), red (600–700 nm), or far-red
(700–750 nm) light. Certain Arabidopsis mutants
have lost the capacity to respond to blue light (cry
1− ) or to red light (PhyB− ), respectively. The Cry1
gene was isolated and shown to be a flavoprotein with
sequence similarity to DNA photolyase, but it lacked
detectable photolyase activity and it contained a distinct C-terminal extension (Figure 3(a)). On the basis
of these properties, it was concluded that Cry was
the long-sought blue light receptor and was named
cryptochrome 1.88,89 Cryptochromes mediate various
light responses, including the entrainment of circadian
Evolution of vision
clocks in Arabidopsis, Drosophila, and mammals (see
above). Blue-light reception and circadian clocks have
coevolved90 and show some important evolutionary
features.
The Evolutionary Origin of a Novel Gene
Encoding Cryptochrome
One of the fundamental problems in evolutionary
biology concerns the origin of genes with a novel
function. The cryptochromes allow us to address this
question in a profound way.90 There at least two
strong evolutionary selective driving forces for the
evolution of photosensory proteins; sunlight as an
energy source, especially for autophototrophic organisms, and UV avoidance to prevent DNA damage.
Geological studies show that there was little oxygen
in the atmosphere at Precambrian times before the
evolving autophototrophic plankton generated large
amounts of oxygen as a by-product of photosynthesis.
Therefore, there was no protective ozone layer, and
primitive organisms were exposed to heavy doses of
UV irradiation during daytime. UV irradiation causes
DNA damage mainly by inducing cyclobutane pyrimidine dimers and 6-4 pyrimidine-pyrimidone adducts
between adjacent pyrimidine bases on the same DNA
strand which lead to mutations. Bacteria have evolved
DNA repair enzymes called DNA photolyases that are
capable of repairing these photoadducts. Photolyases
are flavoproteins mediating DNA repair in a lightdependent manner.92 They contain a flavin adenine
dinucleotide (FAD) as a catalytic chromophore and a
second chromophore which is either methenyltetrahydrofolate (MTHF) or deazaflavin (7,8-didemethyl-8hydroxy-5-deazariboflavin; 8-HDF) which serves as a
light-harvesting antenna. Only the fully reduced form
(FADH− ) of flavin is catalytically active. During light
activation, light energy is mainly absorbed by the
light-harvesting chromophore. After light absorption
by the light-harvesting chromophore, the excitation
energy is transferred to the fully reduced flavin chromophore by resonance transfer. An electron is then
transferred from high-energy FADH−∗ directly to the
pyrimidine dimer, which splits the cyclobutane ring
and restores the two pyrimidine monomers. There are
two classes of photolyases (types I and II) which repair
cyclobutane pyrimidine dimers (CPD) and another
class involved in the repair of (6-4) photoproducts first
identified in Drosophila.93 The fact that photolyases
are activated by blue light may not be coincidental,
because only blue light reaches substantial depth in
water.
Photolyases and cryptochromes not only share
extensive sequence similarity, but they also have
very similar 3D structures,91 the only difference
is the extended C-terminal part of cryptochromes
(Figure 3(a)). Photolyases are found in almost all
species of prokaryotes and eukaryotes, whereas cryptochromes are present in higher plants and most
animals, but only in a few other eukaryotes and
prokaryotes, suggesting that photolyases are ancestral to cryptochromes. The phylogenetic analysis
(Figure 3(b)) indicates that the classic plant cryptochromes (green) which have lost photolyase activity
have evolved from CPD-photolyases (classes I and
III), whereas DASH (Drosophila, Arabidopsis, Synechocystis, Homo) cryptochromes are more closely
related to class II CPD-photolyases. Interestingly,
DASH cryptochromes have retained photolyase activity, but they can repair only single-stranded DNA, and
not double-stranded DNA substrates, whereas classic
plant and animal cryptochromes have lost DNA repair
activity completely. Animal cryptochromes (pink and
violet) are more closely related to 6-4 photolyases. The
tripartite structure of the phylogenetic tree suggests
that the transition from photolyases to cryptochromes
has occurred three times during evolution and that
DASH cryptochromes are in a transition stage because
they function both as blue light receptor and a DNA
repair enzyme, because they have partially retained
DNA repair activity. Mammalian cryptochromes have
lost their photoreceptor activity and retained only
their function as transcriptional regulators. Whereas
rhodopsin, the most common photoreceptor in animals, is localized in membranes at the periphery of the
photoreceptor cells, cryptochromes are localized in the
nucleus, because they are derived from DNA-repair
enzymes, and they were converted to transcriptional
regulators which have to be active in the nucleus.
Therefore, a gene encoding a DNA-repair enzyme has
evolved into a gene encoding a blue-light photoreceptor, and finally a transcription factor.
Magnetoreception by Cryptochromes
In addition to its function as a blue light/near UV
photoreceptor in growth regulation, flowering, circadian clocks etc., there is some recent evidence that
cryptochromes are also involved in magnetoreception in both animals and plants. A large variety
of animals has the ability to sense the geomagnetic
field and use it as a source of directional information, as a magnetic compass. The biophysical basis of
magnetoreception is not known, but a specific model
for magnetoreception in birds has been proposed.94
Behavioral experiments indicated that the magnetic
compass of migratory birds is an inclination compass,
which is sensitive to the axis, and not to the polarity of the magnetic field lines.95,96 Magnetic compass
11
Advanced Review
E.coli photolyase
MTHF
30% identity
45% similarity
FAD
At cryptochrome
Carboxy terminus
is
bid
op
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Xenopus
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Anthera
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Key
Animal cryptochrome
DASH-related cryptochrome
DASH cryptochrome
CPD photolyase class II
Additional bacterial protein
CPD photolyase class I
CPD photolyase class II
Plant cryptochrome
6-4 photolyase
Dual-type
FIGURE 3 | Phylogenetic tree of the family of cryptochromes/photolyases. Upper panel: Domain structure of cryptochromes and photolyases from
Escherichia coli and Arabidopsis thaliana (At), respectively. The N-terminal domain which binds two chromophores methenyltetrahydrofolate (MTHF)
and flavin adenine dinucleotide (FAD) shown in yellow is highly conserved among all classes of cryptochromes and photolyases. By contrast, the
c-terminal extension (black) is variable and not found in photolyases. Lower panel: Phylogenetic tree of cryptochromes/photolyases. Green: plant
cryptochromes. Pink and violet: two groups of animal cryptochromes. Blue: DASH cryptochromes and some related proteins. Red, brown, and gray:
CPD photolyases classes I, II, and III. An asterisk (*) marks dual-type belonging to either class I CPD or (6-4) photolyases capable of DNA repair, but
they also possess photoreceptor activity. CPD, cyclobutane pyrimidine dimer; DASH, Drosophila, Arabidopsis, Synechocystis, Homo. (Reprinted with
permission from Ref 91. Copyright 2011 Annual Reviews Inc)
12
Evolution of vision
orientation is dependent on the wavelength of ambient
light. Orientation is possible in blue and green light,
but under yellow-orange-red light the birds are disoriented. Neurophysiological experiments also indicate
that magnetoreception depends on light and an intact
retina.97 The model proposed by Schulten et al.94
assumes that magnetoreception involves radical-pair
processes that are governed by anisotropic hyperfine
coupling between (unpaired) electrons and nuclear
spins. They show theoretically, that the Earth’s magnetic field strength (∼0.5 G) can produce significantly
different reaction yields for different alignments of
the radical pairs with the magnetic field. However, to
function as magnetic compass sensors absorption of
blue light must produce radical pairs with lifetimes in
the range of milliseconds. Using cryptochrome 1a from
the migratory garden warbler, it can be shown that
these flavoproteins indeed form radicals with lifetimes
in the range of msec, when excited by blue light.98
In Arabidopsis, it has been shown that an
increase in intensity of the ambient magnetic field
enhances hypocotyl growth inhibition under blue light
when cryptochromes serve as photoreceptors, but not
under red light when phytochromes are the active photoreceptors. In Cry− mutants lacking cryptochromes
no increase in growth inhibition is observed.99
In Drosophila cryptochromes are also implicated magnetoreception. In a binary-choice T-maize
behavioral assay in which a magnetic field is produced on one side, whereas the other side has no
exogenous magnetic field, the flies can be entrained
to find their food on one side of the maize under
full-spectrum light (300–700 nm), but not if the UV
A/blue parts of the spectrum (<420) are blocked.
Cryptochromes (Cry− ) deficient mutants do not show
any responses to the magnetic field under full spectrum
lights indicating that Cry+ is required for magnetoreception. These findings are supported by a separate
line of evidence:100 In response to blue light Cry
causes a slowing down of the circadian clock of the
wake–sleep cycle, ultimately leading to arrhythmic
behavior. Exposure of flies to magnetic fields, with
a maximum effect at 300 μT enhances the slowing
down of the clock. This effect of the magnetic field
was only present in blue light, but absent in red-light.
No effects of the magnetic field were found in Cry−
mutants. These genetic experiments clearly implicate
cryptochromes in magnetoreception.
With respect to the biophysical basis of
magnetoreception the widely held view that radical
pairs generated by three trytophane residues in
cryptochromes mediate Cry’s ability to sense magnetic
fields, does not seem to be correct. Mutating the
tryptophane residues to phenylalanine did not affect
the ability of transgenic flies to respond to the
magnetic field.101 However, recent data indicate
that magnetically sensitive light-induced reactions in
cryptochrome are consistent with its proposed role as
a magnetoreceptor.102
Rhodopsins
Rhodopsins are membrane proteins with seven helical transmembrane domains which form an internal
pocket in which the chromophore retinal is bound.
Retinal is bound to the ε-amino group of a specific
lysine residue in the α-helix VII forming a protonated Schiff base. The action of light involves specific
isomerization of a double bond in the chromophore
leading from 11-cis retinal to all-trans retinal in mammalian photoreceptor proteins, and from all-trans to
13-cis retinal in bacteriorhodopsin. This isomerization
induces a conformational change of the protein leading to signal transduction. Earlier work on retinylidene
proteins has been comprehensively reviewed by Spudich et al.47 The extensive work on structure and
function of rhodopsin by Hubbell et al.103 The Handbook of Photosensory Receptors104 is also a most
valuable source of information.
Rhodopsins can be subdivided into two large
categories, the microbial rhodopsins, a heterogeneous
group which includes archaeal light-driven ion pumps
and sensory rhodopsins and halorhodopsins, bacterial
rhodopsins from uncultured marine bacteria, fungal
opsins, rhodopsins from cyanobacteria and dinoflagellates, and channelopsins from green algae. The second
large group comprises the animal rhodopsins which
are G-protein-coupled receptors (GPCRs), which
include mammalian rhodopsin, one of the most extensively studied photoreceptor proteins.
Microbial Rhodopsins
Microbial rhodopsins are phylogenetically and
functionally diverse105 (Figures 4 and 5). An unrooted
phylogenetic tree shown in (Figure 4) reflects this
heterogeneity.
Archaeal Rhodopsins
Primarily archaeal rhodopsins are ion pumps, either
proton pumps or Cl− pumps capable of converting
light energy into chemical energy. The resulting proton gradient is used to drive ATP synthesis. These
type I rhodopsins are found in all three domains of
life suggesting that they existed before the separation
of archaea, eubacteria, and eukaryotes. Therefore,
light-driven proton transport as a means of procuring
energy may have predated the evolution of photosynthesis in cyanobacteria. In fact, the phylogenetic
13
Advanced Review
FIGURE 4 | Unrooted phylogenetic tree of microbial rhodopsins. Source: Hiroshi Suga.
Proteorhodopsin Halorhodopsin
(proteobacteria) (haloarchaea)
Sensory rhodopsin Itransducer complex
(haloarchaea)
Anabaena
sensory rhodopsin
(cyanobacteria)
Chlamydomonas
CSRA
Chlamydomonas
CSRB
D
D
D
D
E
E
E
Y
Y
S
H
H
H+
125
Cl–
250
300
110
130
Cytoplasm
Photosensory
signal
(physiological
function unknown)
Membrane
currents
His-kinase
Flagellar
motor
P
Regulator
P
Regulator
Flagellar
axoneme
FIGURE 5 | Structure and function of various microbial rhodopsins.105 Proteorhodopsin, halorhodopsin, sensory rhodopsin of Anabaena and the
two channelopsins function as ion channels, whereas sensory rhodopsin I is coupled to a transducer complex, which transmits the light signal via a
histidine kinase and phosphorylates a regulator of the flagellar motor. (Reprinted with permission from Ref 105. Copyright 2005 Wiley-VCH)
14
Evolution of vision
analysis suggests that cyanobacteria have acquired
rhodopsins by horizontal gene transfer because Nostoc resides with archaeal rhodopsins and Gloeobacter with the Dinoflagellates which probably took
up a cyanobacterium as an endosymbiont which
became ‘domesticated’ and turned into a chloroplast (see Russian Doll Hypothesis). In H. salinarum,
four types of rhodopsins have been characterized:
Bacteriorhodopsin which is a light driven proton
pump absorbing maximally at λmax 568 nm,106 which
generates a positive outside membrane potential.
Halorhodopsin is a chloride transporter107 which
hyperpolarizes the membrane by chloride uptake
rather than proton ejection. Sensory rhodopsins, SRI
and SRII, are phototaxis receptors controlling the
cells motility in response to changes in light intensity and wavelength.108 SRI has a λmax at 587 nm
and induces positive phototaxis (attraction) to orange
light and is repelled by near-UV wavelength. SRII
appears to have only a repelling function.48 It absorbs
maximally in the spectral peak of sunlight and, therefore, is optimally tuned to efficiently detect the light
and guide the cells to darkness under conditions in
which photo-oxidative damage might occur. Sensory
rhodopsins interact with the HtrI and HtrII transducer molecules109,110 that control a histidine protein
kinase cascade modulating the flagellar motors.108
In cells devoid of Htr transducer SRI is reconverted
from a sensory rhodopsin into a light-driven proton
pump.
Eubacterial Proteorhodopsins
More recently, rhodopsins were also discovered
in proteobacteria, mainly in marine environments
by shot gun random sequencing. In the γ proteobacterium SAR86 from surface water of Monterey Bay a rhodopsin functioning as a proton pump
was found.111 In contrast, in the cyanobacterium
Anabaena (also called Nostoc) it has a photosensory
function.112 Also in the deep-sea at 75 m depth a proteobacterial rhodopsin was found with a blue-shifted
absorption spectrum (λmax = 490 nm).
This blue-shift can be explained by the fact
that only blue light penetrates deeply into the water.
