We would like to thank Ms. Rui Chen for the dissection and images of the Drosophila CNS shown in the figures.

1. Equipment and Materials
<ol>
	<li>Prepare the required components (listed in Table of Materials) of the electrophysiology rig to perform suction electrode recordings of the Drosophila CNS. 
	NOTE: Prior to experimentation, it is necessary to construct chambers for dissection of the Drosophila CNS and to be used for bathing the ganglia in saline during recordings. A step-by-step outline of chamber construction is provided below.</li>
	<li>Prepare the larval chamber.
	<ol>
		<li>Melt the black wax using a hot plate.</li>
		<li>Pour 2 mL of melted wax into a chamber that has a maximum volume of 2 - 2.5 mL.</li>
		<li>Let the wax cool and harden for approximately 2 h.</li>
		<li>Use a razor blade to carve a hole in the hardened wax that will hold a volume of 500 &#181;L, which has a surface area of approximately 0.5 cm2 and a depth of approximately 0.25 cm.</li>
	</ol>
	</li>
</ol>
2. Equipment and Software Configuration
NOTE: The setup of the extracellular recording is briefly described below.
<ol>
	<li>Prepare equipment for dissection and recording.
	<ol>
		<li>Position the tissue preparation, microscope, suction electrode, micromanipulators, light source inside the Faraday cage to reduce noise and eliminate extraneous electrical fields (Figure 2).</li>
		<li>Connect (preferably solder) ground and positive wires onto shielded alligator clips that will be connected to the bath and microelectrode holder, respectively. 
		NOTE: Soldered and exposed wires can be covered with aluminum foil to reduce noise.</li>
		<li>Connect the ground and positive wires to input 1 of the AC/DC differential amplifier.</li>
		<li>Connect the data acquisition system to the AC/DC differential amplifier by connecting a Bayonet Neill-Concelman (BNC) cable from input 1 of the data acquisition system to the channel 1 output of the amplifier. 
		NOTE: It is recommended to incorporate a 50/60 Hz noise eliminator between the data acquisition software and the amplifier. The use of a BNC T-connector is required.</li>
		<li>If desired, include an audio monitor into the setup by connecting input 1 of the audio monitor to input 1 of the data acquisition software.</li>
		<li>Fill the 10 mL syringe with saline and connect it to the pressure port of the electrode holder. 
		NOTE: Ensure that no air bubbles exist between the syringe, the Ag/AgCl pellet, and the electrode opening.</li>
	</ol>
	</li>
	<li>Prepare the software for dissection and recording. 
	NOTE: The outline of methods for software setup is based on the acquisition/analysis software listed in the Table of Materials which will digitize the raw electrical output and convert the data to spike frequency. However, other software can be used and multiple recording conversions can be used.
	<ol>
		<li>Open the acquisition/analysis software.</li>
		<li>Click on &#34;setup&#34; from the main toolbar and select &#34;channel settings,&#34; which will open a dialogue box.</li>
		<li>Reduce the number of total channels to three.</li>
		<li>For Channel 1, select a range of 100 mV.&#160;</li>
		<li>For Channel 2, click on the &#34;calculation&#34; tab that will open a drop-down menu. Select &#34;cyclic measurements,&#34; which will open a second dialogue box.</li>
		<li>Select channel 1 as the source. Then on the drop down menu measurement, select unit spike at events. Finally, in the detection settings area, from the drop down menu select simple threshold.&#160;</li>
		<li>To convert the electrical activity into a rate plot expressed in hertz, the 3rd channel must be set in arithmetic mode: On the input window, type in &#34;1000*smoothsec(ch2,2)&#34;. Then select as the units drop down menu hertz. 
		NOTE: Successful recordings can be performed with multiple different recording setups, but &#34;Unit spike at events and simple threshold&#34; is likely the easiest, since it allows for adjustment of the threshold above the background activity for each individual recording. Recordings under the setting of &#34;Custom-Source Input&#34; increase reproducibility of the data assuming the background activity is not different between recordings.</li>
		<li>Close dialogue boxes to return to the main screen. The y-axis should be expressed in Hz and the x-axis in time.</li>
	</ol>
	</li>
</ol>
3. Dissect and Prepare the Larval Drosophila CNS
NOTE: Methods for larval CNS dissection are clearly illustrated in Hafer and Schedl27, but these previously published methods reduce the length of the descending neurons that are important for measuring spike frequency. Here, an additional method is outlined to excise the larval CNS that maintains long, intact descending neurons.
<ol>
	<li>Saline preparation.
	<ol>
		<li>Prepare the dissection and recording saline to the following in mM28: 157 NaCl, 3 KCl, 2 CaCl2, 4 2-[4-(2-hydroxyethyl)piperazin-1-yl]ethanesulfonic acid (HEPES); titrate pH to 7.25.</li>
	</ol>
	</li>
	<li>Dissect the larval Drosophila CNS.
	<ol>
		<li>Identify and extract a wandering third-instar Drosophila melanogaster from the culture vial and place into 200 &#181;L of saline (Figure 3A).</li>
		<li>Grasp the mouth hooks with a pair of fine forceps and grasp the abdomen of the maggot with a second pair of forceps (Figure 3B). 
		NOTE: Do not apply too much pressure to the abdomen as this can result in tearing of the thin cuticular layer of the maggot.</li>
		<li>Gently pull the mouth hooks and abdomen in different directions to separate the caudal end of the maggot from the head region and expose the viscera of the maggot (Figure 3C). 
		NOTE: The CNS will be intertwined with the trachea and digestive tract. Do not cut the CNS away from any tissue to prevent cutting the descending peripheral nerves.</li>
		<li>Tease the CNS out of the digestive tract and trachea with forceps (Figure 3D).</li>
