The authors would like to thank the following for funding this work: generous start-up funds from the Department of Biomedical Engineering in Washington University, a McDonnell Center for Systems Neuroscience grant, a Office of Naval Research grant (Grant#: N000141210089) to B.R.

1. Odor Preparation and Delivery
<ol>
 <li>Dilute odor solutions in mineral oil by volume to achieve the desired concentration level. Store a 20 ml mixture of mineral oil and the odorant in a 60 ml glass bottle. Insert two syringe needles into a rubber stopper (gauge 19), one from the bottom and the other from the top, to provide an inlet and an outlet line. Seal the glass bottle with this rubber stopper and attach a custom designed activated carbon filter to the inlet line (Figure 1A).</li>
 <li>The carbon filter is made using two 6 ml syringes. Cut the syringes in half and discard the plunger end. Fill each of the remaining pieces with cotton and activated charcoal before connecting them together using heat-shrink tubing.</li>
 <li>Connect the outlet line of the odor bottle (using polyethylene tubing, I.D. 0.86 mm) to the plastic tube (Nalgene FEP Tubing, I.D. 5.8 mm) that supplies a constant airflow across the antenna (Figure 1B).</li>
 <li>Direct carbon-filtered, dehumidified air (carrier gas; flow-rate, 0.75 L/min) towards the locust using a plastic tube placed within a few cm of the locust antenna.</li>
 <li>For odor stimulation, displace a constant volume (0.1 L/min) of the static headspace above the odor solution in the bottle. This is achieved by injecting an equal amount of dehumidified air into the bottle using a Pico-pump (WPI, PV-820). The vapors from the odor bottle are directed through the outlet line onto the airflow tube (Figure 1B).</li>
 <li>Remove the delivered odor vapors by placing a vacuum funnel ~10 cm behind the locust antenna.</li>
</ol>
2. Preparing Locust Antenna for Single Sensillum Recording
<ol>
 <li>Select a young-adult locust of either sex with fully-grown wings, but prior to the mating stage from a crowded colony. To restrain the locust, first amputate its legs. Seal the amputation sites with tissue adhesive (Vetbond, 3M). Secure the locust to a custom designed chamber using a small piece of electrical tape draped around its thorax (Figure 2A).</li>
 <li>Under a dissection microscope, make a shallow groove in the wax platform (Figure 2A) for antenna placement. Place the antenna into the groove and stabilize it using batik wax at the two ends of the antenna (Figure 2B; an electrowaxer is used for melting the wax).</li>
 <li>Insert a ground electrode (chlorided silver wire) into the thorax (~1 cm away from head). Use batik wax to both seal the incision site and hold the ground wire in place (Figure 2A).</li>
</ol>
3. Single Sensillum Recording to Monitor Odor-evoked Responses from Olfactory Receptor Neurons (ORNs)
<ol>
 <li>Place the stabilized locust antenna under a stereomicroscope (Leica M205C) on a vibration isolation table (TMC) (Figure 3A). Make sure that the base of the recording sensillum is clearly visible (Figure 3B).</li>
 <li>Use a micropipette puller (Sutter P-1000) to fabricate glass electrodes (impedance 3-10 M&Omega; when filled with locust saline9, tip diameter 1-3 &mu;m) using a borosilicate glass capillary tube (O.D. 1.2 mm, I.D. 0.69 mm).</li>
 <li>Place the glass electrode into a micropipette holder that is attached to a motorized micromanipulator (Sutter MP-285). Gently insert the electrode into the base of a sensillum (Figure 3B). Note that each sensillum can contain 3-50 ORNs in locusts10.</li>
 <li>Amplify the signal (10,000 times) using an A.C. amplifier (Grass P-55). Filter the signal between 0.3 - 10.0 kHz and acquire at a 15 kHz sampling rate (Figure 3D) using a data acquisition system (LabView, PCI-MIO-16E-4 DAQ cards; National Instruments).</li>
</ol>
4. Locust Dissection Procedure for Antennal Lobe and Mushroom Body Recordings
<ol>
 <li>Follow the restraining procedures as described in section 2.1. Position the locust in a custom-designed chamber as shown in Figure 4A and B.</li>
 <li>To perfuse saline during and after the dissection procedure, build a wax cup around the locust head. The wax cup should start just above the mouth parts, and extend beyond the compound eyes encompassing the region between the two antennae as shown in Figure 4C.</li>
 <li>To allow the antennae to pass through the wax cup, create small tunnels on both sides using plastic (polyethylene) tubing (5 mm long, I.D. 0.86 mm). Ensure that the plastic tubing can slide through the wax cup. The latter is achieved by using a rubber gasket that tightly wraps around the plastic tube but is attached to the wax-cup (Figure 4C).</li>
 <li>Using epoxy resin, attach the base of the antennae to the bottom end of the plastic tubing. This step ensures that the antennae are held in place even after the surrounding cuticle is removed.</li>
 <li>Keep the wax cup filled with saline solution9 from this point onwards. Begin by removing a central rectangular region between the two antennae (longer side of the rectangle aligned with the anteroposterior axis). Subsequently, remove cuticle in neighboring regions without disturbing the compound eyes and cuticle at the base of the antenna (Figure 4D).</li>
 <li>Using fine forceps, gently remove air sacs and fat bodies surrounding the brain. At the end of this step, the locust brain should be clearly seen (Figure 4D). Notice that the brain regions that process the olfactory information (with light yellow pigmentation) are situated between the two antennae.</li>
