Optogenetics: (back to) basics

Optogenetics Primer
A brief overview of optogenetics provided by Nature Methods

Optogenetics is a relatively novel set of research techniques that allow fast and directed control of specific cellular functions.[1] Essentially, optogenetics is the use of cells embedded with light sensitive opsins to control their function. The key to understanding what roles cells play and how they affect behaviour is being able to precisely modify and study different cell types without confounding factors, but previous studies undertaken used indefinite methods such as lesions or electrode stimulation that did not allow for confident determination of causality.[2] Extreme, millisecond-level temporal specificity greatly improves our ability to perturb neuronal systems and devices like the optrode greatly improve our ability to observe them. With optogenetics, we can fundamentally understand and fluently speak the language of cells instead of having to go through translators, and it can all be done in freely moving mammals. There are two basic steps involved, first, a specimen has to be infused with the tools of interest, and second, a light delivery mechanism has to be installed to affect those tools.[3] Both steps encompass a wide variety of choices, giving scientists a veritable grab bag of neuromodulation abilities to suit any experimental needs.

1 Optogenetic Tools

The roster
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A simplified version of the basic properties of each optogenetic tool class [3]

Opsins are at the heart of the optogenetic toolkit, and opsin genes are split into two kinds: microbial opsins (type I) and animal opsins (type II). Both types of opsin proteins require retinal, which serves to trigger conformational changes within the opsin when it becomes photoisomerized after absorbing a photon.[2] In humans, retinal bound to type II opsins (known as rhodopsins) form the basis for vision. However, microbes have evolved diverse opsins in place of eyes to sense and respond to electromagnetic radiation. Some of the wide variety of uses that microbes put opsins to include maintaining ionic homeostasis, and modulation of flagella to direct phototaxis.[4] These light sensitive proteins are introduced into animal cells and exploited to exert precise spatial and temporal control, essentially allowing fast, specific, “loss of function” or “gain of function”. They can be directed to individual neuronal types within tissue, and activated and deactivated on physiologically relevant timescales. For example, it is possible to use microbial opsins embedded in neuronal membranes to cause light-induced ion flow, whether it be inward cation currents causing depolarization and creation of action potentials, or inward flow of chloride ions causing hyperpolarization and inhibition of APs.[2] There are four major types of single component optogenetic tools in use today, each containing a plethora of subtypes, allowing varied experiments to be performed.

Structures of opsins
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The chemical structures and interactions of the microbial opsin family of tools [23]

1.1 Channelrhodopsin (where things get exciting)

There are two types of rhodopsins obtained from the single celled green alga known as Chlamydomonas reinhardtii, Channelrhodopsin-1, a light-gated proton channel[5] and Channelrhodopsin-2, a light-gated cation channel.[6] Boyden et al. first used Channelrhodopsin-2 as a single-component cation channel in 2005[7] and subsequently kicked off the optogenetic revolution.[1] Using a flash of blue light, (450-490nm) they found that ChR2 causes an inward flow of ions within 50µs. Coupled with the fact that ChR2 can be safely and reliably be inserted into mammalian neurons, this method allows for precise control of neuronal depolarization.[7] This ability facilitates the generation of spike trains with any desired pattern, including patterns seen in natural brain function.

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A graph showing the recent boom in optogenetics related papers, after the first successful implantation into mammalian neurons in 2005 [1]

1.1.1 Channelrhodopsin2 variants

Since the use of wild-type ChR2 in 2005, there have been numerous variants developed with slightly differing kinetics, enabling further specificity. A popular modification involves the replacement of algal codons in ChR2 with mammalian ones (known as humanized ChR2 or hChR2), improving expression levels.[8] Another modification of ChR2 is the introduction of the H134R mutation, which causes a twofold increase in the ion currents, but consequently results in a twofold decrease in channel-closure.[9] This same type of trade-off is found with the T159C and L132C mutations of ChR2, to differing degrees.[10][11] Another interesting property that has been induced in channelrhodopsin2 is bistable behaviour, wherein activation occurs by blue light as usual, but off kinetics are greatly increased, up to 30 minutes for some variants. Manual termination of the current is possible, with the use of yellow light in the 560-590nm range.[12] These special mutations are designated step-function opsins or SFOs, and there are two significant differences between them and more traditional ChR2 variants. For one, they are much more sensitive to light, due to more channels remaining open for longer and secondly they allow asynchronous neuronal activation, which may be preferable in some cases.[3] Lastly, the off kinetics of ChR2 can be greatly increased (from ~10ms to ~4ms) at a nominal current loss by replacing glutamate 123 with either threonine or alanine, aptly dubbed ChR E123T/A or ChETA.[13]

