This work was supported by the UNAM-PAPIIT project IA203520. We thank the IFC animal facility for their help with mouse colonies maintenance and the computational unit for IT support, especially to Francisco Perez-Eugenio.

The procedures involving animal use were conducted following local and national guidelines and approved by the corresponding Institutional Animal Care and Use Committee (Institute of Cellular Physiology IACUC protocol VLH151-19). Drd1-Cre transgenic male mice27, 35-40 days postnatal with C57BL/6 background were used in the current protocol. Mice were kept under the following conditions: temperature 22&#177;1 &#176;C; humidity 55%; light schedule 12/12 h with lights off at 7 p.m. and were weaned at postnatal day 21. Weaned pups were housed in same-sex groups of 2-5.&#160;Animals were housed in static housing with micro-barrier tops. Bedding consisted of sterile aspen shavings. Rodent pellets&#160;and RO-purified water were provided ad libitum, except when noted.
1. Surgical procedures
<ol>
	<li>Prepare an LED cannula at the desired length according to the dorsoventral coordinates of the structure of interest (ideally 0.5 mm longer to account for the thickness of the skull, for the dorsolateral striatum 3.5 mm) (Figure 1).
	<ol>
		<li>Cut the glass fiber to a length longer than the final desired size, grind the fiber tip to the target length with rough sandpaper, and finally, polish the fiber tip with fine sandpaper. 
		NOTE: LED cannula is a glass optical fiber of 250 &#181;m diameter attached to an infrared receiver (see Table of Materials).</li>
	</ol>
	</li>
	<li>Pull glass pipettes (1.14 mm outer diameter, 0.53 mm inner diameter, and 3.5 in length) for the nano-injector with a horizontal puller (see Table of Materials) and store them for later. Program the puller in one loop to get a 15-20 &#181;m tip diameter with a long gradual slope taper (4-5 mm).</li>
	<li>Prepare the surgery area by thoroughly disinfecting the stereotaxic apparatus, hood, micro-injector (see Table of Materials), and surrounding surfaces with 70% ethanol. 
	NOTE: A mouse stereotaxic apparatus is essential to inject Adeno Associated Virus (AAVs) precisely and place the LED cannula in the region of interest.</li>
	<li>Wear the appropriate personal protective equipment for the procedure, including a clean lab coat or disposable surgical gown, sterile gloves, face mask, and disposable head cap.</li>
	<li>Place the necessary equipment close to the surgery area, such as sterile surgical tools, cotton tips, solutions, micropipette, pipette tips, capillaries, micro-fill with mineral oil, and marker.</li>
	<li>Fill a pipette for microinjections with mineral oil and place it in the micro-injector. Make sure that the micro-injector is working correctly by ejecting some mineral oil. 
	NOTE: All instruments used during the surgery should be autoclaved and sterile. Aseptic technique should be used.</li>
	<li>Anesthetize animals with gaseous isoflurane 4-5% to induce anesthesia and 1.2% throughout the surgery with 0.5-1 L/min pure oxygen. The surgery begins only after the animal has reached a point of deep anesthesia, assessed by the absence of paw withdrawal after a slight pinch.
	<ol>
		<li>Continuously monitor the breathing rate and temperature of the animal. Maintain the body temperature by a heating pad set at 34 &#176;C.</li>
	</ol>
	</li>
	<li>Apply an ophthalmic ointment. Remove hair from the scalp with a trimer and hair removal cream. Wipe the scalp with cotton swabs having 8% povidone-iodine (see Table of Materials) and 70% ethanol alternated three times each.</li>
	<li>Place the mouse in the stereotaxic apparatus and secure the head, ensuring that the skull is leveled in the mediolateral and anterior-posterior axes.</li>
	<li>Make a 1 cm incision with a scalpel through the scalp at the level of the eyes along the sagittal axis. Retract the skin to expose the skull and clean the periosteum with cotton swabs.</li>
	<li>Clean the cranium surface with saline solution and sterile cotton swabs. Resolve any bleeding at the surface using sterile absorbent eye spears (see Table of Materials) or similar sterile absorbent material.</li>
