This work was supported by the life insurance company "Novalis-Taitbout", Ecole des Neurosciences de Paris (ENP), the Agence Nationale de la Recherche "ANR-09-NEUR-004" in the frame of "ERA-NET NEURON" of FP7 program by the European Commission, "Foundation pour la Recherche Medicale", the Letten Foundation and the Pasteur Foundation.

1. Stereotaxic injections
<ol>
	<li>The virus used in these experiments is a replication-deficient lentiviral vector based on the HIV virus to express Channelrhodopsine2-YFP (yellow fluorescent protein) fusion construct driven by a synapsin promoter6. The plasmid was generously provided by K. Deisseroth's research group (<a href="http://www.stanford.edu/group/dlab/optogenetics/index.html">http://www.stanford.edu/group/dlab/optogenetics/index.html</a>). Lentiviruses can be obtained from commercial sources or produced in the lab according to established protocols7. Average vector titers were in the order of 1010 transduction units/ml. Viral aliquots are stored at -80&deg;C and should be thawed just before pipette loading. Multiple freeze/thaw cycles should be avoided. Never exposure the virus solution directly to dry ice.</li>
	<li>Prepare glass pipettes using borosilicate glass capillaries on standard pipette puller (Sutter P-97 pipette puller). The pulling setting should be properly calibrated to yield a long and rigid pipette. Next, cut the tip of the pipette using a pair of scissors to obtain a tip of 30-40 &mu;m in diameter. Store pulled pipettes in a dust free box.</li>
	<li>Setup the Nanoject II microinjector with the manufacturer's instructions for delivery of 50 nL. Prepare the injection pipette by back-filling with mineral oil and insert it in to the microinjector plunger. Attach the microinjector onto a stereotaxic frame. Partially empty oil from the glass pipette before filling with the virus solution</li>
	<li>With a micropipette, transfer 2 &mu;l of virus solution onto a sterile piece of Parafilm. Gently lower the glass pipette into the drop of virus solution and fill the pipette with 1-2 &mu;l. Make sure that there are no air bubbles in the pipette.</li>
	<li>Anesthetize a mouse with 100 mg/kg Ketamine and 10 mg/kg Xylazine, diluted in sterile saline. Remove the hair from the scalp of the animal using rodent clippers, razor, scissors, or chemical depilitory agent. Disinfect the scalp with three applications of alternating 70% ethanol and betadine. All surgical instruments should also be autoclaved and then disinfected with 70% ethanol. Place the animal in the stereotaxic frame with ear bars and nose bar, assuring that the head is secure. Apply eye ointment to prevent drying of the corneas.</li>
	<li>With a scalpel, cut the scalp from between the eyes to between the ears. Pull the skin aside to expose the skull and anchor the skin with a pair of clamps. Clean the surface of the skull. The stereotaxic apparatus is zeroed with the tip of the glass pipette at bregma. Then the tip is positioned at the injection site (Antero-Posterior, 3.30; Medio-Lateral, &plusmn;0.82, for both right and left hemispheres, respectively). The position of the nose bar has to be adjusted according to the calculation of the stereotaxic coordinates. In our experiment, the nose bar is adjusted until bregma and the injection site are aligned to the same height.</li>
	<li>Identify the injection site. Using a high-speed surgical drill, carefully drill holes into the skull. With a bent syringe needle hooked at the tip, remove the remnants of thin bone to expose the dura mater. Check that the hole is centered at the correct position. Take care not to rupture blood vessels on the surface of the brain while drilling. If blood vessels are ruptured, gently press onto the surface of the brain with a small cotton bud or tissue for several seconds until blood flow stops.</li>
	<li>Lower the pipette and zero the Dorso-Ventral height when the tip of the pipette touches the surface of the brain. Then, lower to the correct Dorso-Ventral depth (Dorso-Ventral, 2.9 from brain surface) and wait 30 seconds for pressure equilibrium. Inject 4 times 50 nl of virus for a total of 200 nl. Wait 30 seconds between each injection.</li>