 <li>In locusts the gut runs below the brain and along the length of the body. To prevent movement of the gut from potentially destabilizing the preparation, gently pull the foregut and cut it using fine scissors. Make a small incision in the abdomen just above the rectum and remove the gut by pulling the hindgut with coarse forceps. To prevent a saline leak, tie the abdomen immediately anterior to the incision site with suture threads.</li>
 <li>Use a small platform made of a thin wire coated with a fine layer of wax to elevate the brain and stabilize it11 during electrophysiology as shown in Figure 4D.</li>
 <li>The insect brain is covered by a thin insulating sheath that needs to be removed prior to experiments. To desheath the brain, gently spread a small amount of an enzyme (0.3-0.4 mg protease, Sigma Aldrich) over the surface of the brain using fine forceps9. After ~5-10 s of enzyme application, rinse the brain thoroughly with saline. Using super-fine forceps very gently pinch and pull the sheath up and subsequently tear it open over the recording locations (AL and MB; shown in Figure 4E, F).</li>
</ol>
5. Multi-unit Recordings from the Antennal Lobe and the Mushroom Body
<ol>
 <li>Place the locust preparation (Figure 5A) under a stereomicroscope suspended from a boom stand placed on a vibration isolation table.</li>
 <li>Maintain a constant saline perfusion rate (about 0.04 L/hr) throughout the experiment. Use a chlorided silver wire immersed in the saline-filled wax cup as the ground electrode.</li>
 <li>For PN recordings, use a 16-channel silicon probe (NeuroNexus Technologies, item #A2x2-tet-3mm-150-150-121-A16, Figure 5B). Prior to each experiment, electroplate the electrodes with gold to achieve impedances in the 200-300 k&Omega; range. Use the circuit shown in Figure 7 for electroplating.</li>
 <li>Position the electrode close to the surface of the antennal lobe and gently insert it into the tissue using a manual micromanipulator (WPI, M3301R) (Figure 5D).</li>
 <li>Advance the electrode in ~10 &mu;m steps. Wait 2-3 min at each step and evaluate the acquired signal quality. In an ideal recording site, extracellular signals will be picked up by multiple recording channels and will have a high signal-to-noise ratio (SNR &gt; 3-5 times noise SDs).</li>
 <li>For KC recordings, use a custom-made twisted wire tetrode (Figure 5C; step-by-step fabrication procedure presented in Section 6). Electroplate these electrodes as discussed in step 5.3. Place the tetrode on the surface of the MB (Figure 5E) as KC somata are restricted to the superficial layer of the MB8.</li>
 <li>Both PN and KC recordings can be made simultaneously from same locust preparation as schematically shown in Figure 5A.</li>
 <li>Wait at least 15 min after finding the recording location to allow stabilization of the electrodes.</li>
 <li>Acquire all extracellular signals at 15 KHz, filter between 0.3-6 KHz, and amplify (10,000 times) using a 16 channel AC amplifier (Biology Electronics Shop; Caltech, Pasadena, CA) (Figure 6A, B).</li>
</ol>
6. Procedures to Make Twisted Wire Electrode for KC Recordings
<ol>
 <li>To design a multi-unit electrode for KC recordings, use insulated nickel chromium wire (RO800, 0.0005" filament)8.</li>
 <li>If eight electrodes are desired then wrap the wire around a 10-15 cm long piece of cardboard 4 times. The ends of the cardboard can be covered with plastic tubing to prevent the edges from cutting the wire. While wrapping, make sure there is little slack, but ensure that the wire is not taut. Handle the wire carefully as it breaks easily.</li>
 <li>Bunch the wires together at the top of the cardboard and use a piece of tape (Time Tape, T-534-RP) to hold them together. Cut the strands at the other end. Remove the wire strands and group the wires at the cut end as well using another piece of the tape.</li>
 <li>Using a titer plate shaker (Thermo Scientific, Model 4625Q), wind the strands together (~72 rev/min for 3 min) to form one twisted wire. The bottom of the taped strands can be clipped to the titer plate rotor and the top can be clipped to a boom stand. During winding, the wire should have little slack, yet not be taut.</li>
 <li>Melt the insulation together with a heat gun (Weller 6966C) to stick the individual strands together and form a single wire. 3-4 slow passes (3-4 sec each) over the length of the wire should be sufficient to heat and fuse the strands together. Then release the wire from the bottom and allow it to relax. Trim the wire near the ends to remove the tape.</li>
 <li>Insert one end of the wire through a 5-6 cm long, glass capillary tube (O.D. 1.0 mm, I.D. 0.58 mm). Tease apart the twisted wire at one end using fine tweezers and remove the coating by very briefly torching the end using a flame. This step must be done carefully; exposing the wire to the flame for too long will cause the strands to melt and tangle.</li>
 <li>Gently separate the 8 strands on the flamed end and solder each strand separately into an 8-pin IC socket. Coat the top of the IC socket with epoxy to hold the strands and glass capillary in place. Also place a small drop of epoxy at the other end of the capillary to hold the wire in place (Figure 5C).</li>
 <li>Finally, cut the tip of the wire at a 45 degree angle with carbide scissors about 0.5 cm from the capillary tip.</li>
</ol>