	<li>Apply a drop of 2.5% hydrogen peroxide with a cotton swab and let it act for a few seconds to make the skull sutures visible and have a better reference. After a few seconds, clean thoroughly with a clean cotton swab.</li>
	<li>With the glass pipette (15 &#181;m final tip diameter), locate bregma and lambda to check that the skull is leveled in the anterior-posterior axis. 
	NOTE: It is recommended to have a stereoscopic microscope or USB microscope to see the tip of the glass pipette. In case it is needed, adjust the height of the mouth holder to level the skull.</li>
	<li>Move the capillary toward the selected anterior-posterior (AP) and medial-lateral (ML) coordinates (dorsolateral striatum AP 1.2 mm, ML 2.28 mm). Paint a reference point in the scalp above the selected coordinates with a sterile marker.</li>
	<li>In the reference point, perform an ~1 mm diameter craniotomy applying gentle pressure to the skull with a sterile rotary tool or dental drill at a low to medium speed with a small round dental drill bit (see Table of Materials).</li>
	<li>Load the capillary with 300-400 nL of Cre-dependent adeno-associated virus (AAV) such as AAV1-dflox-hChR-2-mCherry to express Channelrhodopsin or an AAV to express only the reporter protein (e.g., mCherry) as a control in the region of interest (see Table of Materials). Check that the tip is not clogged, then introduce the glass pipette in the brain at the desired dorso-ventral (DV) coordinates (dorsolateral striatum DV -3.35 mm).
	<ol>
		<li>Inject 200 nL using an automatic injector at a rate of 23 nL/s. Wait for 10 min after finishing the injection, withdraw the glass pipette slowly to avoid spillage. 
		NOTE: It is possible to use a 30 G needle to inject with the appropriate micro-injector.</li>
	</ol>
	</li>
	<li>Clean and dry any residues with cotton swabs.</li>
	<li>Attach the sterile&#160;glass LED cannula to the stereotaxic arm and calibrate the coordinates using bregma as a reference. Insert the cannula very slowly (300 &#181;m/min) to avoid tissue damage and place it 100 &#181;m above the injection site.</li>
	<li>Once the LED cannula is in place, add a drop (100 &#181;L) of tissue adhesive at the edge of the craniotomy.</li>
	<li>Prepare dental cement mixture (see Table of Materials) following the manufacturer&#39;s instructions to fix the fiber to the skull. 
	NOTE: Briefly, use a chilled porcelain dish to have more working time before cement sets. Add 2 scoops of resin clear powder to the porcelain dish, add 4 drops of quick base and 1 drop of catalyst, then mix well. The powder/liquid ratio can be adjusted if a thinner or thicker viscosity is needed.</li>
	<li>Using a sterile brush, apply the dental cement mixture around the cannula connector little by little, building layers until the skull is covered and the connector is securely attached to the skull, leaving the pins completely free. Avoid getting dental cement on the skin of the mouse.</li>
	<li>Allow drying completely.</li>
	<li>Close the skin around the implant using tissue adhesive (see Table of Materials).</li>
	<li>Place the mouse in a recovery cage over a heating pad at 33&#176; C. Monitor for the presence of one or more of the following signs of pain/discomfort: 1) Hunched up, lack or reduction of motor activity, 2) Failure to groom reflected in a unkept dirty coat, 3) Excessive licking or scratching, redness in the incision site, 4) aggressive behavior, 5) anorexia or dehydration, and 6) Lack of nest formation. 
	NOTE: Keep the mouse individually caged during all the procedures to avoid implant detaching. In case of detachment of the cannula, perform euthanasia by injecting 150 mg/kg of sodium pentobarbital followed by decapitation after deep anesthesia is reached.</li>
	<li>Inject subcutaneously (SC) meloxicam 1 mg/kg once daily for three days post-surgery to provide analgesia.</li>
	<li>Wait at least 7 days for complete recovery and 14 days for opsin expression before further procedures. 
	NOTE: Perform a postoperative follow-up every 12 hours for three days, then check animals every day until the day of euthanasia at the end of the experiment.</li>
</ol>