		<li>If necessary, disrupt the blood brain barrier by manually transecting the CNS posterior to the cerebral lobes with Vannas spring scissors. 
		NOTE: This should be done based on the physiochemical properties of the chemical being used. The red line in Figure 4A is the suggested transection point and the transected CNS with intact descending peripheral nerve trunks shown in Figure 4B. The transected CNS is ready for experimentation after completion of step 3.2.5.</li>
	</ol>
	</li>
</ol>
4. Extracellular Recording of Drosophila CNS.
<ol>
	<li>Prepare the CNS for recording.
	<ol>
		<li>Pull a glass pipette electrode from borosilicate glass capillaries to a resistance of 5-15 m&#937;.</li>
		<li>Insert the transected CNS into the wax chamber that contains 200 &#181;L of saline. Clamp an uncoated insect pin with the alligator clip soldered to the ground wire and insert the pin into the saline to complete the circuit.</li>
		<li>Using the micromanipulators, orient the electrode to the caudal end of the transected CNS. Eliminate recording of background noise by adjusting the threshold level in the acquisition/analysis software prior to contacting the peripheral nerve trunks.</li>
		<li>Draw any convenient peripheral nerves into the suction electrode by applying slight negative pressure on the syringe. 
		NOTE: The baseline firing frequency is correlated to the number of neurons drawn into the electrode where more neurons oftentimes result in increased resistance of the electrode.</li>
	</ol>
	</li>
	<li>Begin extracellular recording of CNS descending nerve activity.
	<ol>
		<li>Start the recording on the data acquisition software and monitor the baseline firing frequency. Allow the firing rate to equilibrate for 5 min prior to collecting baseline firing rate data.</li>
		<li>Discard the preparation and recording if the pattern of firing of control treatment is not a bursting pattern similar to that shown in Figure 5A. 
		NOTE: Altered pattern of firing suggests abnormal or unstable activity of the central pattern generator, which can alter neural function.</li>
		<li>After 5 min, add 200 &#181;L of saline + vehicle to bring the total volume of the chamber to 400 &#181;L to begin recording control firing rates.</li>
		<li>After baseline has been established (3-5 min), withdraw 200 &#181;L of saline and add 200 &#181;L of the experimental agent solubilized in saline. 
		NOTE: Dimethyl sulfoxide (DMSO) is the recommended solvent for lipophilic compounds and should not exceed 0.1% v/v.</li>
		<li>Apply drug concentrations in a serial manner (low to high concentrations) and record each concentration for a select period of time (determined by the investigator based on the chemical properties of the drug29). Label this time point of drug application in the acquisition/analysis software by including a comment that includes the drug and the final concentration. 
		NOTE: Firing activity of the CNS will decline by approximately 10-20% after 30-50 min after the dissection and therefore, appropriate untreated controls should be performed, and drug treatments should not exceed this time period.</li>
	</ol>
	</li>
	<li>Analyze the data. 
	NOTE: Multiple analyses can be performed on the collected data, such as changes in action potential bursts29, spike waveform amplitude and time course, and determining mean spike frequencies after drug treatment26,30,31,32. The most common is determining the influence a drug has to the mean spike frequencies over a specified period of time, which is described below.
	<ol>
		<li>Tabulate mean spike frequencies by selecting the entire region of interest (i.e., the drug treatment time period) and determine mean spike frequencies automatically through the &#34;Datapad&#34; located on the main toolbar of the acquisition/analysis software.</li>
		<li>Determine the mean spike frequency of the CNS after drug treatment to the mean spike frequency of the vehicle control (baseline). Calculate the percent change of firing rate after drug treatment by the formula: (Treated Frequency/Baseline Frequency) &#215; 100.</li>
		<li>Use the mean spike frequencies or percent firing of control for each concentration to construct a concentration response curve with standard graphing software.</li>
		<li>Perform statistical analysis (e.g., unpaired t-test) to determine significance between time points, concentrations, or drug treatments. 
		NOTE: There are two primary methods for generating and analyzing concentration-response curves. The first method is one concentration per individual preparation. Here, the spike rate of the single concentration is normalized to baseline spike rate for each preparation. The benefits are reduced run down of the preparation due to shortened recording time, while the pitfalls are increased dissection time because this method will consist of 5-7 individual CNS preparations per concentration, and larger error bars on the averaged data set. The second method is multiple concentrations per individual preparation. Here, the spike rate for each of the 3-5 concentrations are normalized to the same baseline spike rate. The benefits are less dissection time and less variability between replicates for drug concentrations, while the pitfalls are the requirement of a fast-acting drug and unknown impact of previous drug concentration treatments on subsequent chemical treatments.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The details provided in the associated video and text have provided key steps in order to record the activity and spike discharge frequency of the Drosophila CNS ex vivo. The dissection efficacy is the most critical aspect of the method because short or few descending neurons will reduce the baseline firing rate that will result in large variances between replicates. However, once the dissection technique has been mastered, the data collected with this assay are highly reproducible and amendable for a w...