<ol>
	<li>Ache, B. W., Young, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Olfaction:+diverse+species,+conserved+principles.">Olfaction: diverse species, conserved principles.</a> Neuron. 48, 417-430 (2005).</li><li>Laurent, G., Wehr, M., Davidowitz, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+representations+of+odors+in+an+olfactory+network.">Temporal representations of odors in an olfactory network.</a> Journal of Neuroscience. 16, 3837-3847 (1996).</li><li>Stopfer, M., Jayaraman, V., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odor+identity+vs.+intensity+coding+in+an+olfactory+system.">Odor identity vs. intensity coding in an olfactory system.</a> Neuron. 39, 991-1004 (2003).</li><li>Steven de Belle, J., Heisenberg, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Associative+odor+learning+in+Drosophila+abolished+by+chemical+ablation+of+mushroom+bodies.">Associative odor learning in Drosophila abolished by chemical ablation of mushroom bodies.</a> Science. 263, 692-695 (1994).</li><li>Cassenaer, S., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Conditional+modulation+of+spike-timing-dependent+plasticity+for+olfactory+learning.">Conditional modulation of spike-timing-dependent plasticity for olfactory learning.</a> Nature. 482, 47-52 (2012).</li><li>Hallem, E. A., Carlson, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coding+of+odors+by+a+receptor+repertoire.">Coding of odors by a receptor repertoire.</a> Cell. 125, 143-160 (2006).</li><li>Raman, B., Joseph, J., Tang, J., Stopfer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporally+diverse+firing+patterns+in+olfactory+receptor+neurons+underlie+spatiotemporal+neural+codes+for+odors.">Temporally diverse firing patterns in olfactory receptor neurons underlie spatiotemporal neural codes for odors.</a> Journal of Neuroscience. 30, 1994-2006 (2010).</li><li>Perez-Orive, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oscillations+and+sparsening+of+odor+representations+in+the+mushroom+body.">Oscillations and sparsening of odor representations in the mushroom body.</a> Science. 297, 359-365 (2002).</li><li>Naraghi, M., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odorant-induced+oscillations+in+the+mushroom+bodies+of+the+locust.">Odorant-induced oscillations in the mushroom bodies of the locust.</a> The Journal of Neuroscience. 14, 2993-3004 (1994).</li><li>Ochieng, S. A., Hallberg, E., Hansson, B. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fine+structure+and+distribution+of+antennal+sensilla+of+the+desert+locust,+Schistocerca+gregaria+(Orthoptera:+Acrididae).">Fine structure and distribution of antennal sensilla of the desert locust, Schistocerca gregaria (Orthoptera: Acrididae).</a> Cell and Tissue Research. 291, 525-536 (1998).</li><li>Burrows, M., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Synaptic+Potentials+in+the+Central+Terminals+of+Locust+Proprioceptive+Afferents+Generated+by+Other+Afferents+from+the+Same+Sense+Organ.">Synaptic Potentials in the Central Terminals of Locust Proprioceptive Afferents Generated by Other Afferents from the Same Sense Organ.</a> Journal of Neuroscience. 13, 808-819 (1993).</li><li>Pouzat, C., Mazor, O., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Using+noise+signature+to+optimize+spike-sorting+and+to+assess+neuronal+classification+quality.">Using noise signature to optimize spike-sorting and to assess neuronal classification quality.</a> Journal of Neuroscience Methods. 122, 43-57 (2002).</li><li>Mazor, O., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transient+dynamics+versus+fixed+points+in+odor+representations+by+locust+antennal+lobe+projection+neurons.">Transient dynamics versus fixed points in odor representations by locust antennal lobe projection neurons.</a> Neuron. 48, 661-673 (2005).</li><li>Christensen, T. A., Pawlowski, V. A., Lei, H., Hildebrand, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multi-unit+recordings+reveal+context+dependent+modulation+of+synchrony+in+odor-specific+neural+ensembles.">Multi-unit recordings reveal context dependent modulation of synchrony in odor-specific neural ensembles.</a> Nature Neuroscience. 3, 927-931 (2000).</li><li>Pellegrino, M., Nakagawa, T., Vosshall, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+Sensillum+Recordings+in+the+Insects+Drosophila+melanogaster+and+Anopheles+gambiae.">Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae.</a> J. Vis. Exp. (36), e1725 (2010).</li><li>Geffen, M. N., Broome, B. M., Laurent, G., Meister, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+Encoding+of+Rapidly+Fluctuating+Odors.">Neural Encoding of Rapidly Fluctuating Odors.</a> Neuron. 61, 570-586 (2009).</li><li>Ito, I., Ong, R. C., Raman, B., Stopfer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sparse+odor+representation+and+olfactory+learning.">Sparse odor representation and olfactory learning.</a> Nature Neuroscience. 11, 1177-1184 (2008).</li><li>Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Olfactory+network+dynamics+and+the+coding+of+multidimensional+signals.">Olfactory network dynamics and the coding of multidimensional signals.</a> Nature Review Neuroscience. 3, 884-895 (2002).</li><li>Brown, S. L., Joseph, J., Stopfer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Encoding+a+temporally+structured+stimulus+with+a+temporally+structured+neural+representation.">Encoding a temporally structured stimulus with a temporally structured neural representation.</a> Nature Neuroscience. 8, 1568-1576 (2005).</li><li>MacLeod, K., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+mechanism+for+synchronization+and+temporal+patterning+of+odor-encoding+neural+assemblies.">Distinct mechanism for synchronization and temporal patterning of odor-encoding neural assemblies.</a> Science. 274, 976-979 (1996).</li><li>Wehr, M., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relationship+between+afferent+and+central+temporal+patterns+in+the+locust+olfactory+system.">Relationship between afferent and central temporal patterns in the locust olfactory system.</a> The Journal of Neuroscience. 19, 381-390 (1999).</li><li>Moreaux, L., Laurent, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimating+firing+rates+from+calcium+signals+in+locust+projection+neurons+in+vivo.">Estimating firing rates from calcium signals in locust projection neurons in vivo.</a> Frontiers in Neural Circuits. 1, 1-13 (2007).</li><li>Galizia, C. G., Joerges, J., Kuttner, A., Faber, T., Menzel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+semi-in-vivo+preparation+for+optical+recording+of+the+insect+brain.">A semi-in-vivo preparation for optical recording of the insect brain.</a> Journal of Neuroscience Methods. 76, 61-69 (1997).</li><li>Galan, R. F., Sachse, S., Galizia, C. G., Hez, A. V. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Odor-driven+attractor+dynamics+in+the+antennal+lobe+allow+for+simple+and+rapid+olfactory+pattern+classification.">Odor-driven attractor dynamics in the antennal lobe allow for simple and rapid olfactory pattern classification.</a> Neural Computation. 16, 999-1012 (2004).</li><li>Kuebler, L. S., Schubert, M., Karpati, Z., Hansson, B. S., Olsson, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antennal+Lobe+Processing+Correlates+to+Moth+Olfactory+Behavior.">Antennal Lobe Processing Correlates to Moth Olfactory Behavior.</a> Journal of Neuroscience. 32, 5772-5782 (2012).</li><li>Silbering, A. F., Bell, R., Galizia, C. G., Benton, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+Imaging+of+Odor-evoked+Responses+in+the+Drosophila+Antennal+Lobe.">Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe.</a> J. Vis. Exp. (61), e2976 (2012).</li><li>Skiri, H. T., Galizia, C. G., Mustaparta, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Representation+of+Primary+Plant+Odorants+in+the+Antennal+Lobe+of+the+Moth+Heliothis+virescens+Using+Calcium+Imaging.">Representation of Primary Plant Odorants in the Antennal Lobe of the Moth Heliothis virescens Using Calcium Imaging.</a> Chemical Senses. 29, 253-267 (2004).</li></ol>

