This work was supported by the life insurance company "AG2R-La-Mondiale", 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, and the Pasteur Foundation. Sebastien Wagner was supported by the Letten Foundation.

1. Optical Calibration: Measuring LED Power
<ol>
	<li>Attach a LED array to a heatsink actively cooled by a fan and affix this LED/heatsink aparatus to a collimating lens.</li>
	<li>Replace lamp used in brightfield illumination with LED/heatsink/fan/lens apparatus. This apparatus must be carefully positioned so that the collimated LED beam travels along a straight optical path towards the condenser lens. Make sure the heatsink/fan is properly grounded to the system's common ground.</li>
	<li>Drive the LED array with a power supply that can give fast and square pulses of current. This power supply may be controlled by a 5V TTL pulse originating from a pulse generator.</li>
	<li>Center the collimated beam along the light path defined between the field diaphragm and the condensor lens. Ideally, the LED beam will slightly overfill the fully opened field diaphragm. Typically, a more tightly collimated LED beam will fill this aperture less, and will produce more power at the expense of uniformity. In our setup we increased light uniformity by choosing a collimating lens that projected a slightly expanded image of the LED array at its conjugate plane at the consenser diaphragm.</li>
	<li>Achieve Kohler illumination by focusing the condenser so that the image of the field diaphragm (the diaphragm closest to the light source) is focused on the slice chamber (fig. 1). A thin tissue of lens paper can act as a projection screen to visualize the focused image of the field diaphragm at other depths.</li>
	<li>Drill a series of pinholes of known diameter in an opaque material. Place one of these small pinholes over the sensor of an optical power-meter. Place the power meter on the specimen stage and center the power meter over the focused image of the field diaphragm by simply moving the power meter until it gives a maximum reading. Affix the power meter in this position.</li>
	<li>Fully open all apertures (aperature diaphragm and field diaphragm). Systematically move the attached slice chamber/power meter relative to the optical lightpath and calculate the uniformity of optical power within the illuminated area. Build the uniformity plot for your system. If the microscope is properly setup with Kohler illumination centered at the objective's focus, maximum power should be directly underneath the objective, and regions outside of this focus should now receive a known amount of power according to the uniformity plot.</li>
	<li>For each size of pinhole, build a standard curve of optical power vs. pinhole surface area. By adjusting the input current to the LED array, produce this curve at multiple power levels and from each curve calculate the power per mm2. If the LED array is to be used for patching optics, make sure to introduce optical elements necessary for patching (condensers, pinholes and filters) to know how much light is transmitted under patching illumination. </li>
	<li>Flipping the power meter to face the objective, calculate the intensity of illumination at 470nm when the mercury lamp is turned on.</li>
	<li>Add a live slice to the chamber, and rebuild standard curves to determine the light power transmitted through the scattering brain tissue.</li>
</ol>
2.	Slicing Procedure and Electrophysiology
Part A: Slice Preparation
<ol>
	<li>Anesthetize (60 mg/kg Ketamine and 2mg/kg Xylazine) and decapitate the mouse. Dissect the brain in artifical cerebral spinal fluid (ACSF, in mM: 124 NaCl, 3 KCl, 1.3 MgSO4, 26 NaHCO3, 1.25 NaHPO4, 20 glucose, 2 CaCl2; ~310mOsm, pH 7.4 when bubbled with a mixture of 95% O2 and 5% CO25,1), taking care not to damage the olfactory bulbs. Separate the two hemispheres and place on agar with the ventral surface even with one edge (for horizontal sections).</li>
	<li>Glue the agar and dorsal surface of each cortical hemisphere to the vibratome chuck, and slowly fill the bath with ice-cold ACSF. Slice from ventral surface in 300 &mu;m sections, transferring each section to warmed (34-36&deg;C) and oxygenated ACSF, allowing them to recover for 30-45 minutes.</li>
</ol>
Part B: Loose Patch Measurement of the Threshold to Spike
<ol start="3">
	<li>After bringing the slices to room temperature for 30 minutes, gently place a slice in the recording chamber of the microscope chamber under constant perfusion of oxygenated ACSF. 
	At risk of excessively stimulating ChR2 infected neurons, the presence of EYFP-ChR2 can be confirmed under epifluorescence (fig. 2a)</li>
	<li>Pull glass electrodes on a pipette puller (Sutter P-97). Fill this electrode with ACSF. When placed in the ACSF bath the tip resistance should be between 7-10 MOhms.</li>
	<li>Under fluorescent illumination, locate in the slice a healthy ChR2-EYFP neuron with mature morphology. Also locate this neuron's soma under patching optics.</li>
	<li>With light positive pressure passed through the electrode tip, lower the patch electrode towards the identified fluorescent neuron. When membrane contact is made, quickly release positive pressure and apply a small and brief amount of suction through the tip. A giga-ohm seal should be made between the plasma membrane and the walls of the patch electrode.</li>
	<li>Even if a giga-seal is not formed, if the electrode is sufficiently close to a fluorescent neuron spiking activity should produce a measurable local field potential. Activate ChR2 in this neuron by flashing different doses of light. Because light-dose is a function of both LED power and duration, calculate how much light is necessary to evoke an action potential at multiple powers and durations (fig. 2b). Also observe how much spiking occurs under mercury lamp illumination.</li>
</ol>
3. Representative Results: 
On our microscope (Olympus BX51WI), our LED is in line with 2 apertures and a condenser lens, thus maintaining the original lightpath of the factory-installed arc lamp. By closing both the field diaphragm and the aperature diaphragm, we can achieve brightfield contrast sufficient for patch clamp recordings (fig. 1b). With all diaphragms fully open we expose the slice to maximum light power for channelrhodopsin activation (fig. 1a). On our microscope, this patching configuration produces light-density that is approximately three orders of magnitude lower than the maximum full field density (4.1 &mu;W/mm2 versus 6.88 mW/mm2).
We see robust labeling of adult-born olfactory bulb granule and periglomerular neurons weeks after lentiviral infection of migrating neuroblasts in the rostral migratory stream (fig. 2a) A loose-patch recording from a single adult-born ChR2-EYFP expressing granule cell indicates that a 5 ms stimulation at maximum power (6.88 mW/mm2) is sufficient to evoke spiking (fig. 2b). Since expression level varies between cells, the amount of light that passes the threshold to spike will vary and should be described statistically for each cell type of interest. 
<img src="/files/ftp_upload/3379/3379fig1.jpg" alt="Figure 1" /> 
Figure 1. LED array setup for full-field photostimulation and patch-clamp slice electrophysiology. To activate channelrhodopsin (ChR2) we project a collimated beam through open back apertures and condenser optics (a). This configuration can be changed into high-contrast patching optics by fully closing the field diaphragm and modulating the width of the field diaphragm (b). Abbreviations: a. fan, b. heatsink, c. LED array, d. collimating lens, e. mirror, f. field diaphragm, g. aperture diaphragm, h. condenser lens, i. sample stage, j. objective.
<img src="/files/ftp_upload/3379/3379fig2.jpg" alt="Figure 2" /> 
Figure 2. Image of a 300&mu;m horizontal slice of olfactory bulb for patch clamp and whole-field photostimulation (a). Lentivirally infected adult-born granule cells expressing ChR2-EYFP can be seen radiating from the core of the olfactory bulb. The light-dose required to evoke spiking can be found by increasing the duration of the LED flash (b). The threshold for this granule cell was 5ms at full LED intensity (2.43mW/mm2). Scale in (a) = 500&mu;m, scale in (b) = 50ms.