	<li>One minute after the last injection, slowly withdraw the pipette. Repeat points 1.6-1.8 for the other hemisphere. Remove the animal from stereotaxic apparatus, clean the incision and stitch the cut skin using surgical threads. Allow the animal to recover on warming pad before returning to cage. Additionally, animals can be given a non-steroidal anti-inflammatory analgesic (e.g., Carprofen 4mg/kg) immediately after and 3 days following surgery.</li>
</ol>
2. Chronic LED implantation for in vivo optogenetic stimulation in the olfactory bulb
<ol>
	<li>Animals injected with lentiviral particles bilaterally are next chronically implanted with a miniature blue LED (Osram, LED CMS 4.6W blue). Solder the LED pins to a female miniature electrical connector so that the connector is positioned on the opposite side of the LED. Check that there is no electrical short cut and identify the positive and negative pin on the connector. Test the LED by connecting it to a current controller for LED though a proper cable.</li>
	<li>To measure and/or set the absolute light power of the LED, mount the LED on a micromanipulator and position the LED in direct contact with the photodetector of a power meter of a peak light intensity at 470 nm. Drill a pinhole of 3 mm diameter in an opaque PVC board. Affix this pinhole to the power meter and build a standard curve of optical power versus input current to calculate the power per mm2.</li>
	<li>To measure the propagation of light through the brain, cut blocks of fresh brain tissue of 300, 500, 1000, 1500 and 2500 &mu;m thickness in 0-4&deg;C ACSF using a vibratome14. Place a piece of skull with a craniotomy identical to the one performed in live mice under the LED. First, measure the light power without tissue centered on LED. Then, place each block of tissue between the LED and the photodetector of the power meter, and measure the power. Build a curve of tissue thickness versus relative transmission fraction, calculated as the ratio between the power measured through tissue and without tissue.</li>
	<li>Anesthetize a mouse with 100 mg/kg Ketamine and 10 mg/kg Xylazine, diluted in sterile saline. Clean and disinfect the LED with 70% ethanol. Remove the hair from the scalp of the animal and disinfect the scalp with three applications of alternating 70% ethanol and betadine. Then place the animal in the stereotaxic frame. Apply eye ointment to prevent drying of the corneas. Attach the connector to a stereotaxic holder. The position of the nose bar should be adjusted until the surface of the skull above the olfactory bulb is perfectly horizontal and parallel to the surface of the miniature LED.</li>
	<li>With a scalpel, cut the scalp from 3 mm anterior to the eyes to 3 mm posterior to the ears. Pull the skin aside and anchor it with a pair of clamps. Scrape away the membranes at the surface of the skull using a razor blade. Clean and dry the skull. Apply a thin cover of cyanoacrylate adhesive to allow subsequent adhesion of dental acrylic.</li>
	<li>Using a high-speed surgical drill, perform a rectangular craniotomy (1 mm caudal and 3 mm lateral; centered on 5.1 mm caudal, 0 mm lateral from bregma) over the olfactory bulbs. To do this, carefully thin the skull until only a thin layer of bone remains. Carefully remove the remaining bone with the help of a bent syringe needle. Special care should be taken to avoid puncturing the dura matter and rupturing the major blood vessels that lie between the two hemispheres and in the caudal border of the olfactory bulbs. Blood accumulation and coagulation at the brain surface precludes optimal light penetration into the tissue. Rinse the craniotomy and the surface of the brain with sterile saline and then dry the skull.</li>