Recent studies indicate that Anabaena photosensory
rhodopsin interacts with a tetrameric transducer
ASRT (Anabaena Sensory Protein Transducer) that
binds to the promotors of several genes related to the
utilization of light energy.113
the counterion to the retinal Schiff base is conserved in
all fungal rhodopsins homologs except for Cryptococcus, which contains an Ala at this position. Genomic
sequencing of Neurospora revealed the first eukaryotic homolog of archaeal rhodopsin called NOP-1. It
is probably serving a sensory function.114 Neurospora
is non-motile, but in the motile fungus Allomyces
phototaxis of the zoospores was shown to be retinal
dependent.115 However, no animal-type rhodopsin II
has been found in fungi.104
Channelrhodopsins in Green Algae
Positive and negative phototaxis in the green alga
C. reinhardtii is associated with electrical currents
carried by Ca++ and H+ which can be measured
within <30 μs after flash stimulation in the eye spot
(Figure 2). Searching an EST (expressed sequence tag)
data base, two cDNA sequences were discovered
encoding microbial-type rhodopsins, Channelopsin-1
(Chop-1) and Channelopsin-2 (Chop-2).56,57 Expression of the respective RNAs in Xenopus oocytes and
mammalian tissue culture cells produces light-gated
conductance that is highly selective for protons in
ChR1, whereas ChR2 generates a large permeability for monovalent and divalent cations. ChR1 is a
directly light-gated proton channel, and ChR2 is a
directly light-switched cation-selective ion channel. In
contrast to the sensory rhodopsins of haloarchaea
which are coupled to a transducer a histidine kinase
signaling to the flagellar motor (Figure 5), the Channelrhodopsins are not coupled to any transducer,
i.e. the Channelrhodopsins are directly light-gated
ion channels which open in response to a conformational change of the protein after light-induced
isomerization of all-trans retinal to 13-cis retinal. This
makes them an important tool to activate excitable
cells like neurons, by illumination.116 It is possible to trigger behavioral responses in the nematode
Caenorhabditis elegans by Channelrhodopsin-2 simply by illumination.116 Both Channelrhodopsin-2 and
the light-driven Cl-pump halorhodopsin (see Figure 5)
can be used as optogenetic tools, as genetically
encoded switches that enable neurons to be turned
on by bursts of light. Both rhodopsins proved to be
functional in model organisms like C. elegans and
zebrafish, and even to reinstall rudimentary visual
perception in mice. This raises great hopes for gene
therapeutic approaches to overcome blindness.117,118
Rhodopsins in Dinoflagellates
Fungal Rhodopsins
Homologs of archael rhodopsins are also found in
fungi; in ascomycetes like Botrytis and in basidiomycetes such as Ustilago and Cryptococcus. Asp-85,
The phylogenetic tree shown in Figure 4 indicates
that the three partially characterized rhodopsins
from dinoflagellates are closely related to that
of Gloeobacter, a cyanobacterium. As shown for
15
Advanced Review
HOOC
A P A V Q S
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FIGURE 6 | Schematic representation of bovine rhodopsin inserted in the disc membrane. On the basis of crystallographic data.122,127 H1–H7,
transmembrane α-helices; H8, short cytoplasmic α-helix. (Reprinted with permission from Ref 127. Copyright 2005 Wiley-VCH)
Erythropsis, the rhodopsin seems to be localized in the
eye organelle which is thought to be derived from a
secondary chloroplast (see Russian Doll Hypothesis).
As chloroplasts are derived from cyanobacteria, it
is not surprising that the Dinoflagellate rhodopsin
may be derived from cyanobacteria. The functional
analysis by expression in frog oocytes and mammalian
tissue culture cells indicates that the rhodopsin gene
of Oxyrrhis indeed encodes a green-light receptor
(E. Bamberg, unpublished).
Animal Rhodopsins
Animal rhodopsin is the molecule of ultimate
sensitivity because it allows the detection of a single
light quantum.119–121 This may explain why this
molecule has been so successful and is found in all
metazoan phyla, except for the sponges which are
sometimes classified as parazoa, because they do not
form neurons or muscle cells.
High-resolution structures were reported for
bovine122 and squid rhodopsin.123
The structure of bovine rhodopsin is shown
schematically in Figure 6 which is based on X-ray crystallographic studies by Palczewski et al.122 and refined
by Teller et al.124 and Okada et al.125 All known animal rhodopsins carry the same chromophore, 11-cis
16
retinal (or its close analogs) covalently bound to a
specific lysine in the 7th α-helix of its apoprotein
(opsin) via a Schiff base linkage which isomerizes to
all-trans retinal upon illumination. As shown by Palczewski et al.122 ca. 30 amino acid residues contribute
to the binding pocket for 11-cis retinal. The Schiff
base linkage to lysine (K296) is positively charged
and the counterion is glutamate (E113) in α-helix
3, whose presence was predicted 30 years ago.126 A
most impressive amount of work went into the identification of the function of the individual amino acids
by generating site-directed mutations in a rhodopsin
gene that was totally synthesized by Khorana and coworkers (see Hubbell et al.103 ), so that rhodopsin has
become the best known membrane protein.
The seven transmembrane helices form a channel
as in bacteriorhodopsin and in Channelrhodopsin,
but in animal rhodopsins the signal from the lightactivated rhodopsin is not transmitted directly, but
rather amplified by a G-protein coupled enzyme
cascade. Therefore, animal rhodopins are members
of the very large class of GPCRs (Figure 7).
There are two major kinds of animal rhodopsins
with different G-protein coupled enzyme cascades
which are illustrated in Figure 8. They are found in
two different kinds of photoreceptor cells, the ciliary
Evolution of vision
FIGURE 7 | Phylogenetic tree of animal opsins. Maximum likelihood tree. Shaded areas represent cnidarian rhodopsins. Source: Suga et al.77
and the rhabdomeric photoreceptor cells. The ciliary photoreceptor cells (Figure 8(a)) contain c-type
rhodopsins and induce a phosphodiesterase which
hydrolyzes c-GMP, and the decrease in c-GMP levels leads to the closure of cGMP-gated Na+ /Ca2+
channels in the plasma membrane of the photoreceptor cells and therefore, to a hyperpolarization
which can be recorded in an electroretinogram. In
contrast, rhabdomeric photoreceptor cells contain
r-type rhodopsins123 and use a different G-protein
coupled signaling cascade (Figure 8(b)) which activates phospholipase C leading to a rise in phosphoinositol and intracellular Ca2+ and resulting in the
opening of TRP- or TRPL-channels and a depolarization of the membrane potential. It is frequently
stated in the literature that ciliary photoreceptors are
17
Advanced Review
(a)
Discs
Cilium
(b)
Rhabdomere
TRRP
Transient receptor potential
ion-channel
FIGURE 8 | G-protein coupled signal transduction in ciliary and rhabdomeric photoreceptors.134 (a) In ciliary photoreceptors, c-opsin receives the
light signal and transmits it via a trimeric G-protein complex to phosphodiesterase (PDE) which leads to a decrease in cGMP levels and to a closure of
cGMP-gated channels in the plasma membrane. Ciliary photoreceptor form membrane stacks (discs) from the ciliary membrane. By contrast
(b) rhabdomeric photoreceptors transmit the light signal from r-opsins via a different G-protein complex to phospholipase C (PLC) which cleaves
phosphoinositol (PIP2) leading to an increase in intracellular calcium and the opening of Na+ , Ca++ channels and depolarization of the plasma
membrane. Rhabdomeric photoreceptor form membrane stacks from microvilli. (Reprinted with permission from Ref 134. Copyright 1997 Blackwell
Science)
found in vertebrates, whereas invertebrates would
have rhabdomeric receptors. This notion is incorrect;
ciliary photoreceptors which have specializations of
the plasma membrane covering the cilium are found
not only in vertebrates, but also various invertebrate
phyla like Cnidaria, Ctenophores, and Bryozoa, as
well as in embryos of Arthropods and Cephalopods,
and in larvae of Urochordates.128,129 Rhabdomeric
photoreceptors are present in Arthropods, Echinoderms, and Sipunculans. However, both types, ciliary and rhabdomeric coexist in Annelids and Molluscs, Cephalochordates, Nemerteans, Nematodes,
18
Platyhelminthes, and rotifers. In fact, the eye of the
scallop (Pecten) which has both a lens and reflecting
mirror which project onto two retinae, one with ciliary
photoreceptors which hyperpolarize, and the other
with rhabdomeric photoreceptors which depolarize
when illuminated.130 The hyperpolarizing photoreceptor expresses a different kind of rhodopsin which
is coupled to the α subunit of a Go type G-protein. The
hyperpolarizing response in scallops is due to opening
of a cGMP sensitive potassium channel which is different from that of vertebrate cells, i.e. closing of cGMP
sensitive cationic channel.131–133 Thus, it is most likely
Evolution of vision
that the scallop Go− mediated phototransduction
cascade is probably coupled with a guanyl cyclase
to elevate cytosolic cGMP concentration.
Opsin evolution has recently been reviewed.135,136 The phylogenetic tree of the opsin
genes (Figure 7) indicates that the Go− , Gc− , and
Gr− rhodopsins diverged before the divergence of
protostomes and deuterostomes. It is interesting to
note that the mechanism of photo response in the
scallop is shared by some vertebrates: the photoreceptor cells of the parietal eye of a lizard respond
to light by increasing cytosolic cGMP and opening
channels137 suggesting the presence of a similar Go−
mediated phototransduction mechanism. The presence of the three types of photoreceptors in both
proto- and deuterostome species suggests that they
were already there in the last common ancestor. In
the box jellyfish, the larva of Tripedalia seems to have
the most primitive kind of single-celled rhabdomeric
photoreceptors (see below), whereas the adult has
elaborate lens eyes with ciliary photoreceptors. The
electrophysiology and biochemistry of larval photoreceptor cells remains to be done, but the morphology
strongly supports the argument81 that cnidarians, the
most primitive animals with eyes, already possess
both types of photoreceptors ciliary and rhabdomeric.
The phylogenetic tree of animal rhodopsins (Figure 7)
shows that two different subfamilies of opsins are
found in ciliary and rhabdomeric photoreceptors,
respectively, c-opsins and r-opsins. In addition, there
is a third subfamily called Go opsins (coupled to Go
proteins) and a more heterogeneous group including
retinochrome, peropsin, and neuropsin which serve as
photoisomerases.
The analysis of jellyfish opsins77 revealed a
surprising diversity of sensory molecules reflecting
an early evolutionary radiation. The hydrozoan
Cladonema which possesses eyes expresses as many as
18 different opsin genes, whereas Podocoryne which
lacks eyes still expresses two opsins. The conserved
lysine to which the chromophore 11-cis-retinal binds,
is present in all of them suggesting that they are
indeed used for photoreception. However, only 7 of
the 18 opsins are expressed in the eyes of Cladonema,
whereas as 2 genes are expressed ubiquitously, 4 on
the tentacles only and 5 on the manubrium and later
in the gonads. Phylogenetic analysis suggests that the
manubrium-specific genes are derived from the eyespecific opsins and may have been recruited into the
gonads for controlling the spawning process which is
light-controlled.
In contrast to this, large diversity of opsins in
cnidarians, no opsins have been found in marine
and freshwater sponges which use cryptochromes for
vision79,138 and also in choanoflagellates their putative
protist ancestors.77
Genome-scale analyses corroborate the notion
that cnidarian opsins can be classified into three
groups representing the orthologs of the bilaterian
R-, C-, and Go/RGR opsin subfamilies135 prior to the
evolutionary divergence of bilateria. The closest outgroup among the functionally characterized GPCRs
are melatonin receptors. X-ray crystallographic studies indicate GPRCs like the β2 adrenergic receptor have
a structure which is almost superimposable to bovine
rhodopsin.139 Pisani et al.136 propose a maximally
parsimonious scenario of metazoan opsin evolution
leading from the duplication of the melatonin-opsin
ancestor to the eumetazoan stem lineage. Two further
gene duplication events in the eumetazoan stem lineage subsequently separated the R-opsin from the Cplus Go/RGR lineage, and finally the c-opsins from the
Go/RGR opsins. Studies on the box jellyfish Carybdea
rastoni140 have revealed a fourth class of G-protein
coupled rhodopsins which are designated as Gs opsins
and may be typical for the majority of cnidarian
rhodopsins. The Gs-type G-protein mediated phototransduction cascade is used in the highly evolved
ciliary photoreceptor cells of Carybdea and signals
through cAMP rather cGMP. Illumination results in
cAMP levels in the photoreceptor cells. As both the
vertebrate and the box jellyfish phototransduction cascades involve cyclic nucleotide signaling there are clear
similarities between the two. In summary, we can distinguish five different kinds of G-protein coupling; Gt
(transducin) coupling in c-opsins, Gs coupling in box
jellyfish, and Go coupling in scallop ciliary photoreceptors, all of which signal through cyclic nucleotides.
Gq coupling in rhabdomeric photoreceptors in insects,
cephalopods, and human melanopsin which signal
through phosphoinositol. Finally, a fifth class of
rhodopsin molecules which serve as photoisomerases
and are found in humans, amphioxus, squid etc.
As far as the connection of metazoan rhodopsins
to microbial rhodopsins is concerned, we favor a
slightly different view. There is hardly any sequence
conservation which would help to find the origin
of animal rhodopsins. However, there are clearly
similarities which may not be accidental. First, the
seven transmembrane structure with retinal attached
via a Schiff base linkage to a specific lysine residue
in helix 7 is shared. If we compare the rhodopsins
from Cladonema with those of the cyanobacterium Gloeobacter and the three rhodopsins from
dinoflagellates, there may be traces of sequence conservation, which, of course, are very scarce considering
that the split has occurred at Precambrian times.
Besides the homologies in helix 7 (K296, A295,
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Advanced Review
and L293), some homologies can also be detected
in helix 6: Y268 which forms part of the retinal binding pocket and W265, and in helix 4 there is a highly
conserved G174, and in helix 3 a conserved P142.
This indicates that the major evolutionary difference
between microbial and animal opsins is the coupling
to different G-proteins. This is reflected by sequence
homology between the cyclic AMP chemoreceptor of
Dictyostelium and animal rhodopsin.141 There is 22%
amino acid sequence identity between the N-terminal
regions of the two receptors and 32% sequence similarity, if one considers conservative replacements. This
indicates that animal rhodopsins are clearly members of the large family of GPCRs, closely related to
chemoreceptors. This is of particular interest because
Pax6 is not only a master control gene for eye development but also for the nose or antenna, respectively,
with their odor and gustatory receptors. It is difficult
to decide whether animal rhodopsins are derived from
microbial rhodopsins or whether they have evolved
completely independently as G-protein coupled photoreceptors. As all rhodopsins use retinal as a ligand
it seems reasonable to assume that rhodopsin evolved
from a GPCR that acquired the ability to covalently
bind its ligand retinal and thus allowing it to evolve to
a photoreceptor molecule. However, it seems unlikely
to me that the covalent linkage of retinal to a specific
lysine residue in the 7th α-helix could have occurred
several times independently in microbial rhodopsins
and GPRCs. Also the conservation of tryptophane
(W265) and tyrosin (Y268) in α-helix 6 reflect strong
similarities between microbial and animal opsins. This
raises the possibility that Exon shuffling caused by
recombination events between microbial rhodopsin
and different GPCRs led to the rapid evolution of the
four types of metazoan opsins. The distal exons for the
transmembrane domains (TM) 6 and 7, in particular,
TM 7 with the retinal attachment site at lysine 296,
may have originated from a microbial rhodopsin gene
and the more proximal exons including the ERY motif
in helix 3 and the loop between TM 5 and 6 which
serve to bind the G-protein may be derived from the
respective GPCR genes. As more data accumulate it
may be possible to discriminate between these and
other hypotheses.
In addition to its well-known function in
vision, rhodopsin also has role in temperature
discrimination.142 As shown in Drosophila larvae
mutations in the nina A gene encoding rhodopsin,
a GPCR, eliminates thermotactic discrimination
in the comfortable temperature range from 19◦
to 24◦ C. The thermosensory signaling pathway
includes a heterotrimeric guanine-nucleotide-binding
protein (G protein), a phospholipase C, and the
20
transient receptor potential TRPA1 ion channel, as
for vision; however, thermotaxis toward 18◦ C is
light-independent.
