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Stefan Lemke
BIO 385 – Professor Walter
1/30/2013
Channelrhodopsin-2: Structure, Function, and Applications
Photoreception in both vertebrates and invertebrates is mediated by photoreceptors called
rhodopsins, consisting of seven transmembrane-helix proteins (opsins), covalently linked to retinal. Two
types of rhodopsins are differentiated on the basis of primary sequence. Animal rhodopsins, expressed in
animals (including humans), consist of G-protein coupled receptors. Microbial rhodopsins, on the other
hand, are found in archaea, eubacteria, fungi, and algae and are direct-light-activated regulators of
transmembrane ion conductance, including both light driven ion pumps and light-driven ion channels.
In a seminal paper by Nagel et al. (2003), it was demonstrated that Channelrhodopsin-2 (ChR2),
a microbial rhodopsin, is a directly light-gated cation-selective ion channel that can be expressed in
Xenepus oocytes. This discovery began with the detection of two sequences with homology to known
microbial opsins in the genome of Chlamydomonas reinhardtii, a single-cell green algae. These
sequences correspond to Channelrhodopsin-1 (ChR1) and Channelrhodopsin-2 (ChR2), and are
responsible for phototaxic (movement toward light) and photophobic (movement away from light)
behaviors in C. reinhardtii. Nagel et al. suggested that ChR2 could be used as a tool to modulate neural
activity by both increasing cytosolic Ca+ and depolarizing the cell in response to illumination.
This paper aims to discuss the structure and function of ChR2, focusing on the light-gating
mechanisms, electronegative pore, and similarities/differences to bacteriorhodopsin, another microbial
rhodopsin. While the exact structure of ChR2 remains unclear, recent efforts have provided important
information that serves to elucidate the structure and understand the function. I will first discuss the
current understanding of ChR2 structure and function and then go into the application of ChR2 in
optogenetics, both at its origins and today.
Structure and Function
Overall
Muller et al. (2011) performed one of the first studies to elucidate the structure of ChR2. Twodimensional crystals were grown and flattened on electron microscopy grids, allowing for projection
maps to be created, showing the surface topography of the protein. This map suggested two main
findings: (1) ChR2 contains seven transmembrane helices (TMs) and (2) forms a dimer (Figure 1). An
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SDS-PAGE gel was run to examine the dimeric nature of ChR2. As ChR2
remained dimeric in the gel despite the disruption of non-covalent interaction
by SDS, it was suggested that a remarkably stable dimer is formed by ChR2
that is likely to exist in dimeric form in the membrane.
The next structural data on ChR2 was provided by Kato et al. (2012).
As native ChR2 has proven hard to crystalize, Kato et al. formed a chimaera
(C1C2) between ChR1 and ChR2 consisting of ChR1 without its C-terminus
end and with the last two TMs swapped for those from ChR2. This chimaera Figure 1. Projection map
consisted of the 342 amino-terminal residues of the 737 amino acid residues of ChR2. From Muller et
al. (2011)
in the native channel. This was sufficient because the first 300 residues of the
amino-terminal end contain the seven-transmembrane domains, extra-/intra-cellular loops and the lightgating mechanisms. A crystal structure of this chimaera at 2.3 Å was presented to provide a working
model for the molecular function of ChR2 (Figure 2). The main finding for the overall structure was that
the dimeric interactions between the two C1C2 protein domains were found at the N-domains,
extracellular loops 1 (ECL1), and TM3s and TM4s.
Figure 2. Structure
of C1C2 dimer
viewed parallel to
membrane (left,
middle) and from
the extracellular
side (right). N and
C domain, TMs,
extracellular loops
(ECLs) and
intracellular loops
(ICLs) shown.
From Kato et al.
(2012).
Structural comparison with BR
Both Muller et al. and Kato et al. compared their structures (ChR2 and C1C2, respectively) to
bacteriorhosopin (bR), another microbial rhodopsin that acts as a light-driven proton pump. Muller el al.
showed that the arrangement of the seven TMs in both proteins is similar, with the greatest similarities
between TM3-6 (Figure 3). In addition, Muller et al. emphasized that while ChR2 forms a dimer, bR
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arranges itself as a trimer (Figure 3). Kato et al. found two distict differences between bR and C1C2.