	<li>Attach the miniature LED to the connector. Lower the miniature LED on the top of the olfactory bulbs. The main axis of the miniature LED should be parallel to the rostral-caudal axis of the mouse, Clean with sterile saline and dry the skull. Then secure the LED to the skull with a first layer of acrylic (i.e., polycarboxylate cement), followed by a second layer of dental acrylic. The addition of two small screws may be required to fix the LED. The screws should be inserted on the skull (posterior to the LED position) and covered with both acrylics. While the acrylic is still liquid, pull back and glue the free edges of the scalp on the surface of the dental acrylic. Take care not to apply any acrylic on the connector. After the acrylic has hardened, remove the animal from stereotaxic apparatus and let it recover on a warming pad before returning to cage. Animals are left isolated in their cage to recover from the surgery for at least seven days. Additionally, animals can be given a non-steroidal anti-inflammatory analgesic (e.g., Carprofen 4mg/kg) immediately after and 3 days following surgery. LEDs should be tested before implantation and after termination the experiments.</li>
	<li>A light electrical cable is used to tether the animal to a current controller for LED coupled to a pulse generator (Master-8) to control the timing and intensity of the LED. The thin cable allows for complete freedom of movement. Take care to respect the polarity of the connector. Inverting the polarity may break the LED.</li>
</ol>
3. Representative Results:
ChR2-YFP is expressed in transduced neurons approximately 48 hours after injection, and remains stable for several months7. Figure 1 is a representative picture of a sagittal section of a paraformaldehyde-fixed animal 15 days post-injection. Note the dense labeling and the size of the injection site in the RMS. A large number of newborn cells have migrated along the RMS to colonize the different layers of the olfactory bulb (Fig. 1). The fusion protein is expressed in all neuron compartments, including the soma, dendrites, and spines.
Miniature LED implantation allows large scale and inexpensive optical stimulation of a broad region of interest. The animal is tethered to the LED controller with a light electrical cable that allows a complete freedom of movement. By characterizing the light diffusion in the brain tissue and the threshold of ChR2 activation, we estimate that optical activation could be triggered at a depth of ~2mm below the surface of the LED (Fig. 2A). To monitor the functional activation of ChR2-expressing neurons in vivo, the expression of c-fos, a marker of neuronal activity, can be analyzed after repetitive light stimulation (Fig. 2B). As an alternative, in vitro electrophysiological recordings of ChR2-expressing neurons coupled to focal light stimulation can be performed14. Finally, two-photon targeted loose-patch recordings have been used to study the functional activation of ChR2-expressing cortical neurons by similar miniature LED in vivo9.
<img alt="Figure 1" src="/files/ftp_upload/3380/3380fig1.jpg" /> 
Figure 1. ChR2-YFP expression from lentiviral injection into the rostral migratory stream. Representative confocal images of a saggital sections of the olfactory bulb of a mouse 15 days after injections of a lentivirus in the rostral RMS. Note the presence of scattered YFP newborn neurons migrating from the end of the RMS to the different layers of the OB. Most of newborn cells (95%) are granule cells, which soma lie in the Granule Cell Layer (GCL) and dendrites arborize in the External Plexiform Layer (EPL). A small population of newborn neurons migrate up to the most superficial layer of the OB, in the Glomerular Layer (GL). Scale bar, 100&mu;m. Inset, high magnification of a newborn granule cell, showing the membrane expression of ChR2-YFP in the dendrites as well as in spines. Scale bar, 20&mu;m.
<img alt="Figure 2" src="/files/ftp_upload/3380/3380fig2.jpg" /> 
Figure 2. Diagram of implanted LED on the top of the olfactory bulb and light-evoked c-fos expression in ChR2-YFP expressing new granule cells. 
A, Schematic drawing of the implanted LED above the olfactory bulbs. 
B, Representative confocal images showing double-labelled ChR2-YFP+ c-fos+ granule cells one hour after light stimulation of the olfactory bulb. C-fos expression is revealed using a rabbit anti-c-fos primary antibody (Calbiochem, 1:5000) and A568-conjugated anti-rabbit secondary antibody (Invitrogen, 1:1000). Scale bar, 10&mu;m.