The overall goal of this approach is to enable researchers to quickly measure the electrogenesis of the central nervous system (CNS) in the model insect, Drosophila melanogaster. This method is reliable, quick, and cost-efficient to perform physiological and toxicological experimentation. The CNS is essential for life and therefore, the physiological pathways critical for proper neural function have been explored extensively in an effort to understand or modify neural function. Characterization of the signaling pathways within the arthropod CNS has enabled the discovery of several chemical classes of insecticides that disrupt invertebrate neural function to induce mortality while limiting off-target consequences. Thus, the ability to measure the neural activity of insects is of significant interest to the field of insect toxicology and physiology since the nervous system is the target tissue of the majority of deployed insecticides1. Yet, continued growth of fundamental and applied knowledge regarding the insect nervous system requires advanced neurophysiological techniques that are limited in feasibility, since current techniques are labor intensive and require a high expense, insect neural cell lines are limited, and/or there is limited access to the central synapses of most arthropods. Currently, characterization of most insect neural proteins requires the target to be cloned and heterologously expressed for subsequent drug discovery and electrophysiological recordings, as was described for insect inward rectifier potassium channels2, insect ryanodine receptor3, mosquito voltage-sensitive K+ channels4, and others. To mitigate the requirement for heterologous expression and the potential for low functional expression, Bloomquist and colleagues aimed to induce a neuronal phenotype in cultured Spodoptera frugiperda (Sf21) cells as a novel method for insecticide discovery5,6. These methods provide a valid approach for the development of new chemistry, yet they oftentimes create an insurmountable bottleneck for the characterization of pharmacological agents, identifying mechanisms of insecticide resistance, and characterization of fundamental physiological principles. Here, we describe an ex vivo method that enables the recording of electrical activity from a model insect that has malleable genetics7,8,9 and known expression patterns of neural complexes10,11,12 to enable the characterization of resistance mechanisms at the level of the nerve, the mode of action of newly developed drugs, and other toxicological studies.
The fruit fly, D. melanogaster, is a common model organism for defining insect neural systems or insecticide mechanism of action and has been established as a well-suited model organism for the study of toxicological13, pharmacological14,15, neurophysiological16, and pathophysiological17,18,19,20 processes of vertebrates. D.melanogaster is a holometabolous insect that performs complete metamorphosis, including a larval and pupal stage before reaching the reproductive adult stage. Throughout the developmental process, the nervous system undergoes significant remodeling at different life stages, but the larval CNS will be the focus of this methodology. The fully developed larval CNS is anatomically simple with thoracic and abdominal segments that are fused and form the ventral ganglion, which represents an array of repeated and almost identical neuromeric units21,22. Descending motor nerves originate from the caudal end of the subesophageal ganglia and descend to innervate body wall muscles and visceral organs of the larvae. Figure 1 describes the gross anatomy of the larval Drosophila CNS.
The Drosophila blood-brain barrier (BBB) develops at the end of embryogenesis and is formed by subperineurial glial cells (SPG)21. The SPG cells form numerous filopodia-like processes that spread out to establish a contiguous, very flat, endothelial-like sheet that covers the entire Drosophila CNS23. The Drosophila BBB has similarities to the vertebrate BBB, which includes preserving the homeostasis of the neural microenvironment by controlling the entry of nutrients and xenobiotics into the CNS21. This is a prerequisite for reliable neural transmission and function, yet the protection of the CNS by the BBB restricts the permeation of synthetic drugs, most peptides, and other xenobiotics24,25, which introduces potential problems when characterizing potencies of small-molecule modulators. The method uses a simple transection to disrupt this barrier and provide ready pharmacological access to the central synapses. 
The greatest strength of the described methodology is the simplicity, reproducibility, and relatively high-throughput capacity inherent to this system. The protocol is relatively easy to master, the setup requires little space, and only an initial financial input is necessary which is reduced to reagents and consumables. Further, the described method is completely amendable to record the central descending nerve activity of the house fly, Musca domestica26.