No conflicts of interest declared.

Most sensory stimuli evoke combinatorial responses that are distributed across ensembles of neurons. Hence, simultaneous monitoring of multi-neuron activity is necessary to understand how stimulus-specific information is represented and processed by neural circuits in the brain. Here, we have demonstrated extracellular multi-unit recording techniques to characterize odor-evoked responses at the first three processing centers along the insect olfactory pathway. We note that the techniques presented here have been used in ...

<table border="1">
	<tbody>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Electrophysiology Equipment</td>
		</tr>
		<tr>
			<td>A.C. amplifier</td>
			<td>GRASS</td>
			<td>Model P55</td>
			<td>for single sensillum recordings</td>
		</tr>
		<tr>
			<td>Audio monitor (model 3300)</td>
			<td>A-M Systems</td>
			<td>940000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Custom-made 16 channel pre-amplifier and amplifier</td>
			<td>Cal. Tech. Biology Electronics Shop</td>
			<td>&nbsp;</td>
			<td>for AL and MB recordings</td>
		</tr>
		<tr>
			<td>Data acquisition unit</td>
			<td>National Instruments</td>
			<td>BNC-2090</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Fiber optic light</td>
			<td>WPI</td>
			<td>SI-72-8</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Light source 115 V</td>
			<td>WPI</td>
			<td>NOVA</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Manual micromanipulator</td>
			<td>WPI</td>
			<td>M3301R</td>
			<td>for locust brain recordings</td>
		</tr>
		<tr>
			<td>Stereomicroscope1 on boom stand</td>
			<td>Leica</td>
			<td>M80</td>
			<td>for locust brain recordings</td>
		</tr>
		<tr>
			<td>Stereomicroscope2</td>
			<td>Leica</td>
			<td>M205C</td>
			<td>for single sensillum recordings</td>
		</tr>
		<tr>
			<td>Vibration-isolation table</td>
			<td>TMC</td>
			<td>63-500 series</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Motorized micromanipulator</td>
			<td>Sutter Instruments</td>
			<td>MP285/T</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Oscilloscope</td>
			<td>Tektronix</td>
			<td>TD2014B</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Electrodes/Construction Tools</td>
		</tr>
		<tr>
			<td>16-channel electrode</td>
			<td>NeuroNexus</td>
			<td>A2x2-tet-3mm-150-121</td>
			<td>for antennal lobe recordings</td>
		</tr>
		<tr>
			<td>Borosilicate capillary tubes with filament, ID 0.69 mm</td>
			<td>Sutter Instruments</td>
			<td>BF120-69-10</td>
			<td>for making glass electrodes</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instruments</td>
			<td>P-1000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Function generator</td>
			<td>Multimeter Warehouse</td>
			<td>SG1639A</td>
			<td>for gold-plating electrodes</td>
		</tr>
		<tr>
			<td>Gold plating solution (non cyanide)</td>
			<td>SIFCO Industries</td>
			<td>NC SPS 5355</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Impedance tester</td>
			<td>BAK Electronics Inc.</td>
			<td>IMP-2</td>
			<td>for gold-plating electrodes</td>
		</tr>
		<tr>
			<td>Switch rotary</td>
			<td>Electroswitch</td>
			<td>C7D0123N</td>
			<td>for gold-plating electrodes</td>
		</tr>
		<tr>
			<td>Pulse isolator</td>
			<td>WPI</td>
			<td>A365</td>
			<td>for gold-plating electrodes</td>
		</tr>
		<tr>
			<td>Q series electrode holder</td>
			<td>Warner Instruments</td>
			<td>64-1091</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Silver wire 0.010" diameter</td>
			<td>A-M Systems</td>
			<td>782500</td>
			<td>ground electrode</td>
		</tr>
		<tr>
			<td>8 pin DIP IC socket</td>
			<td>Digikey</td>
			<td>ED90032-ND</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Borosilicate capillary tubes with filament, ID 0.58 mm</td>
			<td>Warner Instruments</td>
			<td>64-0787</td>
			<td>twisted wire tetrode construction</td>
		</tr>
		<tr>
			<td>Heat gun</td>
			<td>Weller</td>
			<td>6966C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Rediohm-800 wire</td>
			<td>Kanthal Precision Technologies</td>
			<td>PF002005</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Titer plate shaker</td>
			<td>Thermo Scientific</td>
			<td>4625Q</td>
			<td>twisting wires</td>
		</tr>
		<tr>
			<td>Carbide scissors, 4.5"</td>
			<td>Biomedical Research Instr</td>
			<td>25-1000</td>
			<td>for cutting twisted tetrode wires</td>
		</tr>
		<tr>
			<td>Fine point tweezers</td>
			<td>HECO</td>
			<td>91-EF5-SA</td>