<ol>
	<li>Nissant, A. <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+promotes+synaptic+plasticity+in+the+olfactory+bulb.">Adult neurogenesis promotes synaptic plasticity in the olfactory bulb.</a> Nature Neuroscience. 12, 728-730 (2009).</li><li>Apicella, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pyramidal+Cells+in+Piriform+Cortex+Receive+Convergent+Input+from+Distinct+Olfactory+Bulb+Glomeruli.">Pyramidal Cells in Piriform Cortex Receive Convergent Input from Distinct Olfactory Bulb Glomeruli.</a> Journal of Neuroscience. 30, 14255-14260 (2010).</li><li>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=genetically+targeted+optical+control+of+neural+activity.">genetically targeted optical control of neural activity.</a> Nature. 8, 1263-1263 (2005).</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=where+new+inhibitory+neurons+release+neurotransmitters+in+the+adult+olfactory+bulb.">where new inhibitory neurons release neurotransmitters in the adult olfactory bulb.</a> The Journal of Neuroscience. 30, 17023-17034 (2010).</li><li>Grubb, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+maturation+of+the+first+synapse+in+olfaction:+development+and+adult+neurogenesis.">Functional maturation of the first synapse in olfaction: development and adult neurogenesis.</a> The Journal of neuroscience. 28, 2919-2932 (2008).</li><li>Zhang, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Circuit-breakers:+optical+technologies+for+probing+neural+signals+and+systems.">Circuit-breakers: optical technologies for probing neural signals and systems.</a> Nature reviews. Neuroscience. 8, 577-581 (2007).</li><li>Grossman, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multi-site+optical+excitation+using+ChR2+and+micro-LED+array.">Multi-site optical excitation using ChR2 and micro-LED array.</a> Journal of neural engineering. 7, 16004-16004 (2010).</li><li>Dhawale, A. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-redundant+odor+coding+by+sister+mitral+cells+revealed+by+light+addressable+glomeruli+in+the+mouse.">Non-redundant odor coding by sister mitral cells revealed by light addressable glomeruli in the mouse.</a> Nature neuroscience. 13, 1404-1412 (2010).</li><li>Weick, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+control+of+transplantable+human+ESC-derived+neurons+via+optogenetic+targeting.">Functional control of transplantable human ESC-derived neurons via optogenetic targeting.</a> Stem cells. 28, 2008-2016 (2010).</li><li>Toni, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurons+born+in+the+adult+dentate+gyrus+form+functional+synapses+with+target+cells.">Neurons born in the adult dentate gyrus form functional synapses with target cells.</a> Nature Neuroscience. 11, 901-907 (2008).</li><li>Grubb, M. S., Burrone, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Channelrhodopsin-2+Localised+to+the+Axon+Initial+Segment.">Channelrhodopsin-2 Localised to the Axon Initial Segment.</a> PLoS ONE. 5, e13761-e13761 .</li><li>Tye, K. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amygdala+circuitry+mediating+reversible+and+bidirectional+control+of+anxiety.">Amygdala circuitry mediating reversible and bidirectional control of anxiety.</a> Nature. 471, 358-362 (2011).</li></ol>

