This work was supported by the European Project BRIANBOW (FP7- Young Explorers, The authors would like to thank Dr. Jacopo Tessadori for useful comments on the manuscript, and Silvia Chiappalone for her help in producing the graphics used in the video.

The procedure was done in accordance with the NIH standards for care and use of laboratory animals and was approved by the Tel-Aviv University Animal Care and Use Committee (permit number &#8211; L-14-019).
1. Preparation of Instruments and PDMS
<ol>
	<li>Prepare the wafer (Table of Materials, or order the wafer from a microfabrication lab), a scalpel, and a pair of tweezers &#8211; sterilization is not needed.</li>
	<li>Make poly-D-lysine (PDL) solution according to the following conditions: 4 mg/ml in 0.1 M borate buffer, pH 8, and store at -20 &#176;C.</li>
	<li>Prepare PDMS stencils according to the following procedure:
	<ol>
		<li>Prepare polydimethylsiloxane (PDMS, silicone elastomer) by mixing together the base and the curing agent with a ratio of 10:1, then stir the 2 substances for approximately 5 min.</li>
		<li>Place in vacuum chamber twice for 15 min each time and verify bubbles are gone.</li>
		<li>When the PDMS is ready, prepare spin coater: open nitrogen knobs to allow gas usage, place the silicon wafer (Figure 1A) on spinner, and use the vacuum to prevent it from moving.</li>
		<li>Pour PDMS over wafer and activate the spinner for 1 min at 1,000 rpm (creates approximately 100 &#181;m height).</li>
		<li>Place wafer on a hot plate at 100 &#176;C for 30 min.</li>
		<li>When the PDMS has hardened, outline the stencil&#39;s borders with PDMS, using Pasteur pipette. Place wafer on a hot plate at 100 &#176;C for 30 min.</li>
		<li>When the PDMS borders have hardened, use a scalpel to cut stencils according to the borders and peel off the silicon stencil from the wafer (a squared PDMS containing a single pattern).</li>
	</ol>
	</li>
</ol>
2. Petri Dish and Cover Slip Preparation
<ol>
	<li>Prepare 23 mm square glass cover slips and clean them according to the following order:
	<ol>
		<li>Wash with distilled water, 70% ethanol, and acetone.</li>
		<li>Wash with isopropanol and fast dry with nitrogen.</li>
	</ol>
	</li>
	<li>Place patterned PDMS stencils on coverslip, and gently press to verify that they are firmly attached to the glass surface. Place in vacuum chamber for 15 min.</li>
	<li>Drop 1 ml of PDL on stencil. Insert the coverslip into the vacuum chamber twice for 20 min each time, and leave PDL O/N to dry.</li>
	<li>Prepare Petri dish to create a supporting cell network:
	<ol>
		<li>Cover the surface of the dish (3.5 cm) with 1 ml of PDL for 2 hr in RT. Remove the PDL drop using a pipette.</li>
		<li>Wash with distilled water and leave to dry. Place a very small drop of silicone grease on each corner of the coverslip.</li>
	</ol>
	</li>
	<li>Place the coverslip on the center of the Petri dish (i.e., the PDMS should be facing up) and press gently to verify attachment.</li>
	<li>Gently remove PDMS from coverslip using tweezers. For sterilization, expose to UV illumination for 7 min.</li>
</ol>
3. Multi-Electrode Array (MEA) Preparation
<ol>
	<li>Clean MEA in this order (Table of Materials):
	<ol>
		<li>Wash with water under tap and sonicate in a concentrated, enzymatic detergent 3 times.</li>
		<li>Sonicate in distilled water 3 times.</li>
		<li>Wash with distilled water (in hood, 1,000 &#181;l tip) for 3 times and place under UV for 30 min.</li>
	</ol>
	</li>
	<li>Supporting network preparation:
	<ol>
		<li>Prepare the designed support network mold.
		<ol>
			<li>Pour PDMS into a 12-well plate (22 mm diameter).</li>