			<td>for teasing tetrode wires apart</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Odor Delivery</td>
		</tr>
		<tr>
			<td>6 ml syringe</td>
			<td>Kendall</td>
			<td>1180600777</td>
			<td>for custom designed activated carbon filter</td>
		</tr>
		<tr>
			<td>Brown odor bottles</td>
			<td>Fisher</td>
			<td>08-912-165</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Charcoal</td>
			<td><a href="http://BuyActivatedCharcoal.com" target="_blank">BuyActivatedCharcoal.com</a></td>
			<td>GAC-48C</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Desiccant</td>
			<td>Drierite</td>
			<td>23005</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Drierite gas drying jar</td>
			<td>Fischer Scientific</td>
			<td>09-204</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Heat shrink tubing</td>
			<td>3M</td>
			<td>EPS-200</td>
			<td>odor filter preparation</td>
		</tr>
		<tr>
			<td>Hypodermic needle aluminum hub, gauge 19</td>
			<td>Kendall</td>
			<td>8881-200136</td>
			<td>for providing inlet and outlet lines for odor bottles</td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Mallinckrodt Chemicals</td>
			<td>6357-04</td>
			<td>for odor dilution</td>
		</tr>
		<tr>
			<td>Nalgene plastic tubing, 890 FEP</td>
			<td>Thermo Scientific</td>
			<td>8050-0310</td>
			<td>for carrier gas delivery</td>
		</tr>
		<tr>
			<td>Pneumatic picopump</td>
			<td>WPI</td>
			<td>sys-pv820</td>
			<td>for odor delivery</td>
		</tr>
		<tr>
			<td>Polyethylene tubing ID 0.86 mm</td>
			<td>Intramedic</td>
			<td>427421</td>
			<td>for odor bottle outlet connections and saline profusion tubing</td>
		</tr>
		<tr>
			<td>Stoppers</td>
			<td>Lab Pure</td>
			<td>97041</td>
			<td>for sealing odor bottles</td>
		</tr>
		<tr>
			<td>Time tape</td>
			<td>PDC</td>
			<td>T-534-RP</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Tubing luer</td>
			<td>Cole-Parmer</td>
			<td>30600-66</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Vacuum tube</td>
			<td>McMaster-Carr</td>
			<td>5488K66</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Preparation/Dissection</td>
		</tr>
		<tr>
			<td>100 x 15 mm petri dish</td>
			<td>VWR International</td>
			<td>89000-304</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>18 AWG copper stranded wire</td>
			<td>Lapp Kabel</td>
			<td>4510013</td>
			<td>wire insulation is used as rubber gaskets</td>
		</tr>
		<tr>
			<td>22 AWG stranded hookup wire</td>
			<td>AlphaWire</td>
			<td>1551</td>
			<td>brain platform</td>
		</tr>
		<tr>
			<td>Batik wax</td>
			<td>Jacquard</td>
			<td>7946000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dental periphery Wax</td>
			<td>Henry-Schein Dental</td>
			<td>6652151</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Electrowaxer</td>
			<td>Almore International</td>
			<td>66000</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Epoxy, 5 min</td>
			<td>Permatex</td>
			<td>84101</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hypodermic needle aluminum hub</td>
			<td>Kendall</td>
			<td>8881-200136</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Protease from Streptomyces griseus</td>
			<td>Sigma-Aldrich</td>
			<td>P5147</td>
			<td>for desheathing locust brain</td>
		</tr>
		<tr>
			<td>Suture thread non-sterile</td>
			<td>Fisher</td>
			<td>NC9087024</td>
			<td>for tying the abdomen after gut removal</td>
		</tr>
		<tr>
			<td>Vetbond</td>
			<td>3M</td>
			<td>1469SB</td>
			<td>for sealing amputation sites</td>
		</tr>
		<tr>
			<td>Dumont #1 forceps (coarse)</td>
			<td>WPI</td>
			<td>500335</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dumont #5 titanium forceps (fine)</td>
			<td>WPI</td>
			<td>14096</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dumont #5SF forceps (super-fine)</td>
			<td>WPI</td>
			<td>500085</td>
			<td>desheathing locust brain</td>
		</tr>
		<tr>
			<td>10 cm dissecting scissors</td>
			<td>WPI</td>
			<td>14393</td>
			<td>for removing legs and wings</td>
		</tr>
		<tr>
			<td>Vannas scissors (fine)</td>
			<td>WPI</td>
			<td>500086</td>
			<td>for removing cuticle, cutting the foregut</td>
		</tr>
		<tr>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Saline Profusion</td>
		</tr>
		<tr>
			<td>Extension set with rate flow regulator</td>
			<td>Moore Medical</td>
			<td>69136</td>
			<td>for regulating saline flow</td>
		</tr>
		<tr>
			<td>IV administration set with Y injection site</td>
			<td>Moore Medical</td>
			<td>73190</td>
			<td>for regulating saline flow</td>
		</tr>
	</tbody>
</table>