<table>
	<tbody>
		<tr>
			<td>Drosophila melanogaster (strain OR)</td>
			<td>Bloomington Drosophila Stock Center</td>
			<td>2376</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibration isolation table</td>
			<td>Kinetic Systems</td>
			<td>9200 series</td>
			<td></td>
		</tr>
		<tr>
			<td>Faraday Cage</td>
			<td>Kinetic Systems</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting Microscope on a Boom</td>
			<td>Nikon</td>
			<td>SMZ800N</td>
			<td>Multiple scopes can be used; boom stand is critical</td>
		</tr>
		<tr>
			<td>AC/DC differential amplifier</td>
			<td>ADInstruments</td>
			<td>AM3000H</td>
			<td>The model 1700 can be used instead of the model 3000</td>
		</tr>
		<tr>
			<td>audio monitor</td>
			<td>ADInstruments</td>
			<td>AM3300</td>
			<td></td>
		</tr>
		<tr>
			<td>Hum Bug Noise Eliminator</td>
			<td>A-M Systems</td>
			<td>726300</td>
			<td></td>
		</tr>
		<tr>
			<td>Data Acquisition System (PowerLab)</td>
			<td>ADInstruments</td>
			<td>PL3504</td>
			<td>Multiple PowerLab models can be used.</td>
		</tr>
		<tr>
			<td>Lab Chart Pro Software</td>
			<td>ADInstruments</td>
			<td>N/A - Online Download</td>
			<td></td>
		</tr>
		<tr>
			<td>Fiber Optic Lights</td>
			<td>Edmund Optics</td>
			<td>89-740</td>
			<td>Different light sources can be used, but fiber optics are the most adaptable</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>World Precision Instruments</td>
			<td>M325</td>
			<td></td>
		</tr>
		<tr>
			<td>Microelectrode Holder</td>
			<td>World Precision Instruments</td>
			<td>MEH715</td>
			<td>Different models are acceptable</td>
		</tr>
		<tr>
			<td>BNC cables</td>
			<td>World Precision Instruments</td>
			<td>multiple based on size</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Capillaries</td>
			<td>World Precision Instruments</td>
			<td>PG52151-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Microelectrode Puller</td>
			<td>Sutter Instruments</td>
			<td>P-1000</td>
			<td>Also can use Narashige PC-100</td>
		</tr>
		<tr>
			<td>Black Wax</td>
			<td>Carolina Biological Supply</td>
			<td>974228</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-coated insect pins, size #2</td>
			<td>Bioquip</td>
			<td>1208S2</td>
			<td></td>
		</tr>
		<tr>
			<td>Fince Forceps</td>
			<td>Fine Science Tools</td>
			<td>11254-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas Spring Scissors</td>
			<td>Fine Science Tools</td>
			<td>15000-03</td>
			<td></td>
		</tr>
	</tbody>
</table>

electrophysiological recording of the central nervous system activity of third-instar drosophila melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Spontaneous activity of the descending peripheral nerves arising from the Drosophila central nervous system can be recorded using extracellular suction electrodes with consistent reproducibility. Spontaneous activity of the excised and transected Drosophila CNS produces a cyclical pattern of bursting with 1-2 s of firing with approximately 1 s of near quiescent activity. For example, the CNS is near quiescent (1-2 Hz) for 0.5-1 s, followed by a burst (100-400 Hz) for app...

Watch this Scientific Journal Video about Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster at JoVE.com

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

electrophysiological-recording-central-nervous-system-activity-third

Research

JoVE Journal

Neuroscience

11.4K Views.  Louisiana State University AgCenter. This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

Video: Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the procedure for the visualization of Drosophila embryonic nervous system using antibodies to neuronal proteins. Since the entire embryonic peripheral nervous and central nervous systems are well characterized at the level of individual cells (Dambly-Chaudi&egrave;re et al., 1986; Bodmer et al., 1987; Bodmer et al., 1989), any aberrations to these systems can be easily identified using antibodies to different neuronal proteins. The developing embryos are collected at certain times to ensure that the embryos are in the proper developmental stages for visualization. After collection, the outer layers of the embryo, the chorion membrane and the vitelline envelope that surrounds the embryo, are removed before fixation. Embryos are then incubated with neuronal antibodies and visualized using fluorescently labeled secondary antibodies. Embryos at stages 12-17 are visualized to access the embryonic nervous system. At stage 12 the CNS germ band starts shortening and by stage 15 the definitive pattern of the commissure has been achieved. By stage 17 the CNS contracts and the PNS is fully developed (Campos-Ortega et al. 1985). Thus changes in the pattern of the PNS and CNS can be easily observed during these developmental stages.

Visualization of the Embryonic Nervous System in Whole-mount Drosophila Embryos

We describe the procedure to prepare staged Drosophila embryos for the visualization of the embryonic nervous system during embryogenesis.

The Drosophila embryo is an attractive model system for investigating the cellular and molecular basis of neuronal development. Here we describe the ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

This protocol describes extracellular recording of the action potential responses fired by labellar taste neurons in Drosophila.

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such behavior is spontaneous locomotor activity. Here we describe our protocol that utilizes Drosophila population monitors and a tracking system that allows continuous monitoring of the spontaneous locomotor activity of flies for several days at a time. This method is simple, reliable, and objective and can be used to examine the effects of aging, sex, changes in caloric content of food, addition of drugs, or genetic manipulations that mimic human diseases.

Determination of the Spontaneous Locomotor Activity in Drosophila melanogaster

Drosophila melanogaster are useful in studying genetic or environmental manipulations that affect behaviors such as spontaneous locomotor activity.&#160; Here we describe a protocol that utilizes monitors with infrared beams and data analysis software to quantify spontaneous locomotor activity.&#160;

Drosophila melanogaster has been used as an excellent model organism to study environmental and genetic manipulations that affect behavior. One such ...

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 ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...