Mechanisms of Deep Brain Stimulation Revealed by Optogenetic Deconstruction of Diseased Brain Circuitry

MECHANISMS OF DEEP BRAIN STIMULATION REVEALED BY OPTOGENETIC DECONSTRUCTION OF DISEASED BRAIN CIRCUITRY

A DISSERTATION SUBMITTED TO THE DEPARTMENT OF NEUROSCIENCES AND THE COMMITTEE ON GRADUATE STUDIES OF STANFORD UNIVERSITY IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

Viviana Gradinaru June 2010

This work is licensed under a Creative Commons Attribution- Noncommercial 3.0 United States License. http://creativecommons.org/licenses/by-nc/3.0/us/

This dissertation is online at: http://purl.stanford.edu/rj878dv3879

ii I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Karl Deisseroth, Primary Adviser

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Helen Bronte-Stewart

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Paul Buckmaster

I certify that I have read this dissertation and that, in my opinion, it is fully adequate in scope and quality as a dissertation for the degree of Doctor of Philosophy.

Carla Shatz

Approved for the Stanford University Committee on Graduate Studies. Patricia J. Gumport, Vice Provost Graduate Education

This signature page was generated electronically upon submission of this dissertation in electronic format. An original signed hard copy of the signature page is on file in University Archives.

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Abstract

Deep brain stimulation (DBS) is a powerful therapeutic option for intractable movement and affective disorders (Parkinson’s disease or PD, tremor, dystonia, Tourette syndrome, chronic pain, obsessive compulsive disorder, depression, bipolar). The benefits of DBS are immediate and dramatic, manifested as instantaneous improvements in motor function in the case of PD patients. However, due to the nonspecificity of electrical stimulation, DBS has variable efficacy and can lead to serious side effects. The mechanisms behind the effects of DBS are still highly controversial and there is tremendous interest from both neuroscience and clinical communities to understand and improve DBS.

We have developed a novel technology based on two microbial opsins, Channelrhodopsin (ChR2) and Halorhodopsin (NpHR), that allows to directly and specifically control the activity of distinct cell-types with high temporal precision in well defined brain regions, therefore allowing us to overcome the lack of specificity of electrical DBS.

This study provides the first investigation of the role of specific cell types in ameliorating PD symptoms addressed by effective DBS treatment. The focus of the thesis was twofold: (1) to develop and optimize optogenetic technologies (molecular and hardware) for safe and effective use in behaving mammals; and (2) to employ the above developed optogenetic toolkit to deconstructing diseased brain circuitry, with focus on Parkinson’s disease.

The framework and technological toolbox presented here can be employed across many brain circuits to selectively control individual components and therefore systematically deconstruct intact and disordered brain processes.

Preface

The work presented in this thesis consists of five published studies, as described below. These studies, listed in chronological order except the introductory chapter 1, focused on early optogenetic development (Chapters 1 and 2), using optogenetics to understand diseased brain circuitry (Chapter 3), providing the scientific community with a detailed list of protocols for optogenetic applications (Chapter 4) and advancing the optogenetic tools to new levels of potency and generalizing cellular targeting means for optogenetics (Chapter 5).

CHAPTER 1 – Introduction, based on a published manuscript by Schneider, M.B., Gradinaru, V., Zhang, F., and Deisseroth, K. (2008). Controlling neuronal activity. The American journal of psychiatry 165, 562. This is a very brief introduction to optogenetics and how it relates to application to neurological disorders. The article was written by Bret and Karl; Feng and I contributed the figure panels and text comments.

CHAPTER 2 – Gradinaru, V., Thompson, K.R., Zhang, F., Mogri, M., Kay, K., Schneider, M.B., and Deisseroth, K. (2007). Targeting and readout strategies for fast optical neural control in vitro and in vivo. J Neurosci 27, 14231-14238. This was an article reviewing the early versions of ChR2 and NpHR and introducing: (1) an enhanced ChR2 version for neuronal applications; (2) intracellular and intercellular targeting means for optogenetics; (3) a simultaneous recording-stimulation device, the optrode; (4) the first in vivo demonstration of motor control using optogenetics. This article was written by me and Karl. All the experiments were carried out by me in collaboration with the other authors on the paper.

CHAPTER 3 – Gradinaru, V.*, Thompson, K.R.*, and Deisseroth, K. (2008). eNpHR: a Natronomonas halorhodopsin enhanced for optogenetic applications. Brain cell biology 36, 129-139. *Equal contribution. This study came as a neccesity to develop an NpHR variant that would be well tolerated at high expression levels and functional in the intact mammalian brain. I generated the molecular variants of NpHR and

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performed in vitro and in vivo electrophysiology and Kim performed the toxicity assays and immunocytochemistry; we wrote the paper together with Karl.

CHAPTER 4 – Gradinaru, V.*, Mogri, M.*, Thompson, K.R., Henderson, J.M., and Deisseroth, K. (2009). Optical deconstruction of parkinsonian neural circuitry. Science (New York, NY 324, 354-359. *Equal contribution. This study, which was my main Ph.D. thesis and qualifying proposal, was conceived through collaboration with Karl Deisseroth and Jaimie Henderson. We looked at the mechanisms behind Deep Brain Stimulation in animal models of Parkinson’s disease using optogenetics tools in the development of which I was involved in the first 2 years of my PhD. Most of the experiments were conducted by me and Murtaza, with help from Kim Thompson with immunohistochemistry. The main text was written primarily by me and Karl, with help from Murtaza, who wrote the supplementary information with help from me.

CHAPTER 5 – Zhang, F.*, Gradinaru, V.*, Adamantidis, A.R.*, Durand, R., Airan, R.D., de Lecea, L., and Deisseroth, K (2010). Optogenetic interrogation of neural circuits: technology for probing mammalian brain structures. Nature protocols 5, 439- 456. *Equal contribution. This study came as a neccesity to have a comprehensive collection of optogenetics techniques that the scientific community could access for their own studies. The paper describes the numerous constructs, hardware, and techniques, developed over more than 5 years by all members of the Deisseroth lab and by Antoine Adamantidis in the de Lecea lab. Feng, me, and Antoine contributed equally to the writing and figures in the paper. The sections on VSDI were the exclusive contribution of Remy Durand and Raag Airan; because I did not personally performed VSDI during graduate school, I removed the selected sections from this thesis.

CHAPTER 6 – Gradinaru, V.*, Zhang, F.*, Ramakrishnan, C., Mattis, J., Prakash, R., Diester, I., Goshen, I., Thompson, K.R., Deisseroth, K. Molecular and Cellular Approaches for Diversifying and Extending Optogenetics. Cell. 2010 Apr. *Equal contribution. This study was conceived through collaboration with Karl and Feng. Karl and I wrote the paper with help from Feng and Joanna. I carried out most of the experiments in collaboration with the co-authors on the paper. The paper introduces

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the latest generation of optogenetic tools, which includes highly potent inhibitors that span the visible spectrum, including the infrared border, and promoter-free targeting methods for optogenetics.

Acknowledgements

Stanford has been a wonderful environment for my scientific and personal growth. The Neuroscience program has been truly a home away from home for me, a student from Romania. Being part of the new exciting center, the Bio-X, and supported by wonderful fellowships such as the SGF and SIGF, I had little limits to what I could attempt and achieve. The Deisseroth lab was a playground full of possibilities and working there day by day and witnessing the development of a new area in Neuroscience has been tremendously rewarding and motivating.

I am deeply grateful to my advisor Karl Deisseroth for his scientific and personal guidance. He has been a model for hard work, passion, and excellence. His energy seemed endless and intimidating at times but he nevertheless carries himself with such modesty and candor. I saw the lab growing from a handful of people, mainly graduate students, assembling incubators and rigs, and trying to figure out their way, to a fully mature lab with many highly skilled and intelligent people and amazing resources. I am grateful to Karl for the environment that he provided all of us with so that we can do science and I hope I can follow in his steps and become a collaborative, contributing, member to the scientific community.

The help and support of the entire Deisseroth lab and many members of the Stanford community was crucial to me completing all these projects in the last four years. In particular I would like to thank Feng Zhang (my rotation mentor), Charu Ramakrishna (my molecular cloning collaborator and lab mom), Kim Thompson (with whom I worked with on most of my papers), Leslie Meltzer, Raag Airan, Hsing-Chen Tsai, Ofer Yizhar, Melissa Warden, Lief Fenno, Vikaas Sohal, Mani Roy, Myriam Cordey, and Bret Schneider for their daily advice and support; Kenneth Kay, Young Lee, Sally Pak, and Hong Zeng for their technical support, Cynthia Delacruz for making all of the administrative aspects of research so effortless, and to the Colella Family Fellowship Fund, one of the Stanford Interdisciplinary Graduate Fellowships (SIGFs).

I would also like to thank my thesis and quals committee, Carla Shatz, Helen Bronte-Stewart, Paul Buckmaster, Richard Tsien, and Eric Knudsen, for their mentorship, constructive comments during yearly meetings, and for accommodating my thesis defense despite their

busy schedules.

I want to dedicate this work to my family:

To my Romanian parents, who did not go to college but always thought that education is of utmost importance for their daughter. To my grandmother, for putting little limits to my actions or imagination, while I was growing up in the country-side during my pre-school years.

And to my husband, Andrei, and our son, Theodor, for the opportunity to figure out life together.

I look forward to the trip ahead…

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Table of Contents

ABSTRACT v PREFACE vii ACKNOWLEDGEMENTS xi

CHAPTER 1 Introduction 1

CHAPTER 2 Targeting and readout strategies for fast optical neural 5 control in vitro and in vivo.

CHAPTER 3 Natronomonas halorhodopsin enhanced for optogenetic 19 applications

CHAPTER 4 Optical deconstruction of parkinsonian neural circuitry 32

CHAPTER 5 Optogenetic interrogation of neural circuits: technology for 69 probing mammalian brain structures

CHAPTER 6 Molecular and Cellular Approaches for Diversifying and 101 Extending Optogenetics

BIBLIOGRAPHY 131

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List of Illustrations

FIGURE 1 Optogenetics: Cell-type Specific Neuromodulation Using Light. 1 FIGURE 2 Targeting Modalities. 10 FIGURE 3 Readout modalities in vitro and in vivo. 14 Protein aggregation and photocurrents altered by changes in 21 FIGURE 4 intracellular targeting of NpHR. FIGURE 5 Intracellular targeting of eNpHR. 23 FIGURE 6 Summary of eNpHR functional properties. 25 FIGURE 7 In vivo function of eNpHR at high expression level. 27 FIGURE 8 Direct optical inhibition of local STN neurons. 35 FIGURE 9 Targeting astroglia within the STN. 39 FIGURE 10 Optical depolarization of STN neurons at different frequencies 41 FIGURE 11 Quantification of the tissue volume recruited by optical intervention. 43 FIGURE 12 Selective optical control of afferent fibers in the STN. 45 FIGURE 13 Selective optical stimulation of layer V neurons in anterior primary 48 motor cortex. FIGURE 14 Optogenetic tools. 71 FIGURE 15 Stereotactic implantation of the cannula guide. 76 FIGURE 16 Preparation of an optical fiber for in vivo neural control in rodents. 78 FIGURE 17 Functional expression of microbial opsin genes in the rodent brain. 99 FIGURE 18 Multiple trafficking modules for microbial opsin function in 105 mammalian neurons. FIGURE 19 Trafficking-enhanced projection targeting and topological targeting in 108 vivo. FIGURE 20 Far-red optogenetic inhibition and single-component, bidirectional 113 optical control. FIGURE 21 eBR: novel trafficking-enhanced tool for green inhibition. 115 FIGURE 22 Optogenetics: molecular design for microbial tools. 117

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Chapter 1

Introduction

A problem shared by neuroscience researchers and clinicians alike is the lack of tools for specifically controlling the function of identified neuronal classes within a heterogeneously- populated tissue. In the laboratory, the inability to target stimulation to unique cell-types contributes to the difficulty of elucidating the individual elements of neural circuits. In the clinical setting, electrical stimulation leads to undesired therapeutic side-effects resulting from current spread in deep brain stimulation (DBS).

FIGURE 1 | Optogenetics: Cell-type Specific Neuromodulation Using Light Top: Electrical vs. Optogenetic Neuromodulation: the electrode non-specifically affects all nearby neurons (left) whereas blue or yellow light emitting devices affect only the neurons containing excitatory ChR2 (center) or inhibitory NpHR (right) proteins. (Adapted from Aravanis et al., Journal of Neural Engineering, 4(3):S143, 2007.) Bottom: A prototype implantable light-delivery device based on LED technologies can be directly mounted on the forebrain of laboratory animals used as disease models (left). The use of yet more compact emitters is expected to smooth the path to eventual human use. A neuron expressing

ChR2 and NpHR can be optically controlled to fire precisely-patterned action potentials (right). (Adapted from Zhang et al., Nature, 446(7136):633, 2007.)

Our laboratory has developed a solution using cell-type specific optical neuromodulation by engineering neurons to express two microbial opsins (Figure 1). The first protein, Channelrhodopsin-2 (ChR2), is an algal light-activated cation channel that causes a neuron to depolarize in response to blue light. ChR2’s complement, Natronomonas pharaonis halorhodopsin (NpHR), is a chloride pump that renders a neuron hyperpolarized in response to amber light. These two light-sensitive proteins have been described as “traffic lights” for neural circuits: where as amber light makes neurons “stop”, blue light tells a neuron to “go”. These two proteins can be introduced to specific classes of neurons within a heterogeneous tissue. When exposed to blue and/or amber light, these light-sensitive neurons can fire patterned action-potential trains with millisecond precision, on the timescale of neural signaling. Although just a laboratory tool for now, these proteins may inspire new approaches for cell-type specific neuromodulation, to mitigate undesired side-effects associated with electrical stimulation. By introducing ChR2 and NpHR into disease-relevant cell-types, it will be possible to optically modulate errant neural signals without impacting nearby non-targeted circuits.

Recently, deep brain stimulation of subgenual cingulated area 25 or nucleus accumbens as a treatment for refractory major depression has demonstrated preliminary clinical efficacy. In optogenetic therapy, an implantable NpHR-amber-light stimulator may be used to attenuate the activity of a metabolically hyperactive subgenual cingulate. An analogous ChR2-blue-light stimulator could be used to modulate the nucleus accumbens. By restricting the expression of light-sensitive proteins to the disease-relevant neuronal populations in the nucleus accumbens or subgenual cingulate, it may be possible to avoid non-specific modulation as observed with electrical stimulation. Highly specific modulation of neuronal activity through optogenetics may promise decreased stimulation-related side-effects and increased efficacy.

The optogenetic alternative to electrical deep brain stimulation may one day enable neurosurgeons to selectively control the neurons responsible for disease etiology and symptoms, thereby establishing a new generation of safer and more efficacious DBS treatments. However, safety challenges associated with gene delivery still presents a barrier to clinical translation. In the meantime, FDA-approved clinical trials involving viral and non- viral gene delivery to the human brain are already underway, establishing the safety and gene

delivery efficacy of gene therapy technologies. Despite the numerous safety and ethical challenges, optogenetics holds great promise for the treatment of neurological disease in humans.

Chapter 2

Targeting and Readout Strategies for Fast Optical Neural Control In Vitro and In Vivo

Abstract | Major obstacles faced by neuroscientists in attempting to unravel the complexity of brain function include both the heterogeneity of brain tissue (with a multitude of cell types present in vivo) and the high speed of brain information processing (with behaviorally-relevant millisecond-scale electrical activity patterns). To address different aspects of these technical constraints, genetically-targetable neural modulation tools have been developed by a number of groups (Banghart, Borges et al. 2004; Karpova, Tervo et al. 2005; Lima and Miesenbock 2005; Thompson, Kao et al. 2005; Chambers, Banghart et al. 2006; Tan, Yamaguchi et al. 2006; Gorostiza, Volgraf et al. 2007; Lerchner, Xiao et al. 2007; Szobota, Gorostiza et al. 2007). One approach recently brought to neurobiology, combining both high speed and genetic targeting, is based on a family of fast light-responsive microbial opsins including halorhodopsins (e.g. NpHR) and channelrhodopsins (e.g. ChR2) (reviewed in Zhang et al., 2007b). These microbial opsins are single-component transmembrane conductance regulators encompassing light sensitivity and fast membrane potential control within a single open reading frame, which can be used to achieve fast bidirectional control of specific cell types even in freely moving animals (Adamantidis et al., 2007; Zhang et al., 2007a). While the basic functioning of these tools has been reviewed elsewhere (Zhang, Aravanis et al. 2007), here we describe a collection of targeting and readout strategies designed for rapid and flexible application of the microbial opsin system, and provide pointers to the relevant literature. Combinations of these multiple levels of targeting and readout define an evolving toolbox that may open up new possibilities for basic neuroscience investigation.

Channelrhodopsin-2 (ChR2) and halorhodopsin (NpHR)

Although it had been known that microbial opsins can control membrane potential in their native settings and various reduced systems (e.g. Ehrlich et al., 1984; Kalaidzidis et al., 1998; Nagel et al., 2002, 2003), the surprising ability of these proteins when heterologously expressed to precisely control neurons and the technology for applying these tools in freely- behaving animals have been demonstrated only over the past two years (reviewed in Zhang et al., 2007b). ChR2 was the first microbial opsin brought to neurobiology, where it was initially found in hippocampal neurons that ChR2-expressing neurons can fire blue light-triggered action potentials with millisecond-precision, due to depolarizing cation flux, without addition of chemical cofactors (Nagel, Szellas et al. 2003; Boyden, Zhang et al. 2005; Li, Gutierrez et al. 2005; Nagel, Brauner et al. 2005; Bi, Cui et al. 2006; Schroll, Riemensperger et al. 2006; Zhang, Wang et al. 2006; Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007; Zhang, Wang et al. 2007; Zhang and Oertner 2007). Second, an inhibitory member of the microbial opsin family (Ehrlich et al., 1984; Hegemann et al., 1985; Duschl et al., 1988; Kalaidzidis et al., 1998; Bamberg et al., 1993; Kolbe et al., 2000) was brought to neurobiology; in work stimulated by the finding that the all-trans retinal chromophore required by microbial opsins appears already present within mammalian brains (Zhang et al., 2006), it was found that neurons targeted to express the light-activated chloride pump halorhodopsin from Natronomonas pharaonis (NpHR) can be hyperpolarized and inhibited from firing action potentials when exposed to yellow light in intact tissue and behaving animals (Han and Boyden 2007; Zhang, Wang et al. 2007). Third, genetic tools have been developed for versatile integration of microbial opsins with the enormous resource of Cre-LoxP mouse driver lines suitable for a wide variety of neuroscience investigations (Zhang et al., Society for Neuroscience Abstracts 2007; described below). Finally, integrated fiberoptic and solid-state optical approaches have provided the complementary technology to allow specific cell types, even deep within the brain, to be controlled in freely behaving mammals (Aravanis et al., 2007; Adamantidis et al., 2007; reviewed in Zhang et al., 2007).

What factors limit opsin efficacy? One potential limitation is that for invertebrates like C. elegans and Drosophila the all-trans retinal must be delivered, but this turns out to be straightforward to provide in food or in the environment (Nagel, Brauner et al. 2005; Schroll, Riemensperger et al. 2006; Zhang, Wang et al. 2007). In general, sufficient levels of functional expression (which may depend on gene delivery method, copy number, promoter

strength, and time after initiation of gene expression) must be obtained to either potentiate or inhibit spiking. However, care must be taken not to express the opsins at excessive levels, as overexpression of membrane proteins can lead to toxicity and loss of membrane integrity in a wide variety of systems. We and our colleagues have generated healthy ChR2-EYFP transgenic mice lines with expression throughout the brain using the Thy1 promoter; under these conditions, ChR2 can be used to map synaptically connected neurons in anesthetized mice and in slices (Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007). We note that visualization of NpHR-transduced neurons indicates that intracellular halorhodopsin accumulation is particularly pronounced (Zhang et al., 2007a). Indeed, as we recently have explored the regime of very high levels of expression (strong promoters and high viral-vector multiplicity-of-infection or high gene copy-number) in order to determine upper bounds of effect sizes, we have seen intracellular accumulation and even membrane blebbing toxicity that can be prevented by returning to lower, effective expression levels (Zhang et al., 2007); this first-generation solution soon may be enhanced by second-generation molecular modifications to modulate membrane targeting. NpHR is well tolerated at expression levels that subserve its inhibitory function both in behaving C. elegans and in intact mammalian brain tissue (Zhang, Wang et al. 2007), and NpHR can be introduced developmentally or in adulthood at suitable titers (Figure 2A; Zhang et al., 2007a).

Putting opsins in their place: beyond genetic targeting in networks

The rate limiting step in employing these tools is now directing their activity to the specific locations where electrical control is needed, by controlling either localization of the light or localization of protein expression (e.g. via cell type-specific promoters like the excitatory neuron-specific CaMKIIα promoter; Figure 2C; Aravanis et al., 2007). To rapidly capitalize on the many available mouse lines driving (for example) Cre recombinase under the control of cell type-specific promoters, AAV vectors have been developed (Zhang et al., Society for Neuroscience Abstracts 2007) carrying microbial opsins driven by a strong promoter followed by a floxed-stop sequence that can be injected into Cre-expressing mice. The cell type of interest efficiently recombines the stop sequence out of the genome and allow opsin expression under the strong promoter; the cell type of interest is then susceptible to control by fiberoptic light delivery registered to the virus injection sites using published methods for reliably achieving this crucial co-registration (Adamantidis et al., 2007). These promoter-

based approaches are well-suited to distinguish the roles played by specific neural types in the physiological or pathological functioning of the nervous system.

A potential limitation is that specific promoter-based targeting strategies or mouse Cre lines are not always immediately available or suitable for some experiments. However, in many cases, projection patterns can be employed to achieve optical control over the desired neurons. Contralateral projections in the brain have been targeted by labeling cortical neurons in vivo with ChR2, followed by high-speed circuit mapping with light exposure delivered to brain slices from the contralateral hemisphere (Petreanu, Huber et al. 2007). In this experiment, contralateral axonal projections showed similar target preference in comparison with the ipsilateral projections. These experiments illustrate a targeting principle that may be generalizable. For example, Figure 2B shows selective labeling of hippocampal contralateral projections, in which axons of ChR2-positive hippocampal cells from the virus-injected hemisphere synapse onto ChR2-negative granule cells in the contralateral dentate gyrus. Optical stimulation with blue light of the ChR2-positive axonal projections contralateral to the site of injection can activate postsynaptic ChR2-negative dentate granule cells (Figure 2B, right). In this way the selective impact of fibers travelling from one brain region to another, spatially separated, brain target region can be studied simply by illuminating the target brain region, a principle that could extend to many other circuits (e.g. cortico-thalamic and nigrostriatal projections) and to species in which promoters are less well defined (e.g. primates). However, a limitation of this method is that in some cases insufficient opsin accumulation in the presynaptic terminal may prevent robust spiking responses, which may be overcome with molecular targeting strategies. Another non-promoter-based targeting method would be to use trans-synaptic viral methods to introduce opsins; for example, trans-synaptic tracers based on rabies virus will spread only to directly connected cells, thereby allowing tests of the functional role of neurons one synapse removed from the initially-infected cells (Wickersham, Lyon et al. 2007).

Cell types also can be targeted developmentally. One example involves the targeting of specific cortical layers with in utero electroporation, as demonstrated for ChR2 by Petreanu et al. (2007). Indeed, in utero electroporation at precisely timed embryonic days in mouse can be used to target both ChR2 and NpHR together to individual cortical layers such as layers II&III (E15.5; Figure 2A), layer IV (E13.5), or layers V&VI (E12.5). In utero electroporation also could be used to express ChR2/NpHR in the inhibitory neurons of the striatum or in the

hippocampus (Borrell, Yoshimura et al. 2005; Navarro-Quiroga, Chittajallu et al. 2007). A key advantage of in utero electroporation is that, unlike viral delivery methods, there are no DNA size restrictions, which allows for expression of opsins under control of a wider array of promoters. Promoter-based strategies could also be used to target specific cortical layers, since emerging knowledge of layer-specific markers (Lein, Hawrylycz et al. 2007) may lead to highly specific targeting strategies, and layer V of cortex already has been targeted by generating transgenic mice expressing ChR2-EYFP under control of the Thy1 promoter (Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007). Of course these spatial targeting strategies could be complemented with traditional promoter-based strategies, e.g. with promoter fragments, bacterial artificial chromosome (BAC) transgenics (Heintz 2001), Cre/ROSA26-loxP mice (Srinivas, Watanabe et al. 2001), or Gal4-UAS enhancer expression systems (Perrimon 1998). For example, a combined developmental/promoter-based strategy could allow selective targeting of excitatory (CaMKIIα-expressing) neurons only in Layer 2/3 of cortex.

ChR2 and NpHR are ubiquitously localized membrane proteins when heterologously expressed, and full-field illumination will generate membrane potential changes synchronously across the entire cell. This pattern of activity modulation is suitable for most classes of circuit-level experiments in which neurons simply need to be activated or inhibited in specific temporal patterns. However, subcellular targeting of opsins could be used to modulate activity in localized fashion even with full-field flash illumination; for example, dendritic targeting of ChR2 could preserve physiological information flow from synapse-to- soma-to-axon for studies of subcellular signaling (synapse-to-nucleus signaling, dendritic integration rules, spike timing-dependent plasticity, and other back-propagating action potential effects). To achieve precise optical control within neurons, we have explored the use of intracellular targeting motifs that can be attached to the opsin nucleotide sequences. As one example of this approach, we have attached the PDZ-domain binding sequence ETQV, a motif involved in NMDA receptor clustering at postsynaptic sites, to the C-terminus of ChR2-EYFP (Zito, Parnas et al. 1999; Guerrero, Reiff et al. 2005). When expressed in hippocampal neurons, ChR2-EYFP-ETQV was indeed concentrated at the sites of post-synaptic densities, as indicated by selective colocalization with PSD-95, in contrast to the nonselective localization of ChR2-EYFP (Figure 2D). The postsynaptically recruited protein maintained its ability to induce precisely timed depolarization and action potential firing in hippocampal neurons (not shown). If this synaptic targeting were combined with precise optical guidance

strategies, the causal role of neural population codes, in which members of the population are individual synapses rather than individual neurons, could be studied. Other targeting strategies (for example, to presynaptic terminals, to axon hillocks and nodes, or to intracellular membranes) should also be possible and will facilitate temporally-precise studies of subcellular signaling events as well as network-level optogenetic experiments.

FIGURE 2 | Targeting Modalities A) Developmental targeting: NpHR-EYFP (green) and ChR2- mCherry (red) expression in layers 2&3 of the mouse cortex via timed in utero electroporation (E16 surgery, P27 coronal slicing). A mixture of CMV::NpHR-EYFP and CMV::ChR2-mCherry DNA

(1µg/µl each) was pressure-injected unilaterally into the ventricle and electroporated (5x50ms pulses, 1Hz, 40V; see also (Petreanu, Huber et al. 2007)). Shown are pictures of acute slices taken on a Leica MZ16FA stereomicroscope. B) Anterograde projection-based targeting: unilateral delivery of CaMKIIα::ChR2-mCherry lentivirus (1.5µl, 109 p.f.u./ml) to the adult mouse dentate hilus/CA3 (left, and inset showing DIC image superposition) results in axonal ChR2-mCherry expression in the contralateral hippocampus (middle), with no expression in the granule cell layer (GCL) on either side of the brain (right). Note contralateral ChR2-mCherry positive axonal projections to the molecular layer and subgranular zone. Optical stimulation with 15ms pulses of blue light (HQ470nm/40x, Chroma) of ChR2-positive axonal projections in acute slices from the contralateral hippocampus evoked inward synaptic currents in ChR2-negative dentate granule cells. A Lambda DG-4 optical switch (Sutter Instruments) and 300W Xenon lamp were used for light delivery. Two injection sites were used at AP: 2mm; ML: 1.5mm; DV: -2mm, and AP: 3mm; ML: 3mm; DV: -3mm. C) Promoter-based targeting: ChR2-EYFP (green) was targeted to excitatory hippocampal neurons in culture via CaMKIIα::ChR2- EYFP lentivirus (infected 2 div); immunostaining for GAD67 (red; Chemicon 1:250) in 14 div cultures shows exclusion of ChR2 expression from inhibitory GAD67+ neurons. D) Subcellular targeting: ChR2-EYFP (green) is expressed ubiquitously throughout the cell membrane in 16 div cultures. Adding the PDZ-domain binding motif ETQV to the C-terminus by in-frame cloning led to concentration of ChR2-EYFP at postsynaptic sites, indicated by colocalization with PSD95 staining (red; Affinity Bioreagents 1:200). Pearson’s correlation (Manders et al., 1993) was r=0.78 for ChR2-EYFP-ETQV and r=0.40 for untargeted ChR2-EYFP, unpaired t-test p<.0001, n=22 and 24 dendrites respectively. Single confocal sections through 50µm segments of proximal dendrites were thresholded and correlation coefficients determined using Volocity software.

Integrating multimodal readout technologies with optical control

The fully-equipped neuroscience optical toolbox would include not only ChR2/NpHR optical control, but also multiple reliable readout technologies for answering a broad range of experimental questions, in vitro and in vivo. Indeed, simultaneous optical control and electrophysiological recording of neuronal activity in vivo, even with intraparenchymal stimulation, is now possible with integrated sub-millimeter scale fiberoptics and recording electrodes (Figure 3A), complementing the capabilities of surface illumination described in Arenkiel et al. (2007). In this way local-circuit responses can be measured with temporal precision that matches the millisecond-scale precision of ChR2/NpHR-based optical control, even within deep brain structures in vivo. This rapid feedback not only is experimentally valuable, but also could be important for eventual clinical applications of optical stimulation, so that therapeutic stimulation parameters could be rapidly tuned or timed in patients. A major

limitation in functional neurosurgery and interventional psychiatry is the fact that electrical artifacts from stimulating electrodes prevent precisely simultaneous recording from local neurons, so that it is not clear whether particular brain stimulation paradigms are activating, inhibiting, or desynchronizing the target tissue. Although care must still be taken with regard to potential artifacts from external electronics, the combined optical stimulation/electrical recording approach allows high-temporal-precision electrical assessment of local responses in the absence of locally-generated electrical artifacts, allowing local evoked spikes to be directly visualized (Figure 3A).

However, for some applications slow or stable readouts are preferable. Slower biochemical markers may be ideal for tracking and validating the outcome of optical neural control, particularly in cases where simultaneous imaging of controlled cells is not possible but it is still important to quantify the effect of optical control on the stimulated or downstream population. Activation of immediate-early genes like c-fos and Arc can be measured with biochemical methods (e.g. antibodies or GFP-based reporters) 60 min or more after the presumptive optical stimulation, and other transcription factor activation events (e.g. CREB Ser133 phosphorylation) provide sensitive assays that can detect and stably report changes in membrane potential (Figure 3B). Blue light exposure (470 nm for 10min at 10Hz/25ms) to ChR2 expressing neurons robustly elevated Ser133 pCREB levels (Figure 3B); in this way changes in network activity can be tracked using biochemical traces, even after conclusion of optical control of neural activity, although this method is not precise enough for reporting the firing frequency experienced by any one cell during the stimulation protocol. Even in the presence of synaptic activity blockers, optical activation of NpHR (590 nm for 10min) sufficed to significantly lower Ser133 pCREB levels compared with sham treated neurons (Figure 3B, inset); however, if the conditions are not strong enough to separate two populations, the biochemical trace imaging technique is not suitable for precisely determining the fraction of cells controlled by the optical protocol.

Fast optical readouts are needed as well. The action spectra of ChR2 and NpHR are such that even with single-photon imaging, both tools can be used simultaneously together with the fluorescent calcium indicator Fura-2 (Zhang, Wang et al. 2007), and each are also compatible in principle with several organic voltage-sensitive dye strategies (Zhang, Wang et al. 2006), e.g. the absorbance dye RH-155 for slice physiology (Airan et al., 2007). Moreover, the availability of genetically encoded biosensors that can detect glutamate levels, membrane

voltage, protein synthesis, or intracellular calcium in vitro and in vivo (Reiff, Ihring et al. 2005; Parrish, Wang et al. 2006; Garaschuk, Griesbeck et al. 2007; Heim, Garaschuk et al. 2007) could provide cell type-specific optical readouts for effects of cell type-specific optical control with ChR2/NpHR. A wide range of genetically-targetable activity reporters, such as GCaMPs, pericams, or Camgaroos (Nagai, Sawano et al. 2001; Nakai, Ohkura et al. 2001; Kotlikoff 2007) could be used together with ChR2/NpHR in vitro and in vivo, but care must be taken in choosing and/or developing the right reporters whose excitation spectra will not overlap with the ChR2/NpHR action spectra. For example, the fast troponin C FRET-based biosensor TN-XL (Mank, Reiff et al. 2006) is excited at 430nm and could be a good candidate for pairing with NpHR. The availability of two-photon excitation laser scanning microscopy (Pettit, Wang et al. 1997; Svoboda and Yasuda 2006) relaxes the need for nonoverlapping spectra, and optical techniques including use of multiple excitation spots, defocused beams, and fast scanning over cells may overcome the fact that two-photon microstimulation will only affect a small number of opsins in a typical excitation volume. Conversely, conventional two- photon imaging of circuit responses could be conducted in parallel with single-photon excitation of large numbers of targeted opsins.

A key readout for systems neuroscience-level questions is behavior of freely-moving animals. NpHR and ChR2 have been used in combination to control movement in freely-moving C. elegans (Zhang, Wang et al. 2007) and ChR2 has been used to modulate sleep-wake behavior in mice (Adamantidis et al., 2007); here optical control of locomotion in freely behaving adult mice expressing ChR2 in cortical layer 5 is demonstrated with fiberoptic/laser diode tools (Figure 3D). Timelocked to optical stimulation onset and termination, right motor cortex (area M2)-implanted animals displayed an evoked bias for moving to the left side, a behavior not seen in control (non-CHR2-expressing) mice (Figure 3D; see supplementary movies for experimental and control animals). The lightweight and thin optical fiber allows unrestricted behavior of the subject over long distances, and a commutator allows even behavior involving rotation to occur, an important feature for some disease models like Parkinson’s. In cases where the targeted cells or their dendrites are superficial, cortical-surface LEDs illumination through a cortical window and even through thinned skull (Yoder and Kleinfeld, 2002) can also be used to drive temporally precise motor output patterns (Figure 3E). Light-induced whisker deflections in these intact animals were measured magnetically (Aravanis, Wang et al. 2007) and in control experiments blue light illumination in ChR2-negative mice had no effect.

