Optogenetics Overview

Claire Wells 5/9/2012

Optogenetics: Techniques and Practical Applications in the Khan Lab

Our technology has allowed us to probe the depths of the oceans and the interiors of stars, but one object of particular personal importance to us remains opaque: the human brain. The sophistication of this, the most complex object known to man, defies our attempts to dissect out its circuitry. However, one new technique promises to enable study of the function of specific neuronal populations within the intact brain. Optogenetics is a technique involving the control of neuronal activity using light-sensitive effector molecules targeted to genetically defined populations of neurons which can be carried out in a behaving animal or live brain slices. It has been in use for about a decade but only recently have advancements been made which permit control of neuronal firing rates on biologically relevant timescales. The idea of using light to control neuronal activity is relatively old. As early as 1983, Faber and Grinvald used a photosensitive dye, which formed pores in cellular membranes in response to light, to activate ganglionic neurons with a laser, and were able to identify neurons which were presynaptic to a neuron impaled with an electrode. However this technique also activated axons of passage and so would be of limited utility in the dense neuropil of brain slices, and was quite damaging to cells (Faber and Grinvald 1983). More recently, neurotransmitter caging has been another popular optical technique in which a modified neurotransmitter is bath-applied to a living brain slice in an inactivated form, or microinjected into the brain of a living animal. The alterations to the neurotransmitter may be as simple as the incorporation of an inactivating group joined to the transmitter by a single lightsensitive covalent bond (Adams and Tsien 1993). This is complicated by the observation that a single biologically active molecule may have multiple active sites that are relevant in different

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situations or interactions with different ligands and receptors, that the caged molecule may itself have some biological activity (for instance it may act as an inhibitor of the receptor rather than an agonist), and that it is necessary for the caged molecule to have an extremely high stability to prevent biologically relevant levels of background release (Kao 2006). Upon photostimulation the active neurotransmitter is released from this cage. Patch clamp recording of a single neuron combined with activation of caged glutamate using a small beam of light allows the connections of the recorded neuron to be mapped, albeit imprecisely. This method does not stimulate axons of passage (Callaway and Katz 1993). Callaway and Katz had used light focused by their microscope; however, a laser can create a smaller beam of light that is restricted in three dimensions rather than two. Also, the use of two caging groups instead of one, necessitating the absorption of two photons for neurotransmitter activation, further increases the resolution in three dimensions of this technique, such that it may be used to map receptor densities on a single neuron (Pettit et al. 1997). Despite the utility of such methods, they do not overcome one of the major challenges facing researchers attempting to identify functional circuits within the brain the fact that a population of neurons that have the same function may be spatially dispersed; and conversely, that a group of neurons that are anatomically proximate may have quite different functions and participate in different circuits. Thus previous methods are too imprecise to adequately define the fine circuitry of the central nervous system either the spatial resolution is too low or only a single neuron is selected for electrical stimulation, which is insufficient to elucidate the nature of a circuit. The combination of optical and genetic techniques allows stimulation to be targeted to a precisely defined population of neurons, despite the disparities of their locations in the anatomy, by the use of cell-type-specific promoters. It is even possible to target particular regions of a

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neuron, such as dendrites only, or only the postsynaptic density, by incorporating a localization sequence the cell uses to tag specific proteins to their proper locations in the cellular architecture (Gradinaru et al. 2007). Optogenetics was created by Gero Miesenbock, who suggested in 2001 the possibility of combining light-sensitive receptors with genetically based targeting (Zemelman and Miesenbock 2001), and the following year published the first such study. His lab transfected Xenopus oocytes with mRNA coding for the Drosophila proteins NinaE (an opsin incorporated into R1-R6 photoreceptors sensitive to blue wavelengths), arrestin-2 (responsible for inactivating the rhodopsin G-protein-coupled receptor and allowing regeneration of 11-cis retinal), and the subunit of the heterotrimeric Gq/11 protein coupled to the photoreceptor. The combination of these three proteins was found to be sufficient to render the oocytes sensitive to light. Because invertebrate opsins form a covalent bond with retinal to generate rhodopsin, which after conversion to all-trans retinal remains associated with the opsin and regenerates while participating in this complex, an initial introduction of retinal was sufficient to provide a continuous supply. Since the group incubated their oocytes with all-trans retinal to initiate photosensitivity although it is the 11-cis form that is active, and since vertebrates express background levels of this molecule, no cofactor should need to be provided in vivo. An endogenous G dimer must have been recruited to the transfected subunit to confer functionality. In transfected cultured rat hippocampal neurons, the group was able to achieve firing rates of around 7.5 Hz (in some neurons) with latencies of a few hundred milliseconds to tens of seconds in neurons exposed to about 1.8mW of white light lasting 90 seconds. When a period of illumination was followed by a period of darkness, some spillover activity occurred. Several types of firing patterns were observed (Zemelman et al. 2002). The major drawback of