2. Reach-to-grasp training
<ol>
	<li>On day 7 post-surgery, start the food deprivation protocol28. Weigh mice for three consecutive days to determine their average ad libitum body weight. Then, schedule food restrictions so that the animals receive enough nutrients to maintain approximately 90% and not less than 85% of body weight. 
	NOTE: This is achieved by providing 2.5-3 g of food daily. Monitor animals&#39; weight daily and score for overall well-being observing animals&#39; behavior and appearance, for example, coat and eyes appearance. Use the body condition scoring system from Reference29.</li>
	<li>During the pre-training, training, and testing periods, provide each mouse with 20 pellets (20 mg of dustless chocolate-flavored pellets) daily (see Table of Materials) (eaten during the task or after) besides the standard food pellets.</li>
	<li>Three days before habituation, scatter 0.4 g/animal/day 20 mg of dustless chocolate-flavored pellets in their home cages, so mice get acquainted with the pellets that serve as a reward during the reach-to-grasp task.</li>
	<li>Habituate mice by placing them 10 min in the testing chamber one day before pre-training with pellets scattered on the chamber floor (Figure 1A).</li>
	<li>Allow food daily after training and testing. Keep a similar schedule every day.</li>
	<li>On the first day of pre-training, place the mice in the reach-to-grasp chamber and observe from the front. Place the pellets in front of the chamber close to the opening so that they start consuming the pellets. At this stage, mice are allowed to grab the pellets in any form.</li>
	<li>On day two of pre-training, place the pellets further and further from the opening until getting them to the indentation (1 cm from the opening) so mice can shape their reach-to-grasp movement (Figure 1C).</li>
	<li>Train mice to run to the rear of the cage and return to the cage opening to receive the next food pellet as a strategy to individualize trials. 
	NOTE: This can be achieved by waiting until the mouse is in the rear of the cage before placing a pellet in the indentation for each trial.</li>
	<li>Place pellets to be grasped by either their right or left paw. 
	NOTE: Mice start using preferentially one paw to grasp, which will be used the following days of training and testing.</li>
	<li>Train animals for 6 days in daily sessions lasting 20 trials or until a maximum of 10 min elapse. From day 2 of training, put the mock receiver (dimensions 12 x 18 x 7 mm, 1 g, see Table of Materials), so mice get habituated to the weight while performing the task (Figure 1B). Each day score the number of hit&#160;and missed trials.</li>
	<li>Record behavior with a regular camera and capture 30-60 frames/s from the front of the chamber. Additionally, one can place a mirror under the training chamber at a 45&#176; angle to monitor the animals&#39; posture (Figure 1D,E).</li>
	<li>For post-hoc kinematic analysis (Figure 2), mount a high-speed camera (see Table of Materials) at an angle of 45&#176; to record from the side of the cage. If a 3D analysis is required, place a second high-speed camera to record at a 35&#176; angle from the front of the chamber; both cameras should be placed in the right or left side of the cage depending on animals&#39; sidedness and should capture at the same frame rate and be synchronized7 (Figure 3D,E).</li>
	<li>Set the high-speed cameras to 100 frames/s with a resolution of 376 x 252 pixels or more if possible. Place white Styrofoam walls behind the sides and back of the chamber to reduce background and increase contrast (Figure 1E).</li>
	<li>On test day, replace the mock unit with an infrared receiver for wireless optogenetic stimulation (Figure 1B,C).</li>
	<li>When mice start reaching, turn the LED cannula manually with the remote controller to have a continuous stimulation for the time the behavior is performed and for no longer than 2 s. Programming an automatic stimulation paradigm is preferable. The stimulation device triggers an LED of 470 nm (blue light) with intensity at the tip of 1.0 mW/mm2.</li>
	<li>Collect the videos for further examination, including scoring and kinematic analysis.</li>
</ol>
3. Post-hoc histological confirmation
<ol>
	<li>Upon completion of an experiment, confirm viral expression and LED cannula placement. Anesthetize the animal with a cocktail of ketamine 100 mg/kg and xylazine 10 mg/kg. Once the mouse presents signs of deep anesthesia (step 1.7), perfuse with ice-cold phosphate-buffered saline (PBS) followed by 4% PFA.</li>
	<li>Remove implanted cannula carefully by firmly grasping the connector with forceps and pulling up gently.</li>
	<li>Extract and post-fix the brain for 24 h in 4% PFA23.</li>
	<li>Perform 3-10 min washes with PBS.</li>
	<li>Cut the brain in 50 &#181;m sections using a microtome (see Table of Materials).</li>
	<li>Mount the sections in slides with hard-set mounting media with DAPI to stain nuclei and cover slides.</li>
	<li>After drying, observe the sections under the confocal microscope and verify the implanted cannula location and expression of Ch2R fused with any fluorescent protein.</li>
</ol>