Another interesting nonvisual opsin-mediated
phototransduction is the discharge of nematocytes
in Hydra.143 Bright illumination inhibits nematocyte
discharge. Non-nematocyte neurons located in battery
complexes express opsin, cyclic nucleotide gated
ion channels and arrestin which are all known
components of phototransduction cascades. The
origin of theses phototransduction cascades might
even have preceeded the evolution of eyes in cnidaria.
Color Vision
Vertebrates have basically two types of photoreceptors, rods and cones. The rods serve for highly sensitive
dim-light vision, whereas the cones mediate color discrimination and high visual acuity at higher light intensities (see Yokoyama and Pichaud for review).144,145
The nocturnal Tokay Gecko (Gecko gecko) has a pure
rod retina, whereas the diurnal American chameleon
(Anolis carolinensis) has only cones in its retina. The
respective opsins have a wide range of light absorption (λmax ) between 360 and 560 nm corresponding
to the wide range of light wavelength available in the
environment.
The rods that contain a single rhodopsin receptor
with a λmax of 500 nm are color blind, because they
cannot discriminate between wavelength and light
intensity. It makes no difference whether they have
been hit by 10 light quanta of 500 nm wavelength or
20 quanta of a wavelength for which rhodopsin has
half the sensitivity. A system for color detection must
consist of at least two types of receptors with different
absorption maxima (dichromatic color vision).
Primates including humans have a trichromatic color
detection system with three kinds of cone receptors
having absorption maxima in the blue (420 nm), green
(535 nm), and red (565 nm) range of the spectrum.146
All these light-sensitive receptor molecules are opsins,
i.e. membrane proteins with seven transmembrane
domains and 11-cis retinal attached to a specific lysine
residue in transmembrane helix 7 via a Schiff base. The
three types of color receptors differ only in a relatively
small number of amino acid residues. The 11-cis
retinal has a λmax of 440 nm, but when it is bound to
various opsins it detects a wide range of absorption
maxima from 360 to 560 nm. The absorption maxima
closely correspond to the wavelength to which
the animal is naturally exposed. For example, the
coelacanth fish which lives at a depth of 200 m in the
ocean has only two visual pigments with a λmax around
480 nm, since only blue light can penetrate into deep
Evolution of vision
water.147 In many animals including birds, fish, and
insects, the spectrum extends into the ultraviolet
range from 400 to about 320 nm. Many flowers
have ultraviolet markings which are recognized by
pollinating insects. Some fish and butterflies have
visual pigments extending into the near infrared.
However, wavelength and color are not the
same. What we perceive subjectively is the result of our
wavelength analysis.130 For example, a wavelength of
580 nm is yellow. However, an identical yellow is
produced by an appropriate mixture of green light
(540 nm) and red light (620 nm). The color we see
depends on the relative stimulation of our three cone
types. Also, our red cone receptor has a λmax at 565 nm
which corresponds to a spectral color of yellowishgreen. Nevertheless, we see a red color, because the
color which we see depends on the relative stimulation
of the three types of cones.
Polarization Vision
The study of polarization vision began with Santschi’s
(1923)148 and Frisch’s (1949)149 observations that
insects could use scattered skylight for navigation,
and that it was the polarization of scattered skylight
which served as the most important compass cue.
Mainly through the pioneering work of Wehner150,151
which combines behavioral studies in the field and
sensory neurobiological approaches in the laboratory,
robotics, and computer simulations, we gained deep
insights into the problem of how insects, in this case
the desert ant Cataglyphis, detect the pattern of polarized light in the sky, process it neurally and use it for
navigation. Cataglyphis lives in the Sahara desert and
stays inside its cooler nest until the outside surface
temperature reaches values up to 50◦ C and its predators (lizards) are hiding underground. It leaves the nest
and forages for its prey which are mainly small animals killed by the blazing sun. However, after 20 min
of random foraging it has to return to its nest or it
will be killed by the heat. It does so by homing back
to its nest in a straight line, using optical landmarks
only in the immediate vicinity of its nest. This puts
an extremely strong selective pressure on their navigational skills, because they die when they do not
find their nest in time. Cataglyphis uses a compass for
navigation based on the pattern of polarized light. The
question of how a tiny little brain, as that of the ants,
can solve such a complex navigational problem, has
been solved by a multidisciplinary approach.150,151
The ‘hardware’ for a somewhat simplified polarization map is built into the large compound eyes on the
head of the ant, into the geometrical order in which the
UV-sensitive ommatidia are arranged. The specialized
rhabdomeres of these ommatidia serve as analyzers for
polarized light. The ant does not have to perform any
calculations, it simply has to turn its head in the direction in which the sum of the signals from these special
UV-receptors is maximal, and the animal is orientated
along the sun-meridian. This navigational system can
be considered as a matched-filter system. It deviates
systematically from the real direction, but it is precise
enough for the ant to find its nest. From the directional
deviations the matched-filter system can be deduced.
This is a paradigm for the evolution of behavioral
traits under conditions of extreme selective pressure.
EYE PROTOTYPES
Unicellular Photoreceptor Organelles: The
Russian Doll Hypothesis
Photoreceptor organelles range from a relatively
simple eyespot as in cyanobacteria and in Chlamydomonas (Figure 2) to the most elaborate structures
resembling a human eye in some dinoflagellates like
Erythropsis and Warnovia, but assembled in a single cell (Figure 9). They have been described by
Greuet63,64 and consist of a cornea-like layer, a lenslike structure, a retina-like structure with stacked
membranes (or microvilli) and a pigment cup, all
in one cell (Figure 9).
Greuet proposed that these photoreceptor
organelles are modified chloroplasts which have lost
their capacity for photosynthesis. Therefore, Erythropsis and Warnovia are heterotrophic and use their
photoreceptor organelles (ocelloids) for catching their
prey. Recent work of Hayakawa et al. (Hayakawa
S, Takaku Y, Hwang J S, Horiguchi T, Gehring
W, Ikeo K, Gojobori T, unpubl. data). strongly
supports Greuet’s hypothesis because it shows that
these ocelloids contain DNA presumably a remnant
of chloroplast DNA, and that they function as photoreceptor organelles. Erythropsis and Warnovia are
difficult to culture and therefore, individual cells
have to be collected in the ocean. My collaborator Hiroshi Suga succeeded, however, to establish a
c-DNA library from a single cell, from which a fragment of a rhodopsin gene was sequenced. As shown in
Figure 4 the Erythropsis gene and two other dinoflagellate genes map close to the one of Gloeobacter, a
cyanobacterium indicating that they are of chloroplast
origin. The retina-like structure (Figure 9(h)) consists
of closely stacked membranes or microvilli which are
strongly birefringent in polarized light (Figure 9(f))
indicating that the molecules are highly ordered in
these membranes. In situ hybridization of rhodopsin
c-DNA to Erythropsis (= Erythropsidinium) shows
21
Advanced Review
(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
FIGURE 9 | Eye organelles of the unicellular dinoflagellates
Erythropsis and Warnovia. (a) Erythropsis, (b) eye organelle of
Erythropsis, (c) Warnovia, (d) eye organelle of Warnovia, (e) nucleus
and eye organelle of Warnovia, and (f) birefringence in the retina-like
structure detected by polarized light in Warnovia. (a–f) Source: Makiko
Seimiya and Jean and Colette Febvre. (g) Ultrastructure of the eye
organelle of Warnovia and (h) ultrastructure of the retina-like structure
with regularly stacked membranes and large pigment granules. (g and
h: Reprinted with permission from Ref 64. Copyright 1969)
that rhodopsin is present in the retina-like structure (Hayakawa S, Takaku Y, Hwang J, Horguchi T,
Gehring W, Ikeo K, Gojobori T. unpubl.).151 Therefore, we can conclude that the ocelloid is virtually
a camera-type eye-organelle of endosymbiontic origin. These findings led to the proposal of a Russian
Doll Hypothesis5 which assumes that light sensitivity
first arose in cyanobacteria (first Russian doll), the
earliest known fossils on earth. These cyanobacteria
were subsequently taken up by eukaryotic red algae
(second doll) as primary chloroplasts152 , surrounded
by an outer and an inner bacterial membrane, separated by a proteoglycan layer. Subsequently, the red
algae were taken up by dinoflagellates (third doll) as
secondary chloroplasts surrounded by an additional
third membrane coming from the primary red algal
host. In some species of dinoflagellates like Erythropsis
22
and Warnovia, the secondary chloroplasts were transformed into elaborate photoreceptor organelles as
explained above. Cryptoperidinium which has less
elaborate eyespots, has taken up a third generation chloroplast from a diatome, and has become
autotrophic again. Nevertheless, its eyespots contain
lamellar bodies with stacked membranes which are
highly reminiscent of human rod outer segments with
stacked discs containing rhodopsin.
Although these first three Russian dolls are
strongly supported by phylogenetic data, the fourth
endosymbiontic transfer is highly speculative: Because
dinoflagellates are commonly found as symbionts in
cnidarians, they may have transferred some genes
including some photoreceptor genes to cnidarians representing a fourth Russian doll. This is of course the
most speculative step in the Russian doll model. However, there are some indications that some Endosymbiontic Gene Transfer (EGT) may have occurred
between dinoflagellates and cnidarians. For example,
dinoflagellates and cnidarians exclusively share nematocytes one of the most specialized cell-types which
are not found in any other metazoan phylum. Furthermore, the genes encoding the lamellar body organelles,
which closely resemble the outer segments of human
rod cells, might also have been acquired by metazoans
through EGT. The transfer of genes from mitochondria and chloroplasts to the host nucleus is well
documented. In dinoflagellates, this transfer might
have been facilitated by the fragmentation of the
chloroplast DNA into gene-sized DNA-circles. This
last step of the hypothesis is being tested by comparative sequencing the genomes of dinoflagellates
and cnidarians with eyes and looking for genes that
were possibly transferred. However, the sequencing
of dinoflagellate genomes is hampered by the fact that
they generally have very large genomes, some of which
are considerably larger than the human genome.
A Unicellular Eye Prototype in Cnidarians
A priori the prototypic eye must have evolved from a
single cell. However, most eyes contain at least two
cell-types, sensory photoreceptor cells and pigment
cells. This is also true for cnidaria the most ancestral
animal phylum in which eyes are found. Some
hydrozoan jellyfish like Cladonema have a lens eye
at the base of each tentacle and some box jellyfish like
Tripedalia have a battery of sensory organs on each
of the four sides of the animal. These batteries called
rhopalia contain lens eyes, slit eyes, and a statocyst.
The lens eyes contain both pigment and photoreceptor
cells. However, the eyes of their planula larvae81
are unicellular eye prototypes with pigment granules,
Evolution of vision
(a)
(b)
FIGURE 10 | Unicellular photoreceptors in the planula larva of the
box jellyfish Tripedalia.81 (a) Planula larva and (b) unicellular
photoreceptor with pigment granules, microvilli, and a flagellum.
(Reprinted with permission from Ref 81. Copyright 2003 Royal Society
Publishing)
microvilli which presumably contain rhodopsin or
some other visual pigment, and a steering cilium
for phototaxis (see above; Figure 10). It remains to
be shown whether these unicellular eyes represent
rhabdomeric photoreceptors as is strongly suggested
by their morphology. By contrast, the adult eyes
of Tripedalia and Cladonema contain ciliary-type
photoreceptors and c- or s-rhodopsins.
The Darwinian Eye Prototype
In ‘The Origin of Species’ Charles Darwin had
great difficulties to explain the evolution of organs
of extreme perfection such as the eyes. A priori
it seems absurd to try to explain an eagle’s eye
simply by variation and selection. However, in the
chapter on ‘Difficulties of the theory’ he found a
way out of this dilemma and proposed that all the
highly perfect eyes originated from a single prototypic
photosensory organ consisting of two components
only, a photoreceptor cell (which he called a nerve)
and a pigment cell shielding the light from one side.
Such an eye prototype would allow its carrier to
determine the direction of the incoming light and
confer a selective advantage over organisms which
can only discriminate between light and dark. In fact,
this prototype postulated by Darwin on theoretical
grounds can be found in nature. The planarian
Polycelis auricularia has multiple eyes in the head
region, which consist of one photoreceptor and one
pigment cell only (Figure 11). The two cells arise
from a common precursor by cell differentiation153
(Watanabe et al., unpublished data).
The Darwinian prototype can also be found
in larvae which often reflect a more ancestral
stage of evolution. In the trochophora larva of
Platynereis,84,155 a pair of larval eye spots are found,
which are composed of one pigment cell and one
photoreceptor cell only. As in Drosophila Pax6,
six1/2, and atonal (ath) are expressed in the larval
Platynereis eye anlage. The single photoreceptor cell is
of the rabdomeric type with microvilli and expresses
r-opsin which presumably interacts with the Gq-α
subunit canonical for r-opsins.
In contrast, the prototypic eye of the brachiopod
Terebratalia transversa156 consists of two cells, both
of which are putative photoreceptor cells deploying a
modified enlarged cilium for light perception and have
axonal connections to the larval brain. These cells
express besides Pax6 and otx, c-opsin confirming that
these larval eyes deploy ciliary-type photoreceptors
for directional light detection.
A most interesting kind of larval prototypic eye
has been described in certain marine flatworms. The
larvae of Pseudoceros canadensis have two different
eye-types: The right eye is composed of one cupshaped pigment cell and three sensory cells.157 Each
sensory cell extends an array of straight, cylindrical,
tightly packed microvilli (forming a rhabdomere)
into the pigment cell cup. This clearly represents a
rhabdomeric type of photoreceptor cell. However, the
left eye is composed of one pigment cell and four
sensory cells. Three of these photoreceptor cells are of
the rhabdomeric type as for the right eye; however,
the fourth central sensory cell sends unusually large
arching cilia into the pigment cup between the
three rhabdomeres. This indicates that ciliary and
rhabdomeric (microvillar) characteristics can coexist
in the same eye. This strongly supports the notion
(see animal opsins) that both phototransduction
mechanisms, ciliary and rhabdomeric, were present
from the beginning of metazoan evolution.
The mechanisms underlying this first step
of cellular differentiation leading from a common
precursor cell to a photoreceptor and a pigment
cell can be deduced from evolutionary developmental
studies. The gene locus which serves as a pigment
23
Advanced Review
(a)
(b)
(c)
(d)
FIGURE 11 | Protypic eyes (E) in (a) the planarian Polycelis auricularia, (b and c) higher magnification, (d) histological section across the eye of
Planaria torva. Pc, pigment cell; PcN, pigment cell nucleus; Mv, microvilli; Ph, photoreceptor cell; PhN, photoreceptor cell nucleus. (Reprinted with
permission from Ref 154. Copyright 1897 Royal Society of London)
cell determinant was first identified more than
60 years ago as a mutation in mice by Hertwig158
and later cloned and designated as Microphthalmia
transcription factor (Mitf ). Mitf mutations affect
several different cell types, but mainly pigment cells.
Vertebrates have two kinds of pigment cells, the
melanocytes of the skin, which are derived from the
neural crest, and the retinal pigment epithelial (RPE)
cells of the eye, which are derived from the optic cup,
which represents an evagination of the brain. The Mitf
gene has been cloned159–161 and shown to encode
a basic helix-loop-helix zipper transcription factor.
Arnheiter162 has independently proposed a common
evolutionary origin of pigment cells and photoreceptor
cells.