First, C1C2 has an additional N-terminal and C-terminal domain. It is believed that the N-terminal
domain may explain the dimer formation in ChR2 while the C-terminal domain is involved in cellular
localization. Second, the extracellular ends of TM1 and TM2 in C1C2 are tilted outward by 3.0 and 4.1
Å, respectively (Figure 4). As we will see, the pore of ChR2 is formed between TM1, TM2, TM3, and
TM7, suggesting that this tilt may have implications on the cation permeability of ChR2.
Figure 3. Superimposed
projection map of ChR2 (red)
on bR (gray). From Muller et
al. (2011)
Figure 4. Structural comparison of ChR2 and bR.
From Kato et al. (2012).
Light-gating mechanisms
The light-gating mechanisms of ChR2 are mediated by retinal, a molecule that undergoes a
conformational change in response to the absorption of energy from photons. In the “dark state” of
ChR2, before any light stimulation, the retinal is found in an all-trans conformation and is covalently
linked to Lys 296 on TM7 forming a Schiff base. The changes in ChR2 that occur in response to light, or
the photocycle, were characterized by Ritter et al. (2008) using absorption of UV-visible light and
Fourier Transform Infrared Spectroscopy (FTIR). UV-visible absorbance is used to characterize the
protonation state of the retinal Schiff base. Generally, absorption below 400nm indicates a deprotonated
Schiff base, whereas a red-shifted absorption above 400nm indicates a protonated form. On the other
hand, FTIR allows for analysis of more discrete structural changes in the actual protein.
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Upon light excitation, the dark state of ChR2
(D470, as it absorbs light maximally at 470nm)
becomes a transient P500 intermediate, caused by the
isomerization of all-trans retinal to all-cis retinal
(Figure 5). The P500 quickly changes into another
transient
intermediate,
P390,
indicating
the
deprotonation of the Schiff base. The Schiff base
quickly reprotonates, which results in the conductive
state of the channel, P520. There are two main
structural changes that are believed to coincide with
Figure 5. Proposed photocycle of ChR2. From
the reprotonation of the Schiff base and lead to the Ritter et al. (2008).
opening of the channel. First, Kato et al. used their crystal structure to predict that Asp 292 is the
primary proton acceptor from the Schiff base. This protonation of Asp 292 is believed to result in ionic
“push” of nearby Lys 132 that moves Lys 132’s positive charge away from the channel pore and allows
cations to pass (Figure 6). Secondly, Eisenhauer et al. (2012) showed, also using FTIR, that the
protonated form of Glu 90 prevents cation flow through the channel pore (Figure 7). Therefore, they
hypothesize that the reprotonation of the Schiff base also coincides with the deprotonation and
subsequent solvation of Glu 90 which, combined with the lysine “push,” results in cation permeability.
The closing mechanisms of ChR2 are less well characterized, but are believed to involve changes in the
backbone of the channel, along with changes in hydrogen bonding of Glu 90 and occur on a much
slower timescale.
Figure 6. Interactions between the Schiff base, K292, and
K132. Black lines represent hydrogen bonds. From Kato et
al. (2012).
Figure 7. Comparison of solvation between
protonated and deprotonated Glu 90. From
Eisenhauer et al. (2012)
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Electronegative pore
Channel pore properties of ChR2 have taken longer to elucidate than many other aspects of the
protein. As late as 2011, Muller et al. speculated about the location of the pore channel, whether it
existed at the dimer interface or within each protein domain. Two recent findings, along with knowledge
of the photocycle, have made clear that a pore exists in each protein domain between TM1, TM2, TM3,
and TM7. First, Kato et al. showed that polar/negative amino acid residues form an electronegative pore
that exists on the extracellular side of the channel, consisting of Gln 95, Thr 98, Ser 102, Glu 122, Glu
129, Lys 132, Glu 136, Glu 140, Glu 162, Thr 285, Asp 292, and Asn 297 (Figure 8). Notably, TM2
contains most of the negative residues, and it is suggested that TM2 determines cation conductance and
selectivity.
The second finding was from Richards and Dempski (2012), who performed a sequence analysis
of ChR2 and bR and found eight serine residues within the transmembrane domains of bR that are not
present in ChR2 (C87S, G181S, G224S, and M255S located near the retinal, I197S, Q210S, and V269S
located at the cytoplasmic end of the transmembrane domain, and P234S) (Figure 9). By creating single
serine mutated ChR2 constructs corresponding to homologous residues in bR and expressing them in
Xenopus oocytes, they were able to examine the effects of these missing residues on the channel
properties of ChR2. These mutations were found to decrease cation conductance, perhaps due to the high
Figure 8. Position of serine
mutations. From Richards and
Dempski (2012).