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this initial technique is that it uses a multimeric G-protein-coupled receptor, which is slow to activate and deactivate, limiting temporal resolution. Another difficulty is the variability of response type, which is likely modulated by the different second messenger cascades and different downstream effector ion channels present in different neurons. However, despite the slow kinetics, it is useful in some situations to use G-proteincoupled receptors for optogenetics. Hybrid receptors combining the extracellular domains of rhodopsin with the intracellular domains of G-protein-coupled 1- adrenergic (opto-1AR) and 2-adrenergic (opto-2AR) receptors have been created and are called optoXRs. These receptors allow genetically targeted light-activated control of intracellular signaling cascades. Opto-1AR activates phospholipase C (and therefore creates diacylglycerol and inositol triphosphate), while opto-2AR activates adenylyl cyclase (and leads to rising levels of cAMP). The first increases firing rates in mouse nucleus accumbens, while the second decreases activity. Both were shown to increase phosphorylated CREB (Airan et al. 2009). Another form of optogenetic control uses the introduction of genetically targeted ligandgated ion channels into certain neuronal populations. Through the use of a caged ligand, it is possible to activate these channels only in the presence of light. For example, this technique has been used to study the dopaminergic circuits controlling movement and learning as regards odor preference in Drosophila (Lima and Miesenbock 2005 and Claridge-Chang et al. 2009, respectively). A P2X2 ATP-gated Ca2+ channel with caged ATP was used for these experiments because Drosophila lacks endogenous purinocepters. The drawback of this scheme is that inactivation of the receptor is dependent on the kinetics of diffusion, uptake, and/or degradation of the uncaged ligand.

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Yet another strategy of particular elegance involves the creation of a synthetic protein, such as a receptor, with its ligand (or an antagonist) linked to it near the binding site by a photoisomerizable tether. Thus the ligand will be allowed to interact with the binding site or withheld from such interaction by the isomerization state of the tether, which is controlled by light. Banghart et al. used this method to design a Shaker K+ channel with a cysteine substituted for Glu422 and a tethered MAL-AZO-QA molecule (maleimide-azo-quaternary ammonium where MAL binds cysteine, the AZO group switches from trans to cis and shortens in response to light, and QA blocks the channel pore), and further modified the Shaker channel to display altered voltage-dependence properties so as to avoid interference of voltage in experimental designs. They expressed the modified Shaker, which they have termed SPARK (synthetic photoisomerizable azobenzene-regulated K+ channel), in cultured rat hippocampal neurons incubated with MAL-AZO-QA, and found that action potentials were suppressed in these neurons within about 3 seconds of exposure to 390-nm light and restored within about the same time course of exposure to 500-nm light (Banghart et al. 2004). This technique produces a more reliable time course than Miesenbocks chARGed method, providing temporal resolution medial between the best and worst observed by Miesenbock. It is not ideal to employ a channel that is voltage-gated, yet another system could be designed using different components. The Banghart group used Shaker because it has been extensively described. Another difficulty is the possibility of side-effects arising from interaction of the ligand and tether with endogenous molecules. The most recent advancements in the field came with the incorporation of singlecomponent light-activated ion channels. The first of these microbial channels was discovered in 1971 by Stoeckenius and Oesterhelt and called bacteriorhodopsin. Another such channel, halorhodopsin, was described in 1977, and a third, channelrhodopsin, in 2002 (Deisseroth 2011).

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Prokaryotes and eukaryotes, including fungi and algae, use them to maintain proton and ion gradients, in energy storage, and to modulate the beating of their flagella, among other applications. The channels require only all-trans retinal as a cofactor, which exists endogenously in the mammalian central nervous system in concentrations sufficient for their function. These microbial family opsins are structurally distinct from mammalian family counterparts, in that they are channel receptors rather than G-protein-coupled. They also retain a covalent bond to their retinal cofactor and spontaneously regenerate all-trans retinal (Yizhar et al. 2011), similar to the invertebrate opsins used by Miesenbock (Zemelman and Miesenbock 2002), although the invertebrate opsin relies on 11-cis retinal while the microbial version uses the all-trans form. Not until 2005 were the microbial opsins used in optogenetics, which is perhaps surprising given their fast kinetics, intrinsic light-sensitivity, and the fact that as they do not initiate G-protein second messenger cascades they might be expected to elicit more stereotyped and reliable behavior across different cell types. Boyden et al. demonstrated that channelrhodopsin-2 (ChR2), a cation channel that allows passage of Na+, K+, and Ca2+, may be transfected into cultured rat hippocampal neurons. Though, like Meisenbock, they found that sustained light produced irregular trains of action potentials, they were able to elicit a single spike reliably with latencies of around 8-10 milliseconds with a brief light pulse of around 812mW and 10-15 milliseconds. When exposed to patterns of light pulses, both individual neurons and heterogeneous populations of neurons replicated the pattern with a very high degree of reliability, which increased both with increasing pulse duration and increasing duration of intervals between pulses. When neurons were subjected to pulse series with frequencies of 20 Hz and above, on average less than half the pulses delivered elicited a spike and a slightly higher jitter in spike timing was observed, although the (very low) number of extraneous spikes