Several lines of evidence indicate that Mitf is a
pigment cell determinant. Mitf is strongly expressed
in pigment cells in the RPE160,163 and in loss-offunction mutants the RPE hyperproliferates, and in
the dorsal part the RPE is transformed into a correctly layered, inverted retina. Tachibana et al.164
were able to transform fibroblasts into pigment cells
(melanocytes) by stably transfecting NIH 3T3 cells
with human Mitf c-DNA. In zebrafish, homozygous
nacre mutants carrying a mutation in Mitf, can be
rescued by injecting the wild-type Mitf gene into
24
early embryos and inducing gene expression with a
heat shock promoter. Mitf is positively regulated by
Pax6 and Pax2. In mice, loss-of-function of either
Pax6 or Pax2 alone does not reduce Mitf expression,
but if both Pax6 and Pax2 are nonfunctional Mitf
is no longer expressed.165 As a negative regulator
CHX10, a paired-like homeodomain (HD) transcription factor, has been identified. These observations
have led to the following concept of differentiation
of the retina166,167 : Pax6 first induces the optic vesicle (see below). Subsequently, the differentiation of
these neural cells into neuroretina and RPE is controlled by an interaction between Pax6 and Mitf.
The neuroretina is specified by Pax6 and antagonized
by Mitf. Mitf is a repressor which in turn is negatively regulated by CHX10. CHX10 is induced by
fibroblast growth factor (FGF) in the neuroretina and
prevents Mitf from transforming the neuroretina into
a pigment epithelium. Interestingly, Mitf is highly conserved in evolution and has been found in ascidians,168
Caenorhabditis,169 Drosophila170 and even in Tripedalia jellyfish.171 Therefore, we propose that the
first step of cell differentiation in eye evolution leading
to the formation of photoreceptor versus pigment cells
is based on an interaction between Pax6 and Mitf (see
Figure 12).
Evolution of vision
Light
receptor
Rhodopsin
PAX A,B
Pax6
Mitf
Pax6
Cell differentiation
Photoreceptor cell
Pigment cell
Prototypic eye
Ommatidium
Insect
compound eye
Arca
Compound eye
Lens
Vitreous
Body
Retinal pigmentepithelium
Distal
Retina
Proximal
Retina
Neuroretina
Mirror
Pecten
mirror eye
Eyelid
Iris
Lens
Cornea
Vitreous
Body
Neuroretina
Pigmentepithelium
Vertebrate
Lens eye
Sclera
Cephalopod eye
Lens eye
FIGURE 12 | General scheme of eye evolution. The first step in eye evolution is the evolution of a light receptor molecule which in all metazoa is
rhodopsin. In the most ancestral metazoa, the sponges, a single Pax gene, but no opsin gene has been found. In the larva of the box jellyfish
Tripedalia, a unicellular photoreceptor has been described. The adult jellyfish forms complex lens eye with ciliary photoreceptor cells, which form
under control of PaxB, a putative precursor of Pax6. However, the eyes of the hydrozoan jellyfish Cladonema are controlled by PaxA. We propose that
the prototypic eye consisting of just two cells a photoreceptor cell and a pigment cell originated from a unicellular photoreceptor by a first step of cell
differentiation. This cellular differentiation led to formation of a photoreceptor cell and a pigment cell under the genetic control by Pax6 and Mitf,
respectively. As true innovations are rare in evolution all the more complex eye-types arose monophyletically from one of these Darwinian prototypes
leading to a large diversity of eye-types by divergence, parallel evolution, and convergence.
DEVELOPMENT AND EVOLUTION
OF THE DIFFERENT EYE TYPES
Pax6 and the Monophyletic Origin
of the Various Eye Types
The leading neo-Darwinists like Ernst Mayr assumed
that the various eye types found in the different animal
phyla evolved independently 40–60 times in the course
of evolution.3 This concept of a polyphyletic origin
of the eyes was mainly based on morphological
and embryological arguments. The compound eye
of insects has an entirely different structure from
the camera-type eye of vertebrates. The camera-type
eyes of cephalopods and vertebrates were taken as
the perfect example of convergent evolution, because
they are similar in structure, but their embryological
25
Advanced Review
development is different; the vertebrate eye develops
from an evagination of the brain, whereas the
cephalopod eye forms as an invagination of the
skin. However, this notion of a polyphyletic origin
of the various eye types was hardly compatible
with Darwin’s idea of an ancestral prototype. As
he pointed out in ‘The Origin of Species’1 this
prototype cannot be explained by selection as an
accelerating driving force. Selection can only become
effective once a minimal function is established. The
prototypic eye has to function at least to a small
extent before selection can set in. Thus, the Darwinian
prototype must have arisen as a purely stochastic
event. Therefore, the probability of this happening
40–60 times is extremely low.
More recent developmental genetic experiments
open up an entirely different perspective on eye
evolution (Figure 12) and argue strongly for a
monophyletic origin of the various eye types.4,5,172,173
It began with the discovery of the Drosophila homolog
of the mammalian Pax6 gene. Pax6 was first cloned
in the mouse174 and shown to correspond to the
mutation Small eye (Sey)175 in mice and to Aniridia
in humans.176 Surprisingly, the homolog of Pax6 of
Drosophila turned out to encode the eyeless (ey)
gene, which had been discovered as a mutation as
early as 1915 by Hoge.177 The fact that both the
mammalian and the insect Pax6 genes are essential
for eye development, suggested that the generally
accepted hypothesis of a polyphyletic origin of the
various eye types might not be correct. This led to
the idea that Pax6 might be a master control gene for
eye development in both insects and vertebrates. The
term master control gene was proposed by Weintraub
et al.178 for the muscle determining gene MyoD and
extended by Lewis179 to the homeotic genes which in
a combinatorial fashion determine segmental identity
along the anteroposterior axis in Drosophila and mice.
One way of demonstrating the function of a master
control gene is to construct an inducible gain-offunction mutation and express the master control gene
ectopically. For the homeotic Antennapedia (Antp)
gene, the ectopic expression leads to a homeotic
transformation of the antenna into a leg.180 The
decisive experiment for ey is shown in Figure 13.173
Embryonic eyeless c-DNA was ectopically expressed
in the antennal, wing and leg discs by means of
the yeast gal4 system.181 By using different enhancer
lines, ectopic eyes can be induced on the legs, wings,
halteres, and antennae of the transgenic flies. This
clearly demonstrated the role ey as a master control
gene in switching on the entire cascade of target genes
required for eye morphogenesis. These ectopic eyes
are functional in that the photoreceptor cells can
26
generate a normal electroretinogram and establish
functional synapses.182 Of course, eye morphogenesis
cannot be induced in any tissue of the fly at any
stage of development, but eye induction is possible
in all imaginal discs up to a certain stage of
differentiation. The master control gene first has to
interact with subordinate control genes to repress the
resident developmental program and to install the eye
program. If the cells have proceeded too far along
their developmental pathway they become locked in
(fully determined) and ectopic ey expression has no
longer any effect.
While mammals have a single Pax6 gene,
Drosophila and other holometabolous insects have
two Pax6 paralogs, which were designated as eyeless
(ey) and twin of eyeless (toy).183 The Toy protein
shares 91% sequence identity in the paired domain
(PD) and 90% in the HD with human and murine
Pax6 proteins, as compared to 95 and 90% for EY.
Outside of these highly conserved domains Toy is
more similar to the mammalian protein, particularly
in its overall length and at the C-terminus where it
shares a transcriptional activation domain with other
Pax6 proteins that is absent in Ey. In Drosophila,
it is more widely expressed than Ey and the loss-offunction phenotype is headless rather than eyeless. It
is often assumed that a master control gene for eye
development should be expressed specifically in the
eyes, but this notion is incorrect. The master control
gene not only has to induce eye formation, but the
eye has to be properly located in the body plan (see
below) and connected to the brain. Therefore, Pax6
is also expressed in most of the head region including
parts of the brain where the proper neuronal connections have to be made. Despite the fact that toy has a
more mammalian-like expression pattern, it is not the
more ancestral gene, because ey and toy arose by gene
duplication at exactly the same time. As shown in
Figure 12 toy can also induce ectopic eyes, but it does
so by first activating ey which in the course evolution
became intercalated underneath toy into the eye developmental program. It requires a functional ey gene for
inducing eyes.184 Gene duplication, diversification,
and intercalation into a developmental pathway are
an important evolutionary mechanism4,166 for generating biodiversity. Therefore, toy became the master
control gene on the top of the hierarchy. This is
in line with the observation that toy is expressed
before ey, at the blastoderm stage when the body
plan is laid down, whereas ey is expressed only later
during germband extension. Obviously, the gene on
the top of hierarchy has to be expressed first in
development.
Evolution of vision
GAL 4
(a)
Genomic enhancer
GAL 4
Tissue-specific expression of GAL 4
(b)
eyeless embryonic cDNA
UAS
Transcription of eyeless in antennal,
leg and wing imaginal discs
(c)
FIGURE 13 | Induction of ectopic eyes in Drosophila by targeted expression of the Pax6 homologs eyeless (ey ) and twin of eyeless (toy ).173
(a) Experimental design: targeted expression of ey c-DNA using a genomic enhancer to induce the yeast Gal4 transcription factor in various imaginal
discs. Gal4 binds to the upstream activating sequences (UAS) and drives the expression of ey into the respective areas of the eye-antennal, wing, and
leg discs, respectively. (b) Ectopic eyes induced by ey. (c) Ectopic eyes induced by toy. Source: Urs Kloter and Georg Halder
The mammalian Pax6 gene can functionally
substitute for the Drosophila eyeless gene: The
ectopic expression of mouse Pax6 in Drosophila
induces ectopic compound eyes173 and the reciprocal
experiment of injecting ey and toy mRNA from
Drosophila into Xenopus embryos at the two cell
stage also leads to the induction of ectopic eye
structures.185 Of course, these are frog (vertebrate)
eyes since we have only injected the master control
gene of Drosophila into frog embryos and not the
entire gene cascade necessary to form a Drosophila
compound eye.
So far, we have been considering only the
compound eyes of Drosophila for which a single
master control gene is sufficient for ectopic induction.
However, for the induction of the ocelli on the top of
the fly head the combinatorial interaction between at
least two master control genes, toy and orthodenticle
(otd)166,186 is required as found for homeotic genes.
These gene regulatory pathways for compound eyes
and ocelli differ significantly (Figure 14).
The genetic cascade specifying the eye developmental pathway has been elucidated in some detail.
The master control gene toy first activates ey and
together they activate sine oculis (so) which encodes
a homeobox protein,187,188 which forms a dimer with
eyes absent (eya).189,190 Eya has dual function and acts
as a transcription factor in the nucleus and as a protein
phosphatase in the cytoplasm.191 The gene dachshund
(dac) encodes a nuclear protein which is required for
differentiation of ommatidia, but it is also required
for leg development.192,193 Toy also activates optix
(opt) another member of the sine oculis (six) gene
family and the Pax gene eyegone (eyg) which is also
required for eye development and functions neither
upstream nor downstream of toy and ey, but it cooperates with these two genes in eye morphogenesis.194
A regulatory scheme for the top of eye developmental pathway is presented in Figure 14(a). It involves a
large number of positive and autoregulatory feed back
loops.
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Advanced Review
(a)
(b)
Pax6
toy
otd
Pax6
toy
Initiation
ey
eyg
hh
eya
Auto
regulation
so
eya
Feedback
loops
dac
so
optix
?
Compound eye development
Development of ocelli
FIGURE 14 | Initiation of the developmental pathway for
compound eye and ocelli development. Pax6 homologs toy and ey are
on top of the hierarchy, so and eya as well as dac are second-order
transcription factors connected by various feedback loops to the master
control genes ey and toy. In ocellar development, an additional master
control gene otd is required. toy, twin of eyeless ; ey, eyeless ; eyg,
eyegone ; optix ; so, sine oculis ; eya, eyes absent ; dac, dachshund ; otd,
othodenticle ; hh, hedgehog.
In order to verify the results obtained by gain-offunction mutants, the corresponding loss-of-function
mutants have to be examined. Knocking out ey or toy
does not block the development of compound eyes
completely, and even the ey− , toy− double mutant
which is essentially headless still forms a couple of
red ‘balls’ of ommatidia in the thorax. Only in triple
mutants lacking all three Pax genes ey, toy, and eyg
eye structures are completely lacking.166
Pax6 has been shown to be involved in eye
development in all bilaterian phyla that were analyzed
so far and in all the major eye-types including
the camera-type eye of vertebrates, the compound
eye of insects, the camera-type eye of cephalopods,
the mirror-eye of Pecten, and most importantly the
primitive larval eyes of annelids, the trochophora larva
of Platynereis,84 the brachiopod larva,156 and the
ascidian (urochordate) larva.195 Originating from a
common prototype under the control of Pax6 and a set
of second-order transcription factors like sine oculis
(so/six), eyes absent (eya), and dachshund (dac) which
are also highly conserved in evolution, the various
eye-types have evolved. There are no functional
constraints associated with a given transcription factor
to control eye development. A transcription factor
can control any gene which has the appropriate cisregulatory sequences. If a gene like Pax6 is always
involved in eye development, this is for evolutionary
historical reasons, because it was previously involved
in the genetic control of eye development in the
last common ancestor and continues to be involved
during further evolution. The fact that transcription
factors can control any set of target genes or any
28
FIGURE 15 | Phylogenetic tree of the six /so gene family. The
phylogenetic analysis shows that a given transcription factor can
regulate any developmental program as long as the target genes
possess the appropriate cis -regulatory elements. The six1 and six3
subclasses (blue) both regulate eye development in Drosophila as well
as in vertebrates, whereas the six4 subclass (red) controls muscle
development. h, human; m, mouse; D, Drosophila, zf, zebrafish, c,
ciona; mf, medaka; e, C. elegans.
developmental programme, is clearly shown by the
three subclasses of six genes all of which contain a
homeobox: The six1 and six3 subclasses are both
involved in eye development, both in vertebrates and
insects, whereas the six4 subclass controls muscle
development (Figure 15). This indicates that with
the same tool box of transcription factors, a large
variety of eye-types can be made depending on the
downstream target genes which are selected.
There appeared to be some exceptions to the
rule that Pax6 is consistently expressed in the eye primordia, in particular, in C. elegans and in sea urchins
which do not have image forming eyes. C. elegans has
retained the Pax6 gene even though it does not have
any eyes, but eyes are found in some ancestral marine
nematodes. However, C. elegans has a nose and a
small brain which are under Pax6 control, so that the
selective pressure to retain the Pax6 gene is still maintained. By contrast, the rhodopsin genes have been
lost, because their main function is photoreception,
and as C. elegans lives underground or inside rotting
fruits there is no selective pressure to maintain vision,
which is generally true for animals living in the dark,
e.g. in caves.
Evolution of vision
Gene categories specifically enriched at the larval stage
Transcription factors
Cell proliferation
Cell differentiation
Cell fate commitment
Central nervous system development
Peripheral nervous system development
Axonogenesis
Photoreceptor cell fate commitment
Bolwigorgan development
Optic placode formation
Crain development
Cell-cell adhesion
Proteinkinase cascade
Receptor activity
Rtk signaling pathway
Jnk cascade
Notch signaling pathway
Male courtship behavior
Larval stage (L3)
109 PS (98 genes)
expressed 1.5 more in leg discs where eyeless is
ectopically expressed as compared to wild type
leg discs.