Figure 9. Depiction of
hydrophilic and electronegative
pore lining surface. From Kato et
al. (2012).
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affinity for serine residues to form inter- and intra-helical hydrogen bonds, resulting in a reduction in
pore size. The authors concluded that the cation selectivity of the channel does not solely exist in the
extracellular pore, but further along the channel, and that these missing serines play a large role.
ChR2 and bR
We can now appreciate the differences between ChR2 and bR and ask ourselves, why is ChR2
permeable to cations while bR is a proton pump? Three main differences can help us preliminarily
explain this difference, further work will be needed to affirm and extend these conclusions. First of all,
the light-gated structural changes are much greater in ChR2 compared to bR. As mentioned above, Asp
292 is the proton acceptor from the deprotonation of the Schiff base in ChR2. This leads to a cascade of
structural changes that allow cation permeability. However, in bR the primary proton acceptor is a water
molecule, which does not result in the same structural changes. Incidentally, this also accounts for
ChR2’s blue-shifted absorption spectrum (max absorbance at a wavelength of 470), as compared to that
of BR (max absorbance at a wavelength of 568 nm). The absorption spectrum is determined by the
energy difference between the ground and first excited state of retinal. This is determined by the distance
between the protonated Schiff base and its counter ion, which for ChR2 is 1 Å closer to the Schiff base
compared to BR. The second difference accounting for cation permeability in ChR2, is the outward
shifted TM1 and TM2. As emphasized by Kato et al., TM2 is important in cation permeability due to the
presence of negative amino acid residues. An outward shift of these TMs enlarges the pore and is likely
to influence cation permeability. Lastly, the missing serines studied by Richards and Dempski are a
distinct difference between ChR2 and bR that have been shown to prevent cation permeability if
reintroduced. These differences can begin to explain the differences in permeability between ChR2 and
bR.
However, due to the strong homology between ChR2 and bR, investigations about a possible
proton pump mechanism in ChR2 were undertaken. Feldbauer et al. (2009) were the first to characterize
the function of ChR2 as both a light-driven proton pump and cation channel. They reconstituted ChR2 in
a planar membrane with no ion gradient or electrical potential difference. In response to illumination, no
current flow is expected. However, small outward photocurrents were observed, providing evidence for
the proton-pumping ability of ChR2. Nack et al. (2012) further elucidated the kinetics of this
mechanism. Nack et al. expressed ChR2 in yeast and examined proton currents using spectrophotometry
and an optical pH indicator. They found that proton release and uptake in ChR2 matches the rise and
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decay of the P520 intermediate (Figure 10). As the P520 state
represents the cation permeable state of ChR2, an intimate mechanistic
link between the two functions of ChR2 is suggested. Interestingly,
proton release by ChR2 occurs with the re-protonation of the retinal
Schiff base (rise of the P520 state) and not with the deprotonation as is
seen in BR, suggesting a mechanistic difference between the two
proteins. Therefore, it seems as if ChR2 is a leaky proton pump,
meaning that it allows cation flow, in addition to pumping protons, in
Figure 10. Proton pumping
one of its photointermediates. This suggests a similar function to bR. photocycle of ChR2. From Nack
et al.the
(2012).
However, we must keep in mind the mechanistic differences seen between
proteins, in regards to
both cation conductance and proton pumping.
Application
Optogenetics
The application of ChR2 to modulate neuronal activity, termed optogenetics, began with Boyden
et al. (2005). By injecting adeno-associated virus (AAV), containing ChR2 and yellow fluorescent
protein in rat CA3/CA1 hippocampal neurons, illumination of ChR2-positive neurons with blue light
induced rapid depolarizing currents. These exciting results showed consistency both in the same neuron
and across neurons. Additionally, spike frequency was dependent on light intensity, allowing for
unprecedented control of neural activity (Figure 11). Since its genesis, optogenetics has evolved in
incredible ways. The advances in the functional use of ChR2 far outreach its structural discoveries. We
will look at two recent papers showing how far optogenetics has come, and some of the incredible
applications it posseses.
Figure 11. Data from some of the first optogenetic experiments. From Boyden et al. (2005).