			<li>When the PDMS is hardened take out the mold and using a scalpel, cut a hole in the middle to create a ring shape.</li>
			<li>Place a designed support network mold on the center of the MEA (Figure 2A) and cover the rest of the surface with PDL for 2 hr at RT.</li>
		</ol>
		</li>
		<li>Remove PDL using pipette and wash with distilled water.</li>
		<li>Remove the mold and leave to dry (Figure 2A).</li>
	</ol>
	</li>
	<li>Align stencil to MEA in the following way:
	<ol>
		<li>Place stencil on designated micro manipulator. Use an inverted microscope to accurately align patterned structure to electrodes, and lower the stencil until it is placed on the MEA surface (Figure 2B). 
		Note: Contact with a well cleaned MEA provides competitive adhesion between the PDMS and the MEA itself.</li>
		<li>Lift the micromanipulator and if needed, use tweezers to apply a small amount of pressure above the PDMS to prevent it detaching from the MEA.</li>
		<li>Gently press stencil to MEA surface, and use microscope to verify it is firmly attached, and well aligned. (Figure 2C-D).</li>
	</ol>
	</li>
	<li>Place MEA in vacuum chamber for 15 min; put a 1 ml drop of PDL on stencil.</li>
	<li>Insert into vacuum chamber 2 times for 20 min each. Leave PDL to dry O/N in the incubator.</li>
	<li>Before plating, remove the PDMS stencils from MEAs and wash using sterile water. Place under UV illumination for 7 min.</li>
</ol>
4. Dissection and Culture
<ol>
	<li>Prepare cultures as described in Herzog et al. 2011 21.</li>
</ol>
5. Plating
<ol>
	<li>Calculate number of cells needed for plating, using a hemocytometer (optimal density according to the size of the circuit as discussed in the result).
	<ol>
		<li>Make sure the counting chamber (hemocytometer) is clean and place a cover slip on it (use alcohol to clean).</li>
		<li>Dilute 10 &#181;l of the cell suspension in 190 &#181;l of plating medium (dilution 1:20).</li>
		<li>Load 10 &#181;l of the diluted cells onto the edge of the counting chamber and slowly pipette the cells out allowing the chamber to fill itself.</li>
		<li>Using an inverted microscope, visualise the haemocytometer grid. Determine the number of cells in the chamber by direct counting (healthy cells should be round). Count the cells within the large square without those crossing the edges.</li>
		<li>Calculate the concentration of cells: 
		Total number of cells/1000 &#181;l = Total cells counted &#215; dilution factor &#215; 104 (area of the hemocytometer). 
		Note: For example, if the dilution factor was 20 and the total cells counted were 100: 
		100 &#215; 20 &#215; 104 = 0.1 x 107 cells/1,000 &#181;l or 1 x 106/100 &#181;l. In order to plate 0.75 x 106 cells, take 75 &#181;l out of the cells.</li>
	</ol>
	</li>
	<li>Take the number of cells needed for plating per single MEA or Petri dish, resuspend the cells to prevent aggregation, and plate them at the center, on top of the patternate area (if needed, it is possible to dilute the cells in the medium). For the MEA, place a 100 &#181;l drop in the middle. For cover slip, place a 1,000 &#181;l drop in the middle.</li>
	<li>Incubate the plated cells at 37 &#176;C for 40 min. Add plating medium to the plated cells, up to 1 ml per MEA and 2 ml per cover slip.</li>
	<li>Keep at 37 &#176;C and, every 4 days, dilute with fresh growth medium enriched with 0.5 Pen-Strep, 2% B-27 and 0.75% glutamax.</li>
	<li>From 6/7 DIV, after seeing connections between the islands, dilute medium with 10 &#181;l/ml FUDR (25 mg deoxyuridine + 62.5 mg uridine in 12.5 ml MEM (Minimum Essential Medium Eagle)) or any other anti-mitotic agent to prevent glial overgrowth.</li>
</ol>

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The authors have nothing to disclose.