FIGURE 3 | Readout modalities in vitro and in vivo. A) Electrophysiology in vivo: simultaneous optical stimulation and electrical recording from cortex in an intact Thy1::ChR2-EYFP mouse using an extracellular tungsten electrode (1MΩ, ~125µm outer diameter) glued to an optical fiber (~200µm outer diameter, ThorLabs); the ~0.5mm offset (top right) ensures illumination of recorded neurons. The top- right illumination cone picture was generated by placing the electrode & fiber ensemble into a 1cm thick block of 1% agar (electrode superficial as shown); the geometrical dispersion is greatly amplified by scattering in brain tissue as described (Aravanis et al., 2007). FC/PC-coupled fiberoptic is shown (middle) coupled to a 473 nm laser diode from CrystaLaser through a FC/PC adapter. Thy1::ChR2- EYFP (line 18) mice (Wang et al., 2007, Arenkiel et al., 2007) have strong ChR2 expression in layer 5 of neocortex (left). Recordings in anesthetized mice showed reliable cortex activation with pulsed blue light (10Hz/25ms), without stimulation artifacts at either the onset or offset of the stimulus. pClamp 10 and Digidata 1322A board (Axon Instruments) were used to collect data and generate pulsed light. The recorded signal was bandpass filtered at 300Hz low/5 kHz high (1800 Microelectrode AC Amplifier, A- M Systems). B) Post-hoc biochemical imaging modality. Cultured hippocampal neurons (13-14 div, transduced 2 div with ChR2-EYFP and NpHR-EYFP lentiviruses under the CaMKIIα promoter) were preincubated ~2hr with D-AP5 (25 µM) and NBQX (10 µM) and then stimulated with light in the

continued presence of D-AP5/NBQX. Blue light delivery (HQ470nm/40x, Chroma) for 10 min at 10Hz/25ms to ChR2-expressing neurons significantly elevated Ser133 pCREB compared with sham- treated neurons (unpaired t-test p<.0001, n=60 and 72 cells respectively). Inset: Activation of NpHR with 10 min of continuous amber light (593nm/20x, Semrock) significantly lowered pCREB levels compared with sham-treated neurons (unpaired t-test p<.0001, n=80 and 75 cells respectively). Cumulative probability histograms show nuclear pCREB intensity normalized to the sham condition mean. For quantification of pCREB immunofluorescence (Upstate 1:500), five confocal images (40X/1.3NA oil) were acquired per stimulated region. Single optical sections through the nucleus were analyzed using Volocity software and nuclear regions were defined by DAPI labeling. Imaging and quantification were conducted blind to treatment condition. Optical stimulation was conducted on an Olympus IX71 inverted scope using a 20x/0.75NA objective (7 mW/mm2 light intensity measured with Newport 1815-C power meter). C) Real-time activity imaging modality. ChR2 and NpHR action spectra are shown superimposed with Fura2-Ca2+ indicator excitation spectrum and RH-155 voltage indicator absorption band. Spectral separation allows for simultaneous all-optical control and readout. D) Directional control of mammalian locomotion with fiberoptic light delivery in freely moving animals. Blue light stimulation of the right secondary motor cortex (M2) in Thy1::ChR2-EYFP adult mice induced repetitive rotations to the left that terminated immediately with conclusion of optical stimulation (1.96 ± 0.24 rotations/10s). Light was delivered from the 473nm laser via optical fiber through a fiber guide cannula (Plastics1) targeted to AP: 1mm; ML: 0.5mm; DV: -0.5mm. 30Hz/15ms pulses were generated using a function generator (Agilent 33220A). Cranioplastic cement was used to stabilize the fiber guide and a dummy cannula (a) was placed while animals were not being tested. To ensure stability of the fiber during testing in moving animals, an internal cannula adapter (b) was glued to the stripped optical fiber (c), which was inserted into the fiber guide cannula fixed to the skull (d); see also Adamantidis et al., 2007). A custom aluminum rotating optical commutator (bottom; Doric Lenses) was used to release torsion in the fiber caused by the animal’s rotation. E) Whisking control with LEDs in sedated Thy1::ChR2-EYFP animals. Optical excitation of vibrissal motor cortex was performed either through a small craniotomy (top traces) or through thinned skull (bottom trace;) with a blue LED (LEDtronics; skull thinned ~80% with standard drill as in (Yoder and Kleinfeld 2002)). Whisker deflections were measured as before (Aravanis et al., 2007); a 1mg rare-earth magnetic particle (Magcraft) was attached to the vibrissa and a magnetoresistive sensor (Honeywell) was used to measure deflections.

Summary

Interesting integrated applications of these targeting and readout methods could go beyond simply driving specific populations of cells, to driving specific cells in different brain regions (e.g. cortex and hippocampus) with precise relative timing (synchrony or asynchrony with

well-defined phase shifts). Moreover, the motor behavioral studies (Figure 3D) suggest that specific cortical layers in a region could be turned on or off with high temporal precision in behaving subjects, a possibility unapproachable with electrode-based, pharmacological, or genetic methods. These experiments are within reach now, but there are also many refinements of microbial opsin technology still pending that could enhance the power of the approach. For example, the independence (in mammals) from exogenous cofactors is a positive aspect, but it remains possible that some mammalian cells will not have sufficient retinoids to fully support opsin function; in these cases, co-introduction of enzymes governing rate-limiting steps in retinoid uptake and metabolism (Wang et al., 2007; Yang and O'Tousa, 2007) might be helpful in driving maximally-efficacious opsin function.

The subcellular-targeting approach (Figure 2D) suggests additional strategies to target axons, somata, and synaptic terminals. Other molecular modifications may result in usefully-shifted spectral properties (e.g. redshifted opsins for better tissue penetration and multi-color optical activation experiments; Luecke et al., 2001; Shimono et al., 2003), greater or lesser Ca2+ permeance, or improved conductance properties (Nagel et al., 2005); we find in hippocampal neurons that H134R mutation in ChR2 expressed under the Synapsin-I promoter, gives rise to a 2.37 ± 0.07 (p< 0.005) fold enhancement in steady-state and 1.73 ± 0.10 (p<0.005) fold enhancement in peak current (n=5; comparison with wild-type ChR2). It is important to remember that in principle ChR2 and NpHR will contribute to small, brief and reversible elevations in intracellular Ca2+ and Cl-, respectively; likewise native excitatory transmission also elevates intracellular Ca2+ via Ca2+-permeable glutamate receptors and voltage-activated Ca2+ channels, and native inhibitory transmission typically also elevates intracellular Cl-. Since elevations in Ca2+ can drive intracellular biochemical changes and elevations in Cl- can modulate the effects of endogenous GABAergic neurotransmission, these factors should be considered in experimental design.

Findings obtained from microbial opsin work in animal models may ultimately inform novel clinical approaches, even without introduction of opsins into patients. Clinically compatible, noninvasive interventions that are likely to modulate neural activity, including radiation, ultrasound, and magnetic methods, all can be depth targeted in various ways (for example by using stereotactically-guided accumulation of energy at sub-centimeter resolution, delivered from multiple calculated trajectories as with the radiation-based Cyberknife, or depth focused as with high intensity focused ultrasound/HIFU). Energy can be depth targeted in other ways

as well, for example with fiberoptics or implanted transducers (Adamantidis et al., 2007). In principle many focusing methods could be used to directly transduce energy, or even to activate custom stereotactically- or endovascularly-delivered micro-“antennas” that locally modulate radiation sensitivity, ultrasound response, and magnetic susceptibility, at specific circuit nodes in the clinical setting that had previously been implicated with optogenetic animal models (Zhang et al., 2007b). But work in animal models will be a primary focus of the field, and in itself can promote understanding of the circuit basis of neuropsychiatric diseases like narcolepsy (Adamantidis et al., 2007).

Looking toward the future, plants and many fungi and microbes are highly dependent on light and therefore have developed highly specialized light sensing proteins that will likely continue to provide novel and powerful tools for perturbing and interrogating the intact nervous system. Optical activation of intracellular signaling via modified opsins (Kim et al., 2005) and blue- light-activated adenylate cyclases (PACs) from the flagellate Euglena gracilis (Iseki, Matsunaga et al. 2002; Ntefidou, Iseki et al. 2003) can trigger rapid and reversible increases in cAMP levels in vitro and modulate behavior in Drosophila in vivo (Schroder-Lang, Schwarzel et al. 2007). Phototropins, photo-regulated protein kinases (Huala, Oeller et al. 1997; Briggs, Beck et al. 2001), might also provide useful tools for biochemistry studies of phosphorylation events within a cell that could complement ChR2/NpHR modulation of biochemical signaling, along with other light-activated compounds yet to be identified. Perhaps only a small fraction of the potential opsins and other tools identified in macrogenetic approaches (e.g. Venter et al, 2004) will be immediately useful, as the experience with the comparatively unstable Halobacterium salinarum halorhodopsin (HsHR) revealed (Zhang et al., 2007a), and the broad action spectra of these naturally occurring proteins may limit the number of independently- addressable optical control channels achievable, but identification of new players may refine the microbial opsin approach and molecular optimizations have the potential to incrementally further unlock latent utility. Together, these novel tools, targeting techniques, and readout technologies will extend the utility of the microbial opsins in bidirectional, reversible, and spatiotemporally precise control of activity in targeted neurons.

Chapter 3

eNpHR: a Natronomonas halorhodopsin enhanced for optogenetic applications

Abstract | Temporally precise inhibition of distinct cell types in the intact nervous system has been enabled by the microbial halorhodopsin NpHR, a fast light-activated electrogenic Cl- pump. While neurons can be optically hyperpolarized and inhibited from firing action potentials at moderate NpHR expression levels, we have encountered challenges with pushing expression to extremely high levels, including apparent blebbing and intracellular accumulations. We therefore sought to molecularly engineer NpHR to achieve strong expression without these cellular side effects. We found that high expression correlated with endoplasmic reticulum (ER) accumulation, and that under these conditions NpHR colocalized with ER proteins containing the KDEL ER retention sequence. We screened a number of different putative modulators of membrane trafficking and identified a combination of two motifs, an N–terminal signal peptide and a C-terminal ER export sequence, that markedly promoted membrane localization and ER export defined by confocal microscopy and whole-cell patch clamp. The modified NpHR displayed increased peak photocurrent in the absence of aggregations or toxicity, and potent optical inhibition was observed not only in vitro but also in vivo thalamic single-unit recording. The new enhanced NpHR (eNpHR) allows safe high-level expression in mammalian neurons, without toxicity and with augmented inhibitory function, in vitro and in vivo.

Introduction

A subset of naturally-occurring microbial opsin genes, originally characterized in non-neural systems, encode light-sensitive transmembrane ion conductance regulators (Ehrlich, Schen et al. 1984; Kalaidzidis, Kalaidzidis et al. 1998; Nagel, Szellas et al. 2003; Zhang, Prigge et al. 2008). If successfully adapted as a neuroscience technology, these proteins could be enormously significant, since controlling the membrane potential of targeted cell types with high temporal resolution may

allow elucidation of cellular codes underlying neural circuit computation and behavior. Three functionally distinct classes of these microbial opsin genes have now been introduced to and adapted for neurobiology (VChR1, NpHR, and ChR2; discussed below). Among other important properties, all three operate on the millisecond timescale and can function in mammalian neurons without addition of exogenous chemical cofactors, since the chromophore for these proteins, all-trans retinal, appears to be already present at sufficient levels in mammalian brains (Zhang et al., 2006). Moreover, light and gene delivery challenges have been overcome, as integrated genetic, fiberoptic, and solid-state optical approaches have provided complementary technology to allow specific cell types, deep within the brain, to be controlled in freely behaving mammals (Adamantidis, Zhang et al. 2007; Aravanis, Wang et al. 2007; Gradinaru, Thompson et al. 2007; Zhang, Aravanis et al. 2007b).

First to be brought to neuroscience, the channelrhodopsin ChR2 allows blue light-induced action potentials to be triggered with millisecond-precision in neurons (Boyden, Zhang et al. 2005), due to depolarizing cation flux through a light-gated pore (Nagel et al., 2003); this approach has since been shown to be versatile in many experimental systems (Li, Gutierrez et al. 2005; Nagel, Brauner et al. 2005; Bi, Cui et al. 2006; Deisseroth, Feng et al. 2006; Ishizuka, Kakuda et al. 2006; Schroll, Riemensperger et al. 2006; Zhang, Wang et al. 2006; Petreanu, Huber et al. 2007; Zhang and Oertner 2007; Zhang, Wang et al. 2007a), including generation of transgenic mouse lines (Arenkiel, Peca et al. 2007; Wang, Jiao et al. 2007) and probing neural codes underlying complex behavioral state transitions important in neuropsychiatric disease (Adamantidis, Zhang et al. 2007). Second, we found that neurons targeted to express the light-activated chloride pumping halorhodopsin from Natronomonas pharaonis (NpHR) can be hyperpolarized and inhibited from firing action potentials when exposed to yellow light; because of the excitation wavelength difference, ChR2 and NpHR can be coexpressed for bidirectional control and integrated with imaging and behavior (Gradinaru, Thompson et al. 2007; Han and Boyden 2007; Zhang, Wang et al. 2007a) even in intact tissue and behaving animals (Gradinaru, Thompson et al. 2007; Han and Boyden 2007; Zhang, Wang et al. 2007a), and may turn out to be versatile across a range of in vitro and in vivo applications (Gradinaru, Thompson et al. 2007; Han and Boyden 2007; Zhang, Wang et al. 2007a). Third, a yellow light-activated channelrhodopsin gene was discovered and tested in mammalian neurons (VChR1; (Zhang, Prigge et al. 2008) that opens the door to combinatorial excitation experiments when used together with ChR2, described below. The properties of this third microbial tool also allow for deep penetration of redshifted excitation light, use of well-tolerated low-energy photons for excitation, and improved integration with existing Ca2+ indicators.

Technical challenges still remain, including refining optical and cell type-specific targeting strategies, as well as tuning activation wavelengths and ion permeabilities for different classes of experiments. One major challenge in adapting tools across large evolutionary distances (e.g. to mammals from prokaryotes and simple eukaryotes such as Volvox carteri, Natronomonas pharaonis, and Chlamydomonas reinhardtii), is that expressing heterologous membrane proteins in mammalian cells can lead to poor folding, assembly, and trafficking. We have previously reported that at high expression levels, NpHR (codon optimized for mammalian expression) forms aggregates that could cause cellular toxicity (Gradinaru, Thompson et al. 2007), and noted that this problem could be alleviated by returning to moderate expression levels, but this was not an ideal solution because large photocurrents are useful for efficient inhibition in a variety of experiments, especially for in vivo applications. Therefore, we report here on a strategy to increase the efficiency of NpHR membrane targeting to maximize photocurrents without aggregations or toxicity, even for high expression levels under strong promoters, in vitro and in vivo.

FIGURE 4 | Protein aggregation and photocurrents altered by changes in intracellular targeting of NpHR. A) Table of screened intracellular targeting strategies using N- and C- terminal peptides fused to NpHR. B) The fraction of neurons containing one or more NpHR aggregates was determined in cultured hippocampal neurons transduced with quantitatively titer-matched viral CaMKIIα::NpHR-EYFP constructs, and allowed to express for 10 days. Compared to the unmodified NpHR (orange), some constructs (grey) had little effect on aggregates while others (blue) partially reduced aggregate incidence. Notably, for one construct

(*) aggregates and toxicity were virtually abolished (only a single possible aggregate observed in >400 neurons). Data shown are relative to wild-type NpHR, and number of neurons sampled for each construct is shown in parenthesis after the construct label. C) Functionality was assessed by whole-cell patch clamp (for details see Methods and Figure 6B). All the constructs were functional as indicated by photocurrents that were comparable to the original NpHR, but one construct, the optimal construct from (B) (*) gave rise to significantly higher photocurrent per cell than all other variants (eNpHR). Data shown are relative to wild-type NpHR, and number of neurons sampled for each construct is shown in parenthesis after the construct label.

Results

In order to reduce the incidence of intracellular aggregates observed with NpHR at high expression levels, we first attempted to regulate distribution of the NpHR protein within the cell by using signal peptides from either ChR2 (Nagel, Szellas et al. 2003) or the α and β subunits of the nicotinic acetylcholine receptor (nAChR; (Isenberg and Meyer 1989; Bocquet, Prado de Carvalho et al. 2007), PDZ binding motifs (Zito, Parnas et al. 1999; Weick, Groth et al. 2003; Guerrero, Reiff et al. 2005), and actin binding motifs (Petrecca, Miller et al. 2000) (Figure 4A). Although for most of the NpHR variants this strategy considerably reduced the number of cells with aggregates while maintaining photocurrents (Figure 4B and 4C), aggregates were still observed and when present were as large as with wild-type NpHR. Next, after extensive analysis of the formation, distribution, and evolution of the NpHR aggregates in cultured neurons (data not shown) we observed that the localization of the aggregates appeared similar to ER localization in the soma and dendrites. Indeed, immunocytochemistry confirmed that NpHR aggregates colocalized with proteins containing the KDEL signal for ER retention (Figure 5B, top row), while NpHR itself does not contain a KDEL motif or other known ER retention signals (e.g. the KKAA- or RSRR-class signals).

Transport along the secretory pathway, with ER export being the first step in the pathway, is crucial for surface expression of integral membrane proteins. Although some proteins can exit the ER by bulk flow, ER export of membrane proteins can be impaired if the protein is either misfolded or lacks specific export signals (Li, Takimoto et al. 2000; Ellgaard and Helenius 2003). Because NpHR is functional in mammalian neurons even in vivo (Zhang, Wang et al. 2007a) we hypothesized that aggregate formation might be due to lack of an ER export signal rather than frank misfolding, and therefore sought to determine if adding different ER export signals to the NpHR sequence would abolish aggregate formation. C-terminal ER export signals have been shown to be important for efficient processing and surface expression of many membrane proteins (Farhan, Reiterer et al. 2008). Additionally, previous work has found that when the C-terminal ER export signals on Kv1.4

(VXXSL) or Kir2.1 (FCYENEV) are either mutated or deleted, the resulting protein forms large spheroidal intracellular accumulations similar to NpHR aggregates, and corresponding channel activity is reduced due to lower protein levels in the plasma membrane (Levitan and Takimoto 2000; Ma, Zerangue et al. 2001; Stockklausner, Ludwig et al. 2001). Moreover, suggesting that functionality can be transferred solely with these motifs, the FCYENEV sequence accelerated surface expression and increased current levels for the lobster shal potassium channel (Kv4) when added to the C-terminus (Zhang and Harris-Warrick 2004). Indeed, in the course of our modification screen we found that adding FCYENEV to the NpHR C-terminus along with the signal peptide from the β subunit of the nAChR to the NpHR N-terminus prevented aggregate formation (Figure 4B, asterisk), with such markedly improved properties that we named the resulting tool eNpHR (enhanced NpHR; Figure 5A) and studied its behavior further (summarized in Figures 5-7).

FIGURE 5 | Intracellular targeting of eNpHR. A) Primary structure of the selected construct (eNpHR) showing addition of the N-terminal signal peptide derived from nAChR and the C-terminal ER export signal derived from Kir2.1. Expression here was driven by the CaMKIIα promoter and visualized by fusion to EYFP. B) Top row: Untargeted NpHR (green) colocalized with somatic ER (KDEL ER protein staining in red;

overlap indicated in yellow and by arrows) and also notably aggregated in ER-rich regions of the dendrites (overlap indicated in yellow and by arrowheads). Bottom row: Little colocalization of eNpHR with somatic ER staining could be found, and indeed pronounced accumulations of ER staining in eNpHR dendrites were not observed. Right column: representative images of neuronal populations expressing NpHR and eNpHR. The neurons were infected with quantitatively titer-matched CaMKIIα-NpHR-EYFP or CaMKIIα-eNpHR-EYFP.

First, we found that in contrast to NpHR, eNpHR did not colocalize with ER proteins (Figure 5B), or show any evidence of toxicity even at high expression levels. eNpHR appeared to be present in somatic Golgi structures., which is typical for transmembrane proteins as they are packaged for transport. ( for Golgi colocalization with transmembrane proteins, such as Kir2.1: (Ma, Zerangue et al. 2001; Horton and Ehlers 2003; Hofherr, Fakler et al. 2005). Notably, eNpHR showed more surface membrane localization as defined by whole-cell photocurrents (Figure 6). Indeed, presumably due to increased export from the ER and increased membrane localization, eNpHR displayed significantly higher photocurrents compared to NpHR in cultured neurons infected with quantitatively titer-matched virus levels and allowed to express for the same amount of time (Figure 6: NpHR: 38.9 ± 6.8 pA; eNpHR 68.1 ± 7.2 pA; mean ± s.e.m; unpaired t-test p = 0.008), while membrane resistance was indistinguishable in the two groups (NpHR: 113.5 ± 13.9 mΩ; eNpHR: 116.8 ± 13.9 mΩ; unpaired t-test p = 0.87).

To assess eNpHR function in the setting of high in vivo expression levels, we injected highly concentrated and quantitatively titer-matched virus for both NpHR and eNpHR under the strong CaMKIIα promoter into the CA1 region of adult mouse hippocampus. While NpHR showed aggregates after 10 days of expression in vivo, eNpHR did not show any aggregates or affect cellular integrity, confirming the tolerability and improved targeting of the engineered protein even at these very high expression levels (Figure 7A, top row); enhanced membrane localization of eNpHR in vivo was also evident in dendrites (Figure 7A, bottom row). To confirm functionality, we conducted in vivo recordings using a combined optical fiber/electrode “optrode” previously described (Gradinaru, Thompson et al. 2007), and found that activity of single units in adult mouse thalamus expressing eNpHR (Figure 7B, confocal image) could be readily and reversibly inhibited with yellow light (Figure 7B, middle) in vivo; no such effect was seen in non-transduced tissue (Figure 7B, bottom).

FIGURE 6 | Summary of eNpHR functional properties. Summary of electrophysiological properties of NpHR and eNpHR in cultured hippocampal neurons. Top: Representative confocal images in cultured hippocampal neurons revealed that like a typical membrane protein, eNpHR did not appear to fill cytoplasm like NpHR (left) (right). NpHR and eNpHR were expressed for 10 days in cultured hippocampal neurons. Insets: magnified views of selected regions. Bottom: 593nm light (yellow bar) induced outward photocurrents (top right: sample traces in voltage clamp), with eNpHR evoking significantly stronger photocurrents per cell than NpHR (left bar graph; NpHR: 38.9 ± 6.8 pA; eNpHR: 68.1 ± 7.2 pA; unpaired t-test p = 0.008). Viral titers were quantitatively matched across groups (Methods). Membrane input resistance was similar for all neurons patched (right bar graph; NpHR: 113.5 ± 13.9 mΩ; eNpHR: 116.8 ± 13.9 mΩ; unpaired t-test p = 0.87). Values plotted are mean ± s.e.m; n = 12 for NpHR, n = 10 for eNpHR. Bottom right: Illumination with yellow light as expected sufficed to inhibit spiking induced by current injection in eNpHR+ neurons.

Discussion

In this study we have identified and corrected a major limiting factor in the application of optogenetic inhibition (similar results have been obtained with our colleagues at Duke University; Guoping Feng, personal communication). We traced the problem associated with high NpHR expression back to a membrane trafficking complication (Figures 4, 5), tested a large number of possible solutions (Figure 4), and validated the efficacy of the best strategy both in vitro (Figures 4- 6) and in vivo (Figure 7). We found that eNpHR completely abolished accumulations and blebs seen at very high expression levels with the original NpHR, apparently in part by allowing normal export of NpHR from the ER (assessed by confocal imaging) and by driving increased surface membrane expression (validated by quantified photocurrents). At this point we recommend use of eNpHR for all applications, and certainly those involving high expression levels in mammalian neurons, including transgenic mouse line generation (Guoping Feng, personal communication) and viral transduction approaches. We also anticipate that these modifications may enhance the expression of other microbial opsins at high levels and over long durations, pointing to the likely utility of generating similarly enhanced versions of ChR2 and VChR1.

The altered properties of eNpHR as described here clearly do not simply represent a subtle quantitative change in performance, but rather a distinct step in the development of this optogenetic technology. Future improvements could incrementally further advance eNpHR function, perhaps including the Golgi export signals from Kir2.1 (Stockklausner and Klocker 2003; Hofherr, Fakler et al. 2005), subcellular localization motifs (as in (Gradinaru, Thompson et al. 2007), and mutations that shift wavelength dependence, kinetics, light sensitivity, and ion selectivity. For example, blueshifting ChR2 and redshifting eNpHR and VChR1 will improve the ease with which combinatorial experiments are conducted, and a roadmap for the key residues likely to be involved and the type of changes likely to be helpful in this regard has been described (Zhang et al., 2008).

Finally, it has been noted (Zhang, Prigge et al. 2008) that VChR1 and ChR2 (representing yellow light excitation and blue light excitation respectively) when used together will allow combinatorial tests of the importance of specific activity patterns in interacting cell types. For example, a principal cell population can be recruited with VChR1/yellow light, in the presence or absence of precisely patterned activity in a candidate modulatory cell type driven by added blue light/ChR2. Combinatorial experiments are important to consider now for eNpHR as well, given its improved functionality in vitro and in vivo. For example, coexpression of eNpHR and ChR2 in the same cell type could allow testing the necessity and sufficiency of that specific cell type in neural circuit or animal behavior.

FIGURE 7 | In vivo function of eNpHR at high expression level. A) Confocal images showing NpHR and eNpHR expression in rodent hippocampal CA1. NpHR aggregations were clearly visualized after 10 days of strong expression of high titer virus (left) while eNpHR showed no signs of aggregates or toxicity (right) with quantitatively matched viral titers (Methods). Lower panels: magnified views of the dendritic layer. Compared to NpHR, eNpHR revealed not only absence of aggregates but also more membrane localization in distal processes in vivo. B) Simultaneous optical stimulation and electrophysiology in living mice demonstrates eNpHR potency in vivo: single unit recordings in deep brain structures. Yellow illumination delivered by the optrode method (Gradinaru et al., 2007) in vivo inhibited electrical activity in thalamus previously transduced with eNpHR by lentiviral stereotactic injection (middle trace). Top trace: same thalamic region, recording without illumination. Bottom trace: control recording 1mm ventral and anterior from the eNpHR injection site. As expected, in non-transduced tissue, light did not inhibit recorded spikes. Confocal image: eNpHR

expression in the thalamus (same animal). Inset: expanded view of a spike from unit (*) represented in the top trace.

The interdependecy of different functional groups within a complex neuronal network could be probed by expressing eNpHR and ChR2 in different cell types and determining if the functional significance of cell population A excitation is expressed through or “read out” via a downstream cell population B; this hypothesis could be tested by reversibly inhibiting cell population B (yellow light/eNpHR) in the presence of population A activation (blue light/ChR2). In this way the causal neural codes underlying circuit computation and behavior may be slowly assembled, with the long term goal of understanding how neural system properties emerge from component dynamics, both in health and disease.

Methods

DNA constructs: All NpHR variants were produced by PCR amplification of the NpHR-EYFP construct previously published (Zhang, Wang et al. 2007a) and cloned in-frame into the AgeI and EcoRI restriction sites of a lentivirus carrying the CaMKIIα promoter according to standard molecular biology protocols. All constructs were fully sequenced to check for accuracy of the cloning procedure. The map for eNpHR is available online at www.optogenetics.org.

Lentivirus preparation and titering: Lentiviruses for cultured neuron infection and for in vivo injection were produced as previously described (Zhang, Wang et al. 2007a). Viral titering was performed in HEK293 cells that were grown in 24-well plates and inoculated with 5-fold serial dilutions in the presence of polybrene (8 µg/µl). After 4 days, cultures were resuspended in PBS and sorted for EYFP fluorescence on a FACScan flow cytometer (collecting 20,000 events per sample) followed by analysis using FlowJo software (Ashland, OR). The titer of the virus was determined as follows: [(% of infected cells) × (total number of cells in well) × (dilution factor)]/ (volume of inoculum added to cells) = infectious units/ml. The titer of viruses for culture infection was 105 i.u. /ml. The titer of concentrated virus for in vivo injection was 1010 i.u. /ml.

Hippocampal cultures: Primary cultured hippocampal neurons were prepared from P0 Spague- Dawley rat pups. The CA1 and CA3 regions were isolated, digested with 0.4 mg/mL papain (Worthington, Lakewood, NJ), and plated onto glass coverslips precoated with 1:30 Matrigel (Beckton Dickinson Labware, Bedford, MA) at a density of 65,000/cm2. Cultures were maintained in a 5% CO2 humid incubator with Neurobasal-A media (Invitrogen Carlsbad, CA) containing

1.25% FBS (Hyclone, Logan, UT), 4% B-27 supplement (Gibco, Grand Island, NY), 2 mM Glutamax (Gibco), and FUDR (2 mg/ml, Sigma, St. Louis, MO).

In vitro electrophysiology: Hippocampal cultures grown on coverslips were transduced at 4 div with titer-matched viruses for all CaMKIIα-NpHR-EYFP constructs (final dilution 104 i.u. /ml in neuronal growth media) and allowed to express for 10 days. Whole-cell patch clamp recordings were performed as previously described (intracellular solution: 129 mM K-gluconate, 10 mM

HEPES, 10 mM KCl, 4 mM MgATP, 0.3 mM Na3GTP, titrated to pH 7.2; extracellular solution, tyrode: 125 mM NaCl, 2 mM KCl, 3 mM CaCl2, 1 mM MgCl2, 30 mM glucose, and 25 mM HEPES, titrated to pH 7.3). Light (7 mW/mm2) was delivered from a 300W DG-4 lamp (Sutter Instruments, Novato, CA) through a 593nm ± 20nm filter (Semrock, Rochester, NY) and a 20X/0.45NA air objective (Olympus, Center Valley, PA).

Immunostaining and aggregate count: Primary hippocampal cultures grown on coverslips were infected at 4 div with titer matched virus (final dilution 104 i.u. /ml in neuronal growth media). At 14 div cultures were fixed for 30 min with ice-cold 4% paraformaldehyde and then permeabilized for 30 min with 0.4% saponin in 2% normal donkey serum (NDS). Primary antibody incubations were performed overnight at 4°C using a monoclonal marker of endoplasmic reticulum recognizing endogenous ER-resident proteins containing the KDEL retention signal(KDEL 1:200, Abcam, Cambridge, MA). For detection we used Cy3-conjugated secondary antibodies (Jackson Laboratories, West Grove, PA) in 2% NDS for 1 hour at room temperature. Close-up images of neurons were taken on a Leica confocal microscope using a 63X/1.4NA oil objective. The percentage of cells with aggregates was estimated by an unbiased count over multiple fields and coverslips.

Stereotactic injection into the rodent brain: Adult C57BL/6 mice were housed according to the Laboratory Vertebrate Animal protocols at Stanford. All surgeries were performed under aseptic conditions. The animals were anesthetized with intraperitoneal injections of ketamine (80 mg/kg)/xylazine (15-20 mg/kg) cocktail (Sigma). The head was shaved, cleaned with 70% ethanol and betadine and then placed in a stereotactic apparatus (Kopf Instruments, Tujunga, CA; Olympus stereomicroscope). Ophthalmic ointment was applied to prevent eye drying. A midline scalp incision was made and then a small craniotomy was performed using a drill mounted on the stereotactic apparatus (Fine Science Tools, Foster City, CA). The virus was delivered using a 10µl syringe and a thin 34 gauge metal needle; the injection volume and flow rate (1µl at 0.1µl/min) was controlled

with an injection pump from World Precision Instruments (Sarasota, FL). After injection the needle was left in place for 10 additional minutes and then slowly withdrawn. The skin was glued back with Vetbond tissue adhesive. The animal was kept on a heating pad until it recovered from anesthetic. Buprenorphine (0.03 mg/kg) was given subcutaneously following the surgical procedure to minimize discomfort. For hippocampal slice imaging: 1µl of concentrated lentivirus (1010 i.u. /ml) carrying NpHR or eNpHR under the CaMKIIα promoter was microinjected into the CA1 region of the left and right adult mouse hippocampus, respectively (anteroposterior, -2.0 mm from bregma; lateral, ± 1.5 mm; ventral, 2 mm). For in vivo electrophysiology: 1µl of eNpHR (1010 i.u. /ml) virus was injected in the adult mouse thalamus (anteroposterior -1.8, mm from bregma; lateral, 1.5 mm; ventral, 3.5 mm).

Slice preparation and confocal imaging: For preparation of brain slices, mice were sacrificed 10 days after viral injection. Acute coronal brain slices (250 µm) were prepared in ice-cold cutting solution (64 mM NaCl, 25 mM NaHCO3, 10 mM glucose, 120 mM sucrose, 2.5 mM KCl, 1.25 mM

NaH2PO4, 0.5 mM CaCl2, 7 mM MgCl2, and equilibrated with 95% O2/5% CO2) using a vibratome (VT1000S, Leica). The slices were then fixed for one hour in 4% paraformaldehyde, washed with PBS and mounted on microscope slides. Single confocal optical sections through the CA1 region or thalamus were acquired using a 40X/1.4NA oil objective on a Leica confocal microscope.

In vivo multiunit recordings: Simultaneous optical stimulation and electrical recording in living mice was done as described previously (Gradinaru, Thompson et al. 2007) using an optrode composed of an extracellular tungsten electrode (1MΩ, ~125µm) tightly attached to an optical fiber (~200µm, ThorLabs, Newton, NJ) with the tip of the electrode deeper (~0.3mm) than the tip of the fiber, to ensure illumination of the recorded neurons. The fiberoptic was coupled to a 561 nm laser diode from CrystaLaser (Reno, NV). Single unit recordings were done in animals anesthetized with intraperitoneal injections of ketamine (80 mg/kg)/xylazine (15-20 mg/kg) cocktail (Sigma). pClamp 10 and a Digidata 1322A board (Axon Instruments, Sunnyvale, CA) were used to both collect data and generate light pulses through the fiber. The recorded signal was band pass filtered at 300Hz low/5 kHz high (1800 Microelectrode AC Amplifier, A-M Systems). For precise placement of the fiber/electrode pair, stereotactic instrumentation (Kopf; Olympus stereomicroscope) was used. Immediately after recordings the animal was sacrificed and brain slices were prepared as described above to check for opsin expression and accurate placement of the optrode.

Chapter 4

Optical Deconstruction of Parkinsonian Neural Circuitry

Abstract | Deep brain stimulation (DBS) is a therapeutic option for intractable neurological and psychiatric disorders, including Parkinson’s disease and major depression. Due to the nonspecificity of electrodes, it has been challenging to elucidate the relevant target cell types or underlying mechanisms of DBS. We employed optogenetics and solid-state optics to systematically drive or inhibit an array of distinct circuit elements in freely moving Parkinsonian rodents, and found that therapeutic effects within the subthalamic nucleus can be accounted for by direct selective stimulation of afferent axons projecting to this region. In addition to providing insight into DBS mechanisms, these results demonstrate an optical approach for dissection of disease circuitry, and define the technological toolbox needed for systematic deconstruction of disease circuits by selectively controlling individual components.