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Stuber et al. (2011) used optogenetics to investigate neural
circuits implicated in reward, specifically the Nucelus Accumbens
(NAc), part of the mesolimbic dopamine pathway, thought to be involved
in the rewarding effects of drugs of abuse. Stuber et al. were able to
stimulate specificly glutamatergic afferent projections to the NAc from
both the basolateral amygdala (BLA) and the medial prefrontal cortex Figure 12. Optogenetic
(mPFC). To do this, an AAV dependent on a Camk2a promoter, was technique depiction. From
injected in either the BLA or the mPFC of adult mice, leading to the
Stuber et al. (2011).
specific trasfection of gluatamatergic neurons in either the BLA or the mPFC. Subsequent applications
of light to the BLA only depolarized projections from either the BLA or the mPFC (Figure 12). The
mice were then underwent one hour self-stimulation sessions in which active “nosepokes” provided 5ms
of photostimulation to projections from the BLA or mPFC. Selective activation of BLA, but not mPFC,
glutamatergic inputs to the NAc promoted nosepoking (Figure 13). This is consistent with the
hypothesized role of BLA inputs in facilitating responding to cues, and of mPFC inputs in suppressing
inappropriate actions. Furthermore, it was also shown that selective activation of projections from the
BLA led to EPSC with greater amplitude. This may suggest that BLA projections release glutamate,
while mPFC projections do not (Figure 13).
Figure 13. Comparison of nose pokes and EPSC from BLA-to-NAc and mPFC-to-NAc. From Stuber et
al. (2011).
Packer et al. (2012) used two photon optogenetics to push the limits of neuronal modulation with
light. By activating membrane opsins with two lower-energy, longer-wavelength photons, incredible
precision, on the level of single dendrites and dendritic spines is possible, as lower-wavelength photons
scatter less. Unfortunately, ChR2 has a small single-channel conductance and fast kinetics, meaning that
when using two-photon activation of a neuron with ChR2, very high ChR2 expression would be required
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to create action potentials. To make two-photon microscopy and optogenetics commensurable, Packer et
al. created a chimeric opsin by combining ChR1 and VchR1 (from multicellular algae Volvox carteri).
This chimaera, C1V1, has a red-shifter photon absorption (over 1000nm) and slow channel kinetics,
allowing for the use of two-photon optogenetics to induce neural firing.
Transfection of this chimaera, in conjunction with yellow fluorescent protein, allowed Packer et
al. to make three incredible “gestals.” First, they were able to achieve selective activation of cellular
processes down to the dendrite and dendritic spine (Figure 14). Second, they demonstrate the potential
of two-photon optogenetics to map synaptic circuits. Stimulating single neurons allows for an
investigation of the functional relationship (e.g. excitatory or inhibitory synapse) between neurons.
Specifically, one can patch clamp a neuron and stimulate various surrounding neurons and look for a
response in the patched neuron (Figure 15). Third, Packer et al. examined the use of two-photon spatial
light modulation (SLM)-based microscopy. This method allows for the simultaneous activation of up to
15 neurons with consisted firing patterns. These two papers exemplify both the current work and the
incredible potential for optogenetics as a tool to modulate neural activity. Future efforts to bridge our
knowledge of the structure of ChR2 and other microbial rhodopsins with the incredible new
developments in optogenetics will provide immense benefits to both fronts and allow for even greater
variations and applications of ChR2 going forward.
Figure 14. Two-photon stimulation of cellular
processes (top), including individual dendrites
and spines (bottom). From Packer et al. (2012).
Figure 15. Mapping synaptic circuits with two-photon
optogenetics. Photostimulation of neuron i shows a
response in patch clamped neuron ii. From Packer et al.
(2012).
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References
Boyden ES, Zhang F, Bamberg E, Nagel G, Deisseroth K (2005) Millisecond-timescale, genetically
targeted optical control of neural activity. Nat Neurosci 8:1263-1268.
Eisenhauer K, Kuhne J, Ritter E, Berndt A, Wolf S, Freier E, Bartl F, Hegemann P, Gerwert K (2012) In
channelrhodopsin-2 Glu-90 is crucial for ion selectivity and is deprotonated during the
photocycle. J Biol Chem 287:6904-6911.
Feldbauer K, Zimmermann D, Pintschovius V, Spitz J, Bamann C, Bamberg E (2009)
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