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M., Bizzi, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microstimulation+activates+a+handful+of+muscle+synergies.">Microstimulation activates a handful of muscle synergies.</a> Neuron. 76 (6), 1071-1077 (2012).</li><li>Miyazaki, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large+Timescale+interrogation+of+neuronal+function+by+fiberless+optogenetics+using+lanthanide+micro-particles.">Large Timescale interrogation of neuronal function by fiberless optogenetics using lanthanide micro-particles.</a> Cell Reports. 26 (4), 1033-1043 (2019).</li><li>Yang, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wireless+multilateral+devices+for+optogenetic+studies+of+individual+and+social+behaviors.">Wireless multilateral devices for optogenetic studies of individual and social behaviors.</a> Nature Neuroscience. 24 (7), 1035-1045 (2021).</li><li>Kampasi, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fiberless+multicolor+neural+optoelectrode+for+in+vivo+circuit+analysis.">Fiberless multicolor neural optoelectrode for in vivo circuit analysis.</a> Scientific Reports. 6, 30961 (2016).</li><li>Allen, B. D., Singer, A. C., Boyden, E. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+of+designing+interpretable+optogenetic+behavior+experiments.">Principles of designing interpretable optogenetic behavior experiments.</a> Learning &amp; Memory. 22 (4), 232-238 (2015).</li><li>Packer, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Nature Methods. 9, 1202-1205 (2012).</li></ol>

The authors declare no disclosures.

The use of optogenetic manipulation of neuronal populations in well-defined behavioral paradigms is advancing our knowledge about the mechanisms underlying motor control7,23. Wireless methods are especially suitable for tasks that require tests on multiple animals or free movement34,35. Nevertheless, as techniques and devices are refined, it should be the go-to option for any behavioral task combined with...