1.2 Halorhodopsin (hates fun)

While channelrhodopsins provide a fast and precise way to control neuronal excitation, what goes up must come down. To glean the most information, loss of function studies must be performed alongside gain of function ones, which is where halorhodopsin comes in. However, even though photoactivation of halobacterial HR causes chloride (Cl-) influx (and therefore hyperpolarization), it becomes desensitized too easily to be used in rigorous experimentation.[14]

1.2.1 Halorhodopsin variants

Fortunately, a homologous gene found in Natronomonas pharaonis (termed NpHR)[15] was discovered, which better facilitated stable hyperpolarization. A happy coincidence resulted from the yellow light (~590nm) photocurrent peak of NpHR. At that wavelength, ChR2 shows no activation whatsoever, allowing both channels to be independently activated for bidirectional modulation. Moreover, NpHR requires that there be a constant source of light to function, unlike the channelrhopsins, which are excitatory.[3] Further study on the properties of NpHR expressing cells conducted with enhanced yellow florescent protein discovered that the channels were mostly colocalized with the endoplasmic reticulum, so modifications were made to improve the surface membrane localization (namely the addition of an ER export motif), thereby increasing hyperpolarization efficacy.[16] This variant, eNpHR2.0, was superseded by eNpHR3.0 for use in mammals, due to further alterations increasing its loss of function abilities in mammals.[17]

1.3 Bacteriorhodopsin (the old guy in the room, also hates fun)

Bacteriorhodopsin was the first light activated ion pump to be identified, by Oesterhelt and Stoeckenius in 1971[18], and is far and away the best studied. It is used in haloarchaeal membranes as an alternative way of generating energy in low oxygen conditions, as it pumps protons out of the cytoplasm to generate a proton gradient, enabling ATP synthesis.[19] Two type I opsins have been used to great effect as neural activity inhibitors by Chow et al., who found that both Arch (archaerhodopsin-3, obtained from Halorubrum sodomense) and Mac (from Leptosphaeria maculans) can be light modulated (yellow and blue, respectively) to allow almost complete silencing of specific neurons in the awake mammalian brain.[20] However, there is a downside to using proton-motive pumps as neuronal inhibitors, as the long-term effects of significant proton efflux could have far-reaching and unwanted consequences, interfering with optogenetic observations.[3]

1.4 OptoXR (jack of all intracellular signalling)

Type I opsins such as ChR2 and NpHR enable fast and precise control of nuclear membrane potential (and consequently excitation/inhibition) via manipulation of ion flow, but they do not allow timely modulation of intracellular processes. That is where vertebrate rhodopsin takes over, not only is it a type II opsin, but it also serves as a G protein-coupled receptor.[3] This homologous structure is used by way of vertebrate rhodopsin/ligand gated GPCR chimeras to hijack common signaling pathways[21], giving us precise control through light manipulation. These chimeras are known as optoXRs. Neurological modulation tools such as endocannabinoid, adrenergic, serotonergic and dopaminergic receptors are G proteins, which opens up many experimental manipulation possibilities when paired with optoXRs. Due to the spatial and temporal precision afforded to us by optogenetic techniques, using optoXRs can elucidate the causal impact of intracellular signaling in the brain, as illustrated by Airan et al. through manipulation of mice in vivo.[22]

2 General Techniques

Using optogenetic techniques involves special preparation. First, the opsins of choice have to be introduced specifically into the cells of import through the use of viral vectors or transgenic animals, and then an optrode has to be installed to both deliver light and record the resulting data.[3]

2.1 Opsin Targeting

The most popular way to introduce opsins into experimental specimens are viruses, specifically lentiviral vectors or adeno-associated viral vectors.

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A chart describing the promoters and viral vectors used for optogenetic purposes [3]

2.2 Stereotaxic Surgery

This is the method by which the optrode is fixed in the specimen in vivo. The mouse is held in place by the apparatus, while the specific coordinates are dialed in, allowing precise implantation.

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A cartoonized version of stereotaxic surgery showing the apparatus used to implant the optrode

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The poster mouse for optogenetics

2.3 Light Delivery

The light pulses are delivered by either fibre optic cable or a tool known as an optrode. It can take many forms depending on the experimental needs of the researcher, but its key feature is the ability to simultaneously deliver light pulses and record data.