No conflicts of interest declared.

Recent years have seen an explosion in the popularity of optogenetic tools for neuroscience research6. As a result, it is increasingly important to lower the barrier of entry for labs wishing to begin using these new tools. Here we describe how to conduct a simple and low cost retrofitting and calibration of a conventional patch-clamp rig so that it can do full-field optical stimulation of channelrhodopsin-expressing neurons. In particular, we apply this technique to the study of olfactory bulb adult neuroge...

<table border="1">
	<tbody>
		<tr>
			<td>Material</td>
			<td>Type</td>
			<td>Company</td>
			<td>Catalogue No.</td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>100 mg/ml</td>
			<td>Imalg&egrave;ne 1000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>2%</td>
			<td>Rompun</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>&nbps;</td>
			<td>Sigma </td>
			<td>S5886</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>P5405</td>
		</tr>
		<tr>
			<td>MgSO4</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>M1880</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>S5761</td>
		</tr>
		<tr>
			<td>NaHPO4</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>S5011</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>G7021</td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>C7902</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>&nbsp;</td>
			<td>Sigma </td>
			<td>A9539</td>
		</tr>
		<tr>
			<td>Pipette Puller</td>
			<td>P-97</td>
			<td>Sutter</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Glass Capillaries</td>
			<td>1.5 mm O.D./1.17 mm I.D.</td>
			<td>Harvard Apparatus</td>
			<td>GC150T-10</td>
		</tr>
		<tr>
			<td>LED array</td>
			<td>&nbsp;</td>
			<td>Bridgelux</td>
			<td>BXRA-C2000</td>
		</tr>
		<tr>
			<td>Collimating lens</td>
			<td>40 mm beam diameter</td>
			<td>Thor Labs</td>
			<td>LEDC1</td>
		</tr>
		<tr>
			<td>Power supply</td>
			<td>2.8 amp</td>
			<td>A1W Electronik</td>
			<td>HKO2800</td>
		</tr>
		<tr>
			<td>Optical power meter</td>
			<td>&nbsp;</td>
			<td>Thor Labs</td>
			<td>PM 100</td>
		</tr>
		<tr>
			<td>Heatsink </td>
			<td>Silent Boost K8</td>
			<td>Thermaltake</td>
			<td>A1838</td>
		</tr>
		<tr>
			<td>Fan</td>
			<td>Silent Boost K8</td>
			<td>Thermaltake</td>
			<td>A1838</td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>&nbsp;</td>
			<td>Leica</td>
			<td>VT1200S</td>
		</tr>
	</tbody>
</table>

using affordable led arrays for photo-stimulation of neurons

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single cells in response to electrical or pharmacological manipulations1,2. With the invention of methods to optically control genetically targeted neurons (optogenetics), researchers now have an unprecedented level of control over specific groups of neurons in the standard slice preparation. In particular, photosensitive channelrhodopsin-2 (ChR2) allows researchers to activate neurons with light3,4. By combining careful calibration of LED-based photostimulation of ChR2 with standard slice electrophysiology, we are able to probe with greater detail the role of adult-born interneurons in the olfactory bulb, the first central relay of the olfactory system. Using viral expression of ChR2-YFP specifically in adult-born neurons, we can selectively control young adult-born neurons in a milieu of older and mature neurons. Our optical control uses a simple and inexpensive LED system, and we show how this system can be calibrated to understand how much light is needed to evoke spiking activity in single neurons. Hence, brief flashes of blue light can remotely control the firing pattern of ChR2-transduced newborn cells.

Watch this Scientific Journal Video about Using Affordable LED Arrays for Photo-Stimulation of Neurons at JoVE.com

Using Affordable LED Arrays for Photo-Stimulation of Neurons

Adult-born neurons expressing ChR2 can be manipulated in slice electrophysiological preparations in order to examine their contribution towards the function of olfactory neural circuits.

Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single ...

using-affordable-led-arrays-for-photo-stimulation-of-neurons

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18.5K Views.  Institut Pasteur and Centre National de la Recherche Scientifique (CNRS). Adult-born neurons expressing ChR2 can be manipulated in slice electrophysiological preparations in order to examine their contribution towards the function of olfactory neural circuits.

Video: Using Affordable LED Arrays for Photo-Stimulation of Neurons

Scalable Fluidic Injector Arrays for Viral Targeting of Intact 3-D Brain Circuits

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted in neurological and psychiatric disorders--would be greatly facilitated by a technology for rapidly targeting genes to complex 3-dimensional neural circuits, enabling fast creation of "circuit-level transgenics." We have recently developed methods in which viruses encoding for light-sensitive proteins can sensitize specific cell types to millisecond-timescale activation and silencing in the intact brain. We here present the design and implementation of an injector array capable of delivering viruses (or other fluids) to dozens of defined points within the 3-dimensional structure of the brain (Figure. 1A, 1B). The injector array comprises one or more displacement pumps that each drive a set of syringes, each of which feeds into a polyimide/fused-silica capillary via a high-pressure-tolerant connector. The capillaries are sized, and then inserted into, desired locations specified by custom-milling a stereotactic positioning board, thus allowing viruses or other reagents to be delivered to the desired set of brain regions. To use the device, the surgeon first fills the fluidic subsystem entirely with oil, backfills the capillaries with the virus, inserts the device into the brain, and infuses reagents slowly (&lt;0.1 microliters/min). The parallel nature of the injector array facilitates rapid, accurate, and robust labeling of entire neural circuits with viral payloads such as optical sensitizers to enable light-activation and silencing of defined brain circuits. Along with other technologies, such as optical fiber arrays for light delivery to desired sets of brain regions, we hope to create a toolbox that enables the systematic probing of causal neural functions in the intact brain. This technology may not only open up such systematic approaches to circuit-focused neuroscience in mammals, and facilitate labeling of brain regions in large animals such as non-human primates, but may also open up a clinical translational path for cell-specific optical control prosthetics, whose precision may enable improved treatment of intractable brain disorders. Finally, such devices as described here may facilitate precisely-timed fluidic delivery of other payloads, such as stem cells and pharmacological agents, to 3-dimensional structures, in an easily user-customizable fashion.