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unrelated to a pulse and the time course of individual spikes remained constant. Importantly, it was also possible to evoke reliable trains of subthreshold depolarizations with reproducible amplitudes, though this is theoretically more problematic due to the lack of the nonlinear, stereotyped conductance behavior observed with action potentials. This study marked the first emergence of a technology which could allow the artificial generation of patterned impulses on biologically relevant timescales to genetically defined neuronal populations. However the difficulty of controlling expression levels remains problematic, and could account for the observation that some neurons fired action potentials when others were merely depolarized and for the variability in optimal light pulse duration that evoked a response from different neurons (Boyden et al. 2005). Another limitation is the fact that channelrhodopsin is preferentially sensitive to blue light, which has poor tissue penetration (Pulver et al. 2009). A number of alterations have since been made to channelrhodopsin, improving its firing rate up to frequencies of 200 Hz and shifting its preferred excitation wavelength to wavelengths with better tissue penetration. Some mutants even introduce a switch effect in which one wavelength opens the channel and another closes it (Mancuso et al. 2010). Another of the bacterial opsins, halorhodopsin (Halo), allows the influx of chlorine ions and hyperpolarizes rather than depolarizes cells. It is preferentially activated by yellow light and thus can be used in conjunction with channelrhodopsin to instigate or suppress firing in neurons. This improves the complexity of effects that may be achieved by optogenetic methods (Fiala, Suska, and Schluter 2010). Useful modifications to this channel include the addition of an endoplasmic reticulum export sequence (as the wild-type tends to express in the endoplasmic reticulum) and alterations which render it sensitive to red light (~680 nm), giving it improved tissue penetration (Mancuso et al. 2010).

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There are a number of strategies that can be employed to introduce the desired lightsensitive elements into a cell. The chief among them is transduction using viral vectors, such as lentiviruses. Lentiviruses are a subfamily of retrovirses that includes human immunodeficiency virus, among others. Retroviruses are capable of permanently inserting their genetic information into the host genome, and lentiviruses are unique among them in that they can infect both dividing and non-dividing cells. They are also highly efficient. Lentiviruses have been engineered for safety and stability, since they are RNA viruses, and are a popular viral vector for transduction (Dull et al. 1998). Adeno-associated virus, a parvovirus, is also capable of longterm infection, either by insertion into the host genome or by persistence as an episome, and can also infect dividing and non-dividing cells. This virus requires co-infection with another virus, such as adenovirus, to replicate, and so is safe for experimental use. These features make adenoassociated virus another common choice of viral vector (dos Santos Coura and Nardi 2007). However even with these high efficiency vectors, it can be difficult to achieve sufficient expression levels with some promoters, especially in distant axon terminals. One solution designed by Atasoy et al. uses Cre recombinase-dependent expression with a FLEX switch. Cre recombinase is capable of catalyzing inversion or excision of DNA between precise lox sequences. The FLEX switch utilizes two lox sequences which are each placed on either side of the gene of interest. It is necessary that they should show homotypic but not heterotypic recombination. The group used loxP and lox2272 sequences but other combinations are possible. The gene of interest should be placed in the antisense direction so that it is not expressed. Cre recombinase will then catalyze the inversion of the gene followed by excision of the two redundant lox sites, such that one (heterotypic) lox sequence will be left on each side of the gene of interest, which is now in the sense orientation and may be expressed. Because Cre