Introduction

Parkinson’s disease (PD) is a debilitating neurodegenerative disorder resulting from the loss of dopaminergic (DA) neurons in the substantia nigra pars compacta (SNc), leading to abnormal neuronal activity in the basal ganglia (BG). DA depletion in the BG leads to altered activity of the subthalamic nucleus (STN) and globus pallidus pars interna (GPi), which has been linked to clinical deficits in movement initiation and execution (Albin, Young et al. 1989; Alexander and Crutcher 1990; DeLong 1990). Based on these observations from animal models, and the fact that lesions of the BG can be therapeutic in PD, high-frequency (>90 Hz) stimulation (HFS) of the STN (deep brain stimulation or DBS) has emerged as a highly effective treatment for medically-refractory PD (Benabid, Pollak et al. 1994; Limousin, Pollak et al. 1995; Rezai, Machado et al. 2008).

Exactly how DBS exerts its therapeutic effects is a matter of controversy (Dostrovsky and Lozano 2002; Vitek 2002; McIntyre, Savasta et al. 2004; Liu, Postupna et al. 2008) for three major reasons. First, due to the heterogeneity of brain tissue (Gross and Rolston 2008), it is unclear which circuit elements are responsible for the therapeutic effects. Second, HFS is intrinsically a complicated manipulation because target neurons can respond with increased, decreased, or mixed temporal patterns of activity; as a result the magnitude and even the sign of target cell responses to DBS are unknown. Finally, it is difficult to assess the net outcome of DBS on overall activity in the target cells and region, because electrical stimulation creates artifacts that prevent direct observation of local circuit responses during HFS itself. Together these challenges point to the need to understand, improve, and generalize (Mayberg, Lozano et al. 2005; Ressler and Mayberg 2007; Lozano, Mayberg et al. 2008; McNeely, Mayberg et al. 2008) this important treatment modality.

We have developed and employed optogenetics technology based on single-component microbial light-activated transmembrane conductance regulators and fiberoptic/laser diode- based in vivo light delivery (Boyden, Zhang et al. 2005; Adamantidis, Zhang et al. 2007; Aravanis, Wang et al. 2007; Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007; Zhang, Aravanis et al. 2007; Zhang, Wang et al. 2007; Gradinaru, Thompson et al. 2008; Zhang, Prigge et al. 2008). The channelrhodopsins, including VChR1 (Zhang, Prigge et al. 2008) and ChR2 (Boyden, Zhang et al. 2005), encode light-activated cation channels which can be expressed in neurons under cell type-specific promoters. In contrast, the halorhodopsins (e.g. NpHR) are light-activated Cl- pumps, and NpHR-expressing neurons are hyperpolarized and inhibited from firing action potentials when exposed to 590nm light in intact neural tissue (Zhang, Wang et al. 2007; Gradinaru, Thompson et al. 2008). The ChR2/NpHR system is ideally suited to dissect PD circuitry due to three features that map well onto the challenges outlined above. First, optogenetics allows genetically-targeted photosensitization of individual circuit components within the STN area and therefore testing of hypotheses regarding the causal role of individual cell types. Second, inhibition or excitation of target cells by direct hyperpolarization or depolarization can be achieved (McIntyre and Grill 2002), reducing complications such as soma-axon decoupling (McIntyre, Grill et al. 2004) in which cell bodies can be inhibited and axons stimulated by HFS. Third, optogenetics allows simultaneous optical control and electrophysiological recording of local neuronal activity in vivo with integrated fiber-electrode “optrodes,” avoiding electrical stimulus artifacts which may mask crucial neural responses.

In all of this, optogenetics maintains the millisecond temporal precision of electrodes. Therefore, optogenetics in principle could be employed to systematically probe specific circuit elements with defined frequencies of true excitation or inhibition in freely behaving parkinsonian rodents.

FIGURE 8 | Direct optical inhibition of local STN neurons. (A) Cannula placement, virus injection, and fiber depth were guided by recordings of the STN, which is surrounded by the silent zona incerta (ZI) and internal capsule (IC). (B) Confocal images of STN neurons expressing CaMKIIα::eNpHR- EYFP and labeled for excitatory neuron-specific CaMKIIα (right). (C) Continuous 561nm illumination of the STN expressing CaMKIIα::eNpHR-EYFP in anesthetized 6-OHDA rats reduced STN activity; representative optrode trace and amplitude spectrum shown. Mean spiking frequency was reduced from 29 ± 3 Hz to 5 ± 1 Hz (mean ± s.e.m., p < 0.001, Student’s t-test, n = 8 traces from different STN coordinates in 2 animals). (D) Amphetamine-induced rotations were not affected by stimulation of the STN in these animals (p > 0.05, n = 4 rats, t-test with µ = 0). The red arrow indicates direction of pathologic effects, while the green arrow indicates direction of therapeutic effects. The electrical control implanted with a stimulation electrode showed therapeutic effects with HFS (120-130 Hz, 60 µs pulse width, 130-200 µA, p < 0.05, t-test with µ = 0). Percent change of -100% indicates that the rodent is fully corrected. Data in all figures are mean ± s.e.m. ns p > 0.05, * p < 0.05, ** p < 0.01, *** p < 0.001.

Results

Optical inhibition of STN

To first address the most widely-held hypothesis in the field, we asked if direct, reversible, bona fide inhibition of local-circuit excitatory STN neurons would be therapeutic in PD. The STN measures <1mm3 in rats (Hamani, Saint-Cyr et al. 2004), but targeting accuracy can be aided by extracellular recordings during opsin vector introduction, since STN is characterized by a particular firing pattern which is distinguishable from bordering regions (Figure 8A, Figure S1C).

The STN is a predominantly excitatory structure (Smith and Parent 1988) embedded within an inhibitory network. This anatomical arrangement enables a targeting strategy for selective STN inhibition (Figure 8B), in which eNpHR (Gradinaru, Thompson et al. 2008) is expressed under control of the CaMKIIa promoter, selective for excitatory glutamatergic neurons and not inhibitory cells, fibers of passage, glia, or neighboring structures (Aravanis, Wang et al. 2007). In this way true optical inhibition is targeted to the dominant local neuron type within STN.

Optical circuit interventions were tested in rats that had been made hemiparkinsonian by injection of 6-hydroxydopamine (6-OHDA) unilaterally into the right medial forebrain bundle (MFB). Loss of nigral dopaminergic cells following 6-OHDA administration was confirmed by decreased tyrosine hydroxylase levels unilaterally in the substantia nigra pars compacta

(Fig. S1A). These hemiparkinsonian rodents have specific deficits in contralateral (left) limb use and display (rightward) rotations ipsilateral to the lesion, which increase in frequency when the subjects are given amphetamine to facilitate functional evaluation, and decrease in frequency upon treatment with dopamine agonists (Metz, Tse et al. 2005) or following electrical DBS (Figure 8D, right). This amphetamine-induced rotation test is widely used for identifying treatments in hemiparkinsonian rodents, which can be complemented with other behavioral assays such as locomotion, climbing, and head position bias. To directly inhibit the excitatory STN neurons, we delivered lentiviruses carrying eNpHR under the CaMKIIα promoter to the right STN of the hemiparkinsonian rats. CaMKIIα::eNpHR-EYFP expression was specific to excitatory neurons (as shown by CaMKIIα and glutamate expression; Figure 8B, right; Figure S2A), robust (95.73% ± 1.96 s.e.m infection rate assessed in n = 220 CaMKIIα positive cells), and restricted to the STN (Figure 8B, left and middle). To validate the resulting physiological effects of optical control, a hybrid optical stimulation/electrical recording device (optrode) was employed in isoflurane-anesthetized animals to confirm that eNpHR was functional in vivo, potently inhibiting (>80%) spiking of recorded neurons in the STN (Figure 8C; Figure S4A, B; Figure S5A). This cell type-targeted inhibition was temporally precise and reversible, and extended across all frequency bands of neuronal firing (Figure 8C, Figure S7A).

For behavioral rotation assays in the hemiparkinsonian rats, the STN-targeted fiberoptic was coupled to a 561nm laser diode to drive eNpHR. Electrical DBS was highly effective at reducing pathological rotational behavior, but despite precise targeting and robust physiological efficacy of eNpHR inhibition, the hemiparkinsonian animals did not show even minimal changes in rotational behavior with direct true optical inhibition of the local excitatory STN neurons (Figure 8D). In addition, there was no effect on path length and head position bias in response to light during these experiments (see supplementary methods). While muscimol and lidocaine administration to the region of the STN in monkeys and rodents can relieve Parkinsonian symptoms (Levy, Lang et al. 2001), the data in Figure 8 show that the more specific intervention of selectively decreasing activity in excitatory local neurons of the STN appeared not sufficient by itself to affect motor symptoms.

Another possibility is that DBS could be driving secretion of glial modulators which would have the capability to modulate local STN circuitry; this would be consistent with recent findings (Bekar, Libionka et al. 2008) indicating that a glial-derived factor (adenosine)

accumulates during DBS and plays a role in DBS-mediated attenuation of thalamic tremor. Indeed, the STN expresses receptors for glia-derived modulators (Rivkees, Price et al. 1995) which can inhibit postsynaptic currents in the STN (Shen and Johnson 2003). ChR2 presents an interesting possibility for recruitment of glia; when opened by light, in addition to Na+ and K+ ions, ChR2 can also pass trace Ca2+ currents (Nagel, Szellas et al. 2003; Zhang and Oertner 2007) that trigger Ca2+ waves in and activate ChR2-expressing astroglia (Wang and Deisseroth unpublished observations). We employed a GFAP promoter to target ChR2 to local astroglia, validated with GFAP and S100β staining (Figure 9A, Figure S2B). Optrode recordings revealed that blue light stimulation of STN following transduction with GFAP::ChR2 reversibly inhibited neuronal firing in the STN (Figure 9B, Figure S3A), with variable delay on the timescale of seconds. However, recruiting astroglial cells by this mechanism was not sufficient to cause even trace responses in motor pathology in parkinsonian rodents (Figure 9C, Figure S3B). Path length and head position bias were also not affected by light during these experiments (see supplementary methods). While these data do not exclude the importance of local STN inhibition as a contributing factor in DBS response, as not all STN neurons may be affected in the same way by indirect glial modulation, the direct activation of local glial cells appeared not sufficient to treat parkinsonian symptoms, pointing to other circuit mechanisms.

FIGURE 9 | Targeting astroglia within the STN. (A) Confocal images show STN astrocytes expressing GFAP::ChR2-mCherry, costained with GFAP (right). (B) 473nm illumination of the STN expressing GFAP::ChR2-mCherry in anesthetized 6-OHDA rats. Optrode recording revealed that continuous illumination inhibited STN activity with 404 ± 39 ms delay to onset and 770 ± 82 ms delay to offset (n = 5 traces from different STN coordinates in 2 animals), while 50% duty cycle also inhibited spiking, with delay to onset of 520 ± 40 ms and delay to offset of 880 ± 29 ms (n = 3 traces from different STN coordinates in 2 animals) with p < 0.001. (C) Amphetamine-induced rotations were not affected by 50% duty cycle illumination in these animals (right, p > 0.05, n = 7 rats, t-test with µ = 0).

Optogenetic excitation of targeted STN cells

Network oscillations at particular frequencies could play important roles in both PD pathology and treatment. For example, PD is characterized by pathological levels of beta oscillations in the basal ganglia, and synchronizing STN at gamma frequencies may ameliorate PD symptoms while beta frequencies may worsen symptoms. Because simple inhibition of excitatory cell bodies in the STN did not affect behavioral pathology, and since high- frequency stimulation (HFS: 90-130 Hz) is used for electrical DBS, we used ChR2 to drive high-frequency oscillations in this range within the STN. We injected CaMKIIα::ChR2 into the STN (Figure 10A) and used pulsed illumination with a 473nm laser diode to activate excitatory neurons in the STN (Figure 10B, Figure S5B) during behavioral testing in parkinsonian rodents (Figure 10C, Figure S3C). Despite robust effects on high-frequency power of neuronal spike rate in STN of anesthetized animals (Figure S7B), HFS delivered locally to the STN area failed to affect PD behavioral symptoms (path length and head position bias were unchanged by light - see supplementary methods). Animals tested in parallel with beta frequency pulses also showed no behavioral response, indicating that (while not excluding a contributory role) directly generated oscillations within the STN excitatory neurons are not sufficient to account for therapeutic effects.

FIGURE 10 | Optical depolarization of STN neurons at different frequencies. (A) Confocal images of STN neurons expressing CaMKIIα::ChR2-mCherry and labeled for the excitatory neuron specific CaMKIIα marker. (B) Optical HFS (120 Hz, 5 ms pulse width) of the STN expressing CaMKIIα::ChR2-mCherry in 6-OHDA rats recorded with the optrode connected to a 473nm laser diode (representative trace and amplitude spectrum shown). Frequency of spiking increased from 41 ± 2 Hz to 85 ± 2 Hz (HFS vs. pre, n = 5 traces: p < 0.001, t-test, post, n = 3 traces; traces were sampled from

different STN coordinates in 1 animal). (C) Amphetamine-induced rotations were not affected by high (left, 130 Hz, n = 5 rats) or low (middle, 20 Hz, n = 2 rats) frequency optical stimulation.

Measuring the volume of tissue recruited in STN

We have previously measured in cortical and hypothalamic tissue the propagation of blue light in the setting of laser diode-fiberoptic illumination; we observed that substantial tissue volumes (comparable to that of the STN) could reliably be recruited at a light power density sufficient to drive physiologically significant microbial opsin currents (Adamantidis, Zhang et al. 2007; Aravanis, Wang et al. 2007). It was important to repeat and extend these measurements to the PD setting. First, we confirmed that the propagation measurements of blue light (473 nm) in brain tissue represent a lower bound on the volume of tissue recruited, due to reduced scattering of lower-energy photons delivered from the 561nm laser diode; therefore sufficient light power is present to activate opsins within 1.5mm of the fiber for either wavelength of light (Figure 11A). We next extended these findings with a functional assay for tissue recruitment under conditions mimicking our behavioral experiments (Figure 11B,C). After an in vivo optical stimulation paradigm targeted to the CaMKIIα::ChR2 expressing STN in freely moving rats, we performed immunohistochemistry for c-fos, a biochemical marker of neuronal activation. We observed robust c-fos activation in STN (Figure 11B) over a widespread volume (Figure 11C); indeed, as expected from our light scattering measurements and tissue geometry, we found that at least 0.7 mm3 of STN is recruited by light stimulation, closely matching the actual volume of the STN (Figure 11C). Therefore, light penetration was not limiting since the entire STN is recruited by the optical modulation paradigms of Figures 8-10.

FIGURE 11 | Quantification of the tissue volume recruited by optical intervention. (A) Intensity values for 473nm (blue) and 561nm (yellow) light are shown for a 400µm fiber as a function of depth in brain tissue. The dashed line at 1 mW/mm2 (30 mW light source) indicates the minimum intensity required to activate channelrhodopsins and halorhodopsins (Zhang, Aravanis et al. 2007; Zhang, Wang et al. 2007). (B) Confocal images of STN neurons expressing CaMKIIα::ChR2-mCherry and labeled for the immediate early gene product c-fos show robust neuronal activation produced by light stimulation in vivo. Arrowheads indicate c-fos positive cells. Freely moving rats expressing ChR2 in STN (same animals as in Figure 10), were stimulated with 473nm light (20Hz, 5ms pulse width). (C) The STN volume that showed strong c-fos activation was estimated to be at least 0.7 mm3 (dashed lines indicate STN boundaries); robust c-fos activation was observed medial-lateral (1.155 mm), anterior-posterior (0.800 mm), and dorsal-ventral (0.770 mm) on subthalamic slices imaged by confocal microscopy with DAPI counterstain.

Optical control of afferent axons in STN

Therapeutic effects could arise from driving axonal projections that enter the STN, as DBS electrodes will potently modulate not just local cells and their efferents, but also afferent fibers. Optogenetics discriminates these two possibilities, as the lentiviruses transduce somata without transducing afferent axons (Gradinaru, Thompson et al. 2007). To assess the possibility that PD motor behavioral responses are modulated by targeting afferent projections to the STN, we used Thy1::ChR2 transgenic mice (Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007) in which ChR2 is expressed in projection neurons, and we verified that in Thy1::ChR2 line 18, ChR2-YFP is excluded from cell bodies in the STN but is abundant in afferent fibers (Figure 12A).

We conducted optrode recordings in anesthetized 6-OHDA mice (Figure S1B) (Matsuya, Takuma et al. 2007; Tabar, Tomishima et al. 2008) to assess local effects on STN physiology of driving afferent axons selectively, and found frequency-dependent effects (Figure 12B). First, we observed that HFS of afferent fibers to the STN potently reduced STN spiking across all frequency bands; this effect did not completely shut down local circuitry, as low-amplitude high-frequency oscillations persisted during stimulation (Figures 12B; Figures S4C, D; Figures S5C, D). Next, we found that LFS of afferent fibers increased beta-frequency firing in the STN without affecting endogenous bursting (Figure 12B, Figure S7E). We next assessed the impact of these specific interventions on PD behavior in 6-OHDA mice, and for the first time among the optogenetic interventions, we observed marked effects. Driving STN afferent fibers with HFS robustly and reversibly ameliorated PD symptoms, measured by rotational behavior and head position bias (Figure 12C). The HFS effects were not subtle; indeed, in nearly every case these severely parkinsonian animals were restored to behavior indistinguishable from normal, and in every case the therapeutic effect immediately and fully reversed, with return of ipsilateral rotations upon discontinuation of the light pulse paradigm. Notably, treated animals could freely switch directions of movement and head position from left to right and vice versa. In striking contrast with optical HFS, optical LFS (20 Hz) of the same afferent fibers worsened PD symptoms by driving increased ipsilateral rotational behavior (Figure 12C), demonstrating that behavioral effects seen do not result from simply driving unilateral activity. Therefore, in contrast to direct STN cellular interventions, driving STN afferent fibers with HFS and LFS differentially modulated PD symptoms in a manner

corresponding to frequencies of stimulation linked clinically to ameliorated or exacerbated PD symptoms.

FIGURE 12 | Selective optical control of afferent fibers in the STN. (A) Confocal images of Thy1::ChR2-EYFP expression in the STN and DAPI staining for nuclei shows selective expression in fibers and not cell bodies (right). (B) Optical HFS (130 Hz, 5 ms pulse width) of the STN region in an anesthetized Thy1::ChR2-EYFP 6-OHDA mouse with 473nm light inhibited STN large-amplitude spikes (sample trace, top left), while inducing smaller-amplitude high-frequency oscillations (Figure

S4C, D; S5C, D). Optical LFS (20 Hz, 5 ms pulse width) produced reliable spiking at 20 Hz (bottom left). While HFS prevented bursting (top right, p < 0.001, n = 3), LFS had no significant effect on burst frequency by 2 sample t-test (p > 0.05, n = 3 traces) nor on spikes/burst (bottom right, p > 0.05, n = 3 traces). (C) Optical HFS to STN in these animals (left, 100-130 Hz, 5 ms, n = 5 mice) produced robust therapeutic effects, reducing ipsilateral rotations and allowing animals to freely switch directions. In contrast, optical LFS (second left, 20 Hz, 5 ms, n = 5 mice) exacerbated pathologic effects, causing increased ipsilateral rotations. Both effects were reversible (post). Changes were significant by t-test with µ = 0 with both HFS (p < 0.001, n = 5 mice) compared to baseline (light off) and LFS (p < 0.05, n = 5 mice). (F) Contralateral head position bias also showed robust correction with HFS by 2 sample t- test (HFS vs. light off: p < 0.05; n = 2 mice), but not with LFS (LFS vs. light off: p > 0.05, n = 2 mice).

Optical control of layer V motor cortex projection neurons

A diverse array of fibers from widespread brain areas converge on the STN, perhaps underlying the utility of the STN as a focal DBS target. Many of these afferents likely contribute together to the therapeutic effects, and it is unlikely that a single source of fibers completely accounts for the behavioral effects seen. However, we explored these afferents in greater detail to determine the general class of fibers that may be contributory.

Thy1::ChR2 animals display ChR2 expression chiefly in excitatory projection neurons (Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007). Indeed, the inhibitory markers GAD67 and GABA were not detectable in Thy1::ChR2 fibers within STN (Figure 13A, left), effectively ruling out contributions from the GABAergic pallidal projections (LGP/GPe). We also found no localization of major neuromodulatory markers (dopamine and acetylcholine) within the STN Thy1::ChR2 fibers (Figure S2C), ruling out dopaminergic SNr as a relevant fiber origin as well. We next explored possible sources of excitatory fibers, and found no expression of ChR2-YFP in the cell bodies of the excitatory parafascicular or pedunculopontine nuclei, potential contributors of excitatory fibers to the STN. Within neocortex of these mice, however, ChR2-YFP is expressed strongly in excitatory neurons that project to STN (Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007). Since pathologically strong connectivity between STN and primary motor cortex M1 has been suggested to underlie PD circuit dysfunction (Afsharpour 1985; Degos, Deniau et al. 2008), we therefore explored M1 as a possible contributor.

We verified in Thy1::ChR2 M1 the presence of strong and selective ChR2 expression largely restricted to layer V neurons and corresponding apical dendrites (Arenkiel, Peca et al. 2007;

Wang, Peca et al. 2007) but not in cells within other layers (Figure 13A, right). To probe the functional connectivity between these layer V projection neurons and STN in the PD animals, we conducted a separated-optrode experiment in anesthetized animals in which the fiberoptic and recording electrodes were placed in two different brain regions in Thy1::ChR2 animals (Figure 13B). By driving M1 layer V projection neurons and simultaneously recording in both M1 and STN, we found that precise M1 stimulation of this kind potently influenced neural activity in the STN (Figure 13C, Figures S7C, D), and that M1 Layer V neurons could be antidromically recruited by optical stimulation in the STN (Figure S6). While as noted above, many local afferents in the STN region, including from the ZI, are likely to underlie the complex therapeutic effects of DBS, functional influences between M1 layer V and STN could be a significant contributor. Indeed, we found that selective M1 layer V HFS optical stimulation sufficed to ameliorate PD symptoms in a similar manner to STN stimulation in an array of measures ranging from rotational behavior (Figure 13D) to head position bias and locomotion (Figures 13E, F). As with STN stimulation, pathological rotations and head position bias were reduced by optical HFS to M1; in contrast, while not augmenting the pathology, optical 20 Hz (LFS) stimulation to M1 had no therapeutic effect (Figures 13D, E, F), and even at the highest light intensities achievable without epileptogenesis, M1 LFS did not drive or modify rotational behavior. Finally, increased functional mobility with M1 HFS but not LFS was confirmed with quantification of increased distance and speed of locomotion in bradykinetic PD Thy1::ChR2 mice (Figures 13F).

FIGURE 13 | Selective optical stimulation of layer V neurons in anterior primary motor cortex. (A) GAD67 and GABA staining showed no colocalization with Thy1::ChR2-EYFP in STN (left). Apical dendrites of layer V neurons can be seen rising to the pial surface (Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007) (right). (B) Schematic for optical stimulation of M1 with simultaneous recording in STN of Thy1::ChR2 mice. (C) Optical stimulation (473nm) of M1 and simultaneous recording in STN of anesthetized Thy1::ChR2 mice. Optical HFS (130 Hz, 5 ms pulse width) of M1 modulated activity in both M1 and STN. Optical LFS (20 Hz, 5 ms) of M1 produced 20 Hz tonic firing.

(D) Optical HFS (130 Hz, 5 ms pulse width) reduced amphetamine-induced ipsilateral rotations in 6- OHDA Thy1::ChR2 mice (p < 0.01, n = 5 mice) in contrast to optical LFS (20 Hz, 5 ms pulse width, p > 0.05, n = 4 mice); t-test with µ = 0. (E) Contralateral head position bias was corrected in HFS (HFS vs. light off: p < 0.001, n = 4 mice), while LFS had little effect (LFS vs. light off: p > 0.05, n = 3 mice); 2-sample t-test. (F) HFS but not LFS to M1 significantly increased path length (HFS, p < 0.01, n = 2 mice) and climbing (p < 0.05, n = 3 mice); 2-sample t-test. Sample paths before, during, and after HFS are shown (100 seconds each, path lengths noted in cm).

Discussion

A major promise of optogenetics has been the potential for dissection of disease circuitry and treatment mechanisms. Here we demonstrate that this potential can be realized. Systematically targeting different elements of the disease circuit, we implicate direct frequency-dependent effects on afferents to the subthalamic nucleus region as a major direct target of deep brain stimulation in Parkinson’s Disease.

Cortical-STN interactions have been previously considered in PD; indeed, motor cortex activity may be elevated in PD and reduced in PD treatment (Ridding, Inzelberg et al. 1995; Payoux, Remy et al. 2004), and abnormal oscillatory activity may occur between the cortex and the basal ganglia in PD patients during movement (Brown, Oliviero et al. 2001; Levy, Lang et al. 2001; Kringelbach, Jenkinson et al. 2007). Cortical stimulation could restore balance to Parkinsonian circuitry that is overly devoted to one kind of activity to the exclusion of others (Degos, Deniau et al. 2008), by either disrupting the pathological activity pattern (e.g. low-frequency bursting), by promoting throughput of patterns encoding motor behaviors oppositional or compensatory to the lesion pathology, or both. Cortical stimulation in human beings has been a subject of interest and controversy (Fregni, Simon et al. 2005), with some studies showing promising results for PD treatment (Canavero and Paolotti 2000; Canavero, Paolotti et al. 2002; Pagni, Zeme et al. 2003; Drouot, Oshino et al. 2004; Lefaucheur, Drouot et al. 2004; Khedr, Rothwell et al. 2006) and others less supportive (Cilia, Landi et al. 2007; Strafella, Lozano et al. 2007; Arle, Apetauerova et al. 2008). Our data, in implicating deep layer V neurons as sufficient targets in primary motor cortex, may help address these issues by informing the design of cortical interventions with regard to subdural rather than superficial extradural stimulation. Even with subdural stimulation, optimal cortical stimulation in patients will certainly face particular challenges due to broad cortical representation, and identifying the cortical subregion most functionally connected to STN will be important. Clinical

translation of these concepts will benefit from ongoing work in animal models to facilitate rapid mapping of stimulus space for identification of optimal pulse patterns and duty cycles.

It is important to note that these findings do not exclude other important contributions to DBS targets or disease symptoms. Not only are other afferents to the STN potential upstream factors in STN-initiated therapeutic effects, but all STN-initiated effects will be implemented downstream through the interconnected basal ganglia, cortical, brainstem, and thalamic motor pathways with many potential nodes for intervention. Both the disease and treatment are extraordinarily complex; the fact that DBS can improve many PD symptoms, including tremor, rigidity, and bradykinesia, but not others such as speech impairment, depression, and dementia, points to the need for ongoing work to map and functionally interrogate disease circuitry beyond the brain regions investigated here. DBS can also encounter limitations as a therapy even for the symptoms which typically respond. These issues may be linked to cell type-specific responses, suggesting that disease model investigations will be greatly facilitated by the optogenetic approach.

Axon tract modulation with high temporal precision could turn out to be a common theme in DBS (McIntyre, Grill et al. 2004; Mayberg, Lozano et al. 2005; Ressler and Mayberg 2007; Lozano, Mayberg et al. 2008; McNeely, Mayberg et al. 2008), as these collections of fibers represent compact nodes for accessing activity converging from a broader area. Even without detailed knowledge of the relevant neural code, simple alterations in the propagation of activity through white matter tracts or disrupting circuit loops could represent final common pathways for disease and treatment (Airan, Meltzer et al. 2007). However, for PD as well as for other neurological and psychiatric diseases, maintaining high temporal precision of the circuit interrogation technology will be crucial, because as illustrated here, fundamentally different effects are seen when driving the same cell type at different temporal frequencies. Indeed, themes of synchrony and oscillations driven by particular cell and fiber types will likely be common to other brain stimulation-responsive diseases such as depression and epilepsy, underscoring the importance of understanding and generalizing deep brain stimulation.

Supplementary Material

Methods

Immunohistochemistry and imaging: To verify the phenotype of cells and measure c-fos activity, rodents were anaesthetized with 65 mg/kg sodium pentobarbital and transcardially perfused with ice-cold 4% paraformaldehyde (PFA) in PBS (pH 7.4). Brains were fixed overnight in 4% PFA and then equilibrated in 30% sucrose in PBS. 40 µm-thick coronal sections were cut on a freezing microtome and stored in cryoprotectant at 4ºC until processed for immunohistochemistry. Free-floating sections were washed in PBS and then incubated for 30 min in 0.3% Triton X-100 and 3% normal donkey serum (NDS). Slices were incubated overnight with primary antibody in 0.01% Tx100 and 3% NDS (rabbit anti-cfos 1:500, rabbit anti-GFAP 1:500, mouse anti-MAP2 1:500, mouse anti-GAD67 1:500, rabbit anti-GABA 1:200, mouse anti-vGlut1 1:500, mouse anti-vGlut2 1:500, mouse anti-CaMKIIα 1:200, mouse anti-S100β 1:250, rabbit anti-glutamate 1:200, chicken anti-tyrosine hydroxylase 1:500, and goat anti-choline acetyltransferase 1:200). Sections were then washed and incubated with secondary antibodies (1:1000) conjugated to FITC, Cy3 or Cy5 for 3 hrs at room temperature. Following a 20 min incubation with DAPI (1:50,000) sections were washed and mounted on microscope slides with PVA-DABCO.

Confocal fluorescence images were acquired on a scanning laser microscope using a 20X/0.70NA or a 40X/1.25NA oil immersion objective. To determine the volume of c-fos activation, serial stack images covering a depth of 20 µm through multiple medial-lateral, anterior-posterior and dorsal–ventral subthalamic sections were acquired using equivalent settings. The image analysis software calculated the number of c-fos positive cells per field by thresholding c-fos immunoreactivity above background levels and using the DAPI staining to delineate nuclei. To determine the rate of viral transduction we determined the percentage of CaMKIIα-immunoreactive neurons per 40X field that were also eNpHR-YFP positive in multiple serial stack images of subthalamic sections. Large field images of entire slices were collected on a Leica MZ16FA stereomicroscope.

Lentivirus production and transduction: Lentiviral vectors carrying the genes used were constructed using standard cloning techniques. The CaMKIIα::eNpHR construct was produced by PCR amplification of the eNpHR-EYFP construct previously published (Gradinaru, Thompson et al. 2008) and cloned in-frame into the AgeI and EcoRI restriction sites of a

lentivirus carrying the CaMKIIα promoter. The CaMKIIα::ChR2 construct was produced by PCR amplification of the ChR2-mCherry construct and was also cloned in-frame into the AgeI and EcoRI restriction sites of a lentivirus carrying the CaMKIIα promoter. The GFAP::ChR2 construct was produced by replacing the CaMKIIα promoter with the GFAP promoter in the CaMKIIα::ChR2-mCherry construct using the AgeI and PacI restriction enzyme sites. The map for the CaMKIIα::ChR2, CaMKIIα::eNpHR, is available online at www.optogenetics.org.

High titer lentivirus (>109 pfu/mL) was then produced via calcium phosphate co-transfection of 293FT cells with the lentiviral vector, pCMVΔR8.74 and pMD2.G (Zhang, Wang et al. 2007). 24 h post-transfection, 293FT cells were switched to serum-free medium containing 5 mM sodium butyrate; the supernatant was collected 16 h later and concentrated by ultracentrifugation at 50,000 × g with 20% sucrose cushion. The resulting viral pellet was resuspended in phosphate buffered saline at 1/1000th of the original volume.

Adeno-associated virus production and transduction: To ensure that there would be no significant expression leak in non-targeted cell types, we employed a Cre-inducible AAV vector with a double-floxed inverted open reading frame (ORF), wherein the ChR2-EYFP sequence is present in the antisense orientation. Upon transduction, Cre-expressing cells invert the ChR2-EYFP ORF in a stable, irreversible fashion and thereby activate sustained ChR2- EYFP expression under control of the strong and constitutively active elongation factor 1α (EF-1α) promoter (Feng Zhang, unpublished results). To construct Cre-activated recombinant AAV vectors, the DNA cassette carrying two pairs of incompatible lox sites (loxP and lox2722) was synthesized and the ChR2-EYFP transgene was inserted between the loxP and lox2722 sites in the reverse orientation. The resulting double-floxed reverse ChR2-EYFP cassette was cloned into a modified version of the pAAV2-MCS vector carrying the EF-1α promoter and the Woodchuck hepatitis virus posttranscriptional regulatory element (WPRE) to enhance expression. The recombinant AAV vectors were serotyped with AAV5 coat proteins and packaged by the viral vector core at the University of North Carolina. The final viral concentration was 2 x 1012 genome copies (gc) / mL.

Stereotactic injection and cannula placement: Adult rats (female Fisher, 200–300 g) and mice (male and female, C57BL/6 background, 15–30 g) were the subjects of these experiments. Animal husbandry and all aspects of experimental manipulation of our animals

were in strict accord with guidelines from the National Institute of Health and have been approved by members of the Stanford Institutional Animal Care and Use Committee. All surgeries were performed under aseptic conditions. Rodents were anaesthetized using 1.5% isoflurane (for surgeries longer than 1 hr) or i.p. injection (90 mg/kg ketamine and 5mg/kg xylazine for rats; 80 mg/kg and 15-20 mg/kg, respectively, for mice). The top of the animal’s head was shaved, cleaned with 70% ethanol and betadine and then placed in a stereotactic apparatus. Ophthalmic ointment was applied to prevent eye drying. A midline scalp incision was made and then small craniotomies were performed using a drill mounted on the stereotactic apparatus for the 6-OHDA injection in the medial forebrain bundle (rat: -2 AP, 2 ML, -7.5 DV; mouse: -1.2 AP, 1.2 ML, -4.75 DV) and virus injection in the STN (rat: -3.6 mm AP, 2.5 mm ML; mouse: -1.9 mm AP, 1.7 mm ML).

For rodents that were injected with lentivirus in the STN, in vivo extracellular recording was used to accurately determine the location of the STN along the dorsal-ventral axis. The depth was around -7 mm in rats and -4 mm in mice. The concentrated lentivirus (described above) was delivered to the STN using a 10µl syringe and a thin 34 gauge metal needle; the injection volume and flow rate (3 sites with 0.6 µl each at 0.1 µl/min) was controlled with an injection pump. After the final injection, the needle was left in place for 10 additional minutes and then slowly withdrawn.

6-OHDA was then used to lesion the substantia nigra and produce hemi-Parkinsonian rodents. Desipramine (20mg/kg for rats; 10 mg/kg for mice; noradrenergic reuptake inhibitor to prevent damage to noradrenergic terminals) was administered, followed ~30 minutes later by 6-OHDA (8 μg/4 μl for rats; 6 μg/2 μl for mice) with 0.1% ascorbic acid (to prevent degradation of 6-OHDA) into the right medial forebrain bundle (rat: –2 AP , +2 ML, and -7.5 DV; mouse: -1.2 AP, +1.2 ML, and -4.75 DV). The perfusion for the 6-OHDA injection (rat: 4 uL, mouse 2 uL) was at the rate of 1.2 μl/min for 4 min, and the needle was left in situ for an additional 5 minutes.