Pupal stage (P9)
Gene categories specifically enriched at the pupal stage
Structural molecules
Structural constituent of cuticle
460 PS (409 genes)
expressed 1.5 more in ectopic
eyed legs than in
wild type legs.
Muscle contraction
Pigmentation
Eye pigment synthesis
Ommochrome biosynthesis
Neurotransmission
Gene categories specifically enriched at the adult stage
Intracellular signaling cascade
Adult stage
Organelle organization and biogenesis
Transmission of nerve impulse
Synaptic transmission
428 PS (474 genes)
expressed 1.5 more in ectopic
eyed legs than in
wild type legs.
Cell–cell signaling
Cell adhesion
Programmed cell death
Histolysis
Bristle morphogenesis
Photoreceptor maintenance
DrosGenome1 Chips
RMA normalization; P<0.05
Rhabdomere development
FIGURE 16 | Deciphering the eye developmental program by analyzing the transcriptome. Genes that are expressed specifically in eye discs as
compared to leg disc are listed for the larval, pupal, and adult stages. At the larval stage particularly transcription factor genes are expressed which
initiate the eye developmental pathway. At the pupal stage mostly structural genes are activated as well as genes involved in the synthesis of eye
pigments. In the adult animal mainly genes involved in intracellular signaling, neurotransmission, and the visual process are expressed. In total,
approximately 1000 genes are required for eye development and maintenance. Source: Michaut and Gehring, unpublished data.
An interesting case is provided by sea urchins
which do not have image forming eyes. Soon after
the discovery of Pax6 as a master control gene for
eye development, a Pax6 gene was found in Paracentrotus lividus196 which seemed to be an exception
to the rule. The Pax6 gene was also found to be
expressed in the tube feet which are extended between
the spines and serve for locomotion. It has been known
that sea urchins which lack both image-forming eyes
as well as eyespots are negatively phototactic and
respond to both visible (λmax at 450 nm) and ultraviolet components of sunlight. Two recent studies197,198
on Strongylocentrotus purpuratus and Strongylocentrotus droebachiensis clearly show that sea urchins
have their photosensory organs in their tube feet and
their photoreceptor cells express a rhabdomeric-type
opsin (r-opsin) indicating that the two major types
of opsins, r- and c-opsins, were present prior to the
split of proto- and deuterostomes (see above). These
findings also show that you can use Pax6 expression
to detect previously unknown photosensory organs.
For Drosophila, the transcriptome of eye development has been deciphered.199 In the larval eye
imaginal disc mostly a set of transcription factors
is activated which are high up in the gene cascade specifying eye development and include about
100 genes that are expressed specifically in the eye
imaginal disc as compared to a leg imaginal disc
which serves as a reference (Figure 16). During pupal
development approximately 400 genes become active
mostly endoding structural molecules, muscle proteins, proteins involved in neurotransmission and
pigmentation. In the adult fly approximately 500 genes
involved in signaling pathways, organelle biogenesis,
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Advanced Review
neurotransmission, and vision are transcribed. In
total, we estimate about 1000 genes to be involved in
the eye developmental pathway after subtraction of all
the genes which are also expressed in other imaginal
discs, in this case the leg disc.
Similar studies are being carried out in the mouse
and it is becoming increasingly clear that starting
from a highly conserved set of transcription factors
including Pax6, six, eya, dac etc. the regulatory
cascade diverges rapidly indicating that the various
eye-types are made with the same tool box, but
different overlapping downstream target genes.199
This is compatible with the hypothesis of intercalary
evolution presented below.
The evolutionary origin of Pax6 is still largely
unknown. So far Pax genes characterized by a paired
box which encodes a bipartite DNA binding domain
consisting of a PAI and a RED subdomain200–202
have only been found in multicellular metazoans and
exclusively in bilateria. Both in mammals and insects
there are approximately 9 or 10 Pax genes. They nicely
illustrate an evolutionary principle which Jacob has
called ‘tinkering’.203 Pax genes may contain two DNA
binding domains a PD and HD and an octapeptide
sequence to which an auxiliary factor may bind. These
three domains occur in all possible combinations in
the various Pax genes: Only a PD, a PD plus a HD as
in Pax6, a PD plus a HD plus an octapeptide, a PD
and a partial HD, a partial PD and a complete HD.
Obviously, some recombination and deletion events
have generated this diversity. With respect to Pax6 in
Drosophila, it has been shown experimentally that the
HD is dispensable for eye induction.204 As the HD is
also highly conserved in evolution it must be under
selective pressure and fulfil a function other than eye
induction. Since ey and toy are highly pleiotropic, the
HD is likely to be required for some other function.
In sponges no opsin genes have been found, but
a single Pax gene is present. In cnidarians only five Pax
genes Pax A, B, C, D, and E have been found and no
true Pax6 homolog, even though some hydrozoan and
box jellyfish evolved advanced eye-types.205 PaxA and
PaxC correspond to pox neuro in Drosophila, PaxB
to Pax2/5/8, PaxD to Pax3/7, and PaxE to Drosophila
eyegone (eyg). In the box jellyfish Tripedalia, Kozmic
et al.206 have provided evidence that PaxB an ortholog
of vertebrate Pax2/5/8 is a regulator of eye development capable of inducing ectopic eyes in transgenic
flies. PaxB may be interpreted as a precursor of
Pax6. However, in the hydrozoan Cladonema PaxA is
strongly expressed in the photoreceptor cells, whereas
PaxB and PaxE are transcribed in the manubrium,
the feeding organ in which also the gonads develop.
Cladonema PaxA induces ectopic eyes in Drosophila
30
imaginal discs, whereas PaxB and PaxE do not.
Furthermore, the PD of PaxA protein binds directly
to the 5 upstream region of eye-specific opsin genes,
whereas PaxB does not. These data show an involvement of Pax genes in hydrozoan eye development as
in bilaterians, support the monophyletic evolutionary origin of all animal eyes. The distinct classes of
Pax genes, which may have played redundant roles
in early metazoan evolution, were flexibly deployed
for eye development in different animal lineages. In
bilateria this role was completely taken over by Pax6.
The camera-type eye of cephalopods (octopus,
squid, cuttle-fish etc.) has previously been considered
to represent a paradigm for convergent evolution.
However, this notion has to be modified in the light
of the discovery of a highly conserved Pax6 gene207
in squids which shares 67% amino acid sequence
identity with vertebrates. In the PD, squid Pax6
shows 91–95% sequence identity with its homologs in
vertebrates, Drosophila, Nemertine, and sea urchins,
whereas the HD shows 90–98% identity with Pax6
from these species. In situ hybrization to squid
embryos reveal Pax6 in the developing eyes, the optic
lobes of the brain, the arms, and the mantle. The
hybrization to the eyes and optic lobes is expected,
expression in the arm primordia may reflect the
presence of other sensory organs, and the expression in
the mantle is of particular interest, because this is the
site of eye formation in other molluscs like scallops and
ark clams. Ectopic expression of squid Pax6 cDNA in
Drosophila induces the formation of ectopic eyes.
These findings indicate that the cephalopod eye
like all other bilaterian eyes is specified by the Pax6
gene and has originated monophyletically and subsequently evolved by divergent, parallel, and convergent
evolution. It should be emphasized that bivalves and
snails have lense, mirror, and compound eyes, and
more primitive cephalopods like Nautilus have pinhole eyes, whereas only the squids and octopuses have
the most complex camera-type eyes. Therefore, the
molluscs cover the entire spectrum of eye-types, all
going back to a successful Darwinian prototype.
Comparative genomics between the camera-type
eyes of octopuses and humans207–209 show that of
1052 nonredundant genes which are expressed in the
octopus eye 729 (=69%) are in common with the
human eye, which is a surprisingly large fraction.
Of these 1052 genes, 1019 are already found in the
last common ancestor of bilateria, and 875 out of
1052 are conserved between humans and octopus. By
comparing three species of cephalopods with cameratype eyes with molluscs with pin-hole eyes (Nautilus)
and mirror-eyes (scallops); 5707 cephalopod camera
eye-specific candidate genes (5707) were selected
Evolution of vision
for further analysis. From the 1571 genes found in
common between cephalopod and vertebrate cameratype eyes, 156 genes were identified which were
under positive selection. These studies suggest that
both changes in gene expression and in the primary
structure of the respective proteins (through positive
selection) have contributed to the evolution of the
cephalopod camera-tye eye.
Localization of the Eyes in the Body Plan
The criterion that homologous organs have to occupy
the same position in the body plan was disproven
by our experiments on the ectopic induction of compound eyes all over the body plan. Furthermore, the
notion that the master control gene for eye development has to be eye-specific proved to be wrong.
Pax6 has a highly pleiotropic expression pattern; it is
expressed in the entire head region, in the compound
eyes, the ocelli, in the nervous system, a large area of
the brain and in the spinal cord or the ventral nerve
cord, and also in the nose and antenna, respectively.
This is similar to Hox genes which in a combinatorial
fashion specify entire body segments. Therefore, the
eye not only has to be built, but it has to be positioned in the body plan and properly connected to the
nervous system.
The positioning of the eye primordia in the
Drosophila embryo was analyzed by Blanco and
Gehring210 and is summarized in Figure 17. The
toy gene is activated by the maternal-effect genes
bicoid (bcd) and torso (tor) in the anterior portion
of the embryo and its expression domain is defined
by repression from all sides by hunchback (hb),
knirps (kni), and decapentaplegic (dpp). The repressor
defining the posterior border of repression remains to
dpp
hb
tor
bcd
kni
dl
Bcd gradient
FIGURE 17 | Positioning of the eye in head region of the Drosophila
embryo. Proposed model for the onset of toy expression. toy is
activated (arrows) by the maternal effect genes bicoid (bcd ) which
forms a protein gradient and torso (tor ). It is repressed from all sides by
hunchback (hb ), knirps (kni ), and decapentaplegic (dpp ). The posterior
repressor remains to be identified.210
be identified. It is important to note that the same
genes which specify the body plan are also involved in
positioning the eye.
As mentioned above, the ocelli are positioned
on the head of the fly by orthodenticle (otd). Otd/otx
genes are also involved in positioning of the eyes in
many different phyla like vertebrates, annelids and
planarians.
In recently discovered Precambrian Lobopodia
fossils which are considered to be among the ancestors
of the Arthropods, a species called Microdictyon
sinicum has been described which seems to have a
pair of compound eyes in every segment, whereas
the eyes are confined to the head region in Cardiodictyon catenulum.211 Therefore, the eyes may have
been present in the prototypic body segment and
in the course of cephalization confined to the head
region where they are most useful for controlling
locomotion.212 The original formation of a pair of eyes
in every body segment may still be reflected by Pax6
expression in the ventral nerve cord in every thoracic
and abdominal segment in the Drosophila embryo.
Development and Evolution of the Lens
The eye lens of vertebrates is a classical model of
embryonic induction (see Gunhaga for review).213 A
comparison of the genetic control of lens development
between Drosophila und vertebrates reveals a number
of molecular commonalities which were unsuspected
by classical studies (see review by Charlton-Perkins
et al.).214
Conservation of Pax6 Target Sequences
If Pax6 is highly conserved in evolution, the cisregulatory elements to which it binds should also
be conserved. This prediction was tested analyzing the
cis-regulatory element of the chicken δ crystalline gene
encoding a lens-specific protein that is found in birds
and reptiles only, due to their evolutionary relatedness. Previous studies215 have shown the lens-specific
expression of the δ1-crystallin gene is under the control of the D5 enhancer which is only 30 bp long and
contains both a Pax6 and a Sox2 binding site. To test
the idea that Pax6 and Sox2 together with the DC5
enhancer could form a regulatory circuit in a very
distantly related animal, the DC5 enhancer was fused
to GFP and introduced into Drosophila, to study its
pattern of expression.216 The results show that the
DC5 enhancer is not only active in the compound
eye but, remarkably, specifically active in those cells
which are responsible for secreting the liquid lens, i.e.
the cone cells. However, the regulation of the DC5
enhancer is carried out by Pax2 rather than Pax6 in
31
Advanced Review
combination with SoxN the homolog of Sox2. Both
proteins (D-Pax2 and SoxN) bind cooperatively to the
DC5 enhancer and activate the target gene synergistically. As Pax6 and Pax2 proteins derive from the same
ancestor, we can assume that during evolution Pax6
function in vertebrate lens development was retained
by Pax2 in Drosophila. These findings demonstrate
a remarkable evolutionary conservation of regulatory
circuits between chicken and fly.
Intercalary Evolution of Morphogenetic
Pathways
For the evolution of more complex eyes originating
from the Darwinian prototype, we have proposed
a mechanism of intercalation of new genes into the
eye developmental pathway.4,166 The prototypic eye
already possesses the master control Pax6 on the top
of the hierarchy and the genes required for vision
like opsins, G-protein signaling, arrestins etc., but
it lacks, for example, lens protein genes and other
genes required for morphogenesis of more complex
eyes. At least two genetic mechanisms for the recruitment of new genes into a developmental pathway
are known: Gene duplication and subsequent divergence and enhancer fusion. Both have been found
in eye development in Drosophila. Gene duplication,
for example, has occurred in Drosophila Pax6 giving
rise to ey and toy followed by functional divergence.
The toy gene retains most of the orginal master control function in the head including the compound
eyes, the ocelli, the antennae etc. and in the ventral
nerve chord, whereas ey has undergone subfunctionalization and is mostly required for the formation of
compound eyes. The original positive control over the
second-order target gene sine oculis (so) is retained by
both genes and the so enhancer contains binding sites
that are either recognized by toy alone or by both ey
and toy.217 The ey gene has become inserted downstream of toy into the eye morphogenetic pathway.
In vertebrates, another case of gene duplication and
divergence has been described for Mitf in zebrafish.
As mentioned above, vertebrate pigment cells arise
from two embryonic sources; the neural crest provides the pigment cells of the skin, whereas the retinal
pigment cells originate from the optic cup, i.e. an
evagination of the brain. The nacre mutant is a lossof-function of Mitf (or Mitfa) and lacks the neural
crest derived melanophores, although it is expressed
in both the neural crest precursors and in the eye.218
Subsequently, Lister et al.219 have identified a second
Mitf gene, Mitfb which is coexpressed with Mitf a in
the RPE, where it complements the nacre mutation,
but it is not expressed in neural crest melanoblasts
leading to a loss of neural-crest-derived pigment cells.
This loss of neural-crest-derived pigment cells can be
rescued by expressing mitf b the neural-crest-derived
melanoblasts under the mitfa promoter. Therefore,
Mitfa and Mitf b together recapitulate the function of
a single ancestral gene.
The second known genetic mechanism is
enhancer fusion and has been demonstrated by Piatigorsky and Wistow220 for various lens proteins. A variety of lens proteins originated from various enzymes
or small heat shock proteins. By transposing them in
the vicinity of an enhancer which leads to the expression in the lens, such genes can be recruited into
the lens morphogenetic pathway. If such proteins are
transparent to visible light and have the right refractive index and stability, they can serve as a constituent
of the lens. Similarly, one of the major constituents of
the cuticular lens of Drosophila, Drosocrystallin, was
cloned221 and the analysis of its amino acid sequence
clearly indicates that it belongs to a large family of
cuticle proteins and was recruited into the lens.
By intercalation of additional genes into the eye
developmental pathway increasingly complex eyes can
evolve from the prototype supporting Darwin’s theory of eye evolution from a simple prototype to highly
complex eyes.