<ol>
	<li>Lledo, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adult+neurogenesis+and+functional+plasticity+in+neuronal+circuits.">Adult neurogenesis and functional plasticity in neuronal circuits.</a> Nat. Rev. Neurosci. 7, 179-193 (2006).</li><li>Alonso, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Turning+astrocytes+from+the+rostral+migratory+stream+into+neurons:+a+role+for+the+olfactory+sensory+organ.">Turning astrocytes from the rostral migratory stream into neurons: a role for the olfactory sensory organ.</a> J. Neurosci. 28, 11089-11102 (2008).</li><li>Mouret, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Learning+and+survival+of+newly+generated+neurons:+when+time+matters.">Learning and survival of newly generated neurons: when time matters.</a> J. Neurosci. 28, 11511-11516 (2008).</li><li>Bardy, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How,+when,+and+where+new+inhibitory+neurons+release+neurotransmitters+in+the+adult+olfactory+bulb.">How, when, and where new inhibitory neurons release neurotransmitters in the adult olfactory bulb.</a> J. 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No conflicts of interest declared.

Optogenetic tools have recently increased our control over selective neuronal populations, including adult born neurons in the olfactory system. Here we describe how to perform a stereotaxic injection of a lentiviral vector expressing ChR2-YFP in a specific population of adult-born neurons. By carefully monitoring the period post-injection, we are able to analyze and manipulate the activity of a cohort of adult-born neurons with a relative homogeneous age.
Because of their low diffusion spread...

<table border="1">
	<tbody>
		<tr>
			<td>Material</td>
			<td>Company</td>
			<td>Catalogue No.</td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>Merial</td>
			<td>Imalg&egrave;ne 1000</td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Bayer Health Care</td>
			<td>Rompum 0.2%</td>
		</tr>
		<tr>
			<td>Pipette Puller</td>
			<td>Sutter</td>
			<td>P-97</td>
		</tr>
		<tr>
			<td>Glass Capillaries (3.5")</td>
			<td>Drummond Scientific</td>
			<td>3-000-203-G/X</td>
		</tr>
		<tr>
			<td>Nanoject II Microinjector</td>
			<td>Drummond Scientific</td>
			<td>3-000-204</td>
		</tr>
		<tr>
			<td>High speed surgical drill</td>
			<td>Fine Science Tools</td>
			<td>18000-17</td>
		</tr>
		<tr>
			<td>Micro Drill Steel Burrs 0.7mm tip diameter</td>
			<td>Fine Science Tools</td>
			<td>19008-07</td>
		</tr>
		<tr>
			<td>LED CMS blue 4.6 W</td>
			<td>OSRAM</td>
			<td>LBW5SN</td>
		</tr>
		<tr>
			<td>Female electrical connector (2.54 mm)</td>
			<td>Fischer Elektronik</td>
			<td>BL5.36Z</td>
		</tr>
		<tr>
			<td>Current controller for LED (max: 1A)</td>
			<td>A1W Electronik</td>
			<td>HKO-KL50-1000</td>
		</tr>
		<tr>
			<td>Pulse generator</td>
			<td>AMPI</td>
			<td>Master-8</td>
		</tr>
		<tr>
			<td>Optical power meter</td>
			<td>Thor Labs</td>
			<td>PM100</td>
		</tr>
		<tr>
			<td>Cyanoacrylate adhesive</td>
			<td>Loctite</td>
			<td>454</td>
		</tr>
		<tr>
			<td>Polycarboxylate cement (first layer cement)</td>
			<td>Septodont</td>
			<td>Selfast (#0068Q)</td>
		</tr>
		<tr>
			<td>Methacrylate cement (second layer cement)</td>
			<td>GC InternationalCorp.</td>
			<td>Unifast Trad &#x2013; Liquid (#339292) 
			Unifast Trad &#x2013; Powder (#339114)</td>
		</tr>
		<tr>
			<td>Carprofen</td>
			<td>Pfizer</td>
			<td>Rimadyl</td>
		</tr>
	</tbody>
</table>

selective viral transduction of adult-born olfactory neurons for chronic in vivo optogenetic stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo stimulation of ChR2-expressing neurons.

Watch this Scientific Journal Video about Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation at JoVE.com

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3. In this process, called adult neurogenesis, neural stem cells ...

selective-viral-transduction-adult-born-olfactory-neurons-for-chronic

Research

JoVE Journal

Neuroscience

15.4K Views.  Institut Pasteur and Centre National de la Recherche Scientifique (CNRS). Adult-born neurons of the olfactory bulb can be optogenetically controlled using Channelrhodopsin2-expressing lentiviral injection in the rostral migratory stream and chronic photostimulation with an implanted miniature LED.