A fiber guide (rat: C312G, mouse: C313G) was beveled to form a sharp edge (to more easily penetrate brain tissue and reduce tissue movement), and then inserted through the craniotomy to a depth of approximately 400 µm above the STN or the anterior primary motor cortex (mouse: 2 AP, 2 ML, 0.5 DV). One layer of adhesive cement followed by cranioplastic cement was used to secure the fiber guide system to the skull. After 20 min, the scalp was sealed back

using tissue adhesive. The animal was kept on a heating pad until it recovered from anesthesia. Buprenorphine (0.03 mg/kg) was given subcutaneously following the surgical procedure to minimize discomfort. A dummy cannula (rat: C312G, mouse: C313G) was inserted to keep the fiber guide patent.

For the control electrical DBS rodents, a stimulation electrode (MS303/3-B) was implanted in the STN. The procedure above was followed for OHDA injection, in vivo extracellular recording was then used to determine the depth of the STN, and the stimulation electrode was inserted to that depth and secured using one layer of adhesive cement followed by cranioplastic cement. Tissue adhesive was used to reseal the scalp, the animal was kept on a heating pad until recovery from anesthesia and buprenorphine was given to minimized discomfort. A dust cap (303DC/1) was then used to cover the electrode contacts.

In vivo optrode recordings: Simultaneous optical stimulation and electrical recording in a single region in living rodents was done as described previously (Gradinaru, Thompson et al. 2007) using an optrode composed of an extracellular tungsten electrode (1 Ω,M ~125 µm) tightly attached to an optical fiber (~200 µm) with the tip of the electrode deeper (~0.4 mm) than the tip of the fiber, to ensure illumination of the recorded neurons. For stimulation and recording in two distinct regions, small craniotomies were created above both target regions, and a fiber or optrode was placed above one region through one craniotomy and a plain electrode or optrode was placed in the other region through a separate craniotomy (see Figure 13B for diagram; the picture is modified from The Jackson Laboratory website: http://jaxmice.jax.org/support/Thy1-ChR2-9_WholeBrain4x.pdf). Stimulation in the anterior motor cortex was achieved by placing the optical fiber just above the brain surface, activating layer 5 of the cortex; for STN stimulation, the fiber was 300 µm above the STN. The STN was identified using its highly stereotyped firing pattern and the fact that it is surrounded dorso- ventrally by silent regions. The optical fiber was coupled to a 473 nm or 561 nm laser diode (30 mW fiber output) from CrystaLaser. Single unit recordings were done in rats anesthetized with 1.5% isoflurane and mice anesthetized with intraperitoneal injections of ketamine (80 mg/kg)/xylazine (15-20 mg/kg) cocktail. pClamp 10 and a Digidata 1322A board were used to both collect data and generate light pulses through the fiber. The recorded signal was band pass filtered at 300Hz low/5 kHz high (1800 Microelectrode AC Amplifier). For precise placement of the fiber/electrode pair, stereotactic instrumentation was used.

Behavior: For behavior, multimode optical fibers (NA 0.37; rat: 400 µm core, BFL37-400; mouse: 300 µm core, BFL37-300) were precisely cut to the optimal length for maximizing the volume of the STN receiving light. About one week before behavior, an extracellular recording electrode was used to determine the depth of the dorsal border of the STN from the tip of the cannula guide and fibers were cut to be 200-300 µm shorter. For anterior motor cortex stimulation, the fiber was placed above layer 5 (less than a millimeter deep). To ensure stability of the fiber during testing in moving animals, an internal cannula adapter was glued to the stripped optical fiber. To insert the fiber, rodents were briefly placed under isoflurane and the fiber was inserted while the animal was recovering from anesthesia. The internal cannula adapter snapped onto the cannula guide and the bottom half of the plastic portion of a dummy cannula was also used to ensure the adapter remained connected to the top of the cannula guide (Adamantidis, Zhang et al. 2007; Gradinaru, Thompson et al. 2007).

For optical stimulation, the fiber was connected to a 473 nm or 561 nm laser diode (20 mW fiber output) through an FC/PC adapter. Laser output was controlled using a function generator (33220A) to vary the frequency, duty cycle, and intensity. For Thy1::ChR2 animals, the average minimum intensity used to produce therapeutic behavior was 10 mW. A custom aluminum rotating optical commutator (Adamantidis, Zhang et al. 2007; Gradinaru, Thompson et al. 2007) was used to release torsion in the fiber caused by the animal’s rotation.

Motor behavior was assessed using amphetamine-induced rotations, head position bias, climbing, and track length. Animals were accepted for experimental investigation only if amphetamine reliably induced rotations in the ipsilateral direction confirming a 6-OHDA lesion of the substantia nigra. Before and after each stimulation trial, a trial of equal length with the light off was used as a control. Each of these trials was about 3 minutes long making the entire off-on-off sequence 9 minutes long. For amphetamine-induced behavior, amphetamine (rat: 2 mg/kg; mouse: 2.6 mg/kg) was injected 30 minutes before behavioral measurements; the fiber was inserted into the cannula and the rodent placed in an opaque, non-reflective cylinder (rat: diameter 25 cm, height 61 cm; mouse: diameter 20 cm, height 46 cm) 10 minutes before the behavioral experiments. Rotations ipsilateral to the 6-OHDA lesions (clockwise turns) were counted, and contralateral rotations were subtracted. The percentage change calculation considered the change in rotational bias relative to the period without stimulation. Head position bias was determined by counting the number of head tilts (>10º deviation left or right of midline) over time. Each time the rodent rose up and touched

either paw to the wall of the cylinder was counted as an instance of climbing. Track length was measured with Viewer. After the completion of behavior experiments, cannula placement was confirmed by slicing.

For experiments where optical stimulation did not produce a change in the rodent behavior, we collected path length and head position bias data while the rodents were under amphetamine. Continuous 561nm illumination of the STN expressing CaMKIIα::eNpHR-EYFP in 6-OHDA rats did not affect path length (cm/min; light on vs. light off: 757.05 ± 163.11 vs. 785.74 ± 157.56, p = 0.90, n = 4 rats; mean ± s.e.m; 2-sample t-test) or head position bias (% of time to the right; light on vs. light off: 99.92 ± 0.08 vs. 99.75 ± 0.25, p = 0.56, n = 4 rats; mean ± s.e.m; 2-sample t-test). Optical HFS (120 Hz, 5 ms pulse width) of the STN expressing CaMKIIα::ChR2-mCherry in 6-OHDA rats did not affect path length (cm/min; HFS vs. light off: 803.82 ± 129.04 vs. 851.95 ± 166.20, p = 0.83, n = 5 rats; LFS vs. light off: 847.15 ± 141.95 vs. 779.11 ± 104.01, p = 0.74, n = 2 rats; mean ± s.e.m; 2-sample t-test) or head position bias (% of time to the right; HFS vs. light off: 93.97 ± 3.78 vs. 94.20 ± 2.96, p = 0.96, n = 5 rats; LFS vs. light off: 98.50 ± 1.50 vs. 98.50 ± 0.50, p = 1.00, n = 2 rats; mean ± s.e.m; 2-sample t-test). 473nm illumination of the STN expressing GFAP::ChR2-mCherry in 6- OHDA rats also did not affect path length (cm/min; light on vs. light off: 1042.52 ± 113.73 vs. 1025.47 ± 113.63, p = 0.92, n = 4 rats; mean ± s.e.m; 2-sample t-test) or head position bias (% of time to the right; light on vs. light off: 98.16 ± 0.98 vs. 98.98 ± 0.65, p = 0.52, n = 4 rats; mean ± s.e.m; 2-sample t-test).

Optical Intensity Measurements: Light transmission measurements were conducted with blocks of brain tissue prepared from two 300 g Fisher rats and immediately tested. Blocks of tissue 2 mm in thickness were cut in 0–4 ºC sucrose solution using a vibratome. The tissue was then placed in a Petri dish containing the same sucrose solution over the photodetector of a power meter. The tip of a 200 μm diameter optical fiber coupled to a blue or yellow diode laser (473 nm or 561 nm, 30 mW fiber output) was mounted on a micromanipulator. First, the power was measured through the solution. Then, the tip of the fiber was moved down into the tissue in 100 µm increments and the power was measured. When the fiber reached the Petri dish, the power measured was compared to the initial measurement through the solution to confirm the total power output through the fiber. The percent transmission fraction was then calculated as the ratio between the power measured through tissue and the power measured through solution. The power intensity was then calculated by considering the light intensity

spread due to the conical shape of the 30 mW light output from a 400 µm fiber based on the fiber’s numerical aperture of 0.37 (Aravanis, Wang et al. 2007). The fiber output was assumed to be uniform across the diameter of the cone. Measurements were made through grey matter in three blocks of brain tissue for each wavelength with one block each moving anterior- posterior in the thalamus and in the cortex and dorsal-ventral through the thalamus.

Analysis of Electrophysiological Data: Threshold search in Clampfit was used for automated detection of spikes in the multi-unit recording, which was then validated by visual inspection; the spike waveforms displayed by Clampfit were observed to check the quality of spike detection. For traces with multiple spike populations, thresholds were set to capture all the spikes; during bursting, it is likely that multiple neurons were recorded from simultaneously. Bursts were identified in Clampfit; any two consecutive spikes occurring in an interval less than 300 ms were counted as belonging to the same burst and only bursts of at least 3 spikes were included. To quantify the neural activity at different frequencies, spectra for in vivo extracellular recording traces were generated using a wavelet transform after converting the traces into binary spike trains. The trace was then converted into a histogram with a binwidth of 0.5 ms for each of the duration-matched pre-stimulation, stimulation, and post-stimulation epochs. The start and end times for each of the segments, as well as the number of spikes, are listed below. The spike histograms were then convolved with a wavelet to measure the amplitude of the spectra at frequencies below 150 Hz over time. The average amplitude over time for each frequency was then plotted. The wavelet used is reproduced below.

For determining the change in activity of multiple frequency bands, amplitude spectra for multiple duration-matched baseline and stimulation sweeps were calculated as described above. Mean amplitude within each frequency band was determined and the ratio of this value (stimulation/baseline) was calculated. Spike latencies of the M1 response to optical stimulation of the STN were determined by measuring the delay between the first peaks in simultaneous optrode recordings of M1 and STN of a Thy1::ChR2-EYFP 6-OHDA mouse. 20 Hz, 5 ms pulse width of 473nm light was used to activate the STN.

Supplementary Figures

Fig. S1: Substantia nigra lesion and cannula track. Loss of nigral dopaminergic cells following 6- OHDA administration in rat (A) and mouse (B): coronal slices (rat: AP -5.8; mouse AP -3) show decreased tyrosine hydroxylase levels (red) unilaterally in the substantia nigra pars compacta; SNc is outlined by white brackets. Insets below show higher resolution images of the unlesioned (left) and lesioned (right) sides of the substantia nigra. (C) Cannula track is visible in a coronal slice showing correct placement of the cannula above the STN area.

Fig. S2: Additional histological characterization. (A) STN cells expressing CaMKIIα::eNpHR-EYFP (green) label for the excitatory neuron specific glutamate marker (red). (B) STN cells expressing GFAP::ChR2-mCherry (red) costain with the astroglia-specific marker S100β (green). In both (A) and (B) yellow indicates colocalization of the two markers. (C) Representative confocal images of TH stain for dopamine (top) and CHAT stain for acetylcholine (bottom) showed no colocalization with Thy1::ChR2-EYFP expression in the STN.

Fig. S3: Additional behavioral results. (A) Continuous 473nm illumination of the STN expressing GFAP::ChR2-mCherry in an anesthetized 6-OHDA mouse completely inhibited STN activity. (B) and (C): Extension of mouse results. (B) Amphetamine-induced rotations were not affected by 50% duty cycle illumination of the GFAP::ChR2 expressing STN in 6-OHDA mice (n = 1 mouse and 2 sessions). (C) Amphetamine-induced rotations were not affected by high (130 Hz, n = 1 mouse and 2 sessions) or low (20 Hz, n = 1 mouse and 1 session) frequency optical stimulation in the CaMKIIα::ChR2 expressing STN in 6-OHDA mice. (D) and (C): Modulation of inhibitory neurons during behavior.

Although mainly excitatory, STN has about 7-10% percent cells that stain for inhibitory neuronal markers, such as GAD65/67 and parvalbumin (Allen Brain Atlas). To obtain specific expression in either GAD67 or parvalbumin neurons we injected GAD67-Cre and parvalbumin-Cre mice respectively (gift of Sylvia Arber) with a Cre-inducible adeno-associated virus (AAV) vector carrying ChR2-EYFP (Methods). Cre-dependent opsin expression was observed in the STN region, but behavior was unchanged with optical stimulation. (D) Amphetamine-induced rotations were not affected by high (130 Hz, n = 2 mice and 4 sessions) or low (20 Hz, n = 1 mouse and 2 sessions) frequency optical stimulation in 6-OHDA GAD67-Cre mice. (E) Amphetamine-induced rotations were not affected by high (130 Hz, n = 2 mice and 2 sessions) or low (20 Hz, n = 2 mice and 2 sessions) frequency optical stimulation in 6- OHDA parvalbumin-Cre mice.

Fig. S4: Additional electrophysiological results. Isolation of large amplitude (A) and small amplitude (B) units from the trace in Figure 8C and corresponding power spectra. Red lines represent average waveforms for all superimposed spikes that occurred during 70s of baseline activity (n = 428 spikes for small amplitude unit and n = 205 spikes for large amplitude unit). Both small and large amplitude units showed decreased activity during light that returned to normal baseline levels after stimulation. (C)

Response of STN to optical stimulation of STN in the Thy1::ChR2-EYFP 6-OHDA mouse at 90Hz. The STN is initially excited but activity is reduced in the emergent stationary state measured by loss of the large amplitude spikes evident during the baseline; nevertheless, significant low amplitude activity persists throughout the stimulation. (D) High time resolution trace of the STN response to optical stimulation of STN in the Thy1::ChR2-EYFP 6-OHDA mouse at 130Hz (see Figure 12B for full trace). Again, the STN initially responds with a spike followed by low amplitude activity throughout stimulation. Changes in amplitude of the local circuit responses can reflect either altered recruited cell number or altered excitability of recruited cellular elements. While optrode recordings cannot report on the precise cell types involved in generating activity, by eliminating the electrical stimulation artifact these recordings provide a window into the amplitude and timing properties of local circuit electrical responses arising from local excitatory or inhibitory cell types and fibers in the STN region that could not be achieved with electrical stimulation.

Fig. S5: High-temporal resolution optrode traces. (A) Single unit activity in CaMKIIα::eNpHR-EYFP expressing STN with continuous 561 nm light illumination in an anesthetized 6-OHDA rat (corresponding to trace in Figure 8). (B) Neuronal activity in CaMKIIα::ChR2-mCherry expressing STN with high frequency optical stimulation (120 Hz, 5 ms pulse width, 473 nm) in an anesthetized 6- OHDA rat (corresponding to trace in Figure 10). (C) and (D) Activity in the STN region in an anesthetized Thy1::ChR2-EYFP 6-OHDA mouse in response to high (HFS, 130 Hz, 5 ms pulse width) and low (LFS, 20 Hz, 5 ms pulse width) frequency optical stimulation using 473 nm light. Note the low amplitude of activity in the HFS trace.

Fig. S6: Latency of M1 response to optical stimulation of STN. (A) Response of M1 to optical stimulation of STN in the Thy1::ChR2-EYFP 6-OHDA mouse at 20Hz. (B) While stimulating STN with light, simultaneous recordings of light-induced activity in the STN (top trace) and M1/L5 (bottom trace) revealed short latency differences between the first peaks consistent with antidromic spiking. (C) Individual latency differences between the first peak in STN and M1/L5 for 16 stimulation bouts revealed minimal jitter (S.D. = 0.032 ms) consistent with antidromic spiking in the well-known M1- STN projection.

Fig. S7: Changes in frequency characteristics of neuronal activity produced by optical stimulation. (A) Activity in all frequency bands was reduced by continuous 561nm illumination of the STN expressing CaMKIIα::eNpHR-EYFP in anesthetized 6-OHDA rats (n = 5 sweeps). Frequency bands are defined as: delta 1-3 Hz; theta 4-8 Hz; alpha 9-12 Hz; beta

13-30 Hz; gamma 31-80 Hz; high frequency (HF) 81-130. (B) Optical HFS (120 Hz, 5 ms pulse width) of the STN expressing CaMKIIα::ChR2-mCherry in 6-OHDA rats reduced activity for frequencies between 4 and 80 Hz, while increasing activity in the HF band (n = 3). (C) Activity change in M1 (left, n = 4) and STN (right, n = 4) produced by optical HFS (130 Hz, 5 ms pulse width) stimulation of M1 of 6-OHDA Thy1::ChR2 mice. Delta activity in both M1 and STN was reduced. (D) Activity change in M1 (left, n = 4) and STN (right, n = 4) produced by optical LFS (20 Hz, 5 ms) stimulation of M1 of 6-OHDA Thy1::ChR2 mice. Beta, gamma, and HF activity in both M1 and STN was increased. (E) Optical LFS (20 Hz, 5 ms pulse width) of the STN in 6-OHDA Thy1::ChR2 mice increased activity in the beta, gamma, and HF bands (n = 3). (F) Spike counts for duration-matched baseline and optical stimulation segments for each experiment type. Optical stimulation of the STN expressing CaMKIIα::GFAP-mCherry and optical HFS in 6-OHDA Thy1::ChR2 mice abolished spiking activity, reducing activity across all frequencies to zero (not shown). Error bars are s.e.m.; t- test with µ = 100 used for statistics, * p < 0.05.

Epilogue

Given previous deep brain stimulation (DBS) research for Parkinson’s disease (PD), it is important to acknowledge that rather than excluding the importance of basal ganglia (BG) nuclei manipulations for the therapeutic effect, our results point to a complementary mechanism through which this can occur. Our results do recapitulate the impact of both high and low frequency stimulation for PD, with HFS of afferents into STN area being therapeutic while LFS exacerbates pathology. We have also observed interesting after-effects, similar to the ones observed for PD patients: the therapeutic window extended beyond light offset, and the longer the light exposure the longer the after-effect. The relevance of the cortical activity and input to the STN in regards to the Parkinsonian pathology has been previously explored by multiple groups, as already mentioned in this chapter. Nevertheless, while lesions sparing the cortical fiber input to the STN were possible, the converse was not. Optogenetics has now allowed us to specifically target these axons for manipulation and revealed their critical contribution to the therapeutic effect of deep brain stimulation. As we state in this chapter, our “findings do not exclude other important contributions to DBS targets or disease symptoms,” such as the role of basal ganglia nuclei.

A great deal of further study is indeed critical to understanding the whole truth about DBS for PD. For example, electrical and optical DBS cold be combined in occlusion experiments: one can perform high-frequency electrical stimulation of the STN while simultaneously inhibiting firing in selective neuronal elements of the BG (elements potentially involved in mediating therapeutic effects) via optical DBS. One potential result is that electrical DBS will not cause any additional therapeutic effect on top of optical modulation. If this occlusion occurs, then it is likely that electrical DBS acts, directly or indirectly, through the same cell population as optical DBS.

Recording in other nuclei of the BG with STN stimulation is very valuable, and we are excited about the prospect of measuring activity in the brain, during stimulation, with decreased or no stimulation artifact. In addition to recording the activity of single units or field potentials, it would be greatly advantageous to look at the activity changes due to optical stimulation of extended populations of neurons. Although biochemical markers, such as c-fos, can give readout for activity over large brain regions, an in vivo approach would complete our experiments. Positron emission tomography or PET scans done in PD patients showed increased activity in the thalamus, supplementary motors area, and motor cortex and decreases in the vestibular cortex during stimulation, consistent with increased thalamic output onto the cortex (Ceballos-Baumann et al., 2001; Perlmutter et al., 2002). Recently, optogenetic fMRI (Lee et al., 2010) has been developed in our lab. Both microPET and optofMRI can be used to look at the activity of populations of neurons in both the basal ganglia and motor cortex of optically stimulated and non-stimulated rodents. Correlating neuronal activity brain-wide with behavior should offer unique insight into the relevant structures for PD therapy.

Both STN lesion and STN-DBS can have neuroprotective effects on the dopaminergic population in the substantia nigra par compacta (SNc) of parkinsonian rats (Piallat et al., 1996; Chen et al., 2000; Maesawa et al., 2004). Also, glutamic acid decarboxylase (GAD; the enzyme responsible for synthesis of the inhibitory neurotransmitter GABA) gene therapy in the excitatory glutamatergic neurons of the STN is neuroprotective for nigral dopamine neurons and can rescue parkisonian behavioral phenotype (Luo et al., 2002). It will be interesting to investigate whether long term optical modulation of specific cell-types in the basal ganglia can have neuroprotective effects on the dopaminergic neurons in SNc.

High-frequency stimulation of the pedunculopontine nucleus (PPN, which is also hyperactive in PD) has been also shown to be an effective clinical intervention for PD with potentially fewer side effects (reviewed in Kringelbach et al., 2007 and (Breit et al., 2004). Optical DBS could help understand why PPN is an effective clinical target and whether its modulation is superior to that of STN.

Finally, together with ameliorating motor symptoms, DBS also causes side-effects on cognitive and motivational parameters, with DBS treated patients developing depression or cognitive deficits. Interestingly, STN also controls activity in the ventral palladium, where associative and limbic circuits converge through the core or the shell of the nucleus accumbens respectively, suggesting a role for STN beyond motor control (Temel et al., 2005) which will require a separate investigation that might nevertheless be helped by the results of our studies. For example, it might be useful to compare the results for DBS vs. Optical DBS in reaction time tasks and impulsivity in rodents. Cognitive or depression (forced swim test) studies could be performed as well. Optogenetic DBS, as pioneered in this work, could provide insight into both the motor and cognitive/emotional valences of the STN.

Chapter 5

Optogenetic interrogation of neural circuits: technology for probing deep mammalian brain structures

Abstract | Elucidation of the neural substrates underlying complex animal behaviors depends on precise activity control tools, as well as compatible readout methods. Recent developments in optogenetics have addressed this need, opening up new possibilities for systems neuroscience. Interrogation of even deep neural circuits can be conducted by directly probing the necessity and sufficiency of defined circuit elements with millisecond-scale, cell type- specific optical perturbations, while suitable readouts include electrophysiology, optical circuit dynamics measures, and freely-moving behavior in mammals. Here we detail strategies for delivering microbial opsin genes to deep mammalian brain structures in vivo, along with protocols for integrating the resulting optical control with compatible readouts (electrophysiological, optical, and behavioral). The procedures described here, from initial virus preparation to systems-level functional readout, can be completed within 4 to 5 weeks. Together, these methods may help provide circuit-level insight into the dynamics underlying complex mammalian behaviors in health and disease.

Introduction

Understanding the circuit-level functional organization of the brain will have important implications for both basic and clinical neuroscience. In particular, optical manipulation of activity in neural circuits with light-sensitive rhodopsins such the Chlamydomonas channelrhodopsin-2 (ChR2)(Boyden, Zhang et al. 2005; Zhang, Wang et al. 2006), Volvox channelrhodopsin-1 (VChR1)(Zhang, Prigge et al. 2008), Natronomonas halorhodopsin (NpHR)(Zhang, Wang et al. 2007) (Figure 14a), and synthetic rhodopsin/GPCR

chimeras(Airan, Thompson et al. 2009) (optoXRs; Figure 14b) may help illuminate both normal circuit function and major disease mechanisms(Bi, Cui et al. 2006; Deisseroth, Feng et al. 2006; Adamantidis, Zhang et al. 2007; Petreanu, Huber et al. 2007; Zhang, Aravanis et al. 2007; Alilain, Li et al. 2008; Huber, Petreanu et al. 2008; Arenkiel and Peca 2009; Gradinaru, Mogri et al. 2009; Hira, Honkura et al. 2009; Lima, Hromadka et al. 2009; Petreanu, Mao et al. 2009; Sohal, Zhang et al. 2009; Tsai, Zhang et al. 2009). Here we illustrate the critical steps for implementation of this optogenetics approach(Deisseroth, Feng et al. 2006; Zhang, Aravanis et al. 2007) to probe the function of deep brain circuit elements in mammals. We first review the basic optogenetic control tools, followed by discussion of optimization of expression, targeting, and readout technology. We then present the necessary materials and outline a detailed series of experimental protocols for integrating optogenetic neuronal manipulation with in vivo and ex vivo readout methods, including electrophysiological and optical measures.

Temporally precise genetically-encoded control

Classical neuronal manipulation techniques (electrical, pharmacological, and genetic interventions) either simultaneously affect surrounding cells and processes in addition to the target population, or have slow kinetics and poor reversibility, severely limiting the strength of conclusions that can be drawn. To overcome these spatial and temporal limitations, microbial and chimeric-vertebrate opsin genes have been developed recently to control highly-defined electrical (Figure 14a) and biochemical (Figure 14b) activity with cell-type selectivity, high temporal precision, and rapid reversibility. Since most neurons in the brain are not naturally light-sensitive, selective expression of opsin genes in targeted neural populations makes it possible to specifically control activity in these populations, and the resulting fast on-off kinetics make it possible to evoke or inhibit neural activity within milliseconds, on a timescale relevant to physiological brain functions.

At present, light-activated cation channels from two distinct algal species, channelrhodopsin-2 from Chlamydomonas reinhardtii (ChR2)(Nagel, Szellas et al. 2003; Boyden, Zhang et al. 2005; Zhang, Wang et al. 2006) and channelrhodopsin-1 from Volvox carteri (VChR1)(Zhang, Prigge et al. 2008) (Figure 14a, left and middle), have proven to be a powerful pair of tools for controlling neural activity. ChR2 is maximally activated by blue light at 470 nm whereas VChR1 remains significantly light sensitive even at 589 nm, a wavelength at which

(importantly) ChR2 is no longer responsive (Figure 14c). Both ChR2 (deactivation time constant ~12 ms) and VChR1 (deactivation time constant ~120 ms) are able to transduce trains of millisecond-duration light flashes into defined spike trains up to 30-50 Hz(Boyden, Zhang et al. 2005; Zhang, Wang et al. 2006; Adamantidis, Zhang et al. 2007). Conversely, the chloride-pumping halorhodopsin from Natronomonas pharaonis (NpHR)(Zhang, Wang et al. 2007) has been shown to hyperpolarize neurons upon illumination with yellow light (Figure 14c). Because of sufficient spectral separation, NpHR and ChR2 can be simultaneously expressed in the same neurons to enable bidirectional optical control of neural activity(Zhang, Wang et al. 2007). Moreover, both use all-trans retinal as the chromophore, which is abundantly present in mammalian brain tissue, and therefore these optogenetic tools do not require addition of chemical cofactors in vivo.

FIGURE 14 | Optogenetic tools. a. Naturally occurring light-responsive effectors and their microbial sources: ChR2 from Chlamydomonas reinhardtii, VChR1 from Volvox carteri, and NpHR from Natronomonas pharaonis; useful light wavelengths for each are indicated. ChR2 and VChR1 are cation conducting channels and NpHR is a chloride pump. Adapted from Hausser and Smith, Nature 2007. b. Engineered synthetic rhodopsins for optical control of well-defined intracellular biochemical signaling. The intracellular loops of bovine rhodopsin have been replaced with the intracellular loops of G protein- coupled receptors (GPCRs) to yield light-activated chimeric GPCRs. Green light illumination leads activation of the downstream Gq and Gs signaling pathways. c. Action spectra. The absorbance wavelength of the voltage sensitive dye (VSD) RH 155 is sufficiently separated from the light-sensitive range of all rhodopsins, therefore making it possible to integrate VSD imaging with optogenetic modulation. d. Viral vectors for introducing microbial opsin genes into the brain. Top and middle: Lentiviral and AAV vectors can be used to deliver a cell-specific promoter along with the opsin gene and its fluorescent marker. Bottom: Cre-dependent AAV vector carries an inverted opsin fusion gene in the antisense orientation. Upon transduction in Cre recombinase-expressing cells, the opsin fusion gene will be irreversibly inverted and enable cell-specific gene expression. Part a was modified with permission from Nature(Zhang, Wang et al. 2007) (Nature © 2007; Macmillan Publishers Ltd)

Optimizing expression and function

For all of these proteins, expression issues will determine crucial aspects of performance. As one example, because of the inactivation properties of ChR2, higher-expressing cells will be able to follow fast spike frequencies for longer periods of time, as these cells will be able to employ larger pools of remaining non-inactivated ChR2. As another example, without the proper targeting sequences, high levels of microbial protein expression in mammalian cells can result in aggregation of misfolded proteins in the Golgi, endoplasmic reticulum, and other intracellular compartments. Since the conductance of these individual molecules is relatively low (picosiemens or less for ChR2, and even lower for NpHR as a pump, due to 1:1 coupling between photon absorption and ion flux), it is critical to maximize the number of molecules that are properly integrated into the cytoplasmic membrane, in addition to optimizing transcription of the opsin genes. For this reason, we have been constructing enhanced versions of halorhodopsin (eNpHR) by optimizing membrane trafficking sequences to increase efficiency of membrane targeting(Gradinaru, Thompson et al. 2008; Zhao, Cunha et al. 2008). eNpHR exhibits higher levels of membrane expression and more robust photocurrents, and similar protein-level modifications could be applied to optimize other microbial light-activated proteins for enhanced photocurrents.

Molecular engineering is also leading to expanded and refined function of these microbial proteins by altering spectral properties, conductance, or kinetics. One point mutation (H134R in ChR2) has been shown to result in 2-3x enhanced cellular photocurrents (but at the expense of slowed deactivation). Several C128 point mutants of ChR2 exhibit profound bistability, converting a brief pulse of light into a stable step in membrane potential(Berndt, Yizhar et al. 2009); transduced cells are (at steady-state) responsive to light at >100x lower light levels due to delayed exit of these proteins from the open state(Berndt, Yizhar et al. 2009). These C128 mutants are still activated by blue (470nm) light, but photocurrents elicited by the opening of ChR2(C128A) and ChR2(C128S) can be effectively terminated by a pulse of green (542nm) light. These engineered step function opsin (SFO) genes can be expressed to optically sensitize cells to native patterns of input, by providing a sustained (step) depolarization of the target cell’s membrane potential that can increase excitability. In many settings this approach will be preferable to simply driving user-defined trains with possibly inappropriate spike times. Related experimental leverage in terms of more natural control of native spike timing may also result from use of the optoXRs (Figure 14), which achieve temporally precise modulation of intracellular biochemical activity rather than direct control of spiking; of course, in some settings direct control of spike timing is the desired effect, in which case ChR2 or VChR1 are employed.

Genetic strategies for targeting expression to specific neural populations

Opsin genes can be selectively expressed in defined subsets of neurons in the brain using a variety of expression targeting strategies(Luan and White 2007; Wickersham, Lyon et al. 2007; Luo, Callaway et al. 2008). Here we focus on those techniques that have been shown to be effective in achieving functional expression in vivo.

Viral expression systems: Unlike many other genetic targeting strategies requiring the use of transgenic animal models, viral vectors(Dittgen, Nimmerjahn et al. 2004) based on lentivirus and adeno-associated virus (AAV) can be used to target opsin gene expression in a wide range of experimental subjects ranging from rodents to primates. Specifically, high titer lentivirus and AAV-based vectors (>109 transducing units (TU)/mL for lentivirus and >1012 genome copies (gc)/mL for AAV vectors) can be easily produced in Biosafety Level (BL)2-certified tissue-culture facilities in one to two weeks, or alternatively obtained through a number of virus production facilities (e.g. Viral Vector Core at University of North Carolina and Virapur,

Inc.). These transduction methods have been shown to induce high levels of functional opsin gene expression in neurons for several months.

While most common AAV (Figure 14d, top) and lentivirus (Figure 14d, middle) vectors carry strong ubiquitous or pan-neuronal promoters, some more specific promoter fragments retain cell type-specificity, allowing selective targeting in animals where transgenic technology is not accessible. Additionally, viruses are capable of mediating high levels of opsin gene expression by introducing multiple gene copies into each target cell, an important function for overcoming the low transcriptional activity of some cell-specific promoters. In general for rodent brains, opsin gene expression reaches optimal functional levels within three weeks after AAV injection, and within two weeks after lentivirus injection. We have found that in order the reach high steady-state levels of expression in distal axonal processes, longer periods of expression (> 6 weeks) may be necessary.

Electroporation: Specific cell types can also be targeted developmentally with in utero electroporation(Gradinaru, Thompson et al. 2007; Navarro-Quiroga, Chittajallu et al. 2007), for example at precisely timed embryonic days in mouse to target cortical layers II & III (E15.5), layer IV (E13.5), or layers V&VI (E12.5). In utero electroporation also can be used to express opsin genes in the inhibitory neurons of the striatum or in the hippocampus(Borrell, Yoshimura et al. 2005; Navarro-Quiroga, Chittajallu et al. 2007). Additionally, unlike viral delivery methods, in utero electroporation can be used to deliver DNA of any size, therefore permitting the use of larger promoter segments to achieve higher cell-type specificity.

Transgenic mice: Transgenic technologies can be used to restrict gene expression to specific subsets of neurons in mice or rats. Using either short transgene cassettes carrying recombinant promoters or bacterial artificial chromosome-based transgenic constructs, microbial opsin genes can be functionally expressed in subsets of neurons in intact circuits. Several transgenic mouse lines carrying ChR2 under the Thy-1 promoter, that have proven useful for a wide array of experiments, have been generated without any noticeable behavioral or reproductive defects(Arenkiel, Peca et al. 2007; Wang, Peca et al. 2007; Zhao, Cunha et al. 2008).

Conditional expression systems: Although cell-specific promoters are effective at restricting gene expression to subsets of genetically-defined neurons, some promoters have weak transcriptional activity. Therefore when used to direct the expression of microbial opsin genes, many cell-specific promoters are unable to achieve the level of opsin gene expression

necessary to mediate effective action potential firing or blockade. To amplify the transcriptional activity in a cell-specific manner, conditional AAV expression vectors(Atasoy, Aponte et al. 2008; Kuhlman and Huang 2008; Zhang 2008) (Figure 14d, bottom) have been developed recently to capitalize on the numerous cell-specific Cre-driver transgenic mouse lines that have been made available by individual labs and collective projects such as GENSAT (www.gensat.org). These conditional AAV expression vectors carry transgene cassettes that are activated only in the presence of Cre, and the use of strong ubiquitous promoters to drive the Cre-activated transgene selectively amplifies opsin gene expression level only in the cells of interest.

Circuit-specific cell targeting based on neuronal projection patterns: Neurons identified by a given genetic marker can still be quite diverse, either receiving innervations from or sending axonal projections to distinct brain regions. For example, some of the tyrosine hydroxylase (TH)-expressing dopaminergic (DA) neurons in the midbrain innervate reward-related brain structures such as the nucleus accumbens (NAcc), while other DA neurons project to motor control centers such as the striatum, and spatial separation between different DA neuron populations is not complete. However, since rhodopsins are trafficked to cellular processes including axons, the axonal processes may become light sensitive. Therefore it may be possible to selectively control a connection-defined neural pathway through focal injection of viral vectors followed by photostimulation of axon terminals in the target downstream brain structure. For example, to study the medial prefrontal cortical afferents in the amygdala, it is possible to inject a viral vector carrying ChR2-EYFP into the medial prefrontal cortex and implant the stimulation optical fiber into the amygdala. It is even possible to combinatorially interrogate the role of multiple subsets of afferent fiber bundles using multiple optogenetic proteins with distinct spectral sensitivities (e.g. using yellow light to activate VChR1- expressing afferents independent of ChR2-expressing afferents within the same neural tissue).