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recombinase cannot act on the lox sequences that have been chosen if they are heterotypic, no further rearrangement will be possible. The group used the common CAG promoter to drive expression of the Cre-activated ChR2 gene. Genetic selectivity is achieved by targeting Cre to a specific neuronal population with a particular promoter. This can be done by transduction with a viral vector, but Atasoy et al. used transgenic mice. This technique has proven popular among optogenetic researchers and a number of transgenic mice lines are available expressing Cre under the control of various promoters. It results in higher expression levels and more uniform expression throughout the cell, including distant processes (Atasoy et al. 2008). Other Cre strategies exist as well. One involves the placement of a stop codon flanked by lox sites between the promoter and the gene of interest. Cre will excise the stop codon and allow expression of the gene (Zeng and Madisen 2012). The issue of variable expression levels across a population of cells would be solved by the use of transgenic animal lines rather than transduction with viral vectors. Many lines of transgenic mice, and a few rat lines, expressing ChR2 in various neuronal subsets have been created. Zhao et al. demonstrated the use of four such lines of transgenic mice stably expressing a modified variant of ChR2 known as ChR2(H134R) (2011). This variant has a point mutation at amino acid 134, which was changed from a histidine to a more basic arginine, and was originally developed by Nagel et al. in 2005. It has been found to confer increased firing rates, increased sensitivity to low levels of light (as little as .140 mW for pulse durations of 10 ms), sensitivity to shorter pulse durations (less than 5 ms), and improved performance under constant illumination. This position is implicated in proton exchange with the retinal Schiff base and is therefore important for the opening and closing of the channel. It has been proposed that this mutation improves function by altering the intermediate conformations of the channel to favor the open

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state (Pulver et al. 2009). Zhao et al. found that their transgenic mouse lines were able to sustain reliable firing rates at pulse frequencies of up to 80 Hz (in the VGAT line; up to 50 Hz in the ChAT and 20 Hz in the TPH2 and Pvalb lines), and under constant illumination conditions. Some extra spikes were observed, mainly during low frequency stimulation, but few missed spikes. The group developed a number of lines out of which only a few demonstrated stable expression, and even in these there were some abnormalities or irregularities in the expression levels for certain cell types, which might be related to differential activity of the promoters used in different cells that express the targeted enzymes (Zhao et al. 2009). Nonetheless, as the number of available stable transgenic lines increases, including many selective Cre-expressing lines, research in this area will be facilitated. Another method of delivery of optogenetic proteins is by in utero electroporation. Electroporation refers to the phenomenon in which the application of an electric field causes a lipid bilayer membrane to form small pores that allow the passage of large molecules, including DNA and RNA. If the field is not too strong, the membrane can then recover its integrity. This technique may be performed in utero on embryos and can be targeted to specific brain areas, such as particular layers of cortex or deeper structures like the basal ganglia or hippocampus. Some of the advantages of this method are that it is simpler than the creation of transgenic animal lines, allows the passage of any size of DNA or RNA molecule, and can be combined with promoters to further improve targeting (Gradinaru et al. 2007). The main remaining difficulties with optogenetic techniques lie in delivering light stimulation and recording postsynaptic activity. Light delivery has been accomplished using a flexible fiber optic cable implanted just above the brain region of interest, which can be combined with an electrode for measurement of activity (Airan et al. 2009). It is also possible to

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implant a glass window in the skull of an experimental animal, or to render the skull transparent with a dental resin, though there are issues with light penetration into deep tissues with these methods, and a depth of a few hundred micrometers is about the limit (Mancuso et al. 2010). Frequently recording is achieved by means of a patch pipette or electrode, but the full potential of optogenetics will not be realized by use of such limited means of measuring activity. Several optical tools, that may be genetically targeted, are currently being developed (Mancuso et al. 2010). Nikai, Ohkura, and Imoto created a Ca2+ probe called GCaMP, in which a circular enhanced green fluorescent protein is attached to the M13 subunit of myosin light chain kinase (which binds calmodulin) on one end and to calmodulin on the other. When Ca2+ binds this construct, it undergoes a conformational change that alters its fluorescence. This allows the realtime measurement of intracellular Ca2+ increases over timescales of a few hundred milliseconds (Nikai Ohkura and Imoto 2001); however, there are many reasons why a cell might have increased Ca2+ levels, including release from intracellular compartments, so this is not an absolutely reliable indicator of neuronal activity. Also, Ca2+ perturbances due to subthreshold electrical events are small and may pass unnoticed by the probe (Mancuso et al. 2010). Cl- influxes, correlated with inhibitory activity, can be sensed by Clomeleon, an indicator that consists of a yellow fluorescent protein linked to a cyan fluorescent protein. Cl- quenches the fluorescence of the yellow but not the cyan fluorescent proteins, such that in the presence of Clthe color of the indicator changes from yellow to cyan. Variants of this ensemble improve the signal-to-noise ratio considerably and offer heightened binding affinity and dynamic range. However these proteins are pH-sensitive, and pH is known to fluctuate when a cell is firing trains of action potentials (Mancuso et al. 2010).

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Finally, some genetic probes have been designed that respond to changes in membrane potential. One prominent example, Mermaid, includes the relevant domain from a voltagesensitive phosphatase and two relatively pH-insensitive fluorescent proteins found in corals, mUKG and mKO. This indicator can reliably report action potentials at frequencies up to 50 Hz in mouse cells, but the signal-to-noise ratio does not allow detection of subthreshold events at this time. Although voltage sensors are the most direct route of measuring postsynaptic activity, they have one major drawback, which is that they unavoidably alter the membrane capacitance. This means that they cannot be expressed at high levels without altering the function of the neuron (Mancuso et al. 2010). In the ideal optogenetic setup, the consequences of light-evoked activity in a specific neuronal population would be imaged in real time across a large population of cells using genetically targeted light-emitting reporters such as those described above. The last component of such a strategy is the imaging technology employed. Traditionally, single-photon confocal microscopy has been used to image fluorescence. The main components of such a microscope are a laser; a dichromatic mirror, which allows light longer than a certain wavelength to pass through it and reflects light shorter than this wavelength; two scanning mirrors which target the laser beam along the x and y axes; an objective; a screen with a confocal pinhole, which excludes light that originates from a part of the sample that is not in focus; and a detector. The scanning mirrors focus the light on a particular part of the sample and the speed at which they can change position limits the field of view or sampling rate for the image. They can be replaced with Acoustic Optical Deflectors (AOD), in which a high frequency sound wave is passed through a special crystal. The frequency of sound waves passed through the crystal alter its refractive index, and so can be used to control the angle at which it deflects a light beam. Thus the light can

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be manipulated faster (producing a full image 30 times per second in one setup) than with a scanning mirror that must be physically moved to change the angle of deflection. However, the crystal will deflect light differentially depending on its wavelength, and so reduces the possible resolution. The best resolution of this type of microscope is about 0.2 micrometers in the x and y axes, and about 0.5 in the z axis (Prasad, Semwogerere, and Weeks 2007). The major disadvantage of this technology is that the pinhole excludes signal photons that originate from the point of interest but have become scattered in their journey through the tissue, and this becomes especially pronounced for imaging of relatively deeper tissues, limiting the usefulness of confocal microscopy to a few tens of micrometers. In fact, half of the signal photons may be scattered for every 50-200 micrometers of depth. Attempting to compensate by intensifying the laser beam leads to increased phototoxicity and bleaching. Also, a relatively large portion of the sample is generally subject to excitation, while only a small portion of it is actually being viewed, which also increases photodamage and bleaching (Svoboda and Yasuda 2006). Two-photon excitation microscopy is a technology which improves upon these limitations. It uses two low energy photons arriving almost simultaneously at the fluorophore to excite it sufficiently to release a higher energy fluorescence photon. It is a nonlinear imaging technique, in that excitation relies on the second power of the light intensity. Because the light intensity of a laser is highest at the focal point, and decreases quadratically as distance from this point increases, excitation is limited to a small volume around the focal point. Thus the volume of tissue that can be excited at any given time can be as small as 0.1 cubic micrometers. Also, because the excitation wavelengths used are in the far red range, they have improved tissue penetration and reduced phototoxicity. The design of the microscope is quite similar to that of a confocal microscope, except that a pinhole is unnecessary because only the point of interest is

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being excited at any given time, and therefore all fluorescence photons which reach the objective are signal photons. Instead, light passing through the back-aperture of the objective (fluorescence photons which are likely quite scattered) must be refocused by an additional element. It is also necessary to use a laser which can emit pulses of light at frequencies around 100 MHz to improve the probability that two photons will arrive at the fluorophore nearly simultaneously. These pulses can be specifically shaped in ways that improve signal-to-noise ratio and decrease bleaching. Disadvantages of two-photon imaging include the fact that while overall phototoxicity and bleaching is lessened by two-photon techniques, these effects are greater within the focal volume than for confocal microscopy. Additionally, the effective tissue depth accessible by twophoton methods is around 800 micrometers or less (Svoboda and Yasuda 2006). Ranganathan and Koester have used a technique called dithered random-access scanning, which utilizes two-photon microscopy. In mice injected with a Ca2+-sensitive fluorescent dye, they have used this method to reliably record action potential activity across a large population of neurons. In this technique, each soma is sampled in three or five different locations, rather than the traditional one location with single-point random-access imaging, because Ca2+ levels are not uniform across a cell body. Based on previously made characterizations of Ca2+ influxes associated with an action potential, the group developed an algorithm that extracts spike information from the raw data generated by scanning. The algorithm is capable of employing different parameters for each individual neuron, which is important since different types of neurons have different Ca2+ responses to action potentials. This method detected action potentials with an error rate of 3 missed spikes in 100, and .0023 false positives per second in vitro, with a temporal resolution of 1 millisecond. Its major drawbacks are that it cannot penetrate deep tissues and is therefore limited to surface structures or living brain slices, and that

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high intensity stimulation induces photodamage while low intensities suffer from a low signal-tonoise ratio. Therefore there is only a small window that will allow efficient detection of signals (Ranganathan and Koester 2010). However it is techniques such as this, that combine fast high resolution scanning with computer programs designed to extract signal information, that will allow the determination of behavior in neuronal populations that have been optogenetically manipulated. Undoubtedly these techniques would be of immense use in the Khan lab at UTEP, in which I work. The goal of our research is to elucidate the precise circuits that regulate feeding and glucose homeostasis, so that better drug targets or other treatment stratagems may be proposed for diabetes and morbid obesity. Consequently we are focusing on the hypothalamus, the homeostatic control center of the brain, and primarily on the lateral hypothalamic area (LHA). Although there are a few studies that have been done using optogenetics to explore the circuitry of the hypothalamus, they have mainly investigated sleep/wakefulness. I have only been able to identify one study which employed optogenetics to probe the circuitry of feeding, and so the area is wide open for investigation. The main brain center of importance for feeding is the arcuate nucleus (ARC) of the hypothalamus. The two main neuronal populations present in this nucleus are Agouti generelated protein (AGRP)- and pro-opiomelanocortin (POMC)-expressing cells. AGRP neurons coexpress neuropeptide Y (NPY) and -aminobutyric acid (GABA), while POMC is the precursor of alpha melanocyte-stimulating hormone (MSH) and subsets of these cells coexpress cocaine and amphetamine regulated transcript (CART), glutamate, or GABA. Both populations are sensitive to leptin (a satiety or anorexigenic hormone), insulin, and ghrelin (an appetitestimulating or orexigenic hormone). AGRP neurons induce feeding, while POMC neurons

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suppress it. These cell groups send projections to many areas, including the LHA, parabrachial nucleus (PBN), and paraventricular nucleus (PVH), and tend to co-project (Wu and Palmiter 2010). The mechanisms by which AGRP neurons stimulate feeding are diverse. AGRP neurons have inhibitory GABAergic synapses with POMC neurons, and AGRP itself also acts as an antagonist of the melanocortin receptors 3 and 4 (MC3R and MC4R). Rapid ablation of AGRP neurons results in starvation that can be rescued by injection of benzodiazepines, particularly to the PBN. There is also evidence to suggest that such starvation is via a mechanism independent of MCRs, since MCR blockade does not reverse the effect. In fact injection of GABAA agonists into the PBN stimulates feeding. Together these results indicate that AGRP neuron-mediated inhibition of this area is crucial for normal feeding behavior, and is GABAergic in nature rather than being controlled by MCRs (Wu and Palmiter 2010). The LHA is another important feeding center. Two main neuronal populations contribute to this effect: hypocretin/orexin (HCRT)- and melanin-concentrating hormone (MCH)expressing neurons. Injections of both HCRT and MCH have been shown to increase eating, and both populations receive inputs from AGRP and POMC neurons. HCRT is found only in the LHA, while MCH is also found in the zona incerta (Elmquist, Elias, and Saper 1999). The LHA also contains other peptide transmitters, including galanin, neurotensin, CART, corticotropinreleasing hormone (CRH), and dynorphin; and some LHA neurons coexpress glutamate or GABA with these peptides. Many LHA neurons are sensitive to leptin, glucose, insulin, and ghrelin. The LHA has been implicated in arousal state and in reward mechanisms as well, and may integrate these functions with feeding behaviors. Interestingly, although both HCRT and

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MCH are orexigenic, the first promotes wakefulness while the second promotes sleep. Clearly there is great complexity among LHA subpopulations (Berthoud and Munzberg 2011). The LHA projects to the PVH, nucleus accumbens (NAc) shell, ARC, ventromedial and dorsomedial nuclei of the hypothalamus (also involved in feeding), ventral tegmental area (VTA), striatum, and lower brain structures like the nucleus of the solitary tract and the dorsal motor nucleus of vagus; as well as to higher brain areas like the hippocampus, amygdala, and cortex. The NAc shell sends GABAergic input back to the LHA (Berthoud and Munzberg, 2011). It seems that the LHA is involved in basal ganglia inhibition/permission of motor tasks related to eating. MCH has been found to inactivate the NAc shell to promote feeding possibly involving both direct and indirect D1 and D2 pathways (Gutierrez et al. 2011). Aponte, Atasoy, and Sternson have conducted the only optogenetic study of feeding behavior to date. They have transfected ChR2 fused to the tdtomato fluorophore into AGRP or POMC neurons using the FLEX Cre recombinase method described previously with agrp-cre or pomc-cre transgenic mice. Light was delivered through a flexible fiber optic cable implanted just above the ARC. For agrp-cre mice, light was delivered in 20Hz pulses for 1 second with 3 second intervals for 1 hour, which was similar to the firing patterns previously observed for these neurons in living brain slices upon application of orexigenic hormones. Despite the mice being already sated, they found that such excitation elicited feeding behavior with latencies between 6 and 30 minutes, depending on the number of neurons expressing ChR2. Recruitment of about 300 neurons was necessary for an effect, and at about 800 the maximum effect was observed. The total amount of food consumed was also positively correlated with the number of expressing neurons, and with the frequency of pulses applied. Additionally, continuous stimulation was necessary to maintain feeding; mice that ceased to be excited five minutes after the onset of