Motor skilled behavior is present during most movements performed by us, and it is known to be affected in several brain disorders1,2,3,4,5,6. Implementing tasks that allow studying the development, learning, and performance of skilled movements is crucial to understanding the motor function&#39;s neurobiological underpinnings, especially in models of brain injury, neurodegenerative and neurodevelopmental disorders2,7,8,9,10,11,12,13. Reaching for and retrieving objects is done routinely in everyday life actions, and it is one of the first motor skills acquired during early development and then refined through the years5,6. It comprises a complex behavior that requires sensory-motor processes such as the perception of the object&#39;s features, movement planning, action selection, movement execution, body coordination, and speed modulation7,14,15,16. Thus, unimanual high dexterity tasks require the participation of many brain structures of both hemispheres16,17,18,19,20,21,22. In mice, the single pellet reach-to-grasp task is characterized for several phases that can be controlled and analyzed separately7,13,23. This feature allows to study the contribution of specific neuronal subpopulations at different stages of acquisition and behavior performance and provides a platform for detailed studies of motor systems13,23,24. The movement occurs in a couple of seconds; thus, high-speed videography should be used for kinematic analysis in distinct stages of the skilled motor trajectory7,25. Several parameters can be extracted from the videos, including body posture, trajectory, velocity, and type of errors25. Kinematic analysis can be used to detect subtle changes during wireless optogenetic manipulation7,23.
Using miniaturized light-emitting diodes (LEDs) to deliver light via a wireless head-mounted system makes it possible to have remote optogenetic control while the animal performs the task. The wireless optogenetic controller accepts single-pulse or continuous trigger commands from a stimulator and sends infrared (IR) signals to a receiver connected to the miniaturized LED23,26. The current protocol combines this wireless optogenetics approach with high-speed videography of a dexterity task to dissect the role of specific neuronal populations during the performance of fine motor behavior23. Since it is a unimanual task, it allows for assessing the participation of structures in both hemispheres. Traditionally, the brain controls the body movement in a highly asymmetric manner; however, high dexterity tasks require careful coordination and control from many brain structures, including ipsilateral nuclei and differential contribution of neuronal subpopulations within nuclei10,20,21,22,23. This protocol shows that subcortical structures from both hemispheres control the trajectory of the forelimb23. This paradigm can be suitable to study other brain regions and models of brain disease.

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in vivo wireless optogenetic control of skilled motor behavior

Fine motor skills are essential in everyday life and can be compromised in several nervous system disorders. The acquisition and performance of these tasks require sensory-motor integration and involve precise control of bilateral brain circuits. Implementing unimanual behavioral paradigms in animal models will improve the understanding of the contribution of brain structures, like the striatum, to complex motor behavior as it allows manipulation and recording of neural activity of specific nuclei in control conditions and disease during the performance of the task. 
Since its creation, optogenetics has been a dominant tool for interrogating the brain by enabling selective and targeted activation or inhibition of neuronal populations. The combination of optogenetics with behavioral assays sheds light on the underlying mechanisms of specific brain functions. Wireless head-mounted systems with miniaturized light-emitting diodes (LEDs) allow remote optogenetic control in an entirely free-moving animal. This avoids the limitations of a wired system being less restrictive for animals' behavior without compromising light emission efficiency. The current protocol combines a wireless optogenetics approach with high-speed videography in a unimanual dexterity task to dissect the contribution of specific neuronal populations to fine motor behavior.

The reach-to-grasp task is a paradigm widely used to study shaping, learning, performance, and kinematics of fine skill movement under different experimental manipulations. Mice learn to execute the task in a couple of days and achieve more than 55% accuracy reaching a plateau after 5 days of training (Figure 2A,B). Similar to what has been previously reported, a percentage of animals do not perform the task appropriately (29.62%), and those should be excluded from further a...

Watch this Scientific Journal Video about In Vivo Wireless Optogenetic Control of Skilled Motor Behavior at JoVE.com

In Vivo Wireless Optogenetic Control of Skilled Motor Behavior

The present protocol describes how to use wireless optogenetics combined with high-speed videography in a single pellet reach-to-grasp task to characterize the neural circuits involved in the performance of skilled motor behavior in freely moving mice.

In Vivo Wireless Optogenetic Control of Skilled Motor Behavior

Fine motor skills are essential in everyday life and can be compromised in several nervous system disorders. The acquisition and performance of these ...

in-vivo-wireless-optogenetic-control-of-skilled-motor-behavior

Research

JoVE Journal

Neuroscience

3.2K Views.  National University of Mexico. The present protocol describes how to use wireless optogenetics combined with high-speed videography in a single pellet reach-to-grasp task to characterize the neural circuits involved in the performance of skilled motor behavior in freely moving mice. 

Video: In Vivo Wireless Optogenetic Control of Skilled Motor Behavior

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2 avoidance, proboscis extension and giant-fiber mediated startle response2-4. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-&beta;, encoded by the norpA gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.

Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster

Genetically encoded optogenetic tools enable noninvasive manipulation of specific neurons in the Drosophila brain. Such tools can identify neurons whose activation is sufficient to elicit or suppress particular behaviors. Here we present a method for activating Channelrhodopsin2 that is expressed in targeted neurons in freely walking flies.

A  growing number of genetically encoded tools are becoming available that allow non-invasive  manipulation of the neural activity of specific neurons in ...

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated proteins and subsequent manipulation of neural activity by light. Channelrhodopsins (ChRs) are light-gated cation-channels and when fused to a fluorescent protein their expression allows for visualization and concurrent activation of specific cell types and their axonal projections in defined areas of the brain. Via stereotactic injection of viral vectors, ChR fusion proteins can be constitutively or conditionally expressed in specific cells of a defined brain region, and their axonal projections can subsequently be studied anatomically and functionally via ex vivo optogenetic activation in brain slices. This is of particular importance when aiming to understand synaptic properties of connections that could not be addressed with conventional electrical stimulation approaches, or in identifying novel afferent and efferent connectivity that was previously poorly understood. Here, a few examples illustrate how this technique can be applied to investigate these questions to elucidating fear-related circuits in the amygdala. The amygdala is a key region for acquisition and expression of fear, and storage of fear and emotional memories. Many lines of evidence suggest that the medial prefrontal cortex (mPFC) participates in different aspects of fear acquisition and extinction, but its precise connectivity with the amygdala is just starting to be understood. First, it is shown how ex vivo optogenetic activation can be used to study aspects of synaptic communication between mPFC afferents and target cells in the basolateral amygdala (BLA). Furthermore, it is illustrated how this ex vivo optogenetic approach can be applied to assess novel connectivity patterns using a group of GABAergic neurons in the amygdala, the paracapsular intercalated cell cluster (mpITC), as an example.

Ex Vivo Optogenetic Dissection of Fear Circuits in Brain Slices

Optogenetic approaches are widely used to manipulate neural activity and assess the consequences for brain function. Here, a technique is outlined that&#160;upon in vivo expression of the optical activator Channelrhodopsin, allows for ex vivo analysis of synaptic properties of specific long range and local neural connections in fear-related circuits.

Optogenetic approaches are now widely used to study the function of neural populations and circuits by combining targeted expression of light-activated ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a ...

In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

The quest to understand how neural circuits process information in order to drive behavioral output has been greatly aided by recently-developed optical methods for manipulating and monitoring the activity of neurons in vivo. These types of experiments rely on two main components: 1) implantable devices that provide optical access to the brain, and 2) light-sensitive proteins that change neuronal excitability or provide a readout of neuronal activity. There are a number of ways to express light-sensitive proteins, but stereotaxic injection of viral vectors is currently the most flexible approach because expression can be controlled with genetic, anatomical, and temporal precision. Despite the great utility of viral vectors, delivering the virus to the site of optical implants poses numerous challenges. Stereotaxic virus injections are demanding surgeries that increase surgical time, increase the cost of studies, and pose a risk to the animal's health. The surrounding tissue can be physically damaged by the injection syringe, and by immunogenic inflammation caused by the abrupt delivery of a bolus of high-titer virus. Aligning injections with optical implants is especially difficult when targeting small regions deep in the brain. To overcome these challenges, we describe a method for coating multiple types of optical implants with films composed of silk fibroin and Adeno-associated viral (AAV) vectors. Fibroin, a polymer derived from the cocoon of Bombyx mori, can encapsulate and protect biomolecules and can be processed into forms ranging from soluble films to ceramics. When implanted into the brain, silk/AAV coatings release virus at the interface between optical elements and the surrounding brain, driving expression precisely where it is needed. This method is easily implemented and promises to greatly facilitate in vivo studies of neural circuit function.