A number of plant and microbial proteins and several viral vectors with unique anterograde- or retrograde-transporting properties(Maskos, Kissa et al. 2002; Sugita and Shiba 2005; Wickersham, Lyon et al. 2007) may be engineered with recombinases to activate gene expression in subpopulations of neurons with cell type- and circuit-specificity. For example, expression of fusion proteins containing Cre and either wheat germ agglutinin (WGA) or tetanus toxin fragment C (TTC) in the cell bodies of one brain region will allow the recombinase to be trans-synpatically delivered to the postsynaptic or presynaptic neurons in

another brain region respectively. Similarly, retrograde-transporting viral vectors(Callaway 2008; Boldogkoi, Balint et al. 2009) such as rabies virus(Wickersham, Lyon et al. 2007) or herpes simplex virus 1(Lima, Hromadka et al. 2009) can be used to deliver recombinases or transgene cassettes in a retrograde fashion. When combined with conditional expression systems, either Cre-dependent transgenic mice or viral vectors, this strategy may allow circuit- specific gene expression in a variety of mammalian animal models not limited to mice. Moreover, microbial protein expression can also be restricted to specific intracellular compartments and locations by fusing to targeting motifs and protein domains(Gradinaru, Thompson et al. 2007; Lewis, Mao et al. 2009).

FIGURE 15 | Stereotactic implantation of the cannula guide. a. After mounting the animal into the stereotaxic frame, make a first incision to open the skin above the skull. The skin is gently pulled to the side to reveal the cranial sutures. b. After quickly wiping the skull with hydrogen peroxide, the bregma and lambda can be easily identified (marked spots). c. A thin needle is used to align the skull. d. A dental drill is used to create a small craniotomy at the desired location on the skull, without puncturing the dura. The dura is later removed using a fine forcep to minimize damage to the cortex. e. A cannula guide is implanted on the skull through the craniotomy. e. f.. Metabond and dental cement are used to secure the cannula guide to the skull. g. Use Vetbond and suture to close the incision around the cannulation site. h. An internal cannula guide connected to a pump is inserted through the cannula guide and is used to infuse virus into the target area in the brain. i. The animal is allowed to rest in a recovery cage after surgical implantation. Surgery was conducted according to established animal care guidelines and protocols at Stanford University.

Readouts for optogenetic control of intact circuits

Several classes of circuit readout can be made compatible with optogenetic control. First, voltage sensitive dye (VSD) imaging is an effective approach for monitoring the electrical activity of large populations of neurons ex vivo and in vivo with high temporal sensitivity. A second class of readout involves in vivo control integrated with either or both of electrical recording and behavioral measures. Targeting light-stimulation and electrical-recording devices to reach the site of rhodopsin gene delivery poses an experimental challenge that can be readily overcome with the appropriate protocol. We have developed a fiber optic-based optical neural interface (ONI) that meets this challenge(Aravanis, Wang et al. 2007). The ONI uses stereotactically implanted (Figure 15a-j) cannula guides for targeted virus infusion and light delivery into desired brain structures (Figure 16a).

Stereotactic targeting of brain structures: Stereotactic surgeries can be used to target opsin gene expression to genetically defined and spatially restricted neuronal populations in the brain, and can also be used to place a light delivery mechanism to target the transduced cells using a brain atlas(Paxinos and Franklin 2001) as described below. There are two major methods for stereotactically delivering lentiviral or AAV vectors to the brain. In the first approach, primarily applicable for in vitro acute slice electrophysiology or imaging studies, the viral vectors can be injected through a glass needle, targeted to the brain region of interest via a stereotaxic frame; the use of fine glass needles fabricated from glass capillaries minimizes the extent of tissue damage. (A small diameter Hamilton syringe can be used in

place of the glass needle; the Hamilton syringe can be mounted onto a micro-pump such as the WPI UltraMicroPump III directly attached to a stereotaxic frame).

FIGURE 16 | Preparation of an optical fiber for in vivo neural control in rodents. a. Diagram of the optical neural interface (ONI), consisting of a stereotactically implanted cannula, an optical fiber, and a solid state laser controlled by a signal generator. The fiber is prepared with the appropriate length to illuminate the target brain region. b. All of the tools and parts (fiber with FC connection, dummy cannula, internal cannula, cannula guide, dental drill, super glue, diamond scribe, and fiber stripper) needed to manufacture ONI fibers. c. d. A drill is used to produce a small bore on the center of a cannula cap. e. f. The steel tubing is removed from an internal cannula. The plastic adapter is used as the fiber guide. g. A fiber stripped is used to remove the plastic cladding from a fiber to reveal the fiber core. h. The bare fiber core is threaded through the bore on the cannula cap. i. j. The plastic adapter from the internal cannula is also placed on the fiber. k. l. Super glue is applied to the fiber to secure the plastic adapter against the plastic cladding on the fiber. The plastic adapter is held tightly against the fiber cladding for several minutes to allow the superglue to harden. m. After superglue has hardened, the fiber is inserted through a cannula guide with the right projection length for the target brain region. n. The cannula guide is securely clipped into the plastic adapter. o. A diamond scribe is used to remove the excess fiber from the tip of the cannula guide. p. The finished ONI fiber is allowed to protrude from the tip of the cannula guide by ~0.5 mm. Part a was modified with permission(Zhang, Aravanis et al. 2007) (Nature © 2007; Macmillan Publishers Ltd).

The second approach utilizes a stereotactically implanted cannula guide not only to deliver virus but also to guide an optical fiber to the same brain area of interest (Figure 16a). In this approach, the use of a single cannula guide for both viral vector delivery as well as optical fiber targeting ensures the co-registration of transduced brain area and light illumination. The use of cannula guides and infusion cannulae is a well-suited method for infusion of compounds into specific brain areas. Cannula systems consist of three major components: cannula guide, injection (or internal) cannula, and dummy cannula. The projection length of each component can be customized for each component depending on the brain region of interest. Depending on the experimental application, the material composition and physical dimensions of each component can also be varied; for example silica instead of steel cannulae are more suitable for functional magnetic resonance imaging, and larger diameter cannula guides can be used with bigger rodents such as rats to accommodate the insertion of a larger diameter optical fiber to increase the volume of illumination. The cannula guide is chronically implanted onto the skull of each experimental subject (Figure 15h). A dummy cannula (stylet and screw cap) is temporarily inserted into the cannula guide between experiments to prevent clogging and infection. During behavioral experiments, an optical fiber can be inserted through the cannula guide to illuminate the light-sensitive neurons. Additionally, bilateral

cannula systems are suitable for targeting multiple brain regions within the same experimental subject.

Choice of light sources: There are several factors to be considered, including the intensity of light needed to reliably activate a sufficient volume of tissue, and the ability to deliver high frequencies of light flashes to the experimental sample. For in vivo setups, it is also important to consider the mechanics and co-registration of the light with the opsin-expressing brain region. For in vitro experiments where the tissue sample is manipulated under a microscope, most conventional light sources such as halogen/xenon arc lamps, LEDs, and lasers can be directly coupled to the microscope’s light path. Light flashes can be generated either via a lamp coupled to a fast shutter (e.g. Lambda DG-4 or Uniblitz shutter) or by directly modulating the intensity of light output using a waveform generator (lasers and LEDs).

For in vivo light delivery in freely-moving animals, high power lasers (~10–15mW output at the tip of a 100um fiber) and LEDs are most suitable. For superficial stimulation of cortical layers, commercially available small LEDs can also be directly mounted above the brain through a cranial glass window (Gradinaru, Thompson et al. 2007; Huber, Petreanu et al. 2008). To target deep brain structures, thin optical fibers can be used to efficiently transmit sufficient powers of light to the target area. Optical fibers are versatile elements that can be fused to recording electrodes (creating an “optrode”(Gradinaru, Thompson et al. 2007) as described below for electrophysiological readout), and coupled to lasers and LEDs via FC/PC connectors. For mice, up to 300µm fibers (bare fiber without plastic cladding) can be used without compromising animal movement, while we find that rats can tolerate up to 400µm fibers. The length of the bare optical fiber can be customized based on the depth of the brain area by removing the precisely suitable length of plastic cladding. Mammalian brain tissue scatters light heavily, but ~10% of initial light power density remains at a distance form fiber tip of 500 µm(Adamantidis, Zhang et al. 2007; Aravanis, Wang et al. 2007).

To avoid fiber breakage resulting from repeated insertion through the cannula guide system, a short fiber segment can be permanently implanted in the target region. During experiments, this short indwelling fiber segment can be coupled to a longer fiber connected to the laser via a custom fiber-to-fiber connector (Doric Lenses). This strategy avoids fiber breakage, minimizes damage to brain tissue compared with implanted cannulas, and reduce the likelihood of infection due to environmental exposure via the cannula. However, the fiber

connector can lead to up to 50% loss of transmitted light, which may be overcome with more powerful laser diodes.

Choice of behavioral subjects: It is important to choose the appropriate species and genetic background according to the behavioral assay. For example, BALB/C animals are more suitable than C57BL6/J animals for anxiety and depression studies whereas C57BL6/J are preferred over BALB/C for locomotor and cognitive studies(Crawley 2007). When using transgenic mouse lines, it is important to make sure the mice are backcrossed (at least for 6 generations)(Crawley, Belknap et al. 1997) to the appropriate genetic background.

Mechanical design considerations for behavioral equipments: During optogenetic behavioral experiments, each animal subject will have an optical fiber tethered to the skull throughout the experiment. Therefore each subject should undergo sufficient habituation for the behavioral setup. Some apparati containing obstacles such as doors, tunnels or closed compartments may need to be modified to allow free passage of the optical fiber. In some experiments, the acute insertion and removal of an optical fiber before and after the behavior test may be stressful for the experimental subject, therefore prior handling including practicing fiber insertion and removal will help alleviate some of the stress. A fiber commutator (Doric Lenses) may be used to relieve tension in the fiber and provide more rotational freedom for the experimental animals.

Summary

We have provided above a compact collection of key technical considerations and protocols for optogenetic targeting of deep brain structures in mammals, as well as protocols for major readout strategies, both in vivo and ex vivo. These methods promise to permit systematic exploration of brain circuits that would otherwise be intractable for this kind of work. Together these protocols are intended to facilitate the truly versatile interrogation of circuit dynamics underlying mammalian behaviors in health and disease.

MATERIALS

Reagents

VIRUS PRODUCTION

Lentiviral plasmids

• - Channelrhodopsin Vectors • pLenti-CaMKIIa-hChR2(H134R)-EYFP-WPRE • pLenti-Synapsin-hChR2(H134R)-EYFP-WPRE • pLenti-CaMKIIa-VChR1-EYFP-WPRE • pLenti-EF1a-hChR2(H134R)-EYFP-WPRE • pLenti-CaMKIIa-hChR2(H134R)-mCherry-WPRE - Halorhodopsin Vectors • pLenti-CaMKIIa-eNpHR-EYFP-WPRE - SFO Channelrhodopsin Vectors • pLenti-CaMKIIa-hChR2(C128A)-EYFP-WPRE • pLenti-CaMKIIa-hChR2(C128S)-EYFP-WPRE • pLenti-CaMKIIa-hChR2(C128T)-EYFP-WPRE CRITICAL: These plasmids can be obtained from the Deisseroth lab directly (www.optogenetics.org) or from the Deisseroth plasmid list at Addgene. University- based virus production services, such as the virus vector cores at the University of North Carolina and University of Pennsylvania, will accept plasmid DNA and produce ready-for-injection virus for a fee. Mammalian Expression Vector plasmids for Transient Transfection - Channelrhodopsin Vectors • pcDNA3.1/VChR1-EYFP • pcDNA3.1/hChR2(H134R)-EYFP • pcDNA3.1/hChR2(H134R)-mCherry - OptoXR Vectors • pcDNA3.1/opto-a1AR-EYFP • pcDNA3.1/opto-b2AR-EYFP

Cre-dependent Adeno-associated Virus (AAV) Vectors • pAAV-EF1a-double floxed-hChR2(H134R)-EYFP-WPRE-hGHpA • pAAV-EF1a-double floxed-eNpHR-EYFP-WPRE-hGHpA CRITICAL: These plasmids can be obtained from the Deisseroth lab directly (www.optogenetics.org) or from the Deisseroth plasmid list at Addgene. University- based virus production services, such as the virus vector cores at the University of North Carolina and University of Pennsylvania, will accept plasmid DNA and produce ready-for-injection virus for a fee.

• 293FT cells (Invitrogen, cat. no. R700-07) • Dulbecco’s Modified Eagle Medium (Invitrogen, cat. no. 12-604Q) • UltraCULTURE serum-free medium (Lonza, cat. no. 12-725F) • Penicillin/Streptomycin/L-Glutamine mixture (Lonza, cat. no. 17-718R) • Sodium Pyruvate Solution (Lonza, cat. no. 13-115E) • Sodium Bicarbonate Solution (Lonza, cat. no. 17-613E) • Phosphate Buffered Saline (PBS) without Ca2+ and Mg2+ (Lonza, cat. no. 17-516F) • Defined Fetal Bovine Serum (HyClone, cat. no. SH30070.03) • Sodium Butyrate (Sigma, cat. no. 19364-1G) • HEPES (Sigma, cat. no. 54457-50G-F) • Sodium Phosphate Dibasic (Sigma, cat. no. 71636-250G) • Sodium Chloride (Sigma, cat. no. 71376-1KG) • Hexadimethrine bromide (Sigma, cat. no. 107689-10G) • Sodium Hydroxide (Sigma, cat. no. 71689-500G) ! CAUTION Strong base, handle and store according to hazardous material protocol.

• Distilled H2O (Lonza, cat. no. 17-724F) • Calcium Chloride 2M stock solution (Quality Biological, cat. no. 351-130-061)

Equipment

• 0.45 µm Low-protein binding filter flask (Millipore, cat. no. SCHVU02RE) • T-75 and T-225 tissue culture flasks (Nunc, cat. nos. 156499 and 159934)

• 4-layer Cell Factory culture flask (Nunc, cat. no. 140004) • Ultracentrifuge tubes (Beckman Coulter, cat. no. 344058) • Beckman Coulter Allegra X-12 bench-top centrifuge • Beckman Coulter L-100 K Preparative Ultracentrifuge • Beckman Coulter SW-28 Ultracentrifuge Rotor

STEREOTACTIC INJECTION AND CANNULA IMPLANTATION

• Anesthetics (Mice: Ketamine 80 mg/kg and Xylazine 12 mg/kg, Sigma, cat. no. K113; Rats: 80 mg/kg and Xylazine 6 mg/kg, Sigma, cat. no. K4138). ! CAUTION ketamine is a controlled substance and should be handled according to relevant rules of the host institutions. • Analgesics (Rats: buprenorphine 0.01-0.05 mg/kg; Mice: buprenorphine 0.05-0.1 mg/kg subcutaneous, Sigma, cat. no. B9275). ! CAUTION buprenorphine is a controlled substance and should be handled according to relevant rules of the host institutions. • Lubricant Eye ointment (Pharmaderm, cat. no. NDC 0462-0211-38) • Sterile PBS (Gibco PBS 10X, cat. no. 70011) • Ethanol (Sigma, cat. no. 459836) ! CAUTION Ethanol is flammable. Avoid exposure to ignition. • Hydrogen peroxide 30% (Sigma, cat. no. 31642) • C&B metabond (Parkell, cat. no. IS380) • Dental cement (Stoelting Co, cat. no. 51459) • Tissue adhesive (Fisher, cat. no. NC9259532) • Paraffin oil (Fisher, cat. no. BP26291) • Betadine (insert vendor and cat. no.) • 8-week old C57BL/6J mice (Jackson Laboratory, cat. no. 000664) ! CAUTION All experiments using animals should be carried out under institutional and national guidelines.

Equipment

• Surgical tools including scissors, forceps, scalpel blades (Fine Science Tools) • Small animal stereotaxic frame (Kopf Instruments, Tujunga, CA. cat. no. 922 for mice, cat. no. 955 for rat) with cannula holder (Kopf Instruments cat. no. 1766-AP).

• Programmable microsyringe pump (WPI, cat. no. SP220I) • 10 µL Hamilton Microsyringe (WPI, cat. no. nanofil) and tubing (Intramedic, cat. no. 4705) • Cannula guide for 200 μm fiber in mice (PlasticsOne, cat. no. C313GS-5/SPC) internal/injector cannula (PlasticsOne, cat. no. C313IS-5/SPC), dummy cannula (PlasticsOne, cat. no. C313DCS-5/SPC). The length of the cannula guide and internal cannula can be customized for desired target depth (recommended length for the cannula guide is 0.5 mm above the target region; internal cannula should have a projection length of 0.5 mm beyond the cannula guide tip; dummy cannula should be flush with the tip of the cannula guide). • Dissection stereomicroscope (1-10X) with stand (Leica MZ6, cat. no. 10445614) • Cotton swabs (Puritan Medical Products, cat. no. 25-806 10WC) • High Speed Micro Drill with Charger (Fine Science Tools, cat. no. 18000-17) • 0.5 mm Micro Drill Stainless Steel Burrs (Fine Science Tools, cat. no. 19007-05) • 0.9 mm Micro Drill Stainless Steel Burrs (Fine Science Tools, cat. no. 19007-05) • 1 mL Syringes with subcutaneous needles (VWR, cat. no. 82002-326) • Surgical suture (Myco Medical, cat. no. SK7772) • Heating blanket (Fine Science Tools, cat. no. 21061 or 21060) • Epoxy glue (Fisher, cat. no. NC9863515)

Reagent Setup

Virus injection set up Fill the syringe, the tubing and the glass needle or injection cannula with paraffin oil. Load the virus solution in the needle or injection cannula using the programmable microsyringe pump.

OPTRODE (INTEGRATED FIBER-ELECTRODE) RECORDING

Equipment

• 473 nm diode laser (Crystal Laser, cat. no. BCL-473-050; 50mW, 473nm, TEMoo CW laser, w/ TTL on/off, analog output intensity modulation, PC connector at laser output) • Optical Fiber (ThorLabs, cat. no. BFL37-200 for 200 µm fiber; cat. no. BFL37-300 for 300 µm fiber; fibers need to have a FC end to connect to the PC adapter on the output of

the laser; length can be specified depending on the experimental setup; recommended to have at least 1 m of slack fiber from the head of the animal to the first fiber fixture). • Fiber Stripping Tool (300µm fiber: ThorLabs, cat. no. T18S31; 200 µm fiber: ThorLabs, cat. No. T12S25) • Electrodes, tungsten, 1MΩ, ~125 µm (AM systems, cat. no. 573220) • Electrode holder (Kopf Instruments, cat. no. 1774) • Amplifier (AM systems, Model 1800 2-Channel Microelectrode AC Amplifier, Cat. no. 700000) • Data recording system (Axon Instruments/Molecular Devices Digidata 1440A or prior Digidata 1320 series, with pCLAMP 10 software)

IN VIVO OPTICAL STIMULATION FOR BEHAVIORAL STUDIES

Equipment

• Arbitrary waveform generator, lasers, and optical fiber (Agilent, cat. No. 33220A) • Light Power Meter (ThorLabs, sensor, cat. no. S130A, and digital console, cat. no. PM100D) • Dust cap (PlasticsOne, cat. no. 303DC/1) • A custom aluminum rotating optical commutator (Doric Lenses, Quebec, Canada) • Thy1::ChR2-EYFP Line 18 transgenic mice (Jackson Laboratory, cat. no. 007612) ! CAUTION all experiments using animals should be carried out under institutional and national guidelines.

Reagent Setup

D-10 Cell Maintenance Medium Prepare 500 mL of D-10 medium by supplementing 500 mL of Dulbecco’s Modified Eagle Medium (DMEM) with 50 mL of fetal bovine serum, 5 mL of Penicillin/Streptomycin/L-glutamine mixture, 5 mL of sodium pyruvate, and 5 mL of sodium bicarbonate. The final mixture can be stored at 4 °C for 1 month. Virus Production Medium Prepare serum-free virus production medium by supplementing 500 mL of UltraCULTURE with 5 mL of Penicillin/Streptomycin/L-glutamine mixture, 5 mL sodium pyruvate, and 5 mL of sodium bicarbonate. The final mixture can be stored at 4 °C for 1 month.

20% Sucrose Solution Prepare 10 g sucrose and then add PBS to a final volume of 50 mL. Filter using 0.22 µm filter and then store at 4 °C.

2X HBS Buffer To 450 mL of distilled H2O, add 5.96 g of HEPES (50 mM), 0.106 g of

Na2HPO4 (1.5 mM), 8.18 g of NaCl (280 mM). The pH should initially be around 5.8. Titrate with NaOH to 7.05 (use 5 M NaOH first and then switch to 1M NaOH). Bring the final volume to 500 mL and then filter with 0.22 µm filter. The solution is stable at room temperature (RT, 22 oC) for 6 months.

PROCEDURES

Production of recombinant lentiviral vectors for mammalian optogenetics gene delivery

1. Cell preparation TIMING 2 d Aspirate the growth medium from one 95% confluent T- 225 flask of 293FT cells. Gently rock the flask to ensure complete coverage of the cell monolayer and incubate at room temperature for 5 minutes, or until all of the cells have detached. Add 5 mL of D-10 to the flask to neutralize trypsin present in the medium and pipet 3 times to break up any cell clumps. CRITICAL STEP It is important to use low passage 293FT cells for the production fo viruses. To make sure the cell are always in the fastest growth phase, never let the cells approach full confluence.

2. Split the 10 mL of cell suspension into four new T-225 flasks. Add 30 mL of fresh room temperature D-10 to each flask. Gently rock the flasks to ensure the cells are evenly

distributed in the flask. Put all four flasks into a 37 °C CO2 incubator for 2 days or until 95% confluent.

3. Harvest the cells from each of four 95% confluent T-225 flasks of 293FT cells as described in Step 1. Transfer all of the cells into 500 mL of room temperature D-10 medium, mix and place the cell and medium mixture into one 4-layer cell factory. Transfer the cell factory to

a 37 °C CO2 incubator for 24 hours.

4. Transfection of cell factory TIMING (15 hr) In a 50 mL conical tube, prepare the following mixture: 690 µg of lentivirus plasmid (e.g. pLECYT, pFCK-hChR2-mCherry),

690 µg of pCMVdeltaR8.74, 460 µg of pVSVg, and 5.7 mL of 2 M CaCl2 solution. Mix

gently and bring the total volume to 23.75 mL with distilled H2O. Mix thoroughly.

5. Add 23.75 mL of 2X HBS to the DNA/CaCl2 mixture. Mix thoroughly and quickly. Then transfer directly into 500 mL of room temperature D10. CRITICAL STEP Mix the

DNA/CaCl2 with 2X HBS quickly and then transfer to room temperature D10 immediately. Short but thorough mixing ensures efficient formation of small calcium phosphate precipitates.

6. Remove the old medium from the 4-layer cell factory and replace with the D10 containing the calcium phosphate transfection mix. Transfer the plate back to the incubator for 15 hours.

7. Remove the transfection medium from the cell factory and gently wash using 200 mL of fresh D10. Add 400 mL of fresh room temperature D10 to the cell factory. Put cells back into incubator for 8 hours. ! CAUTION The medium from the cell factory may contain recombinant viral vectors. Apply bleach to the medium to decontaminate.

8. 24 hours post transfection, remove medium from cell factory and replace with 200 mL of Virus Production Medium supplemented with 5 mM Na-Butyrate. Return to incubator. CRITICAL STEP Cells are easily detached at this stage and should be handled gently. ! CAUTION The medium from the cell factory may contain recombinant viral vectors. Apply bleach to the medium to decontaminate.

9. Virus harvest TIMING 4 hr Sterilize 6 ultracentrifuge tubes by spraying with ethanol and air drying in the tissue culture hood.

10. 48 hours post transfection, collect the virus containing supernatant into four 50 mL conical tubes and centrifuge for 5 minutes at 500xg.

11. Prewash a low-protein binding 0.45 µm filter flask with 30 mL of D10 medium, then filter the virus-containing supernatant.

12. Divide the filtered virus-containing supernatant among the six centrifuge tubes.

13. Pipet 2 mL of 20% sucrose solution to the bottom of the centrifuge tube. CRITICAL STEP It is important to pipet slowly so that the sucrose solution forms a discrete layer.

14. Centrifuge in a Beckman SW-28 rotor for 2 hours at 82,700xg at 4°C.

15. Gently carry the centrifuge tubes back to the tissue culture hood and pour out the supernatant. There should be a tiny semi-transparent pellet at the bottom of each tube that resembles a soft contact lens. Dry the side of each tube with Kimwipe. ! CAUTION supernatant may contain trace virus. Apply bleach to the medium to decontaminate.

16. Add 100 µl of cold PBS to the first tube and resuspend the pellet by swirling and gentle pipetting. CRITICAL STEP Excessive pipetting will degrade the virus.

17. Transfer the medium from the first centrifuge tube into the next tube to resuspend the second pellet. Repeat for the 4 additional tubes.

18. Pipet virus solution into an Eppendorf tube and spin at 1000xg for 5 minutes to remove unsuspended virus debris.

19. Aliquot the supernatant and store at –80 °C for up to one year. PAUSEPOINT The supernatant can be store at –80 °C for up to one year.

Stereotactic injection and cannula placement into the rodent brain (Figure 15) TIMING 1-2 hrs

20. Animal Preparation. C57BL/6 mice (15–30 g) or Fisher rats (~200–300 g) should be housed and handled according to institutional and national guidelines.

21. Perform all surgeries under aseptic conditions. Surgical tools must be clean and sterile. The area should be disinfected with ethanol 70% and the tools must be sterilized by autoclaving or immersion in a disinfectant.

22. Anaesthetize rodents using 1.5% isoflurane (for surgeries longer than 1 hr) or i.p. injection (90 mg/kg ketamine and 5mg/kg xylazine for rats; 80 mg/kg and 15-20 mg/kg, respectively, for mice). CRITICAL STEP Check for the absence of toe pinch reflex. Dispose of excess anesthesia according to institutional regulations. ? TROUBLESHOOTING

23. Apply ophthalmic ointment to prevent eye drying.

24. Use a heating pad to keep body temperature at 35 οC.

25. Place the animal in a stereotaxic frame: fix the first ear (or head) bar and place the animal’s ear canal on the ear bar by applying moderate pressure; place the second ear bar in the opposite ear while gently holding the animal; insert the mouth holder between the jaw of the animal; and finally, attach the nose holder to the animal using low pressure. CRITICAL STEP Correct head position in the stereotaxic frame allows vertical, but no lateral, movement of the head.

26. Shave and clean the head, wipe with 70% ethanol and betadine.

27. Inject saline solution subcutaneously to prevent dehydration during surgery (30 mL per kilogram of body weight or ~0.5 mL for each adult mouse and ~5.0 mL for each adult rats)

28. Make a midline scalp incision of ~1cm using scalpel. Gently pull the skin aside to expose the skull (Figure 15a). Clean the skull with cotton swabs immersed in hydrogen peroxide. CRITICAL STEP The thin membranes above skull will be denatured immediately upon exposure to hydrogen peroxide and will appear as a layer of white foam. Stop the reaction with PBS immediately after application of hydrogen peroxide by wiping with a cotton swap soaked in PBS and remove debris with a scalpel blade.

29. After cleaning the skull with cotton swabs identify bregma (the point of intersection of the sagittal suture with the curve of best fit along the coronal suture) and lambda (midpoint of the curve of best fit along the lambdoid suture)(Paxinos, Watson et al. 1985) (Figure 15b). CRITICAL STEP Align the bregma and lambda to the same dorsal-ventral (or z)

coordinates by adjusting the height of the nose clamp on the stereotaxic frame until the head is flat. Align the x and y tilt of the animal’s head by measuring the x and y coordinates of bregma and lambda.

30. Use a brain atlas to identify the coordinates for the target structure.

31. Move to the targeted area defined by the coordinates (Figure 15c). Slowly make a small craniotomy (slightly larger diameter than the cannula guide) using a drill mounted on the stereotaxic frame (Figure 15d). CRITICAL STEP To minimize damage to the brain, do not drill through the dura and instead use fine forceps to remove the dura. This is crucial for cortical injections. ? TROUBLESHOOTING

32. Thaw virus on ice. Lentivirus can be kept on ice for 6 hours without significantly affecting its titer. AAV can be stored at 4C for at least 1 month without losing significant titer.

33. Place the cannula in the cannula holder on stereotaxic frame and slowly lower the cannula guide into the brain. ? TROUBLESHOOTING

34. CRITICAL STEP Dry the skull with cotton swabs. Apply a thin layer of C&B Metabond where the dental cement will be in contact with the skull, including around the cannula pedestal.

35. Once the C&B metabond hardens, release the cannula from the holder and withdraw the cannula holder without moving the cannula guide (Figure 15e). Secure the cannula pedestal with dental cement and ensure that at least 2 rounds of threads in the cannula pedestal are exposed (Figure 15f).

36. Glue the skin back with Vetbond surgical adhesive (Figure 15g).

37. Virus injection can be performed during surgery (A) or after surgery (B). It may be necessary to titrate the virus (Box 1)

a. Virus injection during surgery (We describe here our protocol using the micropump from WPI and with special attention on the surgical procedure and cannula implantation for virus injection into rodent brain. Virus can also be delivered using glass capillaries to minimize tissue damage(Cetin, Komai et al. 2006).) i. Inject 0.2-0.5 µl of virus into the brain using a 10 µl micro-syringe and a thin 34 gauge metal needle. ii. Control the injection volume and flow rate (0.1 µl/min) with a high precision injection pump (Figure 15h). iii. After injection leave the needle in place for 10 additional minutes to allow the virus to diffuse in the brain. Withdraw the needle slowly afterwards. iv. After completely withdrawing the needle check that it is not clogged by pumping out a small droplet (0.1 µl). b. Virus injection after surgery i. Perform this procedure after the dental cement is dry, while the animal is still anesthetized. ii. Use an internal cannula fitted to the cannula guide’s length (Figure 15i). iii. Connect the internal/injector cannula to the tubing filled with oil (which is connected to the microsyringe), load it with virus solution and place it into the cannula guide. iv. Infuse the virus at a very low rate (0.1 µL/minute) as described above. ? TROUBLESHOOTING

38. Give buprenorphine (0.03 mg/kg) subcutaneously following the surgical procedure to minimize pain discomfort.

39. Let the animal fully recover in a clean cage placed over a heating blanket (Figure 15i). Monitor the recovery of the animal every 10 minutes.

40. Simultaneous optical stimulation and electrical recording in living mice with the optrode method. TIMING 2-3 hr Prepare the optrode by firmly attaching an extracellular tungsten electrode (1MΩ, ~125µm) to an optical fiber (~200µm). TIMING 10 min.

CRITICAL STEP The tip of the electrode must be deeper (>0.3mm) than the tip of the fiber to ensure illumination of the recorded neurons and prevent artifacts(Gradinaru, Thompson et al. 2007). CRITICAL STEP Do not contact the tip of the electrode.

41. Prepare animals by performing craniotomies (A) or use animals that have a cannula implanted in the desired region (B). ! CAUTION All experiments using animals should be carried out under institutional and national guidelines. a. New craniotomy above target region i. Prepare the animal as described in steps 20 – 31, in a stereotaxic frame on an optics table. CRITICAL STEP Use a Faraday cage to shield out electrical noises. b. Existing cannula above target region i. Anesthetize animal as described in steps 22 – 24. ii. Place animal in stereotaxic frame (Step 25) so that the cannula is vertically aligned.

42. Use an electrode holder to mount the optrode (or optrodes for more than one location) on the stereotaxic frame.

43. Lower the optrode to the desired depth through the freshly made craniotomies or through the cannula. CRITICAL STEP Take all precautions not to damage the electrode tip. Avoid touching the skull or the cannula directly with the electrode tip; once the tip is safely inside the cannula it will not get damaged as it is lowered further.

44. Attach a ground wire to the scalp as a reference signal.

45. Connect the optical fiber to a laser diode of desired wavelength and the electrode to an amplifier. For single unit recordings the recorded signal is band pass filtered between 300Hz and 5 kHz high. For field potentials, filter the signal at 300Hz.

46. Use an ADC board (e.g. Digidata) and control software (e.g. pClamp) to collect data and generate light pulses through the fiber. ? TROUBLESHOOTING

47. If desired, sacrifice the animal immediately after recording. Its brain may be extracted and sectioned to verify opsin gene expression and accurate placement of the optrode, or acute slices can be prepared for electrophysiology or VSDI.

Setting up optical stimulation for behavioral assays

48. Preparation of ONI fibers (Figure 16) Prepare the optical neural interface (ONI) fiber from the cannula parts from PlasticsOne. It consists of a screwing cap for securing the fiber to the animal’s head, a fiber guard from the internal cannula adapter, and a bare fiber with length customized based on the depth of the target region. Prepare the screwing cap by drilling a ~2 mm diameter hole centrally on the top of a dust cap to allow the optical fiber to pass through (Figures 16c,d). 49. Remove the metal tubing from the plastic pedestal of an internal cannula (Figures 16e,f).

50. Use a fiber stripper to remove ~ 40 mm of plastic coating from the bare end of an optical fiber (Figure 16g).

51. Thread the fiber through the cap with the cap opening facing away from the FC connector on the fiber (Figure 16h). Thread the fiber through the plastic pedestal so that the snap on part faces away from the FC connector on the fiber (Figure 16i).

52. Apply a very thin layer of glue to the interface of coated/uncoated fiber and slide the plastic pedestal onto the fiber so it is tight against the plastic coating on the fiber (Figures 16j-l). Allow the glue to dry for 10 minutes. CRITICAL STEP do not apply excess glue. Too much glue will set slowly, prevent the cap from fitting over the internal cannula, and the internal cannula from snapping onto the cannula guide.

53. Insert the ONI fiber through a cannula guide (same dimensions as the ones implanted on the experimental animals), and ensure that they snap tightly (Figures 16m,n). Use a diamond knife to cut the fiber so that it projects 0.5 mm from the tip of the cannula guide (Figure 16o,p). It is not necessary to polish the fiber end. CRITICAL STEP It is

important to use a diamond knife so the fiber tip is flat to insure consistent illumination. After cutting check the fiber tip under a stereomicroscope.

54. Connect the FC end of the fiber to the PC connector on the output port of the laser. Connect the laser control box to a waveform generator.

55. Program the waveform generator with the desired stimulation protocol (i.e. pulse frequency, duration, and intensity). CRITICAL STEP Use a power meter to measure the light output at the end of the fiber before and after behavioral testing.