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feeding stopped eating soon after, while mice that were excited for the entire 1 hour period ate more for longer and in multiple bouts. No differences were observed between normal mice and mice in which MCRs were constitutively blocked, indicating that the AGRP-induced feeding is by a melanocortin-independent pathway (Aponte, Atasoy, and Sternson 2010). The story for pomc-cre mice is somewhat different. These mice did not reduce their food intake after stimulation for an hour, even though the mice were well-fed. After 24 hours of continuous excitation, a reduction in food intake by approximately 40% was observed. They determined that these effects were dependent on melanocortin pathways (Aponte, Atasoy, and Sternson 2010). Though feeding has been observed following AGRP injection before, this study marks the first time that such behavior can be conclusively tied to the increased firing of AGRP neurons. Since AGRP neurons express GABA and NPY as well, such results are significant. Injection of melanocortin receptor agonists has also been found to suppress feeding in the past; this study proves that such effects are not naturally relevant, possibly because there is a limit to the amount of ligand that POMC neurons will naturally release onto MCRs, which may easily be exceeded by injection. Additionally, the study shows that the amount of feeding is positively related to the amount of activity in AGRP neurons (proving that they do not simply function as a kind of switch) and that continuous activity is necessary for sustained feeding. This is also the first study to demonstrate a complex behavior initiated by a simple firing pattern in a small set of neurons (Aponte, Atasoy, and Sternson 2010). Because the Khan lab uses rats, implementation of optogenetic techniques will be a bit more difficult than in labs using mice. Mice are more genetically tractable than rats, and so it is easier to create transgenic mice lines than rat lines, and also a much greater number of transgenic

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mice lines selectively expressing optogenetic channels or Cre are already commercially available. Also, because the mouse genome has been sequenced, more is known about the selective expression of genes not related to transmitters and associated enzymes and receptors, which can allow for enhanced specificity of targeting. For example, Shimogori et al. have recently undertaken a detailed examination of the expression patterns of about one thousand genes that are differentially expressed during hypothalamic development in the mouse, an invaluable resource for researchers using genetics to probe hypothalamic function (2010). However, the Khan lab has already conducted, and is conducting, extensive research into the rat hypothalamus and associated areas such as the VTA and hindbrain centers. Over the summer some colleagues and I will be performing a study involving immunocytochemical mapping of a number of transmitters and associated enzymes in the LHA onto a parcellation scheme proposed by Swanson (2005). This work will not be of use in directing optogenetic studies in our lab unless we employ rats. In this case, it will likely be easiest to transduce the animals using viral vectors. Because the structures which interest us are deep in the brain, we will have to use a flexible optic cable to deliver light. If the natural firing patterns of our chosen neurons have not been reported in the literature, it will be of use to first determine these using electrophysiology in live brain slices. We can also perform controls ex vivo to confirm the incorporated channel has not altered the electrical properties or apparent health of the neurons. If we use a channel tagged with a fluorescent reporter such as enhanced green fluorescent protein (EGFP), we can run immunocytochemistry double labeling with antibodies against the targeted proteins to confirm specificity and strength of expression. A great many studies using optogenetics to investigate feeding circuits could be done. Some of the more obvious experiments would involve transducing HCRT neurons in the LHA