multi-unit recording methods to characterize neural activity in the locust (schistocerca americana) olfactory circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Odor-evoked responses of a single ORN to two different alcohols are shown in the Figure 3D. Depending on the recording location (sensilla type, placement of the electrode) multi-unit recordings can be achieved.
A raw extracellular waveform from an AL recording is shown in Figure 6A. Action potentials or spikes of varying amplitudes originating from different PNs can be observed in this voltage trace. Although the locust antennal lobe has excitatory projection ...

Watch this Scientific Journal Video about Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits at JoVE.com

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

multi-unit-recording-methods-to-characterize-neural-activity-locust

Research

JoVE Journal

Neuroscience

26.9K Views.  Washington University in St. Louis. We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Video: Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

Surgical Implantation of Chronic Neural Electrodes for Recording Single Unit Activity and Electrocorticographic Signals

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery. Here we provide useful information for surgeons who are learning the process of implanting electrode systems. We demonstrate the implantation procedure of both a penetrating and a surface electrode. The surgical process is described from start to finish, including detailed descriptions of each step throughout the procedure. It should also be noted that this video guide is focused towards procedures conducted in rodent models and other small animal models. Modifications of the described procedures are feasible for other animal models.

We provide useful information for surgeons who are learning the process of implanting chronic neural recording electrodes. Techniques for both penetrating and surface electrode systems are described in a rodent animal model.

The success of long-term electrophysiological recordings often depends on the quality of the implantation surgery.  Here we provide useful information for ...

Optical Recording of Suprathreshold Neural Activity with Single-cell and Single-spike Resolution

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also propagation of suprathreshold spiking activity involves populations of neurons. Empirical studies addressing cortical function directly thus require recordings from populations of neurons with high resolution. Here we describe an optical method and a deconvolution algorithm to record neural activity from up to 100 neurons with single-cell and single-spike resolution. This method relies on detection of the transient increases in intracellular somatic calcium concentration associated with suprathreshold electrical spikes (action potentials) in cortical neurons. High temporal resolution of the optical recordings is achieved by a fast random-access scanning technique using acousto-optical deflectors (AODs)1. Two-photon excitation of the calcium-sensitive dye results in high spatial resolution in opaque brain tissue2. Reconstruction of spikes from the fluorescence calcium recordings is achieved by a maximum-likelihood method. Simultaneous electrophysiological and optical recordings indicate that our method reliably detects spikes (&gt;97% spike detection efficiency), has a low rate of false positive spike detection (&lt; 0.003 spikes/sec), and a high temporal precision (about 3 msec) 3. This optical method of spike detection can be used to record neural activity in vitro and in anesthetized animals in vivo3,4.

Understanding the function of the vertebrate central nervous system requires recordings from many neurons because cortical function arises on the level of populations of neurons. Here we describe an optical method to record suprathreshold neural activity with single-cell and single-spike resolution, dithered random-access scanning. This method records somatic fluorescence calcium signals from up to 100 neurons with high temporal resolution. A maximum-likelihood algorithm deconvolves the underlying suprathreshold neural activity from the somatic fluorescence calcium signals. This method reliably detects spikes with high detection efficiency and a low rate of false positives and can be used to study neural populations in vitro and in vivo.