56. In vivo stimulation for behavior Insert the ONI fiber through the cannula guide on the animal’s head and tighten the screw cap to secure the fiber. CRITICAL STEP Extra care should be taken during fiber insertion to prevent breakage. CRITICAL STEP For most behaviors, it is necessary to habituate the animals to the experimental setup by practicing fiber insertion/removal several times. The animals should be placed in the behavioral testing room for at least 3 hours prior to behavioral testing. ? TROUBLESHOOTING

57. Begin behavioral testing. Stimulation can be applied by triggering the waveform generator.

58. Verify cannula placement and opsin gene expression by post-behavioral testing via standard histological analysis.

TIMING

Steps 1 – 3, Preparation of cells for transfection: 48 hours Steps 4 – 8, Transfection and culturing of cells to produce lentivirus: 39 hours Steps 9 – 19, Harvesting, concentration, and aliquoting of lentivirus: 4 hours Steps 20 – 39, Stereotactic injection of virus in rodents: 2 hours. It takes at least 2 weeks after injection before sufficient levels of opsins are expressed in the brain. Steps 40 – 47, Simultaneous optical stimulation and electrical recording in vivo: 2 to 3 hours. Steps 48 – 58, Behavioral testing with in vivo optical stimulation: 2 or more hours depending on the specific behavioral experiment.

? TROUBLESHOOTING

BOX 1 CRITICAL STEP Virus Titration 1. It is often necessary to titrate the amount of virus injected. 2. It is recommended several injections are performed with increasing volumes of virus at the same concentration 3. For instance, start at 0.2 µl per injection site with 0.2 µl increments up to 1 µl final injection volume. 4. Assess the efficacy of transduction by performing histological analysis.

ANTICIPATED RESULTS

Virus-mediated cell-specific expression of microbial opsin genes in neural tissues: Both lentiviral and AAV vectors carrying an appropriate cell-specific promoter and the opsin gene of interest can mediate robust expression in targeted cell types (Figure 17a-d)(Aravanis, Wang et al. 2007). Injection of 1 µl of high titer VSVg pseudotyped lentivirus (> 109 pfu/mL) or AAV (> 1012 gc/mL) into the cortex or hippocampus can result in transduction volumes >1 mm3. Prolonged expression of any membrane protein driven by strong ubiquitous promoters

such as CAG can lead to toxicity in neurons. However, when the transgene is driven by a reliable and strong cell-specific promoter, expression can be very well-tolerated, stable and also specific, with less than 10% of transduced cells belonging to non-targeted cell types as identified by immunostaining for the marker protein. Thus far, both the human Synapsin I and murine CaMKIIα promoters have been used in viral systems (lentivirus or AAV) to drive strong pan-neuronal and excitatory neuron-specific expression, respectively, in the rodent brain in vivo. The GFAP promoter can also be used to drive astrocyte-specific expression in a lentiviral vector. Other promising cell-specific promoters include ppHcrt(Adamantidis, Zhang et al. 2007), Sst(Tan, Janczewski et al. 2008), and CCK(Jasnow, Ressler et al. 2009). When using cell-specific Cre transgenic mice, 1 ul of Cre-dependent AAV can be delivered to the target brain area to achieve cell-specific expression in the Cre-expressing cell population.

With both lentivirus and AAV, microbial opsin genes will express stably for at least 6 months after injection; when tagged with a fluorescent protein marker, the transduced cells will exhibit strong membrane-localized fluorescence. Comparatively, ChR2 is better localized to the membrane than NpHR and VChR1 but it may still be difficult to identify the somata of ChR2-EYFP expressing cells in intact tissue. To improve the visualization and identification of expressing cells, the optogenetic protein and the fluorescent protein tag can be expressed as two separated proteins using IRES(Bochkov and Palmenberg 2006) or 2A(Holst, Szymczak- Workman et al. 2006; Tang, Ehrlich et al. 2009) bicistronic linkers. Expressing different rhodopsins as separate proteins also ensures that both will integrate into the plasma membrane in the correct orientation. For most reliable optical control of axon terminals (Figure 17e), it is important to achieve a high level of expression in the axonal membranes; therefore for viral mediated expression it is optimal to wait until four or more weeks after injection before conducting experiments.

Electrophysiological validation of functional expression: Depending on the targeted cell type, the maximal evoked firing frequency will vary (for pyramidal neurons in the neocotex, cells expressing ChR2 can be driven to fire spikes at up to 30 Hz reliably, while other cell types such as fast-spiking interneurons will be able to fire at 100 Hz or more). Spiking properties in targeted cells depend spike and illumination history as well as on membrane expression level and local illumination intensity; for any given cell type and circuit, a detailed characterization should be carried out to determine the efficacy of light evoked spike trains. To track and validate activity modulation, optrode recordings can be performed in vivo (Figure 17e).

Artifacts, although much smaller than for electrical stimulation, can be occasionally observed(Zhang, Laiwalla et al. 2009); when present, such artifacts are correlated with the onset and offset of the light pulse; amplitude depends on light power and can be reduced with proper grounding and use of electrodes with coating extending to the tip and staggered relative to the optical fiber by 300-500µm, as is typically important in any case for proper illumination of the recorded area.

Imaging optogenetically-controlled signals using voltage-sensitive dyes: VSD imaging of optically-evoked activity in neural circuits expressing light-gated ion channels allows for fast, all-optical interrogation and quantification of the spatiotemporal dynamics of ex vivo circuits. In general, brain sections prepared from younger animals (< 6 weeks) yield optimal signals; however older animals may still be used but with reduced signal properties. Brain sections should be prepared with sufficient thickness to preserve network connections in the target circuit (e.g. ~300 µm for mice and ~400 µm for rats). Because of the inherently low signal of VSDI (~0.1% change in absorbance or fluorescence) and high levels of optical noise, averaging individual pixels over multiple trials is often necessary. Along with averaging these pixels over time, several pixels can also be averaged over space (with reference to anatomical landmarks) to increase the signal-to-noise ratio (although resolution decreases). Software provided by Scimedia or other fast camera producers (i.e. Redshirtimaging) can perform these initial tasks. However more advanced post-processing and quantitative analysis methods are often performed in our lab using algorithms written in Matlab (Airan, Meltzer et al. 2007).

Optical stimulation of brain sections from a Thy1::ChR2-YFP transgenic mouse with blue (470 nm) light reveals precise optical changes in the brain tissue (Figure 17d). The signal is strongest in the CA1 region, where ChR2 is expression is highest. Signal is also seen in the entorhinal cortex, subiculum and dentate gyrus as a result of both direct activation of ChR2 and percolating circuit activity.

FIGURE 17 | Functional expression of microbial opsin genes in the rodent brain. a. Fluorescence image superimposed on a bright field image of the motor cortex of a wild-type mouse injected with 2 µl lentiviral vectors carrying the CaMKIIα::ChR2-EYFP construct in Layer V of the anterior M1 cortical area (AP 2 mm; ML 2 mm; DV 2 mm); 2 weeks were allowed for expression before preparing 200 µm slices that were examined under confocal (images are z-projection of single planes). b. The injection produced strong expression in Layer V of the cortex (the dendrites projecting from layer V neurons to the surface of the brain are clearly visible). c. Fibers from Layer V of the cortex travel through the corpus callosum (CC) and striatum. d. Axon terminate in target structures and axon terminii are clearly visible. e. Optrode recording from optical stimulation of the axon terminii in the subthalamic nucleus of a Thy-1::ChR2-EYFP mouse. Optical stimulation of the axon terminals instead of the soma produced robust firing in post-synaptic neurons. Example trace shows 10 pulses of 5 ms blue light (470 nm) flashes delivered at 20 Hz. A single 5 ms light flash evoked extracellular spike (inset). f. Example voltage sensitive dye imaging signal from an acute horizontal hippocampal slice stained with RH 155. Trace shows the voltage changes in the slice resulting from 10 pulses of 10 ms blue light flashes delivered at 20 Hz. Acquisition rate: 200 Hz. The trace is averaged over four acquisition periods and an ROI of 147 µm x 133 µm located in the entorhinal cortex. (Scale bars: a, 1 mm; b, 500 µm; c, 50 µm; d, 25 µm).

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Chapter 6

Molecular and cellular approaches to diversifying and extending optogenetics

Abstract | Optogenetic technologies employ light pulses to control biological processes within targeted cells in vivo with high temporal precision. Here we show that application of molecular trafficking principles can expand the optogenetic repertoire along several long- sought dimensions. Subcellular and transcellular trafficking strategies now permit 1) optical regulation at the far red/infrared border, and extension of optogenetic control across the entire visible spectrum; 2) increased potency of optical inhibition (chloride-mediated photocurrents beyond the nA level) achieved without requiring increased light power, that maintain the step- like switching and resistance to inactivation of earlier tools important for behavioral experiments; and 3) generalizable strategies for targeting cells based not on genetic identity, but on morphology or tissue topology, to allow versatile targeting when promoters are not known or in genetically intractable organisms. Together these results illustrate use of cell- biological principles to expand the versatile set of fast optogenetic technologies suitable for intact-systems biology and behavior.

Introduction

A fundamental goal in biology is fast control of defined cells within functioning tissues. Temporal precision of control is important since cells may carry out fundamentally different computations and deliver different outputs depending on the timing and context of input signals. For example, it is almost meaningless to ask the causal role of “activation” of a neuron type in the brain, since changes in context (Fleischman et al., 2008) or millisecond- scale shifts in timing (Bi and Poo 1998; Lee, Barbarosie et al. 2000; Shi, Hayashi et al. 2001; Guan, Giustetto et al. 2002; Lee, Takamiya et al. 2003; Silberberg, Wu et al. 2004; Fan,

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Fricker et al. 2005; Bellone and Nicoll 2007) can change the magnitude or flip the sign of neuronal action on the circuit. Similarly, pancreatic β-cells execute an intricate computation involving synchronized oscillations in membrane potential and cytoplasmic concentrations of Ca2+, cAMP, and phospholipase C, resulting in pulsatile insulin secretion with precise timing across the population of β -cells that may be important for the action of insulin on target tissues (Tengholm and Gylfe 2009). Traditional genetics delivers cell type-specific control if adequate targeting strategies exist, but genetic approaches lack the temporal precision needed to control events with behaviorally- or environmentally-relevant triggering and timing; moreover, pharmacological control strategies lack cell type-specificity, temporal precision, or both. The need to study molecular and cellular events not only in reduced systems, but also within intact biological systems, has driven recent awareness of the opportunities of fast control.

To enable temporally precise control of specific cell types within behaving animals, fast “optogenetic” (Deisseroth, Feng et al. 2006; Zhang, Aravanis et al. 2007) technologies have been developed involving single-component light-responsive proteins that transduce brief pulses of light into well-defined action potential trains and effector functions in vivo (Zhang, Wang et al. 2007). Using optogenetics, precisely timed gain-of-function or loss-of-function of specified events can be achieved in targeted cells of freely-moving mammals and other animals (Adamantidis, Zhang et al. 2007; Gradinaru, Mogri et al. 2009). For example, we have found that direct light-triggered excitation of cellular electrical activity (depolarization and precisely-timed action potentials) can be achieved via expression of the microbial opsin genes encoding Chlamydomonas reinhardtii channelrhodopsin-2 (ChR2) (Boyden, Zhang et al. 2005) or Volvox channelrhodopsin-1 (VChR1) (Zhang, Prigge et al. 2008). On the other hand, direct light-triggered inhibition of electrical activity (precisely-timed hyperpolarization) can be achieved via expression of the Natronomonas pharaonis halorhodopsin (NpHR) in vivo (Hwang, Zhong et al. 2007); this halorhodopsin was selected for its step-like and highly stable photocurrents compared with other microbial generators of inhibitory current (Zhang et al., 2007). The halorhodopsin achieves inhibition by pumping into cells an ion (chloride) that is normally used to signal neuronal inhibition, and therefore this safe and natural physiological signal is well suited to optogenetically inhibit neurons, but proton pumps can also be used to achieve a similar effect (Chow et al., 2010). Finally, more subtle (but still temporally-precise) optical modulatory strategies are also possible, including changes in the input-output relationships of targeted cells via expression of engineered step function opsins, or “SFOs,”

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that alter excitability (Berndt, Yizhar et al. 2009), and fast selective control of modulatory Gs or Gq signaling using synthetic rhodopsin/G protein-coupled receptor chimeras (optoXRs) (Airan, Thompson et al. 2009). This collection of tools, along with the development of versatile devices to deliver light in vivo (Aravanis et al., 2007; Adamantidis et al., 2007; Gradinaru et al., 2007), together have enabled widespread application of optogenetics.

To further expand the optogenetic toolbox, previously we have carried out genomic screening strategies to successfully identify and validate novel classes of opsins for optogenetic control (Zhang et al., 2007, 2008), and we also have carried out rationally-designed mutagenesis strategies to achieve new classes of opsin functionality (Berndt et al., 2008; Gunaydin et al., 2010). Here we apply a third type of intervention, namely application of molecular trafficking strategies, to derive a novel panel of optogenetic resources that both quantitatively and qualitatively enhance the power of optogenetics, opening fundamentally new avenues of investigation. In particular, tools are developed that allow targeting of cells solely by virtue of their topological relationships within tissue, and that extend the reach of optical control to the infrared border, with effector function enhanced beyond any other known tools, and covering the entire visible spectrum.

Results

Membrane trafficking and microbial opsin genes

Deriving optogenetic tools from multiple classes of microbes promises substantial diversity of triggering and effector functions (Zhang et al., 2008), given the ecological diversity of microbial organisms occupying niches with a broad array of environmental signals of informational or energetic value (Venter, Remington et al. 2004; Rusch, Halpern et al. 2007; Yooseph, Sutton et al. 2007; Williamson, Rusch et al. 2008). Moreover, as a necessary adaptation to small cell volume and genome size, microbes carry out sensation, transduction, and action using highly compact mechanisms, often all encompassed within a single open genetic reading frame (as with the microbial opsins, in which both photon sensation and ion flux effector function are implemented within a single compact protein) (Lozier, Bogomolni et al. 1975; Lanyi and Oesterhelt 1982; Kalaidzidis, Kalaidzidis et al. 1998; Nagel, Ollig et al. 2002; Nagel, Szellas et al. 2003). In contrast, metazoan or vertebrate cells may transduce energy or information with more complex multicomponent signaling cascades that afford greater opportunities for modulation but are much less portable (as with the vertebrate opsins).

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Optogenetic tools from simpler organisms therefore present clear opportunities. However, these proteins may not express or be tolerated well by more complex cells. Indeed, archaeal halorhodopsin (the light-activated electrogenic chloride pump that can be used for optogenetic inhibition in metazoans) displays impaired subcellular localization when expressed at high levels in mammalian neurons (Gradinaru, Thompson et al. 2008; Zhao, Cunha et al. 2008). An early trafficking step, export from the ER, was found to be impaired for this first-generation NpHR, leading to intracellular accumulations that colocalized with the ER marker KDEL (Figure 18A; Figure 18B, left). Fusing the FCYENEV ER export motif from a vertebrate inward rectifier potassium channel to the NpHR C-terminus prevented aggregate formation (Figure 18B, center) and greatly enhanced tolerability at high expression levels.

This second generation enhanced tool (eNpHR, now eNpHR2.0) already has been successfully employed in vivo and in intact tissue in a number of studies (Gradinaru, Mogri et al. 2009; Sohal, Zhang et al. 2009). As with many kinds of native inhibition, optogenetic inhibition could be overcome by strong excitatory activity (Sohal, Zhang et al. 2009). Potential additional molecular modifications for enhancing photocurrents from known and emerging opsin gene family members would include signal peptides, additional ER export motifs, Golgi trafficking signals, transport signals, and other motifs involved in transport of membrane proteins along the secretory pathway to the cell surface (Simon and Blobel 1993; Ellgaard and Helenius 2003). We therefore sought to apply combinatorial membrane trafficking strategies that could be generally applicable in a systematic, principled fashion to candidate microbial membrane proteins for translation to metazoan applications.

Examination of eNpHR2.0-expressing hippocampal neurons revealed the absence of globular ER accumulations, as previously reported, but nevertheless persistent intracellular labeling and poor membrane localization (Figure 18B, center), suggesting that indeed additional modifications subsequent to the ER export step would be important. Examination of primary- sequence differences between two forms of an inward rectifier potassium channel with differential membrane localization (Kir2.1 and Kir2.4) revealed differences not only in C- terminal ER export motifs but also in N-terminal Golgi export signals and in C-terminal trafficking signals (Hofherr, Fakler et al. 2005). Surprisingly, we found that provision of the Golgi export signal did not significantly affect surface expression (not shown), but that addition of the trafficking signal from Kir2.1 either between eNpHR and the EYFP fusion, or at the C-terminus of the fusion protein, dramatically reduced intracellular labeling and

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increased apparent surface membrane expression (Figure 18B, right), and also improved labeling of cellular processes (Figure 18B, right). Indeed, high-resolution confocal imaging (Figure 18C) revealed marked localization in processes, with identifiable labeled membranes spanning intracellular regions apparently devoid of the opsin-EYFP fusion protein, in a pattern never previously observed with NpHR or its derivatives.

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FIGURE 18 | Multiple trafficking modules for microbial opsin function in mammalian neurons (A) Hippocampal neurons showing wildtype NpHR-EYFP aggregation in the ER. NpHR (green) and ER (red; immunostaining of the KDEL marker) are also shown in overlay (yellow) to illustrate colocalization of NpHR aggregates and ER (arrows: ER; arrowheads: ER aggregates). (B) Multi-step modification of wildtype NpHR (left) rescues membrane expression. Addition of the K-channel ER export motif (eNpHR2.0, middle) and trafficking signal (eNpHR3.0, right) respectively reduced ER aggregation and improved membrane expression (particularly in neuronal processes for 3.0). (C) Membrane expression enabled in processes for eNpHR3.0 (confocal images showing membrane- localized EYFP fluorescence in the soma (top) and dendrite (inset, bottom)). (D) eNpHR3.0 enables substantially greater photocurrents than eNpHR2.0. Representative traces (left) showing photocurrents in cells virally transduced with eNpHR3.0 (black) and eNpHR2.0 (gray). Summary plot (right) showing average photocurrent levels in cells expressing eNpHR3.0 (747.2 ± 93.9 pA) and eNpHR2.0 (214.1 ± 24.7 pA; unpaired t-test p < 0.0005; n = 10). Values plotted are mean ± s.e.m. Membrane input resistance was similar for all neurons patched (eNpHR: 193.1 ± 36.6 MΩ; eNpHR3.0: 151.6 ± 28.5 MΩ; unpaired t-test p = 0.37). Light delivery (593nm) indicated by the yellow bar. Output power density: 15.5mW/mm2. (E) eNpHR3.0 enables much greater hyperpolarization than eNpHR2.0. Representative traces (left) showing voltage traces in cells virally transduced with eNpHR3.0 (black) and eNpHR2.0 (gray). Summary plot (right) showing average hyperpolarization levels in cells expressing eNpHR3.0 (101.0 ± 24.7 mV) and eNpHR2.0 (57.2 ± 6.8 mV; unpaired t-test p < 0.0005; n = 10). See also Figure S1.

If improved membrane targeting were indeed achieved with this modification, increased photocurrents would be anticipated to result. We therefore examined photocurrents, using whole-cell patch clamp recordings to quantify bona fide functional plasma membrane localization of halorhodopsin pump molecules. Photocurrents were indeed profoundly increased (to a level more than 20-fold larger than the initially described NpHR currents; mean ± s.e.m photocurrent 747.2 ± 93.9 pA in lentivirally-transduced hippocampal pyramidal neurons under the human synapsin I promoter; n = 10; Figure 18D). At action spectrum peak, hyperpolarizations exceeded 1.1 nA (Figure 20F, right) with only 3.5 mW/mm2 yellow light, more potent than any known optogenetic inhibitor published to date (including chloride or proton pumps that require tenfold greater light power to approach nearly comparable current magnitudes; Gradinaru et al., 2008; Chow et al., 2010). In virally transduced neurons, light- induced hyperpolarizations by >100mV were routinely achievable, at the same modest light power levels (mean hyperpolarization 101.0 ± 24.7 mV; n = 10; Figure 18E). Membrane

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potential changes of this new magnitude represent a functionally distinct advance in optogenetic inhibition, and we accordingly designate this 3rd-generation NpHR as eNpHR3.0 (the Natronomonas pharaonis halorhodopsin was named NpHR in 2005 (Sato et al., 2005), and the first trafficking-enhanced version developed by Gradinaru et al., 2008 is now referred to as eNpHR2.0).

As expected from prior work (Zhang et al., 2007) showing that NpHR photocurrents were step-like and exhibit little inactivation over more than 10 minutes of continuous illumination (it was for this photocurrent stability that NpHR was selected in our initial investigations; Zhang et al., 2007), the eNpHR3.0 photocurrents were also step-like, resistant to inactivation, and highly stable over multiple light pulses and timescales, ranging from the sub-second timescale of action potential disruption, to the minute-scale useful in circuit physiology paradigms and animal behavior, to the many minutes (>10 min) useful in certain long- timescale behavioral experiments (Supplementary Figure S1; Zhang et al., 2007).

Targeting by connection topology enabled by enhanced trafficking

We asked if the robust improved expression were preserved in the mammalian brain in vivo. Figure 19A illustrates that this is the case. We injected lentiviral vectors delivering the novel opsin gene under control of the CaMKIIα promoter to the CA1 region of the hippocampal formation in adult mice, and examined distribution of the expressed EYFP fusion. As in cultured cells, strong expression was observed not only in dendrites but also in axons in vivo with both eNpHR3.0 and eNpHR3.1 (a shorter version of eNpHR3.0 with equivalent functionality but the N-terminal signal peptide removed) (Figure 19A). A major in vivo opportunity for systems neurobiology would be controlling not just a projection from region A to region B, but a cell type itself that has (among its connections) a projection from A to B. This fundamentally distinct result requires multiplexing of optical control with other targeting methods (Figure 19B). Such control would be of great value in systems neurobiology; for example, cortical excitatory pyramidal neurons form a genetically- and anatomically-defined class of cell, but within this class are cells that each project to multiple different areas of the brain (e.g. thalamus, spinal cord, striatum, and other cortical areas) and therefore have fundamentally distinct roles (Migliore and Shepherd 2005; Yoshimura, Dantzker et al. 2005; Lein, Hawrylycz et al. 2007; Molyneaux, Arlotta et al. 2007; Leone, Srinivasan et al. 2008; Petreanu, Mao et al. 2009). It is unlikely that genetic tools will advance far enough to separate

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all of these different cell classes, pointing to the need to inhibit or excite cells defined by connection topology—i.e. by the conformation of their projections (Figure 19B).

FIGURE 19 | Trafficking-enhanced projection targeting and topological targeting in vivo (A) Neural process targeting in vivo. Lentiviral delivery of eNpHR3.1 (shorter version of eNpHR3.0 with the N-terminal signal peptide removed; Methods) driven by the CaMKIIα promoter led to expression in CA1 pyramidal neurons and dendrites (top), as well as in axon terminals (bottom; arrowhead indicates

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subiculum). (B) Trans-synaptic gene activation using wheat germ agglutinin (WGA)-Cre fusion. Schematic depicts two injection sites (one with WGA-Cre fusion gene and another with Cre-dependent opsin virus) and long-range projections; Cre can be trans-synaptically delivered from transduced cells (red) to activate distant gene expression only in synaptically-connected neurons that have received the Cre-dependent virus (green) but not in others (gray). (C) Construct design for the WGA-Cre (top) and Cre-dependent (bottom) AAV vectors. WGA and Cre genes are both optimized with mammalian codons and are inframe fused. (D) Cortico-cortical connection testing. Injection of AAV carrying the EF1α::mCherry-IRES-WGA-Cre cassette in the rat primary somatosensory cortex (S1) led to mCherry fluorescence in neuronal somata at the injection site (upper left), but as expected not in distant motor cortex (M1, lower left). Under these conditions, injection of Cre-inducible AAV5-EF1α::eNpHR3.0- EYFP into the ipsilateral motor cortex (M1) led to opsin expression (EYFP fluorescence) in M1 somata (lower right), but as expected, in S1, eNpHR3.0-EYFP was seen only in projecting axon terminals from these M1 cells (dispersed green puncta) and not in somata (upper right). Optrode recordings in vivo from the M1 site confirmed that eNpHR3.0 abolished neuronal activity when exposed to continuous 560nm light (bottom trace). (E) Interhemispheric dentate gyrus connection testing. The two dentate gyri were injected with AAV2-EF1a-mCherry-IRES-WGA-Cre (left) and AAV8-EF1a-DIO-ChR2-EYFP (right). Lower panels show high-magnification images of boxed regions above. Cells transduced with WGA-Cre expressed mCherry as expected (red), almost entirely within the granule cell layer (GCL) delineated by DAPI fluorescence (blue). WGA-Cre activated Cre-dependent opsin expression (EYFP fluorescence) in the contralateral hilus (arrowhead), a known source of projections to the ipsilateral dentate (note that on the contralateral side, EYFP fluorescence was confined to the hilus as expected; upper right). This projection from the contralateral hilus is known to synapse on ipsilateral dentate granule cells in the molecular layer (Ratzliff et al., 2004), which in turn is visible as a thin green strip of opsin-EYFP expression bordering the GCL (ML, upper left). Optrode extracellular recordings during optical stimulation (30Hz, 5ms, 473nm) confirmed functional in vivo opsin expression in dentate neurons expressing ChR2-EYFP (bottom trace).

One way to achieve this goal would be to capitalize on a different kind of membrane trafficking: to introduce into the local cell-body location a Cre-dependent virus conditionally expressing the microbial opsin gene of choice (Tsai et al., 2009; Sohal et al., 2009; Cardin et al., 2009; Atasoy et al., 2008), and rather than additionally employing a Cre-drive mouse line, to instead introduce into a distant target structure (chosen to define the cells of interest by anatomical connectivity) a virus expressing Cre recombinase fused to a trans-cellular tracer protein, e.g. wheat germ agglutinin (WGA) (Figure 19B) or tetanus toxin- fragment C (TTC) (Kissa, Mordelet et al. 2002; Maskos, Kissa et al. 2002; Sugita and Shiba 2005; Perreault, Bernier et al. 2006; Sano, Nagai et al. 2007; Huang and Goshgarian 2009). Cre recombinase in

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the fusion protein would be transported by presumed endosomal trafficking mechanisms along with the tracer to the local cell-body location if anatomically or synaptically connected, and activate opsin expression in the subset of local cells defined by this connectivity (Figure 19B) (Gradinaru, Thompson et al. 2007; Petreanu, Huber et al. 2007; Gradinaru, Mogri et al. 2009; Petreanu, Mao et al. 2009). Note that this approach does not require any specific promoter fragment or genetic definition of target cells (a clear advantage for use in less-genetically tractable species such as rats and primates); but if needed, such additional genetic refinements can be readily added (for example, both the WGA-Cre and the Cre-dependent opsin could be delivered under control of cell type-specific promoters where available (Figure 19C), creating a specific and versatile means for addressing cells defined at the intersection of connectivity, location, and genetics.

We first validated this concept in the rat (Figure 19D) by devising a strategy to selectively introduce eNpHR3.0 into those primary motor cortex (M1) neurons that are involved in cortico-cortical connections with primary sensory cortex (S1) (Zhang and Deschenes 1997; Colechio and Alloway 2009). To do this, we injected the previously described Cre-dependent AAV, now conditionally expressing eNpHR3.0 (AAV5-EF1α-DIO-eNpHR3.0-EYFP) into motor cortex, and injected a novel WGA-Cre-expressing AAV (AAV2-EF1α-mCherry-IRES- WGA-Cre) remotely into primary somatosensory cortex. Robust and physiologically functional eNpHR3.0-EYFP expression was indeed observed in a distributed subset of the motor cortex neurons (Figure 19D) at 5 weeks after injection despite the remoteness of the Cre recombinase AAV injection, while in control animals without Cre recombinase no expression was observed. Consistent with the anticipated mode of trans-synaptic or trans-cellular transport of Cre, no mCherry-positive cell bodies were observed in motor cortex, and no EYFP-positive cell bodies were observed in S1 sensory cortex (Figure 19D). However, the expected EYFP-eNpHR3.0 axon terminals arising from M1 were clearly present in S1 (Figure 19D, upper right). Simultaneous optrode stimulation/recording (Gradinaru, Thompson et al. 2007) was conducted to validate functionality of eNpHR3.0 under the WGA system; indeed, robust inhibition was readily observed in M1 (trace in Figure 19D), as expected from the intense fluorescence of the XFP-opsin fusion protein. These data indicate that neurons involved in cortico-cortical connections can indeed be addressed and targeted as a cell type, not simply a projection type, defined by their connections.

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To independently validate this targeting technology in a distinct circuit and with a different opsin, we next targeted hippocampal formation dentate gyrus neurons involved in interhemispheric projections (Figure 19E). Within the dentate, the only known monosynaptic contralateral projection arises from the hilar mossy cells which terminate on granule cells of the contralateral dentate, in dendrites of the molecular layer (Figure 19B) (Freund and Buzsaki 1996; Ratzliff, Howard et al. 2004). The WGA-Cre AAV was unilaterally injected into one dentate gyrus while the Cre-dependent AAV was injected into the contralateral dentate gyrus of the same animal. Strikingly, opsin expression was observed only in hilar cells of the contralateral side; indeed, the transsynaptic accumulation of Cre was retrograde and monosynaptic to the contralateral hilar cells, as no EYFP labeling was observed in the contralateral granule cell layer (Figures 19B, E). Moreover, the only EYFP-expressing circuit elements in the ipsilateral dentate, affording highly precise opportunities for optical control, were axonal fibers observed to terminate in the molecular layer of the granule cell layer, precisely as expected for fibers arising from the contralateral dentate hilus (Figures 19B, E). In the present setting, optrode recordings on the Cre-dependent AAV site confirmed the functionality of ChR2 under the WGA system for both cell bodies and axonal projections to the contralateral hemisphere; in line with previous optogenetic studies (Zhang, Aravanis et al. 2007; Zhang, Wang et al. 2007), 470nm light pulses at 30 Hz (5 ms pulse width) delivered through the optical fiber reliably drove local neuronal firing (trace in Figure 19E) in vivo.

Far-red optogenetic control

The utility of these targeting strategies for engineered opsins within intact tissue raises the question of whether additional advantages might be accrued with regard to volume of tissue modulatable in vivo. While membrane trafficking modifications will not shift the action spectrum, the capability to control neurons in the far-red is a long-sought goal of optogenetics, as this will allow use of light that penetrates much more deeply into scattering biological tissues (Shu, Royant et al. 2009) and therefore recruitment of larger volumes for optogenetics (Aravanis et al., 2007; Adamantidis et al., 2007; Gradinaru et al., 2009). The massive photocurrents observed for eNpHR3.0 (~20x those initially reported for NpHR, which itself is capable of blocking spiking in response to 589 nm amber light), suggested optogenetic control with far-red light might be achieved. We therefore explored optical control in the far-red with the trafficking-enhanced eNpHR3.0.

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Even in response to true red (630 nm) light of only 3.5 mW/mm2, we observed potent ~250pA outward photocurrents in virally transduced cells — still more than 6-fold larger than the first- observed NpHR currents with yellow light (Figure 20A). Moreover, we found that these photocurrents evoked by red light could be used to trigger large (>40 mV) hyperpolarizations in hippocampal pyramidal neurons (Figure 20B). We therefore explored even further redshifted light. We continued to observe robust photocurrents in the deep red with 660 nm light, and at the red/infrared border with 680nm light (Figure 20C). At 680 nm the photocurrents (~75 pA) were still larger than peak (yellow light) eNpHR2.0 currents previously reported (Figure 20C). Importantly, at all of the red and far-red wavelengths tested, eNpHR3.0 photocurrents readily blocked action potentials induced by current injection (Figure 20D), validating the extension of optogenetic control channels to far-red light.

One important feature of NpHR is its spectral compatibility with ChR2: the two opsins have largely separable action spectra and operate with similar light power density requirements, allowing bidirectional control of optical activity (Zhang et al. 2007) in vitro or in vivo. Combination vectors (Ryan and Drew 1994) have been employed to drive bidirectional control from a single promoter, employing ChR2 before a T2A or P2A motif followed by NpHR (Han, Qian et al. 2009; Tang, Ehrlich et al. 2009). Channelrhodopsin currents with this method have ranged from 150-240 pA, and halorhodopsin currents from 11-40 pA (Han, Qian et al. 2009; Tang, Ehrlich et al. 2009). Therefore, to explore application of altered membrane trafficking to this challenge, we created a bicistronic vector containing eNpHR3.0. We transfected the resulting construct, hSyn-eNpHR3.0-EFYP-P2A-hChR2(H134R)-mCherry (abbreviated eNPAC), into hippocampal pyramidal neurons. Figure 20E shows that trafficking of both opsin gene products to cellular processes was observed. To verify that independent excitation and inhibition was still possible despite the increased currents from eNpHR3.0, we mapped out the steady-state photocurrent action spectrum in detail for eNPAC and for ChR2(H134R) (Gradinaru, Thompson et al. 2007) and eNpHR3.0 alone (Figure 20F). Maximal eNPAC steady-state excitatory and inhibitory currents were both approximately 60% of that observed when each opsin was expressed individually, yielding maximal bidirectional photocurrents of >550 pA (Figure 20F, G).

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FIGURE 20 | Far-red optogenetic inhibition and single-component, bidirectional optical control (A) 630nm light evokes robust photocurrents in neurons transduced with eNpHR3.0 (representative voltage clamp trace at left). Summary plot comparing eNpHR2.0- and eNpHR3.0-expressing neurons (at right); eNpHR2.0: 42.7 ± 4.5 pA; eNpHR3.0: 239.4 ± 28.7 pA; unpaired t-test, p = 0.00004; n = 10). (B) 630nm illumination evoked robust hyperpolarization (representative voltage clamp trace at left). Summary plot comparing eNpHR2.0- and eNpHR3.0-expressing neurons (right); 15.6 ± 3.2 mV for eNpHR2.0 and 43.3 ± 6.1 mV for eNpHR3.0; unpaired t-test p = 0.00116; n = 10). (C) Summary of outward photocurrents evoked by different wavelengths of red and far red/infrared border illumination. 630nm: 239.4 ± 28.7 pA (left; n = 10), 660nm: 120.5 ± 16.7 pA (middle; n = 4), and 680nm: 76.3 ± 9.1

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pA (right, n = 4). (D) Illumination with red and far red/infrared border light inhibited spiking induced by current injection in neurons expressing eNpHR3.0. Typical current-clamp traces show optical inhibition at 630nm (top left), 660nm (top right), and 680nm (bottom). Power density: 3.5 mW/mm2 (630nm) and 7 mW/mm2 (660nm, 680nm). (E) Confocal images: neurons tranfected with eNPAC. eNpHR 3.0-EYFP (left) and hChR2-mCherry (middle) expressed both on the neuronal membrane and throughout the neurites. (F) Activation spectrum for eNPAC (left), and for ChR2(H134R) (right, blue) and eNpHR3.0 (right, yellow) alone. Maximum eNPAC steady-state excitation was 567 ± 49 pA at 427 nm (n=9), 62% of the value for ChR2(H134) alone (916 ± 185 pA; n=5). Similarly, maximum eNPAC inhibition was 679 ± 109 pA at 590 nm (n = 9), 61% of the value for eNpHR3.0 alone (1110 ± 333 pA; n=4). Output power density for peak current values 3.5-4 mW/mm2. (G) Typical eNPAC-mediated optical driving and inhibition of spiking. Blue light (445nm, 5ms pulses) drove spiking at 20Hz (left) and 10Hz (right), while simultaneous application of yellow light (590nm) inhibited spikes.