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Different types of optrodes used for various experimental designs [24]
1. Deisseroth, K. (2011). Optogenetics. Nature Methods(8), 26-29. doi:10.1038/nmeth.f.324
2. Fenno, L., Yizhar, O., & Deisseroth, K. (2011). The Development and Application of Optogenetics. Annual Review of Neuroscience(34), 389-412. doi:10.1146/annurev-neuro-061010-113817
3. Yizhar, O., Fenno, L. E., Davidson, T. J., Mogri, M., & Deisseroth, K. (2011). Optogenetics in Neural Systems. Neuron(71), 9-34. doi:10.1016/j.neuron.2011.06.004
4. Mattis, J., Tye, K. M., Ferenczi, E. A., Ramakrishnan, C., O'Shea, D. J., Prakash, R., … Deisseroth, K. (2012). Principles for applying optogenetic tools derived from direct comparative analysis of microbial opsins. Nature Methods(9), 159-172. doi:10.1038/nmeth.1808
5. Nagel, G., Ollig, D., Fuhrmann, M., Kateriya, S., Musti, A., Bamberg, E., & Hegemann, P. (2002). Channelrhodopsin-1: A Light-Gated Proton Channel in Green Algae. Science, 296, 2395-2398. doi:10.1126/science.1072068
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7. Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G., & Deisseroth, K. (2005). Millisecond-timescale, genetically targeted optical control of neural activity. Nature Neuroscience, 8(9), 1263-1268. doi:10.1038/nn1525
8. Zhang, F., Wang, L.-P., Boyden, E. S., & Deisseroth, K. (2006). Channelrhodopsin-2 and optical control of excitable cells. Nature Methods, 3(10), 785-792. doi:10.1038/NMETH936
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10. Berndt, A., Schoenenberger, P., Mattis, J., Tye, K. M., Deisseroth, K., Hegemann, P., & Oertner, T. G. (2011). High-effciency channelrhodopsins for fast neuronal stimulation at low light levels. Proceedings of the National Academy of Sciences USA, 108(18), 7595-7600. doi:10.1073/pnas.1017210108
11. Kleinlogel, S., Feldbauer, K., Dempski, R. E., Fotis, H., Wood, P. G., Bamann, C., & Bamberg, E. (2011). Ultra light-sensitive and fast neuronal activation with the Ca2+-permable channelrhodopsin CatCh. Nature Neuroscience, 14(4), 513-518. doi:10.1038/nn.2776
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15. Sato, M., Kubo, M., Aizawa, T., Kamo, N., Kikukawa, T., Nitta, K., & Demura, M. (2005). Role of Putative Anion-Binding Sites in Cytoplasmic and Extracellular Channels of Natronomonas pharaonis Halorhodopsin. Biochemistry, 44, 4775-4784. doi:10.1021/bi047500f
16. Gradinaru, V., Thompson, K. R., & Deisseroth, K. (2008). eNpHR: a Natronomonas halorhodopsin enhanced for optogenetic applications. Brain Cell Biology, 36, 129-139. doi:10.1007/s11068-008-9027-6
17. Gradinaru, V., Zhang, F., Ramakrishnan, C., Mattis, J., Prakash, R., Diester, I., … Deisseroth, K. (2010). Molecular and Cellular Approaches for Diversifying and Extending Optogenetics. Cell(141), 154-165. doi:10.1016/j.cell.2010.02.037
18. Oesterhelt, D., & Stoeckenius, W. (1971). Rhodopsin-like protein from the purple membrane of Halobacterium halobium. Nature: New Biology, 233(39), 149-152.
19. Racker, E., & Stoeckenius, W. (1974). Reconstitution of Purple Membrane Vesicles Catalyzing Light-driven Proton Uptake and Adenosine Triphosphate Formation. Biological Chemistry, 249(2), 662-663.
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21. Kim, J.-M., Hwa, J., Garriga, P., Reeves, P. J., RajBhandary, U. L., & Khorana, H. G. (2005). Light-Driven Activation of beta2-Adrenergic Receptor Signaling by a Chimeric Rhodopsin Containing the beta2-Adrenergic Receptor Cytoplasmic Loops. Biochemistry(44), 2284-2292. doi:10.1021/bi048328i
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24. Zalocusky, K., & Deisseroth, K. (2013). Optogenetics in the behaving rat: integration of diverse new technologies in a vital animal model. Optogenetics, 1, 1-17. doi:10.2478/optog-2013-0001

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