ACKNOWLEDGMENTS
The research contribution of all the members of my team, and the financial support by the University of Basel
and the Swiss National Science Foundation is acknowledged. Special thanks go to Greta Backhaus for processing
the manuscript and Annette Roulier for producing the figures.
REFERENCES
1. Darwin C. The Origin of Species by Means of Natural Selection. 6th ed. London: John Murray; 1882,
143–146.
32
2. Salvini-Plawen L, Mayr E. On the Evolution of Photoreceptors and Eyes. New York: Plenum Press;
1977.
Evolution of vision
3. Mayr E. What Evolution Is. New York: Basic Books;
2001.
4. Gehring WJ, Ikeo K. Pax 6: mastering eye morphogenesis and eye evolution. Trends Genet 1999, 15:
371–377.
5. Gehring WJ. New perspectives on eye development
and the evolution of eyes and photoreceptors. J Hered
2005, 96:171–184.
6. Schopf J. Microfossils of the early Archaean Apex
Chert: new evidence of the antiquity of life. Science
1993, 260:640–646.
7. Brasier MD, Green OR, Jephcoat AP, Kleppe AK, Van
Kranendonk MJ, Lindsay JF, Steele A, Grassineau
NV. Questioning the evidence for Earth’s oldest fossils.
Nature 2002, 416:76–81.
8. Schopf J, Packer B. Early Archaean (3.3 billion to
3.5-billion-year-old) microfossils from Warrawoona
Group, Australia. Science 1987, 237:70–73.
9. Allwood AC, Walter MR, Kamber BS, Burch IW.
Stromatolite reef from the Early Archaean era of Australia. Nature 2006, 414:714–718.
10. Albertano P, Barsanti L, Passarelli V, Gualtieri P. A
complex photoreceptive structure in the cyanobacterium Leptolyngbya sp. Micron 2000, 31:27–34.
11. Konopka RJ, Benzer S. Clock mutants of Drosophila
melanogaster. Proc Natl Acad Sci USA 1971, 68:
2112–2116.
12. Rosbash M. The implications of multiple circadian
clock origins. PLoS Biol 2009, 7:421–425.
13. Williams SB. A circadian timing mechanism in
the cyanobacteria. Adv Microb Physiol 2007, 52:
229–296.
14. Pittendrigh CS. Circadian systems: general perspective and entrainment. In: Aschoff J, ed. Handbook of
Behavioral Neurobiology: Biological Rhythms. New
York: Plenum Press; 1981, 57–80, 95–124.
15. Kondo T, Ishiura M. Circadian rhythms of cyanobacteria: monitoring the biological clocks of individual colonies by bioluminescence. J Bacteriol 1994,
176:1881–1885.
16. Ishiura M, Kutsuna S, Aoki S, Iwasaki H. Expression
of a gene cluster kaiABC as a circadian feedback process in cyanobacteria. Science 1998, 281:1519–1523.
17. Nakajima M, Imai K, Ito H, Nishiwaki T, Murayama
Y, Iwasaki H, Oyama T, Kondo T. Reconstitution of
circadian oscillation of cyanobacterial KaiC phosphorylation in vitro. Science 2005, 308:414–415.
18. Dong G, Golden SS. How a cyanobacterium tells time.
Curr Opin Microbiol 2008, 11:541–546.
19. Taniguchi Y, Katayama M, Ito R, Takai N, Kondo T,
Oyama T. LabA: a novel gene required for negative
feedback regulation of the cyanobacterial circadian
clock protein KaiC. Genes Dev 2007, 21:60–70.
20. Ouyang Y, Andersson CR, Kondo T, Golden SS,
Johnson CH. Resonating circadian clocks enhance fitness in cyanobacteria. Proc Natl Acad Sci USA 1998,
95:8660–8664.
21. Nakamichi N. Molecular mechanisms underlying the
Arabidopsis circadian clock. Plant Cell Physiol 2011,
52:1709–1718.
22. Bunning E. The physiological clock. In: The Heidelberg
Science Library. New York: Springer Verlag; 1967.
23. Ahmad M, Cashmore AR. HY4 gene of A. thaliana
encodes a protein with characteristics of a blue-light
photoreceptor. Nature 1993, 366:162–166.
24. Somers DE, Webb AA, Pearson M, Kay SA. The short
period mutant, toc1-1, alters circadian clock regulation of multiple outputs throughout development in Arabidopsis thaliana. Development 1998,
125:485–494.
25. Brunner M, Schafmeier T. Transcriptional and posttranscriptional regulation of the circadian clock of
cyanobacteria and Neurospora. Genes Dev 2006,
20:1061–1074.
26. Schafmeier T, Káldi K, Diernfellner A, Mohr C,
Brunner M. Phosphorylation-dependent maturation
of Neurospora circadian clock protein from a nuclear
repressor toward a cytoplasmic activator. Genes Dev
2006, 20:297–306.
27. Froehlich AC, Liu Y, Loros JJ, Dunlap JC. White
Collar-1, a circadian blue light photoreceptor, binding to the frequency promoter. Science 2002,
297:815–819.
28. Schafmeier T, Diernfellner AC. Light input and processing in the circadian clock of Neurospora. FEBS
Lett 2011, 585:1467–1473.
29. Glossop NR. Circadian timekeeping in Drosophila
melanogaster and Mus musculus. Essays Biochem
2011, 49:19–35.
30. Yu Q, Jacquier AC, Citri Y, Hamblen M, Hall JC,
Rosbash M. Molecular mapping of point mutations
in the period gene that stop or speed up biological
clocks in Drosophila melanogaster. Proc Natl Acad
Sci USA 1987, 84:784–788.
31. Hardin PE, Hall JC, Rosbash M. Feedback of the
Drosophila period gene product on circadian cycling of
its messenger RNA levels. Nature 1990, 343:536–540.
32. Hamblen MJ, White NE, Emery PT, Kaiser K, Hall
JC. Molecular and behavioural analysis of four period
mutants in Drosophila melanogaster encompassing
extreme short, novel long, and unorthodox arrhythmic
types. Genetics 1998, 149:165–178.
33. Plautz JD, Kaneko M, Hall JC, Kay SA. Independent photoreceptive circadian clocks throughout
Drosophila. Science 1997, 278:1632–1635.
34. Rieger D, Stanewsky R, Helfrich-Förster C. Cryptochrome, compound eyes, Hofbauer-Buchner eyelets,
and ocelli play different roles in the entrainment and
33
Advanced Review
masking pathway of the locomotor activity rhythm in
the fruit fly Drosophila melanogaster. J Biol Rhythms
2003, 18:377–391.
retinal is required for phototaxis signaling by sensory rhodopsins in Halobacterium halobium. Biophys
J 1990, 57:807–814.
35. Hattar S, Liao HW, Takao M, Berson DM, Yau KW.
Melanopsin-containing retinal ganglion cells: architecture, projections, and intrinsic photosensitivity.
Science 2002, 295:1065–1070.
49. Gordeliy VI, Labahn J, Moukhametzianov R, Efremov R, Granzin J, Schlesinger R, Büldt G, Savopol
T, Scheidig AJ, Klare JP, et al. Molecular basis of
transmembrane signalling by sensory rhodopsin IItransducer complex. Nature 2002, 419:484–487.
36. Bellingham J, Chaurasia SS, Melyan Z, Liu C, Cameron MA, Tarttelin EE, luvone PM, Hankins MW,
Tosini G, Lucas RJ. Evolution of melanopsin photoreceptors: discovery and characterization of a new
melanopsin in nonmammalian vertebrates. PLoS Biol
2006, 4:e254.
37. Shigeyoshi Y, Taguchi K, Yamamoto S, Takekida S,
Yan L, Tei H, Moriya T, Shibata S, Loros JJ, Dunlap JC, et al. Light-induced resetting of a mammalian
circadian clock is associated with rapid induction of
the mPer1 transcript. Cell 1997, 91:1043–1053.
38. Jékely G. Evolution of phototaxis. Philos Trans R Soc
London B Biol Sci 2009, 364:2795–2808.
39. Bhaya D. Light matters: phototaxis and signal transduction in unicellular cyanobacteria. Mol Microbiol
2004, 53:745–754.
40. Häder DP. Photosensory behavior in procaryotes.
Microbiol Rev 1987, 51:1–21.
41. Yoshihara S, Ikeuchi M. Phototactic motility in the
unicellular cyanobacterium Synechocystis sp. PCC
6803. Photochem Photobiol Sci 2004, 3:512–518.
42. Song JY, Cho HS, Cho JI, Jeon JS, Lagarias JC, Park
YI. Near-UV cyanobacteriochrome signaling system
elicits negative phototaxis in the cyanobacterium Synechocystis sp. PCC 6803. Proc Natl Acad Sci USA 2011,
108:10780–10785.
43. Kondou Y, Mogami N, Hoshi F, Kutsuna S, Nakazawa M, Sakurai T, Matsui M, Kaneko T, Tabata S,
Tanaka I, et al. Bipolar localization of putative photoreceptor protein for phototaxis in thermophilic
cyanobacterium Synechococcus elongatus. Plant Cell
Physiol 2002, 43:1585–1588.
44. Yoshihara S, Katayama M, Geng X, Ikeuchi M.
Cyanobacterial phytochrome-like PixJ1 holoprotein
shows novel reversible photoconversion between blueand green-absorbing forms. Plant Cell Physiol 2004,
45:1729–1737.
45. Luecke H, Schobert B, Lanyi JK, Spudich EN, Spudich
JL. Crystal structure of sensory rhodopsin II at 2.4
angstroms: insights into color tuning and transducer
interaction. Science 2001, 293:1499–1503.
46. Spudich JL. The multitalented microbial sensory
rhodopsins. Trends Microbiol 2006, 14:480–487.
47. Spudich JL, Yang CS, Jung KH, Spudich EN. Retinylidene proteins: structures and functions from archaea to
humans. Annu Rev Cell Dev Biol 2000, 16:365–392.
48. Yan B, Takahashi T, Johnson R, Derguini F, Nakanishi K, Spudich JL. All-trans/13-cis isomerisation of
34
50. Sasaki J, Spudich JL. Signal transfer in haloarchaeal
sensory rhodopsin-transducer complexes. Photochem
Photobiol 2008, 84:863–868.
51. Rudolph J, Tolliday N, Schmitt C, Schuster SC, Oesterhelt D. Phosphorylation in halobacterial signal
transduction. EMBO J 1995, 14:4249–4257.
52. Nultsch W, Schuchart H. Photomovement of the red
alga Porphyridium cruentum (Ag.) Naegeli II. Phototaxis. Arch Microbiol 1980, 125:181–188.
53. Luecke H, Schobert B, Stagno J, Imasheva ES, Wang
JM, Balashov SP, Lanyi JK. Crystallographic structure
of xanthorhodopsin, the light-driven proton pump
with a dual chromophore. Proc Natl Acad Sci USA
2008, 105:16561–16565.
54. Kateriya S, Nagel G, Bamberg E, Hegemann P.
‘‘Vision’’ in single-celled algae. News Physiol Sci 2004,
19:133–137.
55. Foster KW, Saranak J, Patel N, Zarilli G, Okabe M,
Kline T, Nakanishi K. A rhodopsin is the functional
photoreceptor for phototaxis in the unicellular eukaryote Chlamydomonas. Nature 1984, 311:756–759.
56. Nagel G, Ollig D, Fuhrmann M, Kateriya S, Musti
AM, Bamberg E, Hegemann P. Channelrhodopsin-1:
a light-gated proton channel in green algae. Science
2002, 296:2395–2398.
57. Nagel G, Szellas T, Huhn W, Kateriya S, Adeishvili N,
Berthold P, Ollig D, Hegemann P, Bamberg E.
Channelrhodopsin-2, a directly light-gated cationselective membrane channel. Proc Natl Acad Sci USA
2003, 100:13940–13945.
58. Sineshchekov OA, Jung KH, Soudich JL. Two rhodopsins mediate phototaxis to low- and high-intensity
light in Chlamydomonas reinhardtii. Proc Natl Acad
Sci USA 2002, 99:8689–8694.
59. Berthold P, Tsunoda SP, Ernst OP, Mages W, Gradmann D, Hegemann P. Channelrhodopsin-1 initiates
phototaxis and photophobic responses in Chlamydomonas by immediate light-induced depolarization.
Plant Cell 2008, 20:1665–1677.
60. Suzuki T, Yamasaki K, Fujita S, Oda K, Iseki M,
Yoshida K, Watanabe M, Daiyasu H, Toh H, Asamizu
E, et al. Archaeal-type rhodopsins in Chlamydomonas: model structure and intracellular localization. Biochem Biophys Res Commun 2003, 301:
711–717.
61. Dodge JD. The functional and phylogenetic significance of dinoflagellate eyespots. Biosystems 1983,
16:259–267.
Evolution of vision
62. Francis D. On the eyespot of the dinoflagellate, Nematodinium. J Exp Biol 1967, 47:495–501.
63. Greuet C. Structure fine de lócelle d’Erythropsis pavillardi Hertwig, ptéridinien Warnowiidae Lindemann.
CR Acad Sci (Paris) 1965, 261:1904–1907.
64. Greuet C. Anatomie ultrastructurale des Ptéridiniens
Warnowiidae en rapport avec la differenciation des
organites cellulaires. PhD dissertation, Université de
Nice, 1969.
65. Okamoto OK, Hastings JW. Novel dinoflagellate
clock-related genes identified through microarray analysis. J Phycol 2003, 39:519–526.
66. Hader DP. Polarotaxis, gravitaxis and vertical phototaxis in the green flagellate, Euglena gracilis. Arch
Microbiol 1987, 147:179–183.
67. Ntefidou M, Iseki M, Watanabe M, Lebert M, Häder
DP. Photoactivated adenylyl cyclase controls phototaxis in the flagellate Euglena gracilis. Plant Physiol
2003, 133:1517–1521.
68. Thimann KV, Went FW. On the chemical nature of
the root forming hormone. Proc Koninkl Acad Wetenschap Amsterdam 1934, 37:456–459.
69. Bergman K, Burke PV, Cerdá-Olmedo E, David CN,
Delbrück M, Foster KW, Goodell EW, Heisenberg
M, Meissner G, Zalokar M, et al. Phycomyces. Bacteriol Rev 1969, 33:99–157.
70. Cerdá-Olmedo E. Phycomyces and the biology of light
and color. FEMS Microbiol Rev 2001, 25:503–512.
71. Bergman K, Eslava AP, Cerdá-Olmedo E. Mutants of
Phycomyces with abnormal phototropism. Mol Gen
Genet 1973, 123:1–16.
72. Orejas M, Peláez MI, Alvarez Ml, Eslava AP. A genetic
map of Phycomyces blakesleeanus. Mol Gen Genet
1987, 210:69–76.
73. Campuzano V, Galland P, Eslava AP, Alvarez MI.
Genetic characterization of two phototropism mutants
of Phycomyces with defects in the genes madl and
madJ. Curr Genet 1995, 27:524–527.
74. Sanz C, Rodriguez-Romero J, Idnurm A, Christie JM,
Heitman J, Corrochano LM, Eslava AP. Phycomyces
MADB interacts with MADA to form the primary
photoreceptor complex for fungal phototropism. Proc
Natl Acad Sci USA 2009, 106:7095–7100.
75. Leys SP, Degnan BM. Cytological basis of photoresponsive behavior in a sponge larva. Biol Bull 2001,
201:323–338.