Video: Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation

Studying the Integration of Adult-born Neurons

Neurogenesis occurs in adult mammalian brains in the sub-ventricular zone (SVZ) of the lateral ventricle and in the sub-granular zone (SGZ) of the hippocampal dentate gyrus throughout life. Previous reports have shown that adult hippocampal neurogenesis is associated with diverse brain disorders, including epilepsy, schizophrenia, depression and anxiety (1). Deciphering the process of normal and aberrant adult-born neuron integration may shed light on the etiology of these diseases and inform the development of new therapies.
SGZ adult neurogenesis mirrors embryonic and post-natal neuronal development, including stages of fate specification, migration, synaptic integration, and maturation. However, full integration occurs over a prolonged, 6-week period. Initial synaptic input to adult-born SGZ dentate granule cells (DGCs) is GABAergic, followed by glutamatergic input at 14 days (2). The specific factors which regulate circuit formation of adult-born neurons in the dentate gyrus are currently unknown.
Our laboratory uses a replication-deficient retroviral vector based on the Moloney murine leukemia virus to deliver fluorescent proteins and hypothesized regulatory genes to these proliferating cells. This viral technique provides high specificity and resolution for analysis of cell birth date, lineage, morphology, and synaptogenesis.
A typical experiment often employs two or three viruses containing unique label, transgene, and promoter elements for single-cell analysis of a desired developmental process in vivo. The following protocol describes a method for analyzing functional newborn neuron integration using a single green (GFP) or red (dTomato) fluorescent protein retrovirus and patch-clamp electrophysiology.

A way to study the integration of newborn dentate granule cells in adult animals is described. This technique uses an engineered retrovirus to label newborn neurons, followed by electrophysiological recordings to determine in vivo functional integration.

Neurogenesis occurs in adult mammalian brains in the sub-ventricular zone (SVZ) of the lateral ventricle and in the sub-granular zone (SGZ) of the ...

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the olfactory bulb (OB). The molecular mechanisms of OSN axon growth and targeting are not well understood. Manipulating gene expression and subsequent 
visualizing of single OSN axons and their terminal arbor morphology have thus far been challenging. To study gene function at the single cell level within a specified 
time frame, we developed a lentiviral based technique to manipulate gene expression in OSNs in vivo. Lentiviral particles are delivered to OSNs by microinjection 
into the olfactory epithelium (OE). Expression cassettes are then permanently integrated into the genome of transduced OSNs. Green fluorescent protein expression 
identifies infected OSNs and outlines their entire morphology, including the axon terminal arbor. Due to the short turnaround time between microinjection and 
reporter detection, gene function studies can be focused within a very narrow period of development. With this method, we have detected GFP expression within as 
few as three days and as long as three months following injection. We have achieved both over-expression and shRNA mediated knock-down by lentiviral microinjection. 
This method provides detailed morphologies of OSN cell bodies and axons at the single cell level in vivo, and thus allows characterization of candidate gene function 
during olfactory development.

Lentivirus-mediated Genetic Manipulation and Visualization of Olfactory Sensory Neurons in vivo

We present a lentiviral technique for genetic manipulation and visualization of single olfactory sensory neuron axon and its terminal arborization in vivo.

Development of a precise olfactory circuit relies on accurate projection of olfactory sensory neuron (OSN) axons to their synaptic 
targets in the ...

Flash Photolysis of Caged Compounds in the Cilia of Olfactory Sensory  Neurons

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. Caged compounds are molecules made physiologically inactive by a chemical cage that can be broken by a flash of ultraviolet light. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged compounds for the study of olfactory transduction in dissociated mouse olfactory sensory neurons. The process of olfactory transduction (Figure 1) takes place in the cilia of olfactory sensory neurons, where odorant binding to receptors leads to the increase of cAMP that opens cyclic nucleotide-gated (CNG) channels2. Ca entry through CNG channels activates Ca-activated Cl channels. We show how to dissociate neurons from the mouse olfactory epithelium3 and how to activate CNG channels or Ca-activated Cl channels by photolysis of caged cAMP4 or caged Ca5. We use a flash lamp6,7 to apply ultraviolet flashes to the ciliary region to uncage cAMP or Ca while patch-clamp recordings are taken to measure the current in the whole-cell voltage-clamp configuration8-11.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds. Here, we show how to obtain patch-clamp recordings combined with photolysis of caged cAMP or caged Ca for the study of olfactory transduction in dissociated mouse olfactory sensory neurons.

Photolysis of caged compounds allows the production of rapid and localized increases in the concentration of various physiologically active compounds1. ...