A protocol to grow 2D modular neuronal networks in vitro composed of functionally inter-connected circuits is described. The procedure is based on patterning a cellular adhesive layer. Patterning is achieved with PDMS stencils reproducing the negative feature of the desired network architecture. PDMS stencils define the areas where the cellular adhesive layer is deposited. Once cells are plated, they spontaneously assemble to the coated islands and self-organize into active inter-connected circuits. Recordings o...

Experimental and theoretical evidences support the possibility that the brain operates through coordinated activation of cell assemblies1-5, which can be regarded as dynamic functional units that transiently interact with each other, shaping and underlying different brain states. Functional modularity is also dependent on and associated with the structural modular organization of the brain circuits6,7. How function and structure of brain circuits mutually shape each other is still one of the main open questions in neuroscience. To provide a deeper understanding of this question, it is important to identify optimal experimental frameworks where it is possible to address, at least partially, those issues. Since controlled manipulation of the spatio-temporal dynamics of neuronal networks in in vivo experiments is challenging, the development of in vitro neuronal networks models is of significant interest due to their easy accessibility, monitoring, manipulation and modeling8,9. In recent years, in vitro technologies supported by advanced substrate patterning methods have allowed to induce neuronal networks to develop a range of predefined modular structures3 and to study the functional properties of networks with imposed topologies10. In particular, methods were recently used to organize networks by imposing physical constraints4,11. Indeed, to study the link between structure and function in neuronal networks and to provide a simplified but plausible representation of interacting neuronal assemblies, in vitro systems should provide inter-connected neuronal sub-populations. Widely studied 2D homogenous neuronal cultures do not impose any spatial constraints on the self-organized emergent wiring of the circuits. Therefore a possible approach to shape artificially interconnected cell assemblies is to position different neuronal populations in spatially distinct areas. The distance among these areas does not prevent the inter assemblies connections. This approach, while ensuring a considerable control over network complexity, has been shown to provide a richer repertoire of synchronization models6,7,12.
In order to facilitate a reproducible culturing of modular neuronal assemblies, a protocol to assemble the self-organization of networks into neuronal clusters linked by axons and dendrites is presented and described. The polymeric structure for the physical confinement of neuronal cultures has been created from polydimtheylsiloxane (PDMS). PDMS is an elastomer widely used for biomedical applications owing to its biocompatibility, transparency and permeability to gases13. The PDMS is prepared and excluded from the micromachined SU8 207514,15 structures by spin-coating a liquid PDMS onto a &#34;master&#34; as described previously in Jackman et al.16 The achieved patterned neuronal networks are composed of inter-connected modules of different size and they have been successfully obtained on both coverslips and Micro Electrode Arrays (MEAs)17-20. The density of connections between the modules can change the features of the network synchronization, from a fully synchronized network, typical of uniform cultures, to transient states of synchronization among modules.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>PDMS, Sylgard 184</td>
			<td>Dow Corning</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Nalgene Vacuum Chamber</td>
			<td>Thermo</td>
			<td>5305-0609</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-D-Lysine PDL</td>
			<td>Sigma</td>
			<td>P7886</td>
			<td></td>
		</tr>
		<tr>
			<td>silicone grease - SILICAID 1010</td>
			<td>&#160;aidchim Ltd</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>Spin Coater</td>
			<td>Laurell - Technologies Corporation</td>
			<td>WS-650-23</td>
			<td></td>
		</tr>
		<tr>
			<td>12 well culture plate</td>
			<td>Sigma</td>
			<td>CLS3336</td>
			<td></td>
		</tr>
		<tr>
			<td>5-Fluoro-2&#8217;-deoxyuridine</td>
			<td>Sigma</td>
			<td>F0503</td>
			<td></td>
		</tr>
		<tr>
			<td>Uridine</td>
			<td>Sigma</td>
			<td>U3003</td>
			<td></td>
		</tr>
		<tr>
			<td>silicone grease - SILICAID 1010</td>
			<td>&#160;aidchim Ltd</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>MEA1060-Inv-BC</td>
			<td>Multi Channel Systems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>TC02</td>
			<td>Multi Channel Systems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Pen Strep</td>
			<td>Biological Industries Beit Haemek</td>
			<td>03-033-1c</td>
			<td></td>
		</tr>
		<tr>
			<td>B-27</td>
			<td>Gibco</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr>
			<td>glutaMAX</td>
			<td>Gibco</td>
			<td>35050-038</td>
			<td></td>
		</tr>
		<tr>
			<td>MEM Minimum Essential Medium-Eagle</td>
			<td>Biological Industries Beit Haemek</td>
			<td>01-025-1B</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Electrode Arrays 4Q</td>
			<td>Multi Channel Systems</td>
			<td>60-4QMEA1000iR-Ti-pr</td>
			<td>cleaning manual: http://www.multichannelsystems.com</td>
		</tr>
		<tr>
			<td>silicon wafer</td>
			<td>microchem</td>
			<td>SU8-2075</td>
			<td>Preparation protocol: www.microchem.com</td>
		</tr>
	</tbody>
</table>

design, surface treatment, cellular plating, and culturing of modular neuronal networks composed of functionally inter-connected circuits

The brain operates through the coordinated activation and the dynamic communication of neuronal assemblies. A major open question is how a vast repertoire of dynamical motifs, which underlie most diverse brain functions, can emerge out of a fixed topological and modular organization of brain circuits. Compared to in vivo studies of neuronal circuits which present intrinsic experimental difficulties, in vitro preparations offer a much larger possibility to manipulate and probe the structural, dynamical and chemical properties of experimental neuronal systems. This work describes an in vitro experimental methodology which allows growing of modular networks composed by spatially distinct, functionally interconnected neuronal assemblies. The protocol allows controlling the two-dimensional (2D) architecture of the neuronal network at different levels of topological complexity.
A desired network patterning can be achieved both on regular cover slips and substrate embedded micro electrode arrays. Micromachined structures are embossed on a silicon wafer and used to create biocompatible polymeric stencils, which incorporate the negative features of the desired network architecture. The stencils are placed on the culturing substrates during the surface coating procedure with a molecular layer for promoting cellular adhesion. After removal of the stencils, neurons are plated and they spontaneously redirected to the coated areas. By decreasing the inter-compartment distance, it is possible to obtain either isolated or interconnected neuronal circuits. To promote cell survival, cells are co-cultured with a supporting neuronal network which is located at the periphery of the culture dish. Electrophysiological and optical recordings of the activity of modular networks obtained respectively by using substrate embedded micro electrode arrays and calcium imaging are presented. While each module shows spontaneous global synchronizations, the occurrence of inter-module synchronization is regulated by the density of connection among the circuits.

A SU8-2075 mold on a silicon wafer with a feature thickness of approximately 100 &#181;m was used to shape the PDMS. The pattern was composed of squares of several dimensions, with a side length and distance varying between 200 and 700 &#181;m (Figure 1B). The size of the square was chosen to fit the field of view of a 10X (for islands with a side length &#60;800 &#181;m) and of a 20X objective (for islands with a side length &#60;400 &#181;m). Three parameters, namely cell plating density, distance betw...

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Design, Surface Treatment, Cellular Plating, and Culturing of Modular Neuronal Networks Composed of Functionally Inter-connected Circuits

This manuscript describes a protocol to grow in vitro modular networks consisting of spatially confined, functionally inter-connected neuronal circuits. A polymeric mask is used to pattern a protein layer to promote cellular adhesion over the culturing substrate. Plated neurons grow on coated areas establishing spontaneous connections and exhibiting electrophysiological activity.

The brain operates through the coordinated activation and the dynamic communication of neuronal assemblies. A major open question is how a vast repertoire ...

design-surface-treatment-cellular-plating-culturing-modular-neuronal

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JoVE Journal

Neuroscience

8.4K Views.  Tel-Aviv University. This manuscript describes a protocol to grow in vitro modular networks consisting of spatially confined, functionally inter-connected neuronal circuits. A polymeric mask is used to pattern a protein layer to promote cellular adhesion over the culturing substrate. Plated neurons grow on coated areas establishing spontaneous connections and exhibiting electrophysiological activity.

Video: Design, Surface Treatment, Cellular Plating, and Culturing of Modular Neuronal Networks Composed of Functionally Inter-connected Circuits

Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into diverse subtypes based on their immunochemical, morphological, and physiological properties1-4. Although previous research has revealed much about intrinsic properties of individual types of inhibitory neurons, knowledge about their local circuit connections is still relatively limited3,5,6. Given that each individual neuron's function is shaped by its excitatory and inhibitory synaptic input within cortical circuits, we have been using laser scanning photostimulation (LSPS) to map local circuit connections to specific inhibitory cell types. Compared to conventional electrical stimulation or glutamate puff stimulation, LSPS has unique advantages allowing for extensive mapping and quantitative analysis of local functional inputs to individually recorded neurons3,7-9. Laser photostimulation via glutamate uncaging selectively activates neurons perisomatically, without activating axons of passage or distal dendrites, which ensures a sub-laminar mapping resolution. The sensitivity and efficiency of LSPS for mapping inputs from many stimulation sites over a large region are well suited for cortical circuit analysis.
Here we introduce the technique of LSPS combined with whole-cell patch clamping for local inhibitory circuit mapping. Targeted recordings of specific inhibitory cell types are facilitated by use of transgenic mice expressing green fluorescent proteins (GFP) in limited inhibitory neuron populations in the cortex3,10, which enables consistent sampling of the targeted cell types and unambiguous identification of the cell types recorded. As for LSPS mapping, we outline the system instrumentation, describe the experimental procedure and data acquisition, and present examples of circuit mapping in mouse primary somatosensory cortex. As illustrated in our experiments, caged glutamate is activated in a spatially restricted region of the brain slice by UV laser photolysis; simultaneous voltage-clamp recordings allow detection of photostimulation-evoked synaptic responses. Maps of either excitatory or inhibitory synaptic input to the targeted neuron are generated by scanning the laser beam to stimulate hundreds of potential presynaptic sites. Thus, LSPS enables the construction of detailed maps of synaptic inputs impinging onto specific types of inhibitory neurons through repeated experiments. Taken together, the photostimulation-based technique offers neuroscientists a powerful tool for determining the functional organization of local cortical circuits.

This paper introduces an approach of combining laser scanning photostimulation with whole cell recordings in transgenic mice expressing GFP in limited inhibitory neuron populations. The technique allows for extensive mapping and quantitative analysis of local synaptic circuits of specific inhibitory cortical neurons.

Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

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 ...

Closed-loop Neuro-robotic Experiments to Test Computational Properties of Neuronal Networks

Information coding in the Central Nervous System (CNS) remains unexplored. There is mounting evidence that, even at a very low level, the representation of a given stimulus might be dependent on context and history. If this is actually the case, bi-directional interactions between the brain (or if need be a reduced model of it) and sensory-motor system can shed a light on how encoding and decoding of information is performed. Here an experimental system is introduced and described in which the activity of a neuronal element (i.e., a network of neurons extracted from embryonic mammalian hippocampi) is given context and used to control the movement of an artificial agent, while environmental information is fed back to the culture as a sequence of electrical stimuli. This architecture allows a quick selection of diverse encoding, decoding, and learning algorithms to test different hypotheses on the computational properties of neuronal networks.