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also ...

Identification of Olfactory Volatiles using Gas Chromatography-Multi-unit Recordings (GCMR) in the Insect Antennal Lobe

All organisms inhabit a world full of sensory stimuli that determine their behavioral and physiological response to their environment. Olfaction is especially important in insects, which use their olfactory systems to respond to, and discriminate amongst, complex odor stimuli. These odors elicit behaviors that mediate processes such as reproduction and habitat selection1-3. Additionally, chemical sensing by insects mediates behaviors that are highly significant for agriculture and human health, including pollination4-6, herbivory of food crops7, and transmission of disease8,9. Identification of olfactory signals and their role in insect behavior is thus important for understanding both ecological processes and human food resources and well-being.
	To date, the identification of volatiles that drive insect behavior has been difficult and often tedious. Current techniques include gas chromatography-coupled electroantennogram recording (GC-EAG), and gas chromatography-coupled single sensillum recordings (GC-SSR)10-12. These techniques proved to be vital in the identification of bioactive compounds. We have developed a method that uses gas chromatography coupled to multi-channel electrophysiological recordings (termed 'GCMR') from neurons in the antennal lobe (AL; the insect's primary olfactory center)13,14. This state-of-the-art technique allows us to probe how odor information is represented in the insect brain. Moreover, because neural responses to odors at this level of olfactory processing are highly sensitive owing to the degree of convergence of the antenna's receptor neurons into AL neurons, AL recordings will allow the detection of active constituents of natural odors efficiently and with high sensitivity. Here we describe GCMR and give an example of its use.
	Several general steps are involved in the detection of bioactive volatiles and insect response. Volatiles first need to be collected from sources of interest (in this example we use flowers from the genus Mimulus (Phyrmaceae)) and characterized as needed using standard GC-MS techniques14-16. Insects are prepared for study using minimal dissection, after which a recording electrode is inserted into the antennal lobe and multi-channel neural recording begins. Post-processing of the neural data then reveals which particular odorants cause significant neural responses by the insect nervous system.
	Although the example we present here is specific to pollination studies, GCMR can be expanded to a wide range of study organisms and volatile sources. For instance, this method can be used in the identification of odorants attracting or repelling vector insects and crop pests. Moreover, GCMR can also be used to identify attractants for beneficial insects, such as pollinators. The technique may be expanded to non-insect subjects as well.

Identification of Olfactory Volatiles using Gas Chromatography-Multi-unit Recordings GCMR in the Insect Antennal Lobe

Olfactory cues mediate many different behaviors in insects, and are often complex mixtures comprised of tens to hundreds of volatile compounds. Using gas chromatography with multi-channel recording in the insect antennal lobe, we describe a method for the identification of bioactive compounds.

All  organisms inhabit a world full of sensory stimuli that determine their  behavioral and physiological response to their environment. Olfaction is ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

Using Single Sensillum Recording to Detect Olfactory Neuron Responses of Bed Bugs to Semiochemicals

The insect olfactory system plays an important role in detecting semiochemicals in the environment. In particular, the antennal sensilla which house single or multiple neurons inside, are considered to make the major contribution in responding to the chemical stimuli. By directly recording action potential in the olfactory sensillum after exposure to stimuli, single sensillum recording (SSR) technique provides a powerful approach for investigating the neural responses of insects to chemical stimuli. For the bed bug, which is a notorious human parasite, multiple types of olfactory sensillum have been characterized. In this study, we demonstrated neural responses of bed bug olfactory sensilla to two chemical stimuli and the dose-dependent responses to one of them using the SSR method. This approach enables researchers to conduct early screening for individual chemical stimuli on the bed bug olfactory sensilla, which would provide valuable information for the development of new bed bug attractants or repellents and benefits the bed bug control efforts.

Bed bugs rely on olfactory receptor neurons housed in their antennal olfactory sensilla to detect semiochemicals in the environment. Utilizing single sensillum recording, we demonstrate a method to evaluate bed bug response to semiochemicals and explore the coding process involved.

The insect olfactory system plays an important role in detecting semiochemicals in the environment. In particular, the antennal sensilla which house ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. However, there is piling evidence that olfaction is not solely limited to chemical sensitivity, but also includes temperature sensitivity. Premetamorphic Xenopus laevis are translucent animals, with protruding nasal cavities deprived of the cribriform plate separating the nose and the olfactory bulb. These characteristics make them well suited for studying olfaction, and particularly thermosensitivity. The present article describes the complete procedure for measuring temperature responses in the olfactory bulb of X. laevis larvae. Firstly, the electroporation of olfactory receptor neurons (ORNs) is performed with spectrally distinct dyes loaded into the nasal cavities in order to stain their axon terminals in the bulbar neuropil. The differential staining between left and right receptor neurons serves to identify the &#947;-glomerulus as the only structure innervated by contralateral presynaptic afferents. Secondly, the electroporation is combined with focal bolus loading in the olfactory bulb in order to stain mitral cells and their dendrites. The 3D brain volume is then scanned under line-illumination microscopy for the acquisition of fast calcium imaging data while small temperature drops are induced at the olfactory epithelium. Lastly, the post-acquisition analysis allows the morphological reconstruction of the thermosensitive network comprising the &#947;-glomerulus and its innervating mitral cells, based on specific temperature-induced Ca2+ traces. Using chemical odorants as stimuli in addition to temperature jumps enables the comparison between thermosensitive and chemosensitive networks in the olfactory bulb.