76. Srivastava M, Simakov O, Chapman J, Fahey B,
Gauthier ME, Mitros T, Richards GS, Conaco C,
Dacre M, Hellsten U, et al. The Amphimedon queenslandica genome and the evolution of animal complexity. Nature 2010, 466:720–726.
77. Suga H, Schmid V, Gehring WJ. Evolution and functional diversity of jellyfish opsins. Curr Biol 2008,
18:51–55.
78. Müller WE, Wang X, Schröder HC, Korzhev M,
Grebenjuk VA, Markl JS, Jochum KP, Pisignano
D, Wiens M. A cryptochrome-based photosensory
system in the siliceous sponge Suberites domuncula
(Demospongiae). FEBS J 2010, 277:1182–1201.
79. Wiens M, Wang X, Unger A, Schröder HC, Grebenjuk
VA, Pisignano D, Jochum KP, Müller WE. Flashing
light signaling circuit in sponges: endogenous light generation after tissue ablation in Suberites domuncula.
J Cell Biochem 2010, 111:1377–1389.
80. Müller WEG, Kasueske M, Wang X, Schröder HC,
Wang Y, Pisignano D, Wiens M. Luciferase a
light source for the silica-based optical waveguides
(spicules) in the demosponge Suberites domuncula.
Cell Mol Life Sci 2009, 66:537–552.
81. Nordström K, Wallén R, Seymour J, Nilsson D. A simple visual system without neurons in jellyfish larvae.
Proc Biol Sci 2003, 270:2349–2354.
82. Parkefelt L, Skogh C, Nilsson DE, Ekström P. Bilateral symmetric organization of neural elements in the
visual system of a coelenterate, Tripedalia cystophora
(Cubozoa). J Comp Neurol 2005, 492:251–262.
83. Gehring WJ. The genetic control of eye development
and its implications for the evolution of the various
eye-types. Zoology 2001, 104:171–183.
84. Jékely G, Colombelli J, Hausen H, Guy K, Stelzer
E, Nédélec F, Arendt D. Mechanism of phototaxis in
marine zooplankton. Nature 2008, 456:395–399.
85. Rockwell NC, Lagarias JC. A brief history of phytochromes. Chemphyschem 2010, 11:1172–1180.
86. Yeh KC, Wu SH, Murphy JT, Lagarias JC. A cyanobacterial phytochrome two-component light sensory
system. Science 1997, 277:1505–1508.
87. Yoshihara S, Katayama M, Geng X, Ikeuchi M.
Cyanobacterial phytochrome-like PixJ1 holoprotein
shows novel reversible photoconversion between blueand green-absorbing forms. Plant Cell Physiol 2004,
45:1729–1737.
88. Yu X, Liu H, Klejnot J, Lin C. The cryptochrome blue
light receptors. In: Arabidopsis Book. Vol. 8. NIH
Public Access Author Manuscript; 2010, e0135.
89. Lin C, Robertson DE, Ahmad M, Raibekas AA, Joms
MS, Dutton PL, Cashmore AR. Association of flavin
adenine dinucleotide with the Arabidopsis blue light
receptor CRY1. Science 1995, 269:968–970.
90. Gehring W, Rosbash M. The coevolution of blue-light
photoreception and circadian rhythms. J Mol Evol
2003, 57(Suppl. 1):S286–S289.
91. Chaves I, Pokorny R, Byrdin M, Hoang N, Ritz T,
Brettel K, Essen LO, van der Horst GTJ, Batschauer A, Ahmad M. The cryptochromes: blue light
photoreceptors in plants and animals. Annu Rev Plant
Biol 2011, 62:335–364.
92. Sancar A. Structure and function of DNA photolyase.
Biochemistry 1994, 33:2–9.
35
Advanced Review
93. Todo T, Ryo H, Yamamoto K, Toh H, Inui T,
Avaki H, Nomura T, Ikenaga M. Similarity among
the Drosophila (6-4) photolyase, a human photolyase
homolog, and the DNA photolyase-blue-light photoreceptor family. Science 1996, 272:109–112.
94. Ritz T, Adem S, Schulten K. A model for
photoreceptor-based magnetoreception in birds. Biophys J 2000, 78:707–718.
95. Wiltschko W, Wiltschko R. Magnetic compass of
European robins. Science 1972, 176:62–64.
96. Wiltschko W, Wiltschko R. Magnetic orientation in
birds. J Exp Biol 1996, 199:29–38.
97. Semm P, Schneider T. Magnetic responses in the central nervous system of birds. In: Lieth H, eds. Effects of
Atmospherical and Geophysical Variables in Biology
and Medicine. Progress in Biometerology. Vol. 8. The
Hague: SPB Academic Publishing.
98. Liedvogel M, Maeda K, Henbest K, Schleicher E,
Simon T, Timmel CR, Hore PJ, Mouritsen H. Chemical magnetoreception: bird cryptochrome 1a is excited
by blue light and forms long-lived radical pairs. PLoS
One 2007, 2:e1106.
99. Ahmad M, Galland P, Ritz T, Wiltschko R, Wiltschko
W. Magnetic intensity affects cryptochromedependent responses in Arabidopsis thaliana. Planta
2007, 225:615–624.
100. Yoshii T, Ahmad M, Helfrich-Förster C. Cryptochrome mediates light-dependent magnetosensitivity
of Drosophila’s circadian clock. PLoS Biol 2009,
7:e1000086.
101. Gegear RJ, Foley LE, Casselman A, Reppert SM. Animal cryptochromes mediate magnetoreception by an
unconventional photochemical mechanism. Nature
2010, 463:804–807.
102. Maeda K, Robinson AJ, Henbest KB, Hogben HJ,
Biskup T, Ahmad M, Schleicher E, Weber S, Timmel
CR, Hore PJ. Magnetically sensitive light-induced
reactions in cryptochrome are consistent with its proposed role as a magnetoreceptor. Proc Natl Acad Sci
USA 2012, 109:4774–4779.
103. Hubbell WL, Altenbach C, Hubbell CM, Khorana
HG. Rhodopsin structure, dynamics, and activation:
a perspective from crystallography site-directed spin
labeling, sulfhydryl reactivity, and disulfide crosslinking. Adv Protein Chem 2003, 63:243–290.
104. Dunlap JC, Loros JJ. Neurospora photoreceptors. In:
Briggs WR, Spudich JL, eds. Handbook of Photosensory Receptors. Weinheim, Germany: Wiley-VCH;
2005, 371–388.
105. Spudich JL, Jung KH. Microbial rhodopsins: phylogenetic and functional diversity. In: Briggs WR, Spudich JL, eds. Handbook of Photosensory Receptors.
Weinheim, Germany: Wiley-VCH; 2005, 1–24.
106. Oesterheld D. The structure and mechanism of the
family of retinal proteins from halophilic archaea.
Curr Opin Struct Biol 1998, 8:489–500.
36
107. Váró G. Analogies between halorhodopsin and
bacteriorhodopsin. Biochim Biophys Acta 2000,
1460:220–229.
108. Hoff WD, Jung KH, Spudich JL. Molecular mechanism of photosignaling by archaeal sensory
rhodopsins. Annu Rev Biophys Biomol Struct 1997,
26:223–258.
109. Yao VJ, Spudich JL. Primary structure of an archaebacterial transducer, a methyl-accepting protein associated with sensory rhodopsin I. Proc Natl Acad Sci
USA 1992, 89:11915–11919.
110. Seidel R, Scharf B, Gautel M, Kleine K, Oesterhelt D,
Engelhard M. The primary structure of sensory
rhodopsin II: a member of an additional retinal protein subgroup is coexpressed with its transducer, the
halobacterial transducer of rhodopsin II. Proc Natl
Acad Sci USA 1995, 92:3036–3040.
111. Béjà O, Spudich EN, Spudich JL, Leclerc M, DeLong
EF. Proteorhodopsin phototrophy in the ocean. Nature
2001, 411:786–789.
112. Jung KH, Trivedi VD, Spudich JL. Demonstration of a
sensory rhodopsin in eubacteria. Mol Microbiol 2003,
47:1513–1522.
113. Wang S, Kim SY, Jung KH, Ladizhansky V, Brown
LS. A eukaryotic-like interaction of soluble cyanobacterial sensory rhodopsin transducer with DNA. J Mol
Biol 2011, 411:449–462.
114. Brown LS, Dioumaev AK, Lanyi JK, Spudich EN, Spudich JL. Photochemical reaction cycle and proton
transfers in Neurospora rhodopsin. J Biol Chem 2001,
276:32495–32505.
115. Saranak J, Foster KW. Rhodopsin guides fungal phototaxis. Nature 1997, 387:465–466.
116. Nagel G, Brauner M, Liewald JF, Adeishvili N,
Bamberg E, Gottschalk A. Light activation of
channelrhodopsin-2 in excitable cells of Caenorhabditis elegans triggers rapid behavioral responses. Curr
Biol 2005, 15:2279–2284.
117. Bamann C, Nagel G, Bamberg E. Microbial rhodopsins in the spotlight. Curr Opin Neurobiol 2010,
20:610–616.
118. Lagali PS, Balya D, Awatramani GB, Münch TA, Kim
DS, Busskamp V, Cepko CL, Roska B. Light-activated
channels targeted to ON bipolar cells restore visual
function in retinal degeneration. Nat Neurosci 2008,
11:667–675.
119. Yeandle S, Spiegler JB. Light-evoked and spontaneous
discrete waves in the ventral nerve photoreceptor of
Limulus. J Gen Physiol 1973, 61:552–571.
120. Baylor DA, Lamb TD, Yau KW. Responses of retinal
rods to single photons. J Physiol 1979, 288:613–634.
121. Henderson SR, Reuss H, Hardie RC. Single photon
responses in Drosophila photoreceptors and their regulation by Ca2+ . J Physiol 2000, 524(Pt 1):179–194.
Evolution of vision
122. Palczewski K, Kumasaka T, Hori T, Behnke CA,
Motoshima H, Fox BA, Le Trong I, Teller DC,
Okada T, Stenkamp RE, et al. Crystal structure of
rhodopsin: a G protein-coupled receptor. Science
2000, 289:739–745.
123. Murakami M, Kouyama T. Crystal structure of squid
rhodopsin. Nature 2008, 453:363–367.
124. Teller DC, Okada T, Behnke CA, Palczewski K, Stenkamp RE. Advances in determination of a highresolution three-dimensional structure of rhodopsin,
a model of G-protein-coupled receptors (GPCRs).
Biochem 2001, 40:7761–7772.
125. Okada T, Palczewski K. Crystal structure of rhodopsin: implications for vision and beyond. Curr Opin
Struct Biol 2001, 11:420–426.
126. Honig B, Ebrey T, Callender RH, Dinur U, Ottolenghi M. Photoisomerization, energy storage, and
charge separation: a model for light energy transduction in visual pigments and bacteriorhodopsin. Proc
Natl Acad Sci USA 1979, 76:2503–2507.
138. Plachetzki DC, Degnan BM, Oakley TH. The origins
of novel protein interactions during animal opsin evolution. PLoS One 2007, 2:e1054.
139. Rasmussen SG, DeVree BT, Zou Y, Kruse AC,
Chung KY, Kobilka TS, Thian FS, Chae PS, Pardon E,
Calinski D, et al. Crystal structure of the β2 adrenergic receptor-Gs protein complex. Nature 2011,
477:549–555.
140. Koyanai M, Takano K, Tsukamoto H, Ohtsu K, Tokunaga F, Terakita A. Jellyfish vision starts with cAMP
signaling mediated by opsin G s cascade. Proc Nat
Acad Sci USA 2008, 105:15576–15580.
141. Klein PS, Sun TJ, Saxe CL, 3rd. Kimmel AR, Johnson RL, Devreotes PN. A chemoattractant receptor
controls development in Dictyostelium discoideum.
Science 1988, 241:1467–1472.
142. Shen WL, Kwon Y, Adegbola AA, Luo J, Chess A,
Montell C. Function of rhodopsin in temperature discrimination in Drosophila. Science 2011,
331:1333–1336.
127. Kumauchi M, Ebry T. Visual pigments as photoreceptors. In: Briggs WR, Spudich JL, eds. Handbook of
Photosensory Receptors. Wiley-VCH; 2005, 43–76.
143. Plachetzki DC, Fong CR, Oakley TH. Cnidocyte discharge is regulated by light and opsin-mediated phototransduction. BMC Biol 2012, 10:17–26.
128. Eakin R. Evolutionary significance of photoreceptors:
in retrospect. Integr Comp Biol 1979, 19:647.
144. Yokoyama S. Evolution of dim-light and color vision
pigments. Annu Rev Genomics Hum Genet 2008,
9:259–282.
129. Eakin R. Continuity and diversity of photoreceptors.
In: Westfall J, ed. Visual Cells in Evolution. New York:
Raven Press; 1982, 91–105.
130. Land MF, Nilsson DE. Animal Eyes. Oxford University Press; 2002, 201–221.
131. McReynolds JS, Gorman ALF. Photoreceptor potentials of opposite polarities in the eye of the scallop, Pecten irradians. J General Physiol 1970, 56:
376–391.
132. Fesenko EE, Kolesnikov SS, Lyubarsky AL. Induction
by cyclic GMP of cationic conductance in plasma
membrane of retinal rod outer segment. Nature 1985,
313:310–313.
133. Gomez MD, Nasi E. Blockage of the light-sensitive
conductance in hyperpolarizing photoreceptors of
the scallop. Effects of tetraethylammonium and 4aminopyridine. J Gen Physiol 1994, 104:487–505.
134. Gerhart J, Kischner M. Cells, Embryos, and Evolution. Malden, MA: Blackwell Science; 1997.
135. Feuda R.Hamilton S, Nicholls S, King N, McInerney
JO, Pisani D. Genome-scale analyses of non-bilaterian
metazoans clarify opsin evolution and the origin of
vision in animals Proc Nat Acad Sci USA. In press.
136. Porter ML, Blasic JR, Bok MJ, Cameron EG, Pringle
T, Cronin TW, Robinson PR. Shedding new light
on opsin evolution. Proc Biol Sci 2012, 279:
3–14.
137. Finn JT, Solessio EC, Yau KW. A cGMP-gated cation
channel in depolarizing photoreceptors of the lizard
parietal eye. Nature 1997, 385:815–819.
145. Pichaud F, Briscoe A, Desplan C. Evolution of color
vision. Curr Opin Neurobiol 1999, 9:622–627.
146. Nathans J, Thomas D, Hogness DS. Molecular genetics of human color vision: the genes encoding blue,
green, and red pigments. Science 1986, 232:193–202.
147. Yokoyama S, Zhang H, Radlwimmer FB, Blow NS.
Adaptive evolution of color vision of the Comoran
coelacanth (Latimeria chalumnae). Proc Natl Acad Sci
USA 1999, 96:6279–6284.
148. Santschi F. L’orientation sidérale des fourmis, et
quelques considerations sur leurs différentes possibilités d’orientation. Mém Soc Vaudoise Sci Nat 1923,
4:137–175.
149. von Frisch K. Die Polarisation des Himmelslichtes
als orientierender Faktor bei den Tänzen der Bienen.
Experientia 1949, 5:142–148.
150. Wehner R, Labhart T. Polarization vision. In: Warrant E, Nilsson DE, eds. Invertebrate Vision. Cambridge: Cambridge University Press; 2006, 291–348.
151. Wehner R. The polarization-vision project: championing organismic biology. In: Schildberger K, Elsner
N, eds. Neural Basis of Behavioral Adaptations.