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with ChR2 (or Halo) and observing feeding behavior initiated by their activation (or inhibition). Gutierrez et al. (2011) have claimed that the optogenetic study performed by Adamantidis et al. in 2007, in which mouse HCRT neurons were transfected with ChR2 and the effect of their firing on arousal state examined, found that HCRT firing had no effect on food consumption. However, I was unable to find any statement in this paper which suggested they had monitored food intake. Therefore this experiment still remains to be done. Naturally it could include transduction of another group of animals, this time targeting to MCH neurons in the LHA. Microinjection with viral vectors could ensure that expression was limited to LHA MCH neurons and did not extend to neurons in the zona incerta. Of course further experiments of this nature could be done using promoters associated with any of the transmitters observed in the LHA, including CART, GABA, and glutamate, or with promoters associated with receptors for transmitters from other locations such as AGRP, MSH, and norepinephrine; although care should be taken with some of these that are common in the hypothalamus that the injection site does not extend significantly beyond the bounds of the LHA (immunocytochemical controls could confirm this). Although it seems obvious to use ChR2 or Halo in these experiments, as they are fast single-component ion channels, in some situations it might be preferable to use another method, such as Meisenbocks chARGing technique. Because each of the three components of their system NinaE, arrestin-2, and the G protein subunit was found to be necessary to confer light sensitivity to transfected cells (Zemelman et al. 2002), the LHA could be injected with viral vectors containing each of the related genes tied to different promoters. A functional lightsensitive receptor would then be present only in those neurons that express genes tied to each of the three chosen promoters. This would allow greater specificity of expression in a region of high complexity, as we would then target neurons defined by three genetic parameters rather than just

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one. This would follow from our immunocytochemical experiments identifying neurons that coexpress various transmitters and associated enzymes, or their receptors. Although we would not be able to use this system to drive exact patterns of action potentials, it is likely that whatever patterns of activity we did observe would be natural patterns, since they would utilize the cells own natural ion channels and machinery. We could assess such firing in live brain slices or with an electrode incorporated with the fiber optic cable (an optrode). This could be an excellent tool for dissecting the function of precise subpopulations of neurons receiving input or sending output to various specific locations. For instance, we could determine which subpopulations are more related to feeding and which to arousal state. An additional element in the above experiments could be the inclusion of an electrode inserted into a brain area we believe our targeted neurons innervate. Many could be chosen, including the ARC, PBN, PVH, VTA, and NAc to name a few. After the manner of Tsunematsu et al., we could identify presumed neurons of interest in the impaled area by distinctive characteristics of their action potential waveforms and overall activity patterns (2011). In this way, we could prove that the activity of light-sensitive neurons is indeed associated with altered spike patterns in the downstream population, and we could characterize these perturbances. This would be of great value in determining the functional consequences of activity in these circuits. With living brain slices, it could be possible to probe the interconnections within the LHA using two-photon microscopy. The Khan lab uses a confocal microscope, but it is possible though probably expensive to convert a confocal for two-photon microscopy. All that is required is to remove the confocal screen, to ensure that the laser is capable of emitting light at a sufficiently high frequency (and to replace it if not), and to add an additional focusing element positioned after the back-aperture of the objective, to focus scattered fluorescence photons

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(Svoboda and Yasuda 2006). Using such a setup, we could transduce some cells with either a ChR2 or Halo channel, and others with the Mermaid voltage sensor or Clomeleon Cl- sensor (though Clomeleon is pH-sensitive). With the right software (likely requiring collaboration with someone in computer science and engineering who would write that software for us), we could image the disturbances in firing patterns elicited by activity (or lack thereof) in our lightsensitive neurons in real time. A variety of altered ChR2 and Halo channels are available with different light sensitivities, so we could ensure their excitation wavelengths did not overlap with the emitting wavelength of our reporter protein. Similar experiments could be performed with target structures of the LHA if we could contrive the plane of section such that the projecting axons werent severed. The benefits of two-photon microscopy in this setup are that it would allow good depth penetration in a slice several hundred micrometers thick, and that it would allow excitation to be focused on a specific neuron or on a number in turn, with little to no bleed over onto nearby neurons in three dimensions. Another ex vivo experiment would involve transducing neurons that express peptide transmitters as well as GABA or glutamate with ChR2, again guided by immunocytochemical studies. By raising or lowering the frequency of spikes in these neurons and monitoring their release of transmitter, we could confirm whether the specific transmitter released is determined by the level of activity in the neuron, as well as characterize such release patterns. Various means of measuring transmitter release exist, including enzymatic assays, patch-clamp electrophysiology (Hires et al. 2007), and chromatographic analysis (Gaskins et al. 1995). Recently, several genetically targeted optical reporters have been developed for imaging glutamate release in real time (Hires et al. 2007). Another scheme using quantum dots targeted to a particular neurotransmitter with antibodies has been developed for fluorescent imaging of

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transmitter release. It is not specified in this paper whether the technique was tested in vitro or in vivo but it is implied that it could be used in vivo (Ye, Park, and Kim 2009), and so presumably would serve in live brain slices. The advent of optogenetics promises to spark a fresh frenzy of investigation into functional brain circuitry, and will herald important advancements in the depth and subtlety of our understanding. Though there have thus far been few such experiments focusing on the hypothalamus, and it seems the full potential of optogenetic techniques is yet to be realized, the area is ripe for study.

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Optogenetics Overview

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