Recording Temperature-induced Neuronal Activity through Monitoring Calcium Changes in the Olfactory Bulb of Xenopus laevis

Here we describe a protocol for measuring and analyzing temperature responses in the olfactory bulb of Xenopus laevis. Olfactory receptor neurons and mitral cells are differentially stained, after which calcium changes are recorded, reflecting a sensitivity of some neural networks in the bulb to temperature drops induced at the nose.

The olfactory system, specialized in the detection, integration and processing of chemical molecules is likely the most thoroughly studied sensory system. ...

Quadruple Immunostaining of the Olfactory Bulb for Visualization of Olfactory Sensory Axon Molecular Identity Codes

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory Sensory Neuron (OSN) is a bipolar cell with a single dendrite and a single unbranched axon. An OSN expresses only one Olfactory Receptor (OR) gene, OSNs expressing a given type of OR converge their axons to a few sets of invariant glomeruli in the Olfactory Bulb (OB). A remarkable feature of OSN projection is that the expressed ORs play instructive roles in axonal projection. ORs regulate the expression of multiple axon-sorting molecules and generate the combinatorial molecular code of axon-sorting molecules at the OSN axon termini. Thus, to understand the molecular mechanisms of OR-specific axon guidance mechanisms, it is vital to characterize their expression profiles at the OSN axon termini within the same glomerulus. The aim of this article was to introduce methods for collecting as many glomeruli as possible on a single OB section and for performing immunostaining using multiple antibodies. This would allow the comparison and analysis of the expression patterns of axon-sorting molecules without staining variation between OB sections.

Olfactory sensory neurons express a wide variety of axon-sorting molecules to establish proper neural circuitry. This protocol describes an immunohistochemical staining method to visualize combinatorial expressions of axon-sorting molecules at the axon termini of olfactory sensory neurons.

The mouse olfactory system is often used to study mechanisms of neural circuit formation&#12288;because of its simple anatomical structure. An Olfactory ...

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the pathophysiological basis of these diseases, it is necessary to precisely characterize in which way the processing of information is disturbed between the different neuronal parts of the circuit. Using extracellular in vivo electrophysiological recordings, it is possible to accurately delineate neuronal activity within a neuronal network. The application of this method has several advantages over alternative techniques, e.g., functional magnetic resonance imaging and calcium imaging, as it allows a unique temporal and spatial resolution and does not rely on genetically engineered organisms. However, the use of extracellular in vivo recordings is limited since it is an invasive technique that cannot be universally applied. In this article, a simple and easy to use method is presented with which it is possible to simultaneously record extracellular potentials such as local field potentials and multiunit activity at multiple sites of a network. It is detailed how a precise targeting of subcortical nuclei can be achieved using a combination of stereotactic surgery and online analysis of multi-unit recordings. Thus, it is demonstrated, how a complete network such as the hyperdirect cortico-basal ganglia loop can be studied in anesthetized animals in vivo.

Acute In Vivo Electrophysiological Recordings of Local Field Potentials and Multi-unit Activity from the Hyperdirect Pathway in Anesthetized Rats

In this study, the methodology is presented on how to perform multi-site in vivo electrophysiological recordings from the hyperdirect pathway under urethane anesthesia.

Converging evidence shows that many neuropsychiatric diseases should be understood as disorders of large-scale neuronal networks. To better understand the ...

Combining Optogenetics with Artificial microRNAs to Characterize the Effects of Gene Knockdown on Presynaptic Function within Intact Neuronal Circuits

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a workflow on how to combine artificial microRNA (miR)-mediated RNA interference with optogenetics to achieve selective stimulation of manipulated presynaptic boutons in acute brain slices. The experimental approach involves the use of a single viral construct and a single neuron-specific promoter to drive the expression of both an optogenetic probe and artificial miR(s) against presynaptic gene(s). When stereotactically injected in the brain region of interest, the expressed construct makes it possible to stimulate with light exclusively the neurons with reduced expression of the gene(s) under investigation. This strategy does not require the development and maintenance of genetically modified mouse lines and can in principle be applied to other organisms and to any neuronal gene of choice. We have recently applied it to investigate how the knockdown of alternative splice isoforms of presynaptic P/Q-type voltage-gated calcium channels (VGCCs) regulates short-term synaptic plasticity at CA3 to CA1 excitatory synapses in acute hippocampal slices. A similar approach could also be used to manipulate and probe the neuronal circuitry in vivo.

This protocol provides a workflow on how to combine artificial microRNA-mediated RNA interference with optogenetics to stimulate specifically presynaptic boutons with reduced expression of selective gene(s) within intact neuronal circuits.

The purpose of this protocol is to characterize the effect of gene knockdown on presynaptic function within intact neuronal circuits. We describe a ...