Controlling and analyzing neural circuits in vivo would be facilitated by a technology for delivery of viruses and other reagents to desired 3-dimensional sets of brain regions. We demonstrate customized fluidic injector array fabrication, and delivery of virally-encoded optical sensitizers, enabling optical manipulation of complex brain circuits.

Our understanding of neural circuits--how they mediate the computations that subserve sensation, thought, emotion, and action, and how they are corrupted ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato

Larval zebrafish are emerging as a model for describing the development and function of simple neural circuits. Due to their external fertilization, rapid development, and translucency, zebrafish are particularly well suited for optogenetic approaches to investigate neural circuit function. In this approach, light-sensitive ion channels are expressed in specific neurons, enabling the experimenter to activate or inhibit them at will and thus assess their contribution to specific behaviors. Applying these methods in larval zebrafish is conceptually simple but requires the optimization of technical details. Here we demonstrate a procedure for expressing a channelrhodopsin variant in larval zebrafish somatosensory neurons, photo-activating single cells, and recording the resulting behaviors. By introducing a few modifications to previously established methods, this approach could be used to elicit behavioral responses from single neurons activated up to at least 4 days post-fertilization (dpf). Specifically, we created a transgene using a somatosensory neuron enhancer, CREST3, to drive the expression of the tagged channelrhodopsin variant, ChEF-tdTomato. Injecting this transgene into 1-cell stage embryos results in mosaic expression in somatosensory neurons, which can be imaged with confocal microscopy. Illuminating identified cells in these animals with light from a 473 nm DPSS laser, guided through a fiber optic cable, elicits behaviors that can be recorded with a high-speed video camera and analyzed quantitatively. This technique could be adapted to study behaviors elicited by activating any zebrafish neuron. Combining this approach with genetic or pharmacological perturbations will be a powerful way to investigate circuit formation and function.

Optogenetic techniques have made it possible to study the contribution of specific neurons to behavior. We describe a method in larval zebrafish for activating single somatosensory neurons expressing a channelrhodopsin variant (ChEF) with a diode-pumped solid state (DPSS) laser and recording the elicited behaviors with a high-speed video camera.

Larval  zebrafish are emerging as a model for describing the development and function  of simple neural circuits. Due to their  external fertilization, ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Production and Isolation of Axons from Sensory Neurons for Biochemical Analysis Using Porous Filters

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is required to shape functional neuronal circuits. The protocol described here explains how to use cell culture inserts bearing a porous membrane (filter) to obtain large amounts of pure axonal preparations suitable for examination by conventional biochemical or immunocytochemical techniques. The functionality of these filter inserts will be demonstrated with models of developmental pruning and Wallerian degeneration, using explants of embryonic dorsal root ganglion. Axonal integrity and function is compromised in a wide variety of neurodegenerative pathologies. Indeed, it is now clear that axonal dysfunction appears much earlier in the course of the disease than neuronal soma loss in several neurodegenerative diseases, indicating that axonal-specific processes are primarily targeted in these disorders. By obtaining pure axonal samples for analysis by molecular and biochemical techniques, this technique has the potential to shed new light into mechanisms regulating the physiology and pathophysiology of axons. This in turn will have an impact in our understanding of the processes that drive degenerative diseases of the nervous system.

This article describes a neuronal culture system that can be used to obtain large quantities of pure axonal samples for biochemical and immunocytochemical analyses. This technique will allow an improved understanding of normal axonal physiology and signaling pathways leading to neurodegeneration.