In this paper, an experimental framework to perform closed-loop experiments is presented, in which information processing (i.e., coding and decoding) and learning of neuronal assemblies are studied during the continuous interaction with a robotic body.

Information coding in the Central Nervous System (CNS) remains unexplored. There is mounting evidence that, even at a very low level, the representation ...

Rewiring Neuronal Circuits: A New Method for Fast Neurite Extension and Functional Neuronal Connection

Brain and spinal cord injury may lead to permanent disability and death because it is still not possible to regenerate neurons over long distances and accurately reconnect them with an appropriate target. Here a procedure is described to rapidly initiate, elongate, and precisely connect new functional neuronal circuits over long distances. The extension rates achieved reach over 1.2 mm/h, 30-60 times faster than the in vivo rates of the fastest growing axons from the peripheral nervous system (0.02 to 0.04 mm/h)28 and 10 times faster than previously reported for the same neuronal type at an earlier stage of development4. First, isolated populations of rat hippocampal neurons are grown for 2-3 weeks in microfluidic devices to precisely position the cells, enabling easy micromanipulation and experimental reproducibility. Next, beads coated with poly-D-lysine (PDL) are placed on neurites to form adhesive contacts and pipette micromanipulation is used to move the resulting bead-neurite complex. As the bead is moved, it pulls out a new neurite that can be extended over hundreds of micrometers and functionally connected to a target cell in less than 1 h. This process enables experimental reproducibility and ease of manipulation while bypassing slower chemical strategies to induce neurite growth. Preliminary measurements presented here demonstrate a neuronal growth rate far exceeding physiological ones. Combining these innovations allows for the precise establishment of neuronal networks in culture with an unprecedented degree of control. It is a novel method that opens the door to a plethora of information and insights into signal transmission and communication within the neuronal network as well as being a playground in which to explore the limits of neuronal growth. The potential applications and experiments are widespread with direct implications for therapies that aim to reconnect neuronal circuits after trauma or in neurodegenerative diseases.

This procedure describes how to rapidly initiate, extend and connect neurites organized in microfluidic chambers using poly-D-lysine-coated beads fixed to micropipettes that guide neurite elongation.

Brain and spinal cord injury may lead to permanent disability and death because it is still not possible to regenerate neurons over long distances and ...

Determining Cell-surface Expression and Endocytic Rate of Proteins in Primary Astrocyte Cultures Using Biotinylation

Cell-surface proteins mediate a wide array of functions. In many cases, their activity is regulated by endocytic processes that modulate their levels at the plasma membrane. Here, we present detailed protocols for 2 methods that facilitate the study of such processes, both of which are based on the principle of the biotinylation of cell-surface proteins. The first is designed to allow for the semi-quantitative determination of the relative levels of a particular protein at the cell-surface. In it, the lysine residues of the plasma membrane proteins of cells are first labeled with a biotin moiety. Once the cells are lysed, these proteins may then be specifically precipitated via the use of agarose-immobilized streptavidin by exploiting the natural affinity of the latter for biotin. The proteins isolated in such a manner may then be analyzed via a standard western blotting approach. The second method provides a means of determining the endocytic rate of a particular cell-surface target over a period of time. Cell-surface proteins are first modified with a biotin derivative containing a cleavable disulfide bond. The cells are then shifted back to normal culture conditions, which causes the endocytic uptake of a proportion of biotinylated proteins. Next, the disulfide bonds of non-internalized biotin groups are reduced using the membrane-impermeable reducing agent glutathione. Via this approach, endocytosed proteins may thus be isolated and quantified with a high degree of specificity.

Two biotinylation-based methods, designed for determining the cell-surface expression and endocytic rate of proteins expressed at the plasma membrane, are presented in this report.

Cell-surface proteins mediate a wide array of functions. In many cases, their activity is regulated by endocytic processes that modulate their levels at ...

Chronic Transcranial Electrical Stimulation and Intracortical Recording in Rats

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a closed-loop event-triggered manner. Although many studies are focusing on the possible benefits and side-effects of TES in healthy and pathologic brains, there are still many fundamental open questions regarding the mechanism of action of the stimulation. Therefore, there is a clear need for a robust and reproducible method to test the acute and the chronic effects of TES in rodents. TES can be combined with regular behavioral, electrophysiological, and imaging techniques to investigate neuronal networks in vivo. The implantation of transcranial stimulation electrodes does not impose extra constraints on the experimental design while it offers a versatile, flexible tool to manipulate brain activity. Here we provide a detailed, step-by-step protocol to fabricate and implant transcranial stimulation electrodes to influence brain activity in a temporally constrained manner for months.

This detailed protocol describes transcranial stimulation electrode placement on the temporal bone in order to investigate the short- and long-term effects of transcranial electrical stimulation in freely moving rats.

Transcranial electrical stimulation (TES) is a powerful and relatively simple approach to diffusely influence brain activity either randomly or in a ...

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 ...

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in the brain sense various biological conditions in the periphery and transmit the signals to neurons. Microglia, immune cells in the brain, are involved in synaptic development and plasticity. Therefore, the contribution of microglia to neural circuit construction in response to the internal state of the body should be tested critically by intravital imaging of the relationship between microglial dynamics and neuronal activity.
Here, we describe a technique for the simultaneous imaging of microglial dynamics and neuronal activity in awake mice. Adeno-associated virus encoding R-CaMP, a gene-encoded calcium indicator of red fluorescence protein, was injected into layer 2/3 of the primary visual cortex in CX3CR1-EGFP transgenic mice expressing EGFP in microglia. After viral injection, a cranial window was installed onto the brain surface of the injected region. In vivo two-photon imaging in awake mice 4 weeks after the surgery demonstrated that neural activity and microglial dynamics could be recorded simultaneously at the sub-second temporal resolution. This technique can uncover the coordination between microglial dynamics and neuronal activity, with the former responding to peripheral immunological states and the latter encoding the internal brain states.

Here, we describe a protocol combining adeno-associated virus injection with cranial window implantation for simultaneous imaging of microglial dynamics and neuronal activity in awake mice.

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in ...

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the cytoplasm or the release of intracellular Ca2+ stores. Pirt-GCaMP3-based Ca2+ imaging of primary sensory neurons of the dorsal root ganglion (DRG) in mice offers the advantage of simultaneous measurement of a large number of cells. Up to 1,800 neurons can be monitored, allowing neuronal networks and somatosensory processes to be studied as an ensemble in their normal physiological context at a populational level in vivo. The large number of neurons monitored allows the detection of activity patterns that would be challenging to detect using other methods. Stimuli can be applied to the mouse hindpaw, allowing the direct effects of stimuli on the DRG neuron ensemble to be studied. The number of neurons producing Ca2+ transients as well as the amplitude of Ca2+ transients indicates sensitivity to specific sensory modalities. The diameter of neurons provides evidence of activated fiber types (non-noxious mechano vs. noxious pain fibers, A&#946;, A&#948;, and C fibers). Neurons expressing specific receptors can be genetically labeled with td-Tomato and specific Cre recombinases together with Pirt-GCaMP. Therefore, Pirt-GCaMP3 Ca2+ imaging of DRG provides a powerful tool and model for the analysis of specific sensory modalities and neuron subtypes acting as an ensemble at the populational level to study pain, itch, touch, and other somatosensory signals.

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

This protocol describes the surgical exposure of the dorsal root ganglion (DRG) followed by GCaMP3 (genetically-encoded Ca2+ indicator; Green Fluorescent Protein-Calmodulin-M13 Protein 3) Ca2+ imaging of the neuronal ensembles using Pirt-GCaMP3 mice while applying a variety of stimuli to the ipsilateral hind paw.

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the ...