Optical inhibition in the green and blue enabled by cellular and genomic tools

The known wide action spectrum of the microbial opsins (Figure 20) poses challenges with regard to achieving multiple independent channels of control; interestingly, eNpHR3.0 becomes not only a potent far-red optical control tool, but also the most potent known blue light-driven opsin-based inhibitor (>400 pA at 472 nm; Figure 20F). However, given the remarkable diversity of microbial opsins (Zhang et al., 2008; Chow et al., 2010), it is important to consider that the membrane trafficking strategies delineated here may form a generalizable strategy for adapting diverse microbial opsins with unique properties for optogenetic control purposes. In a final series of experiments, we explored whether these and other enhanced membrane trafficking principles could enable the addition of genetically and functionally distinct components to the optogenetic toolbox.

Indeed, we have found that most microbial opsins tend to traffic poorly without additional molecular engineering (Figures 21, S8). We illustrate this principle first with the best characterized microbial opsin, bacteriorhodopsin (BR) (Stoeckenius and Bogomolni 1982), a green light-activated regulator of transmembrane ion conductance (Marti, Otto et al. 1991). We found that expressed in unmodified form, prominent intracellular accumulations were observed, similar to those seen when the Natronomonas halorhodopsin is expressed at high levels (Figure 21A, top left), and no photocurrents were observed. However, addition of the trafficking signal (TS, as employed for eNpHR3.0) between BR and EYFP substantially improved membrane and process localization (Figure 21A, top right), with smaller persistent

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ER-like accumulations that were eliminated with further C-terminal addition of the ER export signal FCYENEV (Figure 21A, center row). The resulting construct (eBR, doubly-engineered for optimal membrane trafficking) was well tolerated in cultured neurons, with marked membrane localization and process targeting (Figure 21A, center right; bottom row). Validation of functional plasma membrane targeting revealed that eBR could typically deliver ~50pA of outward photocurrent (Figure 21B) and ~10mV hyperpolarizations (Figure 21C) that sufficed to block spiking in hippocampal pyramidal neurons when exposed to the optimal wavelength light of 560nm (Figure 21D), thereby providing another channel for optogenetic control.

FIGURE 21 | eBR: novel trafficking-enhanced tool for green inhibition (A) Confocal images in hippocampal neurons: unmodified BR forms aggregates similar to those of unmodified NpHR (top left). Provision of the TS motif before EYFP decreased aggregate size but did not fully eliminate aggregates (top right). Only when the ER export motif (FCYENEV) was also added to the C-terminus were aggregates abolished, resulting in good membrane targeting throughout the soma (middle panels) and far into processes (bottom panels). (B) 560nm light (green bar) induced outward photocurrents in eBR cells (left: sample trace in voltage clamp), 46.4 ± 7.2 pA (right bar graph). Mean ± s.e.m plotted; n = 12. Membrane input resistance was similar for all neurons patched (131.6 ± 19.5 mΩ). Light power

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density at sample was 7 mW/mm2. (C) Corresponding light-induced hyperpolarizations (left: sample trace in current clamp) were 10.8 ± 1.0 mV (right bar graph). Mean ± s.e.m plotted; n = 12. (D) Illumination with green light (560nm) sufficed to inhibit current injection-induced spiking. See also Figure S9.

We also continued genomic strategies similar to those that allowed our identification of the red-shifted excitatory opsin VChR1 from Volvox carteri (Zhang et al., 2008). Indeed, a number of microbes have been reported to display light sensitivities from violet to near infrared; we accordingly took a broad genomic mining approach in environmental sequencing databases, plant/microbial expressed sequence tag (EST) libraries, and whole genome shotgun (WGS) sequencing repositories to search for new rhodopsins with channel or pump properties and novel light sensitivities. Using the primary amino acid sequences for ChRs, HRs, and BRs as the template sequence, we expanded our search to evolutionarily distant species to sample more sequence varieties. Among other candidate sequences from diverse hosts (Cryptomonas, Guillardia, Mesostigma, Dunaliella, Gloeobacter, etc), one of these from Guillardia theta was different from the previously reported G. theta rhodopsins 1 and 2 (GtR1 and GtR2) (Sineshchekov, Govorunova et al. 2005), and showed high amino acid homology to the Chlamydamonas reinhardtii channelrhodopsin-2 (ChR2). We designated this new protein as G. theta rhodopsin-3 (GtR3, Figure S9A), optimized the codon bias of GtR3 for mammalian expression, and delivered the GtR3-EYFP fusion gene via lentivirus to hippocampal pyramidal neurons. In an emerging theme, GtR3 in its native state showed intracellular accumulations (Figure S9B, left) and no photocurrents. Provision of the TS signal between GtR3 and EYFP only mildly reduced accumulations (Figure S9B, middle), but together with addition of the ER export signal FCYENEV to the C-terminus (Figure S9B, right) accumulations were abolished and increased surface and process localization observed. The resulting GtR3 hyperpolarizes hippocampal neurons (Figure S9C, left) in response to 472nm blue light, albeit with smaller currents than eBR (Figure S9C, right), and could inhibit spiking as well (Figure S9D). We also achieved blue inhibition of spiking with an AR opsin from Acetabularia acetabulum (Tsunoda, Ewers et al. 2006) engineered for improved trafficking; AR generates little current alone but was initially aggregate-free and required only addition of the TS signal between AR and EYFP for functional membrane localization and spike inhibition (Figure S9E), pointing to the potential diversity in strategies that will be indicated for different microbial opsin genes (Figure 22).

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FIGURE 22 | Optogenetics: molecular design for microbial tools (A) General subcellular targeting strategies for adapting microbial opsin genes to metazoan intact-systems biology. (B) Refinement of targeting at the tissue and subcellular levels.

Discussion

Optogenetic approaches previously have demonstrated substantial utility in the control of biological processes and behavior in freely-moving mammals (Adamantidis, Zhang et al. 2007; Airan, Meltzer et al. 2007; Petreanu, Huber et al. 2007; Gradinaru, Mogri et al. 2009; Petreanu, Mao et al. 2009; Sohal, Zhang et al. 2009; Tsai, Zhang et al. 2009) and other animals (Nagel, Brauner et al. 2005; Hwang, Zhong et al. 2007; Lerchner, Xiao et al. 2007; Zhang, Wang et al. 2007; Douglass, Kraves et al. 2008; Wyart, Del Bene et al. 2009), with the high temporal precision that is important for intact-systems biology. However, full versatility in probing the action of distinct cell types will require addition of new capabilities to the optogenetic toolbox. We have found that engineering specific membrane-trafficking capabilities for microbial proteins is an important step in generating cellular control technologies for applications to mammalian intact-systems biology. Indeed, a systematic approach to subcellular trafficking may be applicable to all opsins of non-mammalian origin (Figure 22). These approaches may serve to eliminate pathological targeting associated with

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high expression levels in mammalian cells, reduce accumulations in the intracellular organelles such as endoplasmic reticulum and Golgi, improve targeting to selected intracellular membranes, and ultimately increase photomodulatory potency (Figure 22). It may be possible to construct modified versions of the opsins described here for altered kinetic (Berndt et al., 2009; Gunaydin et al., 2010) or action-spectrum (Zhang et al., 2008) properties, and in some cases chimeric opsins may also be generated with improved expression properties (Lin, Lin et al. 2009). Not all trafficking strategies will be suitable for all microbial opsins, with different motifs required for opsins that encounter trafficking difficulty at different stages; careful subcellular analysis with rational selection of proper modifications together consititue a directed and principled strategy for generating novel optogentic tools from raw genomic information (Figure 22). eNpHR3.0: new quantitative properties and new classes of application We have previously observed that inhibition with NpHR and NpHR2.0, while useful for many applications (Gradinaru, Mogri et al. 2009; Sohal, Zhang et al. 2009), can in some cases be overcome by very strong excitation (Sohal, Zhang et al. 2009). Hyperpolarizations by greater than 100mV (Figure 18) with eNpHR3.0 provide a long-sought step forward in the potency of optical inhibition. The inhibition now provided with eNpHR3.0, more than 20-fold stronger than the initial NpHR, remains tunable with light intensity or duty cycle adjustments, as with any of the optogenetic tools. At the action spectrum peak, hyperpolarizations exceeded 1.1 nA (Figure 20F, right) with only 3.5 mW/mm2 yellow light (earlier chloride or proton pumps require tenfold greater light power to approach comparable current magnitudes; Gradinaru et al., 2008; Chow et al., 2010). With eNpHR3.0, light-induced hyperpolarizations were also massive (Figure 18E), with hyperpolarizations by >100mV below resting potential routinely observed (mean hyperpolarization 101.0 ± 24.7 mV; n = 10; Figure 18E). At the same time, eNpHR3.0 preserved the well-known (Zhang et al., 2007) step-like kinetics, fast recovery, and resistance to inactivation over long timescales of NpHR (Figure S8). eNpHR3.0 is particularly well-suited for in vivo applications, as the most redshifted and potent optogenetic inhibitor to date, but further strategies to enhance potency will no doubt emerge. Of course, inhibition this strong will not be required for every experimental setting; indeed, the earlier and less potent eNpHR2.0 has already found utility across many systems (Gradinaru, Mogri et al. 2009; Sohal, Zhang et al. 2009). For example, optogenetic work on Parkinsonian models (Gradinaru et al., 2009) showed that therapeutically effective deep brain

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stimulation (DBS) in the subthalamic nucleus is likely initiated by action on afferent axons (which may in turn then inhibit downstream or upstream networks); modulating axonal fibers may be the most efficient way to modulate a structure or network when using a point source such as an electrode (or fiber). But there is little downside to employing eNpHR3.0, and inhibition can be readily dialed down if needed by using weaker promoters, shorter expression times, or reduced light levels. However, even more important than this quantitative advance is the functionally significant access to new wavelengths ~100 nm redshifted from previous reports (680nm vs. 589 nm; Figure 20) which enables new classes of experimentation at the infrared border, with deeper-penetrating, reduced-scattering, and safer photons.

Tissue-topology targeting for optogenetics

To enable control of cell types based on synaptic connectivity properties, we leveraged transsynaptic delivery of a Cre recombinase (Figure 19). First, in a genetically less-tractable species (rat), we selectively targeted those primary motor cortex (M1) neurons that are involved in cortico-cortical connections with primary sensory cortex (S1) (Figure 19D), by delivering a Cre-dependent opsin vector into M1 and a trans-synaptically transported WGA- Cre recombinase fusion protein to S1. Second, to independently validate this targeting technology in a distinct circuit, we targeted hippocampal formation dentate gyrus neurons involved in interhemispheric projections (Figure 19E); in this experiment, resulting opsin localization in both local cell bodies and in distant axon terminals precisely fit the pattern expected for the targeted cell type, defined only by its connectivity, without use of cell type- specific promoter fragments or transgenic animals. This approach is best served by vectors that do not directly transduce axon terminals; this may not be the case for all AAV serotypes (Paterna et al., 2004), and indeed in each system this aspect must be validated along with the directionality (antero- or retro-grade) of Cre transport, which may depend on cell-specific endosomal dynamics. However, direct transduction of axons terminals in other cases may be desireable, and can be achieved with HSVs and pseudotyped lentiviruses; however, such an approach (unlike the present strategy) does not allow selection of the cell type postsynaptic to the transduced terminals, and is not as efficient with the high-titer and well-tolerated AAVs that are the vector of choice for many applications.

As shown in Figure 19, the cell-process targeting enabled by membrane trafficking modification allows for control of cells that are defined topologically—that is, by the essential

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character of their connections within tissue. The term “topological targeting” is here used to underscore the deformation-independence of the fundamental character of the connection—an axonal connection can take any path from A to B, and as long as the connection is present, the topological targeting strategy remains valid. Simpler or more complex versions of this topological targeting are also possible, all sharing the common property that no specific promoter/enhancer information is required to achieve cell type-specific targeting. This property is important in genetically less-tractable organisms, but also of substantial value even in animals such as mice, where targeting tools are in many cases inadequate. Of course, genetic targeting strategies may be multiplexed with topological targeting; for example, expression from the Cre-dependent vector and the Cre-fusion vector (Figure 19C) may each be governed by specific genetic targeting sequences if available. Moreover, the availability of multiple channels of optical control for both excitation and inhibition (discussed below) opens the door to combinatorial topological targeting strategies as well (Figure S9).

Avenues for technology development and application

Like ChR2, NpHR, and VChR1, we note that most microbial opsins can benefit from substantial protein engineering before achieving full functionality. Indeed, we and others have previously demonstrated molecular strategies for eliciting from microbial opsins increased light sensitivity (Berndt, Yizhar et al. 2009) increased photocurrent magnitude (Nagel, Szellas et al. 2005; Gradinaru, Thompson et al. 2007; Gradinaru, Thompson et al. 2008; Zhao, Cunha et al. 2008), faster kinetics (Lin, Lin et al. 2009), and bistable switching behavior (Berndt, Yizhar et al. 2009). Other possibilities such as shifted action spectrum (Zhang, Prigge et al. 2008), increased two-photon responsivity (Rickgauer and Tank 2009), and altered ion permeability (e.g. for Ca2+) may also be achievable (indeed, a single amino acid mutation can convert BR into a chloride pump) (Sasaki, Brown et al. 1995).

While ion conductance-regulating opsins have been the most versatile for ready translation (employing a language common across living organisms), biochemical control with light in defined cell types is also possible (but with a different set of approaches, given that microbial signal transduction employs distinct players and principles compared with metazoan signaling). Indeed, optogenetic control of well-defined biochemical signaling pathways was recently achieved both in cultured cells and in freely moving mammals, using the optoXR method for optical control of specified G protein-coupled receptor signaling (Airan,

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Thompson et al. 2009). A photoactivatable adenylyl cyclase has been studied from Euglena, although with high dark activity that limits in vivo application (Schroder-Lang, Schwarzel et al. 2007), and subsequent work on light-sensitive PAS or LOV domains (Levskaya, Weiner et al. 2009; Wu, Frey et al. 2009) as photoisomerizable modules that can be coupled to other proteins may open up new ways to control protein-protein association if these approaches can be made to operate in living organisms. ChR2 itself fluxes low levels of Ca2+ that are sufficient to drive Ca2+ waves (Gradinaru, Mogri et al. 2009), an intracellular messenger with broad biochemical signaling significance across vertebrate cells and tissues.

While not all microbial opsins may express or preserve functionality in metazoan cells, results thus far point to substantial versatility of the optogenetic approach across animal species (Nagel, Szellas et al. 2003; Boyden, Zhang et al. 2005; Li, Gutierrez et al. 2005; Nagel, Brauner et al. 2005; Bi, Cui et al. 2006; Ishizuka, Kakuda et al. 2006; Schroll, Riemensperger et al. 2006; Zhang, Wang et al. 2006; Adamantidis, Zhang et al. 2007; Aravanis, Wang et al. 2007; Arenkiel, Peca et al. 2007; Hwang, Zhong et al. 2007; Petreanu, Huber et al. 2007; Wang, Peca et al. 2007; Zhang, Wang et al. 2007; Zhang and Oertner 2007; Alilain, Li et al. 2008; Douglass, Kraves et al. 2008; Gradinaru, Thompson et al. 2008; Huber, Petreanu et al. 2008; Zhang, Holbro et al. 2008; Zhao, Cunha et al. 2008; Airan, Thompson et al. 2009; Gradinaru, Mogri et al. 2009; Han, Qian et al. 2009; Petreanu, Mao et al. 2009; Pulver, Pashkovski et al. 2009; Tsai, Zhang et al. 2009). Together with fiberoptic (Adamantidis, Zhang et al. 2007; Aravanis, Wang et al. 2007) and integrated fiberoptic-electrode “optrode” assemblies (Gradinaru, Thompson et al. 2007), even cells located deep within large, dense organs can be readily accessed and interrogated in freely moving mammals. In the nervous system, optogenetic tools have been applied to probe the neural circuit underpinnings of information transmission, oscillations, locomotion, awakening, and reward (Adamantidis, Zhang et al. 2007; Airan, Thompson et al. 2009; Cardin, Carlen et al. 2009; Gradinaru, Mogri et al. 2009; Sohal, Zhang et al. 2009; Tsai, Zhang et al. 2009), and to probe the operation of neural circuits important in Parkinson’s Disease, autism, schizophrenia, drug abuse, and depression (Adamantidis, Zhang et al. 2007; Gradinaru, Mogri et al. 2009). The additional resources defined here arise from the application of molecular, cellular, and genomic strategies to fundamentally expand the capabilities of optical control, and as this toolbox rapidly grows, optogenetics may come to play an increasingly potent and versatile role in intact-systems biology for the fast control of defined cells within functioning tissues.

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Supplemental Information (SI)

Supplemental Data

FIGURE S8 | Stable and potent photocurrents for eNpHR3.0 over both short and long timescales relevant to physiology and behavior. (A) Stability and recovery over seconds: representative traces (left) showing photocurrents in cells expressing eNpHR3.0 when exposed to pairs of 10s long yellow light pulses separated in time by (top to bottom): 2.5s, 5s, 10s, 20s. Summary plot for pulses 20s apart showing normalized average photocurrent levels (top right) in cells expressing eNpHR3.0 (P1 = first pulse peak, 1.00 ± 0.09; S1 = first pulse steady state, 0.74 ± 0.07; P2 = second pulse peak, 0.87 ± 0.09; n = 11 cells). Summary plot for pulses 20s apart showing ~50% peak recovery, (P2-S1)/(P1-S1), (bottom right); after 20s, the peak recovers to (45.2 ± 6.6) %. (B) Stability over minutes: timecourse of

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eNpHR3.0 normalized photocurrents for long-term continuous light exposure (n = 11 cells). Values plotted are mean ± s.e.m for all above. (C) Stability over long-term: > 10 min hyperpolarizing currents enabled by eNpHR3.0. Light delivery (593nm) for all above is indicated by the yellow bar. Output power density: 2.5mW/mm2.

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FIGURE S9 | Additional trafficking-enhanced tools for blue inhibition (A) New opsin sequence: G. theta rhodopsin-3 or GtR3. The EST sequence included all seven transmembrane helices; the 5` amino acid sequence was provided from ChR2 (transmembrane motifs: blue bars; conserved residues: yellow; truncation site for signal peptide: *; signal peptide provided from ChR2: gray). (B) Confocal images of hippocampal neurons; note GtR3 formed aggregates as with unmodified NpHR or BR (left). Provision of the TS motif before EYFP decreased the size of the aggregates but did not abolish them (middle). Only when the ER export motif (FCYENEV) was also provided to the C-terminus were aggregates abolished (right). (C) Sample current clamp (left ) and voltage clamp (right) 1traces and summary data show GtR3 function under 472nm light (18.5 mW/mm2). Light induced outward photocurrent summary (left bar graph) and corresponding hyperpolarization summary (right bar graph) for blue light peak. Corresponding photocurrents and hyperpolarization were: 0.5 ± 0.4 pA and 0.12 ± 0.09 mV for yellow light (589nm; 7.5 mW/mm2); 20.0 ± 6.7 pA and 5.6 ± 1.2 mV for blue light (472nm; 18.5 mW/mm2); 1.7 ± 0.9 pA and 0.6 ± 0.3 mV for purple light (406nm; 3 mW/mm2). Mean ± s.e.m plotted; n = 10; input resistance was similar for all neurons (113.5 ± 24.2 MΩ). (D) Illumination with blue light (472nm; 18.5 mW/mm2) sufficed to inhibit spiking induced by current injection in GtR3-expressing neurons. (E) TS-modification enhanced AR function (as AR showed no aggregates the ER export signal was not used). Illumination with blue light (472nm; 18.5 mW/mm2) sufficed to inhibit spiking. Additional molecular engineering is required to establish utility of these two opsin genes.

EXPERIMENTAL PROCEDURES

Opsin Sources: All opsins described here have been optimized for mammalian expression by changing each gene’s codon usage to conform to human codon usage distribution (http://www.kazusa.or.jp/codon/cgi-bin/showcodon.cgi?species=9606). The GenBank accession code for the original AR, BR, and GtR3 sequences are DQ074124, M11720, and EG722553.

DNA constructs: All NpHR variants were produced by PCR amplification of the NpHR- EYFP construct previously published (Zhang, Wang et al. 2007) and cloned in-frame into the AgeI and EcoRI restriction sites of a lentivirus carrying the CaMKIIα or Synapsin-1 promoters according to standard molecular biology protocols. A similar strategy was used for BR and AR. GtR3 was identified through genomic searches. All opsins described here have been optimized for mammalian expression by changing each gene’s codon usage to conform to human codon usage distribution (http://www.kazusa.or.jp/codon/cgi- bin/showcodon.cgi?species=9606), and the optimized sequence was custom synthesized (DNA2.0, Inc., Menlo Park, CA). The GenBank accession code for the original AR, BR, and

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GtR3 sequences are DQ074124, M11720, and EG722553., pAAV-EF1a-mCherry-IRES- WGA-Cre vector was constructed using standard molecular biology protocols. Codons for the WGA and Cre genes were optimized for expression in mammalian cells. The genes were synthesized by DNA2.0 (Menlo Park, CA). Cre was fused in-frame to the C-term of WGA, which in turn was fused to IRES. The mCherry-IRES-WGA-Cre bicistronic expression cassette was designed using the EMCV IRES sequence (Bochkov and Palmenberg 2006). The pAAV-EF1a plasmid backbone is the same as described previously (Sohal, Zhang et al. 2009; Tsai, Zhang et al. 2009). pAAV-hSyn-eNpHR3.0-EFYP-P2A-ChR2H134R-mCherry was constructed with a 120-mer primer:

(5`caagttctgctacgagaacgaggtgggctccggagccacgaacttctctctgttaaagcaagcaggagacgtggaagaaaacccc ggtcccatggactatggcggcgctttgtctgccg 3`) that contained the p2A region with the ER export sequence at the 5' end and 20 bases of the start of hChR2 at the 3' end. First, the ChR2(H134R)-mCherry fragment was amplified using the 120-mer as the forward primer and 5`–atatcgaattctcattacttgtacagctcgt– 3` as the reverse primer. Second, this amplified product was used as a reverse primer along with the forward primer 5`–ccggatccccgggtaccggtaggccaccatgacagagaccctgcct– 3` to fuse eNpHR 3.0-EYFP to ChR2 (H134R)-mCherry with the p2A region interposed. The 3.4 Kb fragment was then purified and cloned into the BamHI and EcoRI sites of the pAAV-hSyn vector. All constructs were fully sequenced for accuracy of cloning; updated maps are available online at www.optogenetics.org.

Lentivirus preparation and titering: Lentiviruses for cultured neuron infection and for in vivo injection were produced as previously described (Zhang, Wang et al. 2007). Viral titering was performed in HEK293 cells that were grown in 24-well plates and inoculated with 5-fold serial dilutions in the presence of polybrene (8 µg/ml). After 4 days, cultures were resuspended in PBS and sorted for EYFP fluorescence on a FACScan flow cytometer (collecting 20,000 events per sample) followed by analysis using FlowJo software (Ashland, OR). The titer of the virus was determined as follows: [(% of infected cells) × (total number of cells in well) × (dilution factor)]/ (volume of inoculum added to cells) = infectious units/ml. The titer of viruses for culture infection was 105 i.u. /ml. The titer of concentrated virus for in vivo injection was 1010 i.u. /ml.

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Hippocampal cultures: Primary cultured hippocampal neurons were prepared from P0 Spague-Dawley rat pups. The CA1 and CA3 regions were isolated, digested with 0.4 mg/mL papain (Worthington, Lakewood, NJ), and plated onto glass coverslips precoated with 1:30 Matrigel (Beckton Dickinson Labware, Bedford, MA) at a density of 65,000/cm2. Cultures were maintained in a 5% CO2 humid incubator with Neurobasal-A medium (Invitrogen Carlsbad, CA) containing 1.25% FBS (Hyclone, Logan, UT), 4% B-27 supplement (Gibco, Grand Island, NY), 2 mM Glutamax (Gibco), and FUDR (2 mg/ml, Sigma, St. Louis, MO).

Calcium phosphate transfection: 6-10 div hippocampal neurons were grown at 65,000 cells/well in a 24-well plate. DNA/CaCl2 mix for each well: 1.5-3 µg DNA (Qiagen 2+ endotoxin-free preparation) + 1.875 µl 2M CaCl2 (final Ca concentration 250 mM) in 15 µl total H20. To DNA/CaCl2 was added 15 µl of 2X HEPES-buffered saline (pH 7.05), and the final volume was mixed well by pipetting. After 20 min at RT, the 30 µl DNA/CaCl22/HBS mixture was dropped into each well (from which the growth medium had been temporarily removed and replaced with 400 µl warm MEM) and transfection allowed to proceed at 37C for 45-60 minutes. Each well was then washed with 3 X 1mL warm MEM and the growth medium replaced. Opsin expression was generally observed within 20-24 hours.

Electrophysiology: Hippocampal cultures grown on coverslips were transduced at 4 div with titer-matched viruses for all constructs (final dilution 104 i.u. /ml in neuronal growth medium) and allowed to express for one week. Whole-cell patch clamp recordings were performed as previously described (intracellular solution: 129 mM K-gluconate, 10 mM HEPES, 10 mM

KCl, 4 mM MgATP, 0.3 mM Na3GTP, titrated to pH 7.2; extracellular Tyrode: 125 mM NaCl,

2 mM KCl, 3 mM CaCl2, 1 mM MgCl2, 30 mM glucose, and 25 mM HEPES, titrated to pH 7.3). For voltage clamp recordings cells were held at -70mV. Light in the visible range was delivered from a 300W DG-4 lamp (Sutter Instruments, Novato, CA) through filters of different wavelength selectivity (Semrock, Rochester, NY) and a Leica 40X/0.8NA water objective. Filters, except for power spectra, (given here as wavelength in nm / bandwidth in nm / output power in mW/mm2) were: 406 / 15 / 3; 472 / 30 / 18.5; 560 / 14 / 7; 589 / 15 / 7.5; 593 / 40 / 15.5; 630 / 20 / 3.5. Far red and near-infrared light delivery: light (7 mW/mm2) for 660nm inhibition was delivered using a light emitting diode and a 40x/.8 NA water objective. Light (7mW/mm2) for 680nm inhibition was delivered using the X-Cite 120W halogen light source through a 680 ± 13nm filter and a 40x/0.8 NA water objective. Light delivery for eNPAC, ChR2(H134R), and eNpHR3.0 power spectra was delivered from a 300W DG-4

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lamp fitted with a Lambda 10-3 filter wheel (Sutter Instruments) with a 10-position wheel for 25-mm filters of different wavelengths and a 40X/0.8NA water objective. Filters (given here as wavelength in nm / bandwidth in nm / output power in mW/mm2) were: 387 / 10 / 3.5; 406 / 15 / 3.5; 427 / 10 / 4.0; 445 / 20 / 4.0; 470 / 22 / 4.0; 494 / 20 / 4.5; 520 / 15 / 4.5; 542 / 27 / 5.0; 560 / 20 / 5.0; 590 / 20 / 3.5; 630 / 20 / 3.5. For Figures 18, 20A-D, 21, and S9, confocal images and whole-cell patch clamp data are from cultured hippocampal neurons either transfected (confocal data) or transduced (patch data) with lentiviral NpHR, BR, GtR3 and AR-based constructs, and allowed to express for one week. Expression was driven by the human Synapsin I promoter and visualized by fusion to EYFP.

Immunohistochemistry: Primary hippocampal cultures were infected at 4 div with titer matched virus (final dilution 104 i.u./ml in neuronal growth medium). At 14 div cultures were fixed for 30 min with ice-cold 4% paraformaldehyde and then permeabilized for 30 min with 0.4% saponin in 2% normal donkey serum (NDS). Primary antibody incubations were performed overnight at 4°C using a monoclonal marker of endoplasmic reticulum recognizing endogenous ER-resident proteins containing the KDEL retention signal (KDEL 1:200, Abcam, Cambridge, MA). Cy3-conjugated secondary antibodies (Jackson Laboratories, West Grove, PA) were applied in 2% NDS for 1 hour at room temperature. Images were obtained on a Leica confocal microscope using a 63X/1.4NA oil objective.

Stereotactic injection into the rodent brain: Adult mice and Long-Evans rats were housed according to the approved protocols at Stanford. All surgeries were performed under aseptic conditions. The animals were anesthetized with intraperitoneal injections of a ketamine (80 mg/kg)/xylazine (15-20 mg/kg) cocktail (Sigma). The head was placed in a stereotactic apparatus (Kopf Instruments, Tujunga, CA; Olympus stereomicroscope). Ophthalmic ointment was applied to prevent eye drying. A midline scalp incision was made and a small craniotomy was performed using a drill mounted on the stereotactic apparatus (Fine Science Tools, Foster City, CA). The virus was delivered using a 10 µl syringe and a thin 34 gauge metal needle; the injection volume and flow rate (1 µl at 0.1 µl/min) was controlled with an injection pump from World Precision Instruments (Sarasota, FL). After injection the needle was left in place for 5 additional minutes and then slowly withdrawn. The skin was glued back with Vetbond tissue adhesive. The animal was kept on a heating pad until it recovered from anesthesia. Buprenorphine (0.03 mg/kg) was given subcutaneously following the surgical procedure to minimize discomfort. For the experiment in Figure 19A, to cover a large area in

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dorsal CA1, 1µl of concentrated lentivirus (1010 i.u. /ml) carrying eNpHR3.1 (a shorter form of eNpHR3.0 with the N-terminal signal peptide, the first 17 amino acids of original NpHR, removed) under the CaMKIIα promoter was microinjected into 2 sites in each hippocampus (site one: anteroposterior -1.5 mm from bregma; lateral, ± 1 mm; ventral, 1.5 mm; site two: AP, -2.5 mm from bregma; lateral, ± 2 mm; ventral, 1.5 mm). For Figure 19D and E, two 12 different adeno-associated viruses (AAVs) (virus titer 2x10 g.c./mL), were stereotactically 12 injected during the same surgery with an injection speed of 0.15ul/min. High-titer (2x10 g.c./mL) AAV was produced by the UNC VectorCore. For Figure 19D, double-floxed cre- dependent AAV5 carrying eNpHR3.0–EYFP (AAV5-Ef1a-DIO-eNpHR3.0-EYFP) was injected into M1, and AAV2-Ef1α-mCherry-IRES-WGA-Cre was injected into S1 of adult Long-Evans rats. 1 µl of virus was delivered at five different sites defined by the following coordinates: M1 injection I: AP, +1 mm from bregma; lateral, 1.5 mm; ventral, 2 mm; M1 injection II: AP, +2 mm; lateral, 1.5 mm; ventral, 2 mm; S1 injection I: AP, -0.3 mm; lateral, 3.4 mm; ventral, 2 mm; S1 injection II: AP, -1.3 mm; lateral, 3 mm; ventral, 2 mm; S1 injection III: AP, -2.12 mm; lateral, 3 mm; ventral, 2 mm. For Figure E, 1 µl of virus was injected bilaterally into the dentate gyrus (DG) of adult BL6 mice. AAV8-EF1a-DIO-ChR2- EYFP was injected in the right DG and of AAV2-EF1a-mCherry-IRES-WGA-Cre was injected in the left DG with the following coordinates: AP, -2.1 from bregma; lateral, ±1.05 mm; ventral, 2.1 mm.

In vivo optrode recordings. To validate opsin functionality in the WGA-Cre system simultaneous optical stimulation and electrical recording in living rodents was conducted as described previously (Gradinaru, Thompson et al. 2007)[54](Gradinaru, Thompson et al. 2007)[54](Gradinaru et al., 2007) using an optrode composed of an extracellular tungsten electrode (1 MΩ, ~125 µm) attached to an optical fiber (~200 µm) with the tip of the electrode deeper (~0.4 mm) than the tip of the fiber to ensure illumination of the recorded neurons. The optical fiber was coupled to a 473 nm (for ChR2) or 560 nm (for eNpHR3.0) laser diode (10 mW fiber output) from CrystaLaser. Optrode recordings were conducted in rodents anesthetized with 1.5% isoflurane and the optode was placed through small craniotomies created above target regions. pClamp 10 and a Digidata 1322A board were used to both collect data and generate light pulses through the fiber. The recorded signal was band pass filtered at 300Hz low/5 kHz high (1800 Microelectrode AC Amplifier). For precise placement of the fiber/electrode pair, stereotactic instrumentation was used.

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Tissue slice preparation: For preparation of brain slices, mice or rats were sacrificed 4 to 5 weeks after viral injection. Rodents were perfused with 20ml of ice-cold PBS, followed by 20 ml of 4% paraformaldehyde. The brains were then fixed overnight in 4% paraformaldehyde, and transferred to 30% sucrose solution for 2 days. Brains were frozen and coronal slices (40 µm) were prepared using a Leica SM2000R cryostat, and preserved in 4ºC in cryoprotectant (25% glycerol, 30% ethylene glycol, in PBS). Slices (DAPI stain 1:50,000) were mounted with PVA-DABCO on microscope slides, and single confocal optical sections (e.g. through dorsal CA1 region, ~1-2.5mm posterior to bregma or the dorsal subiculum, 2.7-3 mm posterior to bregma) were acquired using a 10X air and 40X/1.4NA oil objectives on a Leica confocal microscope.