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Live Images of GLUT4 Protein Trafficking in Mouse Primary Hypothalamic Neurons Using Deconvolution Microscopy

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral tissues. The hypothalamus in the central nervous system (CNS) is the control center for energy and insulin signal response regulation. Chronic inflammation in peripheral tissues and imbalances of certain chemokines (such as CCL5, TNF&#945;, and IL-6) contribute to diabetes and obesity. However, the functional mechanism(s) connecting chemokines and hypothalamic insulin signal regulation still remain unclear.
In vitro primary neuron culture models are convenient and simple models which can be used to investigate insulin signal regulation in hypothalamic neurons. In this study, we introduced exogeneous GLUT4 protein conjugated with GFP (GFP-GLUT4) into primary hypothalamic neurons to track GLUT4 membrane translocation upon insulin stimulation. Time-lapse images of GFP-GLUT4 protein trafficking were recorded by deconvolution microscopy, which allowed users to generate high-speed, high-resolution images without damaging the neurons significantly while conducting the experiment. The contribution of CCR5 in insulin regulated GLUT4 translocation was observed in CCR5 deficient hypothalamic neurons, which were isolated and cultured from CCR5 knockout mice. Our results demonstrated that the GLUT4 membrane translocation efficiency was reduced in CCR5 deficient hypothalamic neurons after insulin stimulation.

This protocol describes a technique for observation of real-time Green Fluorescence Protein (GFP) tagged Glucose Transporter 4 (GLUT4) protein trafficking upon insulin stimulation and characterization of the biological role of CCR5 in the insulin&#8211;GLUT4 signaling pathway with Deconvolution Microscopy.

Type 2 diabetes mellitus (T2DM) is a global health crisis which is characterized by insulin signaling impairment and chronic inflammation in peripheral ...

An In Vitro Model for Studying Tau Aggregation Using Lentiviral-mediated Transduction of Human Neurons

Aberrant aggregation of the protein tau is pathogenically involved in a number of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD). Although mouse models of tauopathy have provided a valuable resource for investigating the neurotoxic mechanisms of aggregated tau, it is becoming increasingly apparent that, due to interspecies differences in neurophysiology, the mouse brain is unsuitable for modeling the human condition. Advances in cell culture methods have made human neuronal cultures accessible for experimental use in vitro and have aided in the development of neurotherapeutics. However, despite the adaptation of human neuronal cell cultures, in vitro models of human tauopathy are not yet widely available. This protocol describes a cellular model of tau aggregation in which human neurons are transduced with lentiviral-derived vectors that code for pathogenically mutated tau fused to a yellow fluorescent protein (YFP) reporter. Transduced cultures produce tau aggregates that stain positively for thioflavin and display markers of neurotoxicity, such as decreased axonal length and increased lysosomal volume. This procedure may be a useful and cost-effective model for studying human tauopathies.

This protocol details a procedure in which human neuronal cultures are transduced with lentiviral constructs coding for mutant human tau. Transduced cultures display tau aggregates and associated pathologies.

Aberrant aggregation of the protein tau is pathogenically involved in a number of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD). ...

Chronic Implantation of Multiple Flexible Polymer Electrode Arrays

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of scientific and clinical development. The use of flexible polymer electrode arrays can support long-lasting recording, but the same mechanical properties that allow for longevity of recording make multiple insertions and integration into a chronic implant a challenge. Here is a methodology by which multiple polymer electrode arrays can be targeted to a relatively spatially unconstrained set of brain areas.
The method utilizes thin-film polymer devices, selected for their biocompatibility and capability to achieve long-term and stable&#160;electrophysiologic recording interfaces. The resultant implant allows accurate and flexible targeting of anatomically distant regions, physical stability for months, and robustness to electrical noise. The methodology supports up to sixteen serially inserted devices across eight different anatomic targets. As previously demonstrated, the methodology is capable of recording from 1024 channels. Of these, the 512 channels in this demonstration used for single neuron recording yielded 375 single units distributed across six recording sites. Importantly, this method also can record single units for at least 160 days.
This implantation strategy, including temporarily bracing each device with a retractable silicon insertion shuttle, involves tethering of devices at their target depths to a skull-adhered plastic base piece that is custom-designed for each set of recording targets, and stabilization/protection of the devices within a silicone-filled, custom-designed plastic case. Also covered is the preparation of devices for implantation, and design principles that should guide adaptation to different combinations of brain areas or array designs.

Described below is a method for implantation of multiple polymer electrode arrays across anatomically distant brain regions for chronic electrophysiological recording in freely moving rats. Preparation and surgical implantation are described in detail, with emphasis on design principles to guide adaptation of these methods for use in other species.

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of ...