In Vivo Targeted Expression of Optogenetic Proteins Using Silk/AAV Films

Here, we present a method for delivering viral expression vectors into the brain using silk fibroin films. This method allows targeted delivery of expression vectors using silk/AAV coated optical fibers, tapered optical fibers, and cranial windows.

The quest to understand how neural circuits process information in order to drive behavioral output has been greatly aided by recently-developed optical ...

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. Learning-induced changes in synaptic transmission are often distributed across many neurons and levels of processing in the brain. Therefore, methods to visualize learning-dependent synaptic plasticity across neurons are needed. The fruit fly Drosophila melanogaster represents a particularly favorable model organism to study neuronal circuits underlying learning. The protocol presented here demonstrates a way in which the processes underlying the formation of associative olfactory memories, i.e., synaptic activity and their changes, can be monitored in vivo. Using the broad array of genetic tools available in Drosophila, it is possible to specifically express genetically encoded calcium indicators in determined cell populations and even single cells. By fixing a fly in place, and opening the head capsule, it is possible to visualize calcium dynamics in these cells whilst delivering olfactory stimuli. Additionally, we demonstrate a set-up in which the fly can be subjected, simultaneously, to electric shocks to the body. This provides a system in which flies can undergo classical olfactory conditioning - whereby a previously na&#239;ve odor is learned to be associated with electric shock punishment - at the same time as the representation of this odor (and other untrained odors) is observed in the brain via two-photon microscopy. Our lab has previously reported the generation of synaptically localized calcium sensors, which enables one to confine the fluorescent calcium signals to pre- or postsynaptic compartments. Two-photon microscopy provides a way to spatially resolve fine structures. We exemplify this by focusing on neurons integrating information from the mushroom body, a higher-order center of the insect brain. Overall, this protocol provides a method to examine the synaptic connections between neurons whose activity is modulated as a result of olfactory learning.

In Vivo Optical Calcium Imaging of Learning-Induced Synaptic Plasticity in Drosophila melanogaster

Here we present a protocol with which pre- and/or postsynaptic calcium can be visualized in the context of Drosophila learning and memory. In vivo calcium imaging using synaptically localized calcium sensors is combined with a classical olfactory conditioning paradigm such that the synaptic plasticity underlying this type of associative learning may be determined.

Decades of research in many model organisms have led to the current concept of synaptic plasticity underlying learning and memory formation. ...

Measuring and Manipulating Functionally Specific Neural Pathways in the Human Motor System with Transcranial Magnetic Stimulation

Understanding interactions between brain areas is important for the study of goal-directed behavior. Functional neuroimaging of brain connectivity has provided important insights into fundamental processes of the brain like cognition, learning, and motor control. However, this approach cannot provide causal evidence for the involvement of brain areas of interest. Transcranial magnetic stimulation (TMS) is a powerful, noninvasive tool for studying the human brain that can overcome this limitation by transiently modifying brain activity. Here, we highlight recent advances using a paired-pulse, dual-site TMS method with two coils that causally probes cortico-cortical interactions in the human motor system during different task contexts. Additionally, we describe a dual-site TMS protocol based on cortical paired associative stimulation (cPAS) that transiently enhances synaptic efficiency in two interconnected brain areas by applying repeated pairs of cortical stimuli with two coils. These methods can provide a better understanding of the mechanisms underlying cognitive-motor function as well as a new perspective on manipulating specific neural pathways in a targeted fashion to modulate brain circuits and improve behavior. This approach may prove to be an effective tool to develop more sophisticated models of brain-behavior relations and improve diagnosis and treatment of many neurological and psychiatric disorders.

This article describes new approaches to measure and strengthen functionally specific neural pathways with transcranial magnetic stimulation. These advanced noninvasive brain stimulation methodologies can provide new opportunities for the understanding of brain-behavior relations and development of new therapies to treat brain disorders.

Understanding interactions between brain areas is important for the study of goal-directed behavior. Functional neuroimaging of brain connectivity has ...