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Bibliography

Adamantidis, A. R., F. Zhang, et al. (2007). "Neural substrates of awakening probed with optogenetic control of hypocretin neurons." Nature 450(7168): 420-424. Afsharpour, S. (1985). "Topographical projections of the cerebral cortex to the subthalamic nucleus." J Comp Neurol 236(1): 14-28. Airan, R. D., L. A. Meltzer, et al. (2007). "High-speed imaging reveals neurophysiological links to behavior in an animal model of depression." Science 317(5839): 819-23. Airan, R. D., K. R. Thompson, et al. (2009). "Temporally precise in vivo control of intracellular signalling." Nature 458(7241): 1025-9. Albin, R. L., A. B. Young, et al. (1989). "The functional anatomy of basal ganglia disorders." Trends Neurosci 12(10): 366-75. Alexander, G. E. and M. D. Crutcher (1990). "Functional architecture of basal ganglia circuits: neural substrates of parallel processing." Trends Neurosci 13(7): 266-71. Alilain, W. J., X. Li, et al. (2008). "Light-induced rescue of breathing after spinal cord injury." J Neurosci 28(46): 11862-70. Aravanis, A. M., L. P. Wang, et al. (2007). "An optical neural interface: in vivo control of rodent motor cortex with integrated fiberoptic and optogenetic technology." J Neural Eng 4(3): S143-56. Arenkiel, B. R. and J. Peca (2009). "Using light to reinstate respiratory plasticity." J Neurophysiol 101(4): 1695-8. Arenkiel, B. R., J. Peca, et al. (2007). "In vivo light-induced activation of neural circuitry in transgenic mice expressing channelrhodopsin-2." Neuron 54(2): 205-18. Arle, J. E., D. Apetauerova, et al. (2008). "Motor cortex stimulation in patients with Parkinson disease: 12-month follow-up in 4 patients." J Neurosurg 109(1): 133-9. Atasoy, D., Y. Aponte, et al. (2008). "A FLEX switch targets Channelrhodopsin-2 to multiple cell types for imaging and long-range circuit mapping." J Neurosci 28(28): 7025-30. Banghart, M., K. Borges, et al. (2004). "Light-activated ion channels for remote control of neuronal firing." Nat Neurosci 7(12): 1381-6. Bekar, L., W. Libionka, et al. (2008). "Adenosine is crucial for deep brain stimulation- mediated attenuation of tremor." Nat Med 14(1): 75-80. Bellone, C. and R. A. Nicoll (2007). "Rapid bidirectional switching of synaptic NMDA receptors." Neuron 55(5): 779-85. Benabid, A. L., P. Pollak, et al. (1994). "Acute and long-term effects of subthalamic nucleus stimulation in Parkinson's disease." Stereotact Funct Neurosurg 62(1-4): 76-84. Berndt, A., O. Yizhar, et al. (2009). "Bi-stable neural state switches." Nat Neurosci 12(2): 229- 34. Bi, A., J. Cui, et al. (2006). "Ectopic expression of a microbial-type rhodopsin restores visual responses in mice with photoreceptor degeneration." Neuron 50(1): 23-33.

131

Bi, G. Q. and M. M. Poo (1998). "Synaptic modifications in cultured hippocampal neurons: dependence on spike timing, synaptic strength, and postsynaptic cell type." J Neurosci 18(24): 10464-72. Bochkov, Y. A. and A. C. Palmenberg (2006). "Translational efficiency of EMCV IRES in bicistronic vectors is dependent upon IRES sequence and gene location." Biotechniques 41(3): 283-4, 286, 288 passim. Bocquet, N., L. Prado de Carvalho, et al. (2007). "A prokaryotic proton-gated ion channel from the nicotinic acetylcholine receptor family." Nature 445(7123): 116-9. Boldogkoi, Z., K. Balint, et al. (2009). "Genetically timed, activity-sensor and rainbow transsynaptic viral tools." Nat Methods 6(2): 127-30. Borrell, V., Y. Yoshimura, et al. (2005). "Targeted gene delivery to telencephalic inhibitory neurons by directional in utero electroporation." J Neurosci Methods 143(2): 151-8. Boyden, E. S., F. Zhang, et al. (2005). "Millisecond-timescale, genetically targeted optical control of neural activity." Nat Neurosci 8(9): 1263-8. Briggs, W. R., C. F. Beck, et al. (2001). "The phototropin family of photoreceptors." Plant Cell 13(5): 993-7. Brown, P., A. Oliviero, et al. (2001). "Dopamine dependency of oscillations between subthalamic nucleus and pallidum in Parkinson's disease." J Neurosci 21(3): 1033-8. Callaway, E. M. (2008). "Transneuronal circuit tracing with neurotropic viruses." Curr Opin Neurobiol 18(6): 617-23. Canavero, S. and R. Paolotti (2000). "Extradural motor cortex stimulation for advanced Parkinson's disease: case report." Mov Disord 15(1): 169-71. Canavero, S., R. Paolotti, et al. (2002). "Extradural motor cortex stimulation for advanced Parkinson disease. Report of two cases." J Neurosurg 97(5): 1208-11. Cardin, J. A., M. Carlen, et al. (2009). "Driving fast-spiking cells induces gamma rhythm and controls sensory responses." Nature 459(7247): 663-7. Cetin, A., S. Komai, et al. (2006). "Stereotaxic gene delivery in the rodent brain." Nat Protoc 1(6): 3166-73. Chambers, J. J., M. R. Banghart, et al. (2006). "Light-induced depolarization of neurons using a modified Shaker K(+) channel and a molecular photoswitch." J Neurophysiol 96(5): 2792-6. Cilia, R., A. Landi, et al. (2007). "Extradural motor cortex stimulation in Parkinson's disease." Mov Disord 22(1): 111-4. Colechio, E. M. and K. D. Alloway (2009). "Differential topography of the bilateral cortical projections to the whisker and forepaw regions in rat motor cortex." Brain Struct Funct 213(4-5): 423-39. Crawley, J. N., J. K. Belknap, et al. (1997). "Behavioral phenotypes of inbred mouse strains: implications and recommendations for molecular studies." Psychopharmacology (Berl) 132(2): 107-24. Crawley, N. J. (2007). What's Wrong with My Mouse?: Behavioral Phenotyping of Transgenic and Knockout Mice, 2nd Edition, Wiley. Degos, B., J. M. Deniau, et al. (2008). "Evidence for a direct subthalamo-cortical loop circuit in the rat." Eur J Neurosci 27(10): 2599-610. Deisseroth, K., G. Feng, et al. (2006). "Next-generation optical technologies for illuminating genetically targeted brain circuits." J Neurosci 26(41): 10380-6. DeLong, M. R. (1990). "Primate models of movement disorders of basal ganglia origin." Trends Neurosci 13(7): 281-5.

132

Dittgen, T., A. Nimmerjahn, et al. (2004). "Lentivirus-based genetic manipulations of cortical neurons and their optical and electrophysiological monitoring in vivo." Proc Natl Acad Sci U S A 101(52): 18206-11. Dostrovsky, J. O. and A. M. Lozano (2002). "Mechanisms of deep brain stimulation." Mov Disord 17 Suppl 3: S63-8. Douglass, A. D., S. Kraves, et al. (2008). "Escape behavior elicited by single, channelrhodopsin-2-evoked spikes in zebrafish somatosensory neurons." Curr Biol 18(15): 1133-7. Drouot, X., S. Oshino, et al. (2004). "Functional recovery in a primate model of Parkinson's disease following motor cortex stimulation." Neuron 44(5): 769-78. Ehrlich, B. E., C. R. Schen, et al. (1984). "Bacterial rhodopsins monitored with fluorescent dyes in vesicles and in vivo." J Membr Biol 82(1): 89-94. Ellgaard, L. and A. Helenius (2003). "Quality control in the endoplasmic reticulum." Nat Rev Mol Cell Biol 4(3): 181-91. Fan, Y., D. Fricker, et al. (2005). "Activity-dependent decrease of excitability in rat hippocampal neurons through increases in I(h)." Nat Neurosci 8(11): 1542-51. Farhan, H., V. Reiterer, et al. (2008). "Signal-dependent export of GABA transporter 1 from the ER-Golgi intermediate compartment is specified by a C-terminal motif." J Cell Sci 121(Pt 6): 753-61. Fregni, F., D. K. Simon, et al. (2005). "Non-invasive brain stimulation for Parkinson's disease: a systematic review and meta-analysis of the literature." J Neurol Neurosurg Psychiatry 76(12): 1614-23. Freund, T. F. and G. Buzsaki (1996). "Interneurons of the hippocampus." Hippocampus 6(4): 347-470. Garaschuk, O., O. Griesbeck, et al. (2007). "Troponin C-based biosensors: A new family of genetically encoded indicators for in vivo calcium imaging in the nervous system." Cell Calcium. Gorostiza, P., M. Volgraf, et al. (2007). "Mechanisms of photoswitch conjugation and light activation of an ionotropic glutamate receptor." Proc Natl Acad Sci U S A 104(26): 10865-70. Gradinaru, V., M. Mogri, et al. (2009). "Optical deconstruction of parkinsonian neural circuitry." Science 324(5925): 354-9. Gradinaru, V., K. R. Thompson, et al. (2008). "eNpHR: a Natronomonas halorhodopsin enhanced for optogenetic applications." Brain Cell Biol 36(1-4): 129-39. Gradinaru, V., K. R. Thompson, et al. (2008). "eNpHR: a Natronomonas halorhodopsin enhanced for optogenetic applications." Brain Cell Biol. Gradinaru, V., K. R. Thompson, et al. (2007). "Targeting and readout strategies for fast optical neural control in vitro and in vivo." J Neurosci 27(52): 14231-8. Gross, R. E. and J. D. Rolston (2008). "The clinical utility of methods to determine spatial extent and volume of tissue activated by deep brain stimulation." Clin Neurophysiol 119(9): 1947-50. Guan, Z., M. Giustetto, et al. (2002). "Integration of long-term-memory-related synaptic plasticity involves bidirectional regulation of gene expression and chromatin structure." Cell 111(4): 483-93. Guerrero, G., D. F. Reiff, et al. (2005). "Heterogeneity in synaptic transmission along a Drosophila larval motor axon." Nat Neurosci 8(9): 1188-96.

133

Hamani, C., J. A. Saint-Cyr, et al. (2004). "The subthalamic nucleus in the context of movement disorders." Brain 127(Pt 1): 4-20. Han, X. and E. S. Boyden (2007). "Multiple-color optical activation, silencing, and desynchronization of neural activity, with single-spike temporal resolution." PLoS ONE 2: e299. Han, X., X. Qian, et al. (2009). "Millisecond-timescale optical control of neural dynamics in the nonhuman primate brain." Neuron 62(2): 191-8. Han, X., X. Qian, et al. (2009). "Informational lesions: optical perturbation of spike timing and neural synchrony via microbial opsin gene fusions." Front Mol Neurosci 2: 12. Heim, N., O. Garaschuk, et al. (2007). "Improved calcium imaging in transgenic mice expressing a troponin C-based biosensor." Nat Methods 4(2): 127-9. Heintz, N. (2001). "BAC to the future: the use of bac transgenic mice for neuroscience research." Nat Rev Neurosci 2(12): 861-70. Hira, R., N. Honkura, et al. (2009). "Transcranial optogenetic stimulation for functional mapping of the motor cortex." J Neurosci Methods 179(2): 258-63. Hofherr, A., B. Fakler, et al. (2005). "Selective Golgi export of Kir2.1 controls the stoichiometry of functional Kir2.x channel heteromers." J Cell Sci 118(Pt 9): 1935-43. Holst, J., A. L. Szymczak-Workman, et al. (2006). "Generation of T-cell receptor retrogenic mice." Nat Protoc 1(1): 406-17. Horton, A. C. and M. D. Ehlers (2003). "Dual modes of endoplasmic reticulum-to-Golgi transport in dendrites revealed by live-cell imaging." J Neurosci 23(15): 6188-99. Huala, E., P. W. Oeller, et al. (1997). "Arabidopsis NPH1: a protein kinase with a putative redox-sensing domain." Science 278(5346): 2120-3. Huang, Y. and H. G. Goshgarian (2009). "Identification of the neural pathway underlying spontaneous crossed phrenic activity in neonatal rats." Neuroscience. Huber, D., L. Petreanu, et al. (2008). "Sparse optical microstimulation in barrel cortex drives learned behaviour in freely moving mice." Nature 451(7174): 61-4. Hwang, R. Y., L. Zhong, et al. (2007). "Nociceptive neurons protect Drosophila larvae from parasitoid wasps." Curr Biol 17(24): 2105-16. Iseki, M., S. Matsunaga, et al. (2002). "A blue-light-activated adenylyl cyclase mediates photoavoidance in Euglena gracilis." Nature 415(6875): 1047-51. Isenberg, K. E. and G. E. Meyer (1989). "Cloning of a putative neuronal nicotinic acetylcholine receptor subunit." J Neurochem 52(3): 988-91. Ishizuka, T., M. Kakuda, et al. (2006). "Kinetic evaluation of photosensitivity in genetically engineered neurons expressing green algae light-gated channels." Neurosci Res 54(2): 85-94. Jasnow, A. M., K. J. Ressler, et al. (2009). "Distinct Subtypes of Cholecystokinin (CCK)- Containing Interneurons of the Basolateral Amygdala Identified Using a CCK Promoter-Specific Lentivirus." J Neurophysiol 101(3): 1494-506. Kalaidzidis, I. V., Y. L. Kalaidzidis, et al. (1998). "Flash-induced voltage changes in halorhodopsin from Natronobacterium pharaonis." FEBS Lett 427(1): 59-63. Karpova, A. Y., D. G. Tervo, et al. (2005). "Rapid and reversible chemical inactivation of synaptic transmission in genetically targeted neurons." Neuron 48(5): 727-35. Khedr, E. M., J. C. Rothwell, et al. (2006). "Effect of daily repetitive transcranial magnetic stimulation on motor performance in Parkinson's disease." Mov Disord 21(12): 2201- 5.

134

Kissa, K., E. Mordelet, et al. (2002). "In vivo neuronal tracing with GFP-TTC gene delivery." Mol Cell Neurosci 20(4): 627-37. Kotlikoff, M. I. (2007). "Genetically encoded Ca2+ indicators: using genetics and molecular design to understand complex physiology." J Physiol 578(Pt 1): 55-67. Kringelbach, M. L., N. Jenkinson, et al. (2007). "Translational principles of deep brain stimulation." Nat Rev Neurosci 8(8): 623-35. Kuhlman, S. J. and Z. J. Huang (2008). "High-resolution labeling and functional manipulation of specific neuron types in mouse brain by Cre-activated viral gene expression." PLoS ONE 3(4): e2005. Lanyi, J. K. and D. Oesterhelt (1982). "Identification of the retinal-binding protein in halorhodopsin." J Biol Chem 257(5): 2674-7. Lee, H. K., M. Barbarosie, et al. (2000). "Regulation of distinct AMPA receptor phosphorylation sites during bidirectional synaptic plasticity." Nature 405(6789): 955-9. Lee, H. K., K. Takamiya, et al. (2003). "Phosphorylation of the AMPA receptor GluR1 subunit is required for synaptic plasticity and retention of spatial memory." Cell 112(5): 631-43. Lefaucheur, J. P., X. Drouot, et al. (2004). "Improvement of motor performance and modulation of cortical excitability by repetitive transcranial magnetic stimulation of the motor cortex in Parkinson's disease." Clin Neurophysiol 115(11): 2530-41. Lein, E. S., M. J. Hawrylycz, et al. (2007). "Genome-wide atlas of gene expression in the adult mouse brain." Nature 445(7124): 168-76. Lein, E. S., M. J. Hawrylycz, et al. (2007). "Genome-wide atlas of gene expression in the adult mouse brain." Nature 445(7124): 168-76. Leone, D. P., K. Srinivasan, et al. (2008). "The determination of projection neuron identity in the developing cerebral cortex." Curr Opin Neurobiol 18(1): 28-35. Lerchner, W., C. Xiao, et al. (2007). "Reversible silencing of neuronal excitability in behaving mice by a genetically targeted, ivermectin-gated Cl- channel." Neuron 54(1): 35-49. Levitan, E. S. and K. Takimoto (2000). "Surface expression of Kv1 voltage-gated K+ channels is governed by a C-terminal motif." Trends Cardiovasc Med 10(7): 317-20. Levskaya, A., O. D. Weiner, et al. (2009). "Spatiotemporal control of cell signalling using a light-switchable protein interaction." Nature. Levy, R., A. E. Lang, et al. (2001). "Lidocaine and muscimol microinjections in subthalamic nucleus reverse Parkinsonian symptoms." Brain 124(Pt 10): 2105-18. Lewis, T. L., Jr., T. Mao, et al. (2009). "Myosin-dependent targeting of transmembrane proteins to neuronal dendrites." Nat Neurosci 12(5): 568-76. Li, D., K. Takimoto, et al. (2000). "Surface expression of Kv1 channels is governed by a C- terminal motif." J Biol Chem 275(16): 11597-602. Li, X., D. V. Gutierrez, et al. (2005). "Fast noninvasive activation and inhibition of neural and network activity by vertebrate rhodopsin and green algae channelrhodopsin." Proc Natl Acad Sci U S A 102(49): 17816-21. Lima, S. Q., T. Hromadka, et al. (2009). "PINP: a new method of tagging neuronal populations for identification during in vivo electrophysiological recording." PLoS ONE 4(7): e6099. Lima, S. Q. and G. Miesenbock (2005). "Remote control of behavior through genetically targeted photostimulation of neurons." Cell 121(1): 141-52. Limousin, P., P. Pollak, et al. (1995). "Effect of parkinsonian signs and symptoms of bilateral subthalamic nucleus stimulation." Lancet 345(8942): 91-5.

135

Lin, J. Y., M. Z. Lin, et al. (2009). "Characterization of engineered channelrhodopsin variants with improved properties and kinetics." Biophys J 96(5): 1803-14. Liu, Y., N. Postupna, et al. (2008). "High frequency deep brain stimulation: what are the therapeutic mechanisms?" Neurosci Biobehav Rev 32(3): 343-51. Lozano, A. M., H. S. Mayberg, et al. (2008). "Subcallosal cingulate gyrus deep brain stimulation for treatment-resistant depression." Biol Psychiatry 64(6): 461-7. Lozier, R. H., R. A. Bogomolni, et al. (1975). "Bacteriorhodopsin: a light-driven proton pump in Halobacterium Halobium." Biophys J 15(9): 955-62. Luan, H. and B. H. White (2007). "Combinatorial methods for refined neuronal gene targeting." Curr Opin Neurobiol 17(5): 572-80. Luo, L., E. M. Callaway, et al. (2008). "Genetic dissection of neural circuits." Neuron 57(5): 634-60. Ma, D., N. Zerangue, et al. (2001). "Role of ER export signals in controlling surface potassium channel numbers." Science 291(5502): 316-9. Mank, M., D. F. Reiff, et al. (2006). "A FRET-based calcium biosensor with fast signal kinetics and high fluorescence change." Biophys J 90(5): 1790-6. Marti, T., H. Otto, et al. (1991). "Bacteriorhodopsin mutants containing single substitutions of serine or threonine residues are all active in proton translocation." J Biol Chem 266(11): 6919-27. Maskos, U., K. Kissa, et al. (2002). "Retrograde trans-synaptic transfer of green fluorescent protein allows the genetic mapping of neuronal circuits in transgenic mice." Proc Natl Acad Sci U S A 99(15): 10120-5. Matsuya, T., K. Takuma, et al. (2007). "Synergistic effects of adenosine A2A antagonist and L- DOPA on rotational behaviors in 6-hydroxydopamine-induced hemi-Parkinsonian mouse model." J Pharmacol Sci 103(3): 329-32. Mayberg, H. S., A. M. Lozano, et al. (2005). "Deep brain stimulation for treatment-resistant depression." Neuron 45(5): 651-60. McIntyre, C. C. and W. M. Grill (2002). "Extracellular stimulation of central neurons: influence of stimulus waveform and frequency on neuronal output." J Neurophysiol 88(4): 1592-604. McIntyre, C. C., W. M. Grill, et al. (2004). "Cellular effects of deep brain stimulation: model- based analysis of activation and inhibition." J Neurophysiol 91(4): 1457-69. McIntyre, C. C., M. Savasta, et al. (2004). "Uncovering the mechanism(s) of action of deep brain stimulation: activation, inhibition, or both." Clin Neurophysiol 115(6): 1239-48. McNeely, H. E., H. S. Mayberg, et al. (2008). "Neuropsychological impact of Cg25 deep brain stimulation for treatment-resistant depression: preliminary results over 12 months." J Nerv Ment Dis 196(5): 405-10. Metz, G. A., A. Tse, et al. (2005). "The unilateral 6-OHDA rat model of Parkinson's disease revisited: an electromyographic and behavioural analysis." Eur J Neurosci 22(3): 735- 44. Migliore, M. and G. M. Shepherd (2005). "Opinion: an integrated approach to classifying neuronal phenotypes." Nat Rev Neurosci 6(10): 810-8. Molyneaux, B. J., P. Arlotta, et al. (2007). "Neuronal subtype specification in the cerebral cortex." Nat Rev Neurosci 8(6): 427-37. Nagai, T., A. Sawano, et al. (2001). "Circularly permuted green fluorescent proteins engineered to sense Ca2+." Proc Natl Acad Sci U S A 98(6): 3197-202.

136

Nagel, G., M. Brauner, et al. (2005). "Light activation of channelrhodopsin-2 in excitable cells of Caenorhabditis elegans triggers rapid behavioral responses." Curr Biol 15(24): 2279-84. Nagel, G., D. Ollig, et al. (2002). "Channelrhodopsin-1: a light-gated proton channel in green algae." Science 296(5577): 2395-8. Nagel, G., T. Szellas, et al. (2003). "Channelrhodopsin-2, a directly light-gated cation-selective membrane channel." Proc Natl Acad Sci U S A 100(24): 13940-5. Nagel, G., T. Szellas, et al. (2005). "Channelrhodopsins: directly light-gated cation channels." Biochem Soc Trans 33(Pt 4): 863-6. Nakai, J., M. Ohkura, et al. (2001). "A high signal-to-noise Ca(2+) probe composed of a single green fluorescent protein." Nat Biotechnol 19(2): 137-41. Navarro-Quiroga, I., R. Chittajallu, et al. (2007). "Long-term, selective gene expression in developing and adult hippocampal pyramidal neurons using focal in utero electroporation." J Neurosci 27(19): 5007-11. Ntefidou, M., M. Iseki, et al. (2003). "Photoactivated adenylyl cyclase controls phototaxis in the flagellate Euglena gracilis." Plant Physiol 133(4): 1517-21. Pagni, C. A., S. Zeme, et al. (2003). "Further experience with extradural motor cortex stimulation for treatment of advanced Parkinson's disease. Report of 3 new cases." J Neurosurg Sci 47(4): 189-93. Parrish, A. R., W. Wang, et al. (2006). "Manipulating proteins for neuroscience." Curr Opin Neurobiol 16(5): 585-92. Paxinos, G. and K. Franklin (2001). "The Mouse Brain in Stereotaxic Coordinates 2nd edn. Academic Press: New York.". Paxinos, G., C. Watson, et al. (1985). "Bregma, lambda and the interaural midpoint in stereotaxic surgery with rats of different sex, strain and weight." J Neurosci Methods 13(2): 139-43. Payoux, P., P. Remy, et al. (2004). "Subthalamic nucleus stimulation reduces abnormal motor cortical overactivity in Parkinson disease." Arch Neurol 61(8): 1307-13. Perreault, M. C., A. P. Bernier, et al. (2006). "C fragment of tetanus toxin hybrid proteins evaluated for muscle-specific transsynaptic mapping of spinal motor circuitry in the newborn mouse." Neuroscience 141(2): 803-16. Perrimon, N. (1998). "New advances in Drosophila provide opportunities to study gene functions." Proc Natl Acad Sci U S A 95(17): 9716-7. Petreanu, L., D. Huber, et al. (2007). "Channelrhodopsin-2-assisted circuit mapping of long- range callosal projections." Nat Neurosci 10(5): 663-8. Petreanu, L., T. Mao, et al. (2009). "The subcellular organization of neocortical excitatory connections." Nature 457(7233): 1142-5. Petrecca, K., D. M. Miller, et al. (2000). "Localization and enhanced current density of the Kv4.2 potassium channel by interaction with the actin-binding protein filamin." J Neurosci 20(23): 8736-44. Pettit, D. L., S. S. Wang, et al. (1997). "Chemical two-photon uncaging: a novel approach to mapping glutamate receptors." Neuron 19(3): 465-71. Pulver, S. R., S. L. Pashkovski, et al. (2009). "Temporal dynamics of neuronal activation by Channelrhodopsin-2 and TRPA1 determine behavioral output in Drosophila larvae." J Neurophysiol 101(6): 3075-88.

137

Ratzliff, A. H., A. L. Howard, et al. (2004). "Rapid deletion of mossy cells does not result in a hyperexcitable dentate gyrus: implications for epileptogenesis." J Neurosci 24(9): 2259-69. Reiff, D. F., A. Ihring, et al. (2005). "In vivo performance of genetically encoded indicators of neural activity in flies." J Neurosci 25(19): 4766-78. Ressler, K. J. and H. S. Mayberg (2007). "Targeting abnormal neural circuits in mood and anxiety disorders: from the laboratory to the clinic." Nat Neurosci 10(9): 1116-24. Rezai, A. R., A. G. Machado, et al. (2008). "Surgery for movement disorders." Neurosurgery 62 Suppl 2: 809-38; discussion 838-9. Rickgauer, J. P. and D. W. Tank (2009). "Two-photon excitation of channelrhodopsin-2 at saturation." Proc Natl Acad Sci U S A. Ridding, M. C., R. Inzelberg, et al. (1995). "Changes in excitability of motor cortical circuitry in patients with Parkinson's disease." Ann Neurol 37(2): 181-8. Rivkees, S. A., S. L. Price, et al. (1995). "Immunohistochemical detection of A1 adenosine receptors in rat brain with emphasis on localization in the hippocampal formation, cerebral cortex, cerebellum, and basal ganglia." Brain Res 677(2): 193-203. Rusch, D. B., A. L. Halpern, et al. (2007). "The Sorcerer II Global Ocean Sampling expedition: northwest Atlantic through eastern tropical Pacific." PLoS Biol 5(3): e77. Ryan, M. D. and J. Drew (1994). "Foot-and-mouth disease virus 2A oligopeptide mediated cleavage of an artificial polyprotein." Embo J 13(4): 928-33. Sano, H., Y. Nagai, et al. (2007). "Inducible expression of retrograde transynaptic genetic tracer in mice." Genesis 45(3): 123-8. Sasaki, J., L. S. Brown, et al. (1995). "Conversion of bacteriorhodopsin into a chloride ion pump." Science 269(5220): 73-5. Schroder-Lang, S., M. Schwarzel, et al. (2007). "Fast manipulation of cellular cAMP level by light in vivo." Nat Methods 4(1): 39-42. Schroll, C., T. Riemensperger, et al. (2006). "Light-induced activation of distinct modulatory neurons triggers appetitive or aversive learning in Drosophila larvae." Curr Biol 16(17): 1741-7. Shen, K. Z. and S. W. Johnson (2003). "Presynaptic inhibition of synaptic transmission by adenosine in rat subthalamic nucleus in vitro." Neuroscience 116(1): 99-106. Shi, S., Y. Hayashi, et al. (2001). "Subunit-specific rules governing AMPA receptor trafficking to synapses in hippocampal pyramidal neurons." Cell 105(3): 331-43. Shu, X., A. Royant, et al. (2009). "Mammalian expression of infrared fluorescent proteins engineered from a bacterial phytochrome." Science 324(5928): 804-7. Silberberg, G., C. Wu, et al. (2004). "Synaptic dynamics control the timing of neuronal excitation in the activated neocortical microcircuit." J Physiol 556(Pt 1): 19-27. Simon, S. M. and G. Blobel (1993). "Mechanisms of translocation of proteins across membranes." Subcell Biochem 21: 1-15. Sineshchekov, O. A., E. G. Govorunova, et al. (2005). "Rhodopsin-mediated photoreception in cryptophyte flagellates." Biophys J 89(6): 4310-9. Smith, Y. and A. Parent (1988). "Neurons of the subthalamic nucleus in primates display glutamate but not GABA immunoreactivity." Brain Res 453(1-2): 353-6. Sohal, V. S., F. Zhang, et al. (2009). "Parvalbumin neurons and gamma rhythms enhance cortical circuit performance." Nature 459(7247): 698-702. Srinivas, S., T. Watanabe, et al. (2001). "Cre reporter strains produced by targeted insertion of EYFP and ECFP into the ROSA26 locus." BMC Dev Biol 1: 4.

138

Stockklausner, C. and N. Klocker (2003). "Surface expression of inward rectifier potassium channels is controlled by selective Golgi export." J Biol Chem 278(19): 17000-5. Stockklausner, C., J. Ludwig, et al. (2001). "A sequence motif responsible for ER export and surface expression of Kir2.0 inward rectifier K(+) channels." FEBS Lett 493(2-3): 129- 33. Stoeckenius, W. and R. A. Bogomolni (1982). "Bacteriorhodopsin and related pigments of halobacteria." Annu Rev Biochem 51: 587-616. Strafella, A. P., A. M. Lozano, et al. (2007). "Subdural motor cortex stimulation in Parkinson's disease does not modify movement-related rCBF pattern." Mov Disord 22(14): 2113- 6. Sugita, M. and Y. Shiba (2005). "Genetic tracing shows segregation of taste neuronal circuitries for bitter and sweet." Science 309(5735): 781-5. Svoboda, K. and R. Yasuda (2006). "Principles of two-photon excitation microscopy and its applications to neuroscience." Neuron 50(6): 823-39. Szobota, S., P. Gorostiza, et al. (2007). "Remote control of neuronal activity with a light-gated glutamate receptor." Neuron 54(4): 535-45. Tabar, V., M. Tomishima, et al. (2008). "Therapeutic cloning in individual parkinsonian mice." Nat Med 14(4): 379-81. Tan, E. M., Y. Yamaguchi, et al. (2006). "Selective and quickly reversible inactivation of mammalian neurons in vivo using the Drosophila allatostatin receptor." Neuron 51(2): 157-70. Tan, W., W. A. Janczewski, et al. (2008). "Silencing preBotzinger complex somatostatin- expressing neurons induces persistent apnea in awake rat." Nat Neurosci 11(5): 538- 40. Tang, W., I. Ehrlich, et al. (2009). "Faithful expression of multiple proteins via 2A-peptide self- processing: a versatile and reliable method for manipulating brain circuits." J Neurosci 29(27): 8621-9. Tengholm, A. and E. Gylfe (2009). "Oscillatory control of insulin secretion." Mol Cell Endocrinol 297(1-2): 58-72. Thompson, S. M., J. P. Kao, et al. (2005). "Flashy science: controlling neural function with light." J Neurosci 25(45): 10358-65. Tsai, H. C., F. Zhang, et al. (2009). "Phasic firing in dopaminergic neurons is sufficient for behavioral conditioning." Science 324(5930): 1080-4. Tsunoda, S. P., D. Ewers, et al. (2006). "H+ -pumping rhodopsin from the marine alga Acetabularia." Biophys J 91(4): 1471-9. Venter, J. C., K. Remington, et al. (2004). "Environmental genome shotgun sequencing of the Sargasso Sea." Science 304(5667): 66-74. Vitek, J. L. (2002). "Mechanisms of deep brain stimulation: excitation or inhibition." Mov Disord 17 Suppl 3: S69-72. Wang, H., J. Peca, et al. (2007). "High-speed mapping of synaptic connectivity using photostimulation in Channelrhodopsin-2 transgenic mice." Proc Natl Acad Sci U S A 104(19): 8143-8. Wang, L. P. and K. Deisseroth (unpublished observations). unpublished observations. Wang, T., Y. Jiao, et al. (2007). "Dissection of the pathway required for generation of vitamin A and for Drosophila phototransduction." J Cell Biol 177(2): 305-16.

139

Weick, J. P., R. D. Groth, et al. (2003). "Interactions with PDZ proteins are required for L-type calcium channels to activate cAMP response element-binding protein-dependent gene expression." J Neurosci 23(8): 3446-56. Wickersham, I. R., D. C. Lyon, et al. (2007). "Monosynaptic restriction of transsynaptic tracing from single, genetically targeted neurons." Neuron 53(5): 639-47. Williamson, S. J., D. B. Rusch, et al. (2008). "The Sorcerer II Global Ocean Sampling Expedition: metagenomic characterization of viruses within aquatic microbial samples." PLoS One 3(1): e1456. Wu, Y. I., D. Frey, et al. (2009). "A genetically encoded photoactivatable Rac controls the motility of living cells." Nature 461(7260): 104-8. Wyart, C., F. Del Bene, et al. (2009). "Optogenetic dissection of a behavioural module in the vertebrate spinal cord." Nature 461(7262): 407-10. Yoder, E. J. and D. Kleinfeld (2002). "Cortical imaging through the intact mouse skull using two-photon excitation laser scanning microscopy." Microsc Res Tech 56(4): 304-5. Yooseph, S., G. Sutton, et al. (2007). "The Sorcerer II Global Ocean Sampling expedition: expanding the universe of protein families." PLoS Biol 5(3): e16. Yoshimura, Y., J. L. Dantzker, et al. (2005). "Excitatory cortical neurons form fine-scale functional networks." Nature 433(7028): 868-73. Zhang, F. (2008). Fast optical neural circuit interrogation technology: development and applications. Larry M. Katz Memorial Lecture, Cold Spring Harbor Laboratory Meeting on Neuronal Circuits. Cold Spring Harbor Laboratory. Zhang, F., A. M. Aravanis, et al. (2007). "Circuit-breakers: optical technologies for probing neural signals and systems." Nat Rev Neurosci 8(8): 577-81. Zhang, F., A. M. Aravanis, et al. (2007b). "Circuit-breakers: optical technologies for probing neural signals and systems." Nat Rev Neurosci 8(8): 577-81. Zhang, F., M. Prigge, et al. (2008). "Red-shifted optogenetic excitation: a tool for fast neural control derived from Volvox carteri." Nat Neurosci 11(6): 631-3. Zhang, F., L. P. Wang, et al. (2006). "Channelrhodopsin-2 and optical control of excitable cells." Nat Methods 3(10): 785-92. Zhang, F., L. P. Wang, et al. (2007a). "Multimodal fast optical interrogation of neural circuitry." Nature 446(7136): 633-9. Zhang, J., F. Laiwalla, et al. (2009). "Integrated device for optical stimulation and spatiotemporal electrical recording of neural activity in light-sensitized brain tissue." J Neural Eng 6(5): 55007. Zhang, Y. and R. M. Harris-Warrick (2004). "An ER export signal accelerates the surface expression of shal potassium channels in pyloric neurons of the lobster stomatogastric ganglion." Pflugers Arch 447(4): 401-4. Zhang, Y. P., N. Holbro, et al. (2008). "Optical induction of plasticity at single synapses reveals input-specific accumulation of alphaCaMKII." Proc Natl Acad Sci U S A 105(33): 12039-44. Zhang, Y. P. and T. G. Oertner (2007). "Optical induction of synaptic plasticity using a light- sensitive channel." Nat Methods 4(2): 139-41. Zhang, Z. W. and M. Deschenes (1997). "Intracortical axonal projections of lamina VI cells of the primary somatosensory cortex in the rat: a single-cell labeling study." J Neurosci 17(16): 6365-79. Zhao, S., C. Cunha, et al. (2008). "Improved expression of halorhodopsin for light-induced silencing of neuronal activity." Brain Cell Biol 36(1-4): 141-54.

140

Zito, K., D. Parnas, et al. (1999). "Watching a synapse grow: noninvasive confocal imaging of synaptic growth in Drosophila." Neuron 22(4): 719-29.

141

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