Fortschritte der Zoologie. Vol. 39. Stuttgart, Jena,
New York: Gustav Fischer; 1994, 103–143.
152. Reyes-Prieto A, Weber AP, Bhattacharya D. The origin and establishment of the plastid in algae and plants.
Annu Rev Genet 2007, 41:147–168.
153. Takeda H, Nishimura K, Agata K. Planarians maintain a constant ratio of different cell types during
37
Advanced Review
changes in body size by using the stem cell system.
Zoolog Sci 2009, 26:805–813.
development of the pigmented epithelium of the vertebrate eye. Pigment Cell Res 2006, 19:380–394.
154. Hesse R. Untersuchungen über die Organe der
Lichtempfindung bei niederen Tieren. II. Die Augen der
Plathelminthen. Zeitung für wissenschaftliche Zoologie 1897, 62:527–582.
168. Yajima I, Endo K, Sato S, Toyoda R, Wada H, Shibahara S, Numakunai T, Ikeo K, Gojobori T, Goding
CR, et al. Cloning and functional analysis of ascidian Mitf in vivo: insights into the origin of vertebrate
pigment cells. Mech Dev 2003, 120:1489–1504.
155. Purschke G, Arendt D, Hausen H, Müller MC. Photoreceptor cells and eyes in Annelida. Arthropod Struct
Dev 2006, 35:211–230.
156. Passamaneck YJ, Furchheim N, Heynol A, Martindale MQ, Lüter C. Ciliary photoreceptorin the cerebral eyes of a protostome larva. Evodevo 2011, 2:6.
157. Eakin RM, Brandenburger JL. Fine structure of the
eyes of Pseudoceros canadensis (Turbellarla, Polydadida). Zoomorphology 1981, 98:1–16.
158. Hertwig P. Neue Mutationen und Kopplungsgruppen bei der Hausmaus. Z. Indukt. Abstammungs-u.
Vererbungsl 1942, 80:220-246.
159. Tachibana M, Hara Y, Vyas D, Hodgkinson C, Fex J,
Grundfast K, Arnheiter H. Cochlear disorder associated with melanocyte anomaly in mice with a transgenic insertational mutation. Mol Cell Neurosci 1992,
3:433–445.
160. Hodgkinson CA, Moore KJ, Nakayama A, Steingrimsson E, Copeland NG, Jenkins NA, Arnheiter H.
Mutation at the mouse microphtalmia locus are associated with defects in a gene encoding a novel basic helixloop-helix zipper protein. Cell 1993, 74:395–404.
161. Hughes MJ, Lingrel JB, Krakowsky JM, Anderson
KP. A helix-loop-helix transcription factor-like gene
is located at the mi locus. J Biol Chem 1993,
268:20687–20690.
162. Arnheiter H. Evolutionary biology. Eyes viewed from
the skin. Nature 1998, 391:632–633.
163. Nakayama A, Nguyen MT, Chen CC, Opdecamp K,
Hodgkinson CA, Arnheiter H. Mutations in microphthalmia, the mouse homolog of the human deafness
gene MITF, affect neuroepithelial and neural crestderived melanocytes differently. Mech Dev 1998,
70:155–166.
164. Tachibana M, Takeda K, Nobukuni Y, Urabe K, Long
JE, Meyers KA, Aaronson SA, Miki T. Ectopic expression of MITF, a gene for Waardenburg syndrome type
2, converts fibroblasts to cells with melanocyte characteristics. Nat Genet 1996, 14:50–54.
165. Bäumer N, Marquardt T, Stoykova A, Spieler D,
Treichel D, Ashery-Padan R, Gruss P. Retinal pigmented epithelium determination requires the redundant activities of Pax2 and Pax6. Development 2003,
130:2903–2915.
169. Rehli M, Den Elzen N, Cassady AI, Ostrowski MC,
Hume DA. Cloning and characterization of the murine
genes for bHLH-ZIP transcription factors TFEC and
TFEB reveal a common gene organization for all MiT
subfamily members. Genomics 1999, 56:111–120.
170. Hallsson JH, Haflidadotti BS, Stivers C, Odenwald W,
Arnheiter H, Pignoni F, Steingrimsson E. The basic
helix-loop-helix leucine zipper transcription factor
Mitf is conserved in Drosophila and functions in eye
development. Genetics 2004, 167:233–241.
171. Kozmik Z, Ruzickova J, Jonasova K, Matsumoto Y,
Vopalensky P, Kozmikova I, Strand H, Kawamura S,
Piatigorsky J, Paces V, et al. Assembly of the cnidarian camera-type eye from vertebrate-like components.
Proc Natl Acad Sci USA 2008, 105:8989–8993.
172. Quiring R, Walldorf U, Kloter U, Gehring WJ. Homology of the eyeless gene of Drosophila to the small eye
gene in mice and Aniridia in humans. Science 1994,
265:795–789.
173. Halder G, Callaerts P, Gehring WJ. Induction of
ectopic eyes by targeted expression of the eyeless gene
in Drosophila. Science 1995, 267:1788–1792.
174. Walther C, Gruss P. Pax-6, a murine paired box gene,
is expressed in the developing CNS. Development
1991, 113:1435–1449.
175. Hill RE, Favor J, Hogan BL, Ton CC, Saunders GF,
Hanson IM, Prosser J, Jordan T, Hastie ND, van
Heyningen V. Mouse small eye results from mutations
in a paired-like homeobox-containing gene. Nature
1991, 354:522–525.
176. Ton CC, Hirvonen H, Miwa H, Weil MM, Monaghan
P, Jordan T, van Heyningen V, Hastie ND, MeijersHeijboesr H, Drechaler M, et al. Positional cloning
and characterization of a paired box- and homeoboxcontaining gene from the aniridia region. Cell 1991,
67:1059–1074.
177. Hoge MA. Another gene in the fourth chromosome of
Drosophila. Am Nat 1915, 49:47–49.
178. Weintraub H, Tapscott SJ, Davis RL, Thayer MJ,
Adam MA, Lassar AB, Miller AD. Activation of
muscle-specific genes in pigment, nerve, fat, liver, and
fibroblast cell lines by forced expression of MyoD.
166. Gehring W, Seimiya M. Eye evolution and the origin of Darwin’s eye prototype. Ital J Zool 2010,
77:124–136.
179. Lewis EB. The 1991 Albert Lasker Medical Awards.
Clusters of master control genes regulate the development of higher organisms. JAMA 1992, 267:
1524–1531.
167. Bharti K, Nguyen MT, Skuntz S, Bertuzzi S, Arnheiter H. The other pigment cell: specification and
180. Schneuwly S, Klemenz R, Gehring WJ. Redesigning
the body plan of Drosophila by ectopic expression
38
Evolution of vision
of the homoeotic gene Antennapedia. Nature 1987,
325:816–818.
181. Brand AH, Perrimon N. Targeted gene expression as
a means of altering cell fates and generating dominant
phenotypes. Development 1993, 118:401–415.
182. Clements J, Lu Z, Gehring WJ, Meinertzhagen IA,
Callaerts P. Central projections of photoreceptor
axons originating from ectopic eyes in Drosophila.
183. Czerny T, Halder G, Kloter U, Souabni A, Gehring
WJ, Busslinger M. Twin of eyeless, a second pax-6
gene of Drosophila, acts upstream of eyeless in the control of eye development. Mol Cell 1999, 3:297–307.
184. Halder G, Callaerts P, Flister S, Walldorf U, Kloter U,
Gehring WJ. Eyeless initiates the expression of both
sine oculis and eyes absent during Drosophila
compound eye development. Development 1998,
125:2181–2191.
185. Onuma Y, Takahashi S, Asashima M, Kurata S, Gehring WJ. Conservation of Pax6 function and upstream
activation by Notch signaling in eye development
of frogs and flies. Proc Natl Acad Sci USA 2002,
99:2020–2025.
186. Blanco J, Seimiya M, Pauli T, Reichert H, Gehring
WJ. Wingless and Hedgehog signaling pathways regulate orthodenticle and eyes absent during ocelli development in Drosophila. Dev Biol 2009, 329:104–115.
187. Cheyette BN, Green PJ, Martin K, Garren H, Hartenstein V, Zipursky SL. The Drosophila sine oculis locus
encodes a homeodomain-containing protein required
for the development of the development of the entire
visual system. Neuron 1994, 12:977–996.
188. Serikaku MA, OT́ousa JE. Sine oculis is a homeobox
gene required for Drosophila visual system development. Genetics 1994, 138:1137–1150.
189. Bonini NM, Leiserson WM, Benzer S. The eyes absent
gene: genetic control of cell survival and differentiation in the developing Drosophila eye. Cell 1993,
72:379–395.
190. Pignoni F, Hu B, Zavitz KH, Xiao J, Garrity PA,
Zipursky SL. The eye-specification proteins So and
Eya form a complex and regulate multiple steps in
Drosophila eye development. Cell 1997, 91:881–891.
191. Jemc J, Rebay I. The eyes absent family of phosphotyrosine phosphatases: properties and roles in developmental regulation of transcription. Annu Rev Biochem
2007, 76:513–538.
192. Mardon G, Solomon NM, Rubin GM. Dachshund
encodes a nuclear protein required for normal eye and
leg development in Drosophila. Development 1994,
120:3473–3486.
Two Pax genes, eye gone and eyeless, act cooperatively
in promoting Drosophila eye development. Development 2003, 130:2939–2951.
195. Glardon S, Callaerts P, Halder G, Gehring WJ. Conservation of Pax-6 in a lower chordate, the ascidian Phallusia mammillata. Development 1997, 124:
817–825.
196. Czerny T, Busslinger M. DNA-binding and transactivation properties of Pax-6: three amino acids in
the paired domain are responsible for the different
sequence recognition of Pax-6 and BSAP (Pax-5). Mol
Cell Biol 1995, 15:2858–2871.
197. Ullrich-Lütera EM, Dupont S, Arboleda E, Hausen
H, Arnone MI. Unique system of photoreceptors in
sea urchin tube feet. Proc Natl Acad Sci USA 2011,
108:8367–8372.
198. Lesser MP, Carleton KL, Böttger SA, Barry TM,
Walker CW. Sea urchin tube feet are photosensory
organs that express a rhabdomeric-like opsin and
PAX6. Proc Biol Sci 2011, 278:3371–3379.
199. Michaut L, Flister S, Neeb M, White KP, Certa U,
Gehring WJ. Analysis of the eye development pathway in Drosophila using DNA microarrays. Proc Natl
Acad Sci USA 2003, 100:4024–4029.
200. Codutti L, van Ingen H, Vascotto C, Fogolari F,
Corazza A, Tell G, Quadrifoglio F, Viglino P, Boelens R, Esposito G. The solution structure of DNA-free
Pax-8 paired box domain accounts for redox regulation of transcriptional activity in the pax protein
family. J Biol Chem 2008, 283:33321–33328.
201. Xu W, Rould MA, Jun S, Desplan C, Pabo CO. Crystal structure of a paired domain-DNA complex at 2.5
A resolution reveals structural basis for Pax developmental mutations. Cell 1995, 80:639–650.
202. Xu HE, Rould MA, Xu W, Epstein JA, Maas RL,
Pabo CO. Crystal structure of the human Pax6 paired
domain-DNA complex reveals specific roles for the
linker region and carboxy-terminal subdomain in
DNA binding. Genes Dev 1999, 13:1263–1275.
203. Jacob F. Evolution and tinkering. Science 1977,
196:1161–1166.
204. Punzo C, Kurata S, Gehring WJ. The eyeless homeodomain is dispensable for eye development in
Drosophila. Genes Dev 2001, 15:1716–1723.
205. Suga H, Tschopp P, Graziussi DF, Stierwald M, Schmid V, Gehring WJ. Flexibly deployed Pax genes in eye
development at the early evolution of animals demonstrated by studies on a hydrozoan jellyfish. Proc Natl
Acad Sci USA 2010, 107:14263–14268.
193. Shen W, Mardon G. Ectopic eye development in
Drosophila induced by directed dachshund expression.
Development 1997, 124:45–52.
206. Kozmik Z, Daube M, Frei E, Norman B, Kos L, Dishaw LJ, Noll M, Piatigorsky J. Role of Pax genes in
eye evolution: a cnidarian PaxB gene uniting Pax2 and
Pax6 functions. Dev Cell 2003, 5:773–785.
194. Jang CC, Chao JL, Jones N, Yao LC, Bessarab DA,
Kuo YM, Jun S, Desplan C, Beckendorf SK, Sun YH.
207. Tomarev SI, Callaerts P, Kos L, Zinovieva R, Halder
G, Gehring W, Piatigorsky J. Squid Pax-6 and eye
39
Advanced Review
development. Proc Natl Acad Sci USA 1997, 94:
2421–2426.
208. Yoshida M, Ogura A. Genetic mechanisms involved in
the evolution of the cephalopod camera eye revealed
by transcriptomic and developmental studies. BMB
Evol Biol 2011, 180–191.
209. Ogura A, Ikeo K, Gojobori T. Comparative analysis
of gene expression for convergent evolution of camera
eye between octopus and human. Genome Res 2004,
14:1555–1561.
210. Blanco J, Gehring WJ. Analysis of twin of eyeless
regulation during early embryogenesis in Drosophila
melanogaster. Gene Expr Patterns 2008, 8:523–527.
211. Hou XG, Aldridge R, Bergstrom J, Siveter DJ, Siveter
D, Feng X-H. The Cambrian Fossils of Chengjiang
China, the Flowering of Early Animal Life 2007.
Oxford: Blackwell.
212. Gehring WJ. Chance and necessity in eye evolution.
Genome Biol Evol 2011, 3:1053–1066.
215. Kamachi Y, Uchikawa M, Tanouchi A, Sekido R,
Kondoh H. Pax6 and SOX2 form a co-DNA-binding
partner complex that regulates initiation of lens development. Genes Dev 2001, 15:1272–1286.
216. Blanco J, Girard F, Kamachi Y, Kondoh H, Gehring
WJ. Functional analysis of the chicken delta1-crystallin
enhancer activity in Drosophila reveals remarkable
evolutionary conservation between chicken and fly.
Development 2005, 132:1895–1905.
217. Punzo C, Seimiya M, Flister S, Gehring WJ, Plaza S.
Differential interactions of eyeless and twin of eyeless with the sine oculis enhancer. Development 2002,
129:825–834.
218. Lister JA, Robertson CP, Lepage T, Johnson SL,
Raible DW. nacre encodes a zebrafish microphthalmiarelated protein that regulates neural-crest-derived pigment cell fate. Development 1999, 126:3757–3767.
219. Lister JA, Close J, Raible DW. Duplicate mitf genes
in zebrafish: complementary expression and conversation of melanogenic potential. Dev Biol 2001,
237:333–344.
213. Gunhaga L. The lens: a classical model of embryonic
induction providing new insights into cell determination in early development. Philos Trans R Soc Lond B
Biol Sci 2011, 366:1193–1203.
220. Piatigorsky J, Wistow GJ. Enzyme/crystallins: gene
sharing as an evolutionary strategy. Cell 1989, 57:
197–199.
214. Chariton-Perkins M, Brown NL, Cook TA. The lens
in focus: a comparison of lens development in
Drosophila and vertebrates. Mol Genet Genomics
2011, 286:189–213.
221. Janssens H, Gehring WJ. Isolation and characterization of drosocrytallin, a lens crystallin gene
of Drosophila melanogaster. Dev Biol 1999, 207:
204–214.
40

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