Fiber-optic Implantation for Chronic Optogenetic Stimulation of Brain Tissue

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard for analyzing patterns of synaptic connectivity, but paired electrophysiological recordings can be both cumbersome and experimentally limiting. The development of optogenetics has introduced an elegant method to stimulate neurons and circuits, both in vitro1 and in vivo2,3. By exploiting cell-type specific promoter activity to drive opsin expression in discrete neuronal populations, one can precisely stimulate genetically defined neuronal subtypes in distinct circuits4-6. Well described methods to stimulate neurons, including electrical stimulation and/or pharmacological manipulations, are often cell-type indiscriminate, invasive, and can damage surrounding tissues. These limitations could alter normal synaptic function and/or circuit behavior. In addition, due to the nature of the manipulation, the current methods are often acute and terminal. Optogenetics affords the ability to stimulate neurons in a relatively innocuous manner, and in genetically targeted neurons. The majority of studies involving in vivo optogenetics currently use a optical fiber guided through an implanted cannula6,7; however, limitations of this method include damaged brain tissue with repeated insertion of an optical fiber, and potential breakage of the fiber inside the cannula. Given the burgeoning field of optogenetics, a more reliable method of chronic stimulation is necessary to facilitate long-term studies with minimal collateral tissue damage. Here we provide our modified protocol as a video article to complement the method effectively and elegantly described in Sparta et al.8 for the fabrication of a fiber optic implant and its permanent fixation onto the cranium of anesthetized mice, as well as the assembly of the fiber optic coupler connecting the implant to a light source. The implant, connected with optical fibers to a solid-state laser, allows for an efficient method to chronically photostimulate functional neuronal circuitry with less tissue damage9 using small, detachable, tethers. Permanent fixation of the fiber optic implants provides consistent, long-term in vivo optogenetic studies of neuronal circuits in awake, behaving mice10 with minimal tissue damage.

The development of optogenetics now provides the means to precisely stimulate genetically defined neurons and circuits, both in vitro and in vivo. Here we describe the assembly and implantation of a fiber optic for chronic photostimulation of brain tissue.

Elucidating patterns of neuronal connectivity has been a challenge for both clinical and basic neuroscience. Electrophysiology has been the gold standard ...

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 ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

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 ...

Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene expression in the mouse brain. Here we describe a technique first introduced by Passini and Wolfe for direct intracranial delivery of virally-encoded transgenes into the neonatal mouse brain. In its most basic form, the procedure requires only an ice bucket and a microliter syringe. However, the protocol can also be adapted for use with stereotaxic frames to improve consistency for researchers new to the technique. The method relies on the ability of adeno-associated virus (AAV) to move freely from the cerebral ventricles into the brain parenchyma while the ependymal lining is still immature during the first 12-24 hr after birth. Intraventricular injection of AAV at this age results in widespread transduction of neurons throughout the brain. Expression begins within days of injection and persists for the lifetime of the animal. Viral titer can be adjusted to control the density of transduced neurons, while co-expression of a fluorescent protein provides a vital label of transduced cells. With the rising availability of viral core facilities to provide both off-the-shelf, pre-packaged reagents and custom viral preparation, this approach offers a timely method for manipulating gene expression in the mouse brain that is fast, easy, and far less expensive than traditional germline engineering.

Here we demonstrate a technique for widespread neuronal transduction by intraventricular injection of adeno-associated virus into the neonatal mouse brain. This method provides a rapid and easy way to attain lifelong expression of virally-delivered transgenes.

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene ...

Optogenetic Stimulation of the Auditory Nerve

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of&#160;the cochlea has been used to evoke responses in the auditory nerve10. Nevertheless it requires higher energies than electrical stimulation10,11 and uncertainty remains as to the underlying mechanism12. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13 or virus-mediated expression of the ChR2-variant CatCh14. We used micro-light emitting diodes (&#181;LEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.

Cochlear implants (CIs) enable hearing by direct electrical stimulation of the auditory nerve. However, poor frequency and intensity resolution limits the quality of hearing with CIs. Here we describe optogenetic stimulation of the auditory nerve in mice as an alternative strategy for auditory research and developing future CIs.

Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted ...