This work was supported by NSF grant IOS 1121692, and by the University of Washington's Research Foundation.

1. Volatile Follection
 <ol>
 <li>In this example, we use volatile samples from M. lewisii flowers - an alpine wildflower native to California. Volatiles are collected using dynamic sorption methods according to Riffell et al.14. Briefly, this method employs a closed loop trapping system where flowers are enclosed in a Teflon bag. Using an inert vacuum pump, the air around the flowers is sucked through a "trap" comprised of a Pasteur pipette filled with Porapak Q matrix. The return air from the pump is filtered by activated charcoal. After a prescribed time period, in our case 24 h, the Porapak Q matrix is eluted with a non-polar solvent, typically hexane, to collect the concentrated extract. The extract is then stored in -80 &deg;C until analysis. If necessary, samples can be concentrated prior to analysis under a stream of nitrogen gas. Unless the sample is already well-characterized, run an aliquot of it through a gas chromatograph-mass spectrometer (GC-MS) to identify volatile components prior to using the sample.</li>
 </ol>
 2. Electrophysiological Preparation
 <ol>
 <li>Cut approximately 1 cm from the end of a 1,000 &mu;l pipette tip. Place a bumble bee (Bombus impatiens) into the base of the pipette tip and gently push towards the opposite end of the tip until only the head is exposed.</li>
 <li>Melt dental wax and mold it around the exposed head, making sure that the wax adheres onto the compound eyes for security and to make the bee's head completely immobile. Be sure not to get any wax onto the antenna of the bee.</li>
 <li>Once the head is secure, make a square, window-like, incision into the head capsule using a razor blade-breaker or the appropriate sized scalpel to cut the cuticle. Using the blade-breaker, start from the dorsal side of the head capsule, immediately behind the antenna and adjacent to one of the compound eyes. Cut a straight line, from one compound eye to the contralateral compound eye. After cutting a straight line to the opposite compound eye, begin making an incision dorsally until the head capsule curves and ends near its thorax. At this point, begin to cut towards the opposite end of the head capsule. Finally, once a line has been cut to the opposite end, cut a line back up to the starting position of the initial incision. It is important to remove the cuticle that is adjacent to the antenna, as this will impede the insertion of the electrode.</li>
 <li>Once the cuticle is cut, use a pair of forceps to remove the bee cuticle, which should then expose the bee's brain and, more importantly, the antennal lobes. Immediately begin superfusing the brain with insect saline, so that the brain does not become dessicated. After the brain is exposed, carefully use a pair of very fine forceps to remove the perineural sheath immediately above the antennal lobes. Be very careful not to puncture the bee's brain with the forceps.</li>
 </ol>
 3. Gas Chromatography with Multi-channel Recording
 <ol>
 <li>The bee "preparation" - fixed in a tube with its brain exposed - is now ready for electrophysiological recording. Place the bee in a clamp, fixed to a magnetic base that is located on an air table.</li>
 <li>Arrange an IV bag, flow controller, and tubing (filled with insect saline) so that the saline continuously superfuses the brain.</li>
 <li>Using a micromanipulator, insertareference electrode, made of tungsten wire, into the bee's eye.</li>
 <li>Using a separate micromanipulator, insert the multi-channel electrode, such as a coiled wire tetrode, or silicon multi-channel electrode (Neuronexus Technologies), into the antennal lobes of the bee. This electrode is connected to a pre-amplifier such as the TDT system's S-3 Z series to the Z-bus bioamp processor of the TDT system. Output from the Gas Chromatogram's detector via a shielded BNC cable can be interfaced with the amplifier and data acquisition system such that both the neural and GC signal is synchronized.</li>
 <li>Wait approximately 30-60 minutes for the neural recordings to stabilize. Once the spontaneous activity and waveform shape of units in the recording channels have become consistent, use an odor syringe to stimulate the bee and observe the response of the recording channels to the odor.</li>
 <li>Record extracellular spikes from neurons by auto-thresholding the recording channels by 3.5 to 5 sigma of the signal on the individual recording channels. Manual thresholding may be required for some channels to avoid contamination from electrical noise. The action potentials from neurons will appear as voltage spikes in the recording channel. When the channel's voltage exceeds the threshold, the system buffers and saves the few milliseconds before and after the crossing of the threshold, thereby taking a snap-shot of the waveform, or spike.</li>
 <li>Immediately next to the air table is the GC. Before injecting the floral extract into the GC, make sure that the method for the temperature ramp of the GC run is correct. In our example, we use a temperature method starting at 50 &deg;C for 4 min followed by an increase of temperature at a rate of 10 &deg;C/min. to 220 &deg;C, at which time we hold the GC for an additional 6 min. We use a DB-5 GC column (J&amp;W Scientific, Folsom, CA, USA), with helium as the carrier gas. The inlet is splitless, with a temperature set to 200 &deg;C. The flame ionization detector temperature is set to 230 &deg;C.</li>
 <li>Inject the extract sample of the floral headspace into the heated injection port of the GC to release the adsorbed volatiles into the GC column. The effluent from the column is split 1:1 between the Flame Ionization Detector (FID) and the bee antenna using a glass "Y" connector (J&amp;W Scientific). Begin recording from the electrode as you inject a sample into the GC.</li>
 <li>After the GC run has finished, let the preparation rest for 5 to 15 min. Then either inject another sample into the GC or stimulate the preparation using single volatile compounds or mixtures of compounds. In this latter method of stimulating the preparation, pulses of air from a constant air stream are diverted through a glass syringe containing a piece of filter paper on which the compounds have been deposited. The odor stimulus was pulsed by means of a solenoid-activated valve controlled by the software.</li>
 <li>If unit activity suddenly stops or changes, check the saline drip and let the preparation rest for 15 min. If the spontaneous activity does not regain its previous level then the preparation should be discarded and another bee used if available.</li>
 <li>After the experiment, fix the brain with 5% formalin for 20 min with the probe still in the tissue. Next, excise the brain and place it in 2% glutaraldehyde for 4 hr, and subsequently do a graded ethanol dehydration series and clear the brain with methyl salicylate. Based on the fixing and clearing of the tissue, the locations in the AL where the electrodes punctured the tissue should be clearly discernible by confocal microscopy.</li>
 </ol>
 4. Data Analysis
 <ol>
 <li>Analyze collected data after the experiment to separate and identify the recorded neural units. Use typical software programs (Offline Sorter, MClust, and SClust) to separate waveforms, or "spikes", based on spike shapes, such as peak or valley amplitude, peak half-width, etc., or reduced measures (principal components)17,18 (Figure 2). Use only those clusters of spikes that separated in three-dimensional space (PC1-PC3) and are statistically different from one another (multivariate ANOVA; p &lt; 0.05) (Figure 2) for further analysis. Please refer to citation #s 17-19 for full description of tetrode recording and spike-sorting methodologies.</li>
 <li>Time-stamp spikes in each cluster, and export these data for analysis using MATLAB or Neuroexplorer (Nex Technologies, Winston-Salem, NC) to create raster plots and firing rate responses (Figures 2, 3A).</li>
 <li>Identify the retention times of volatiles using the simultaneously recorded GC data. Use the retention times of the volatiles, determined by the apex of the peak from the chromatogram, to examine unit responses at those time points.</li>
 <li>To examine individual unit responses through the GC run, bin the number of spikes in 100 msec intervals and examine the time-course of firing rate responses with reference to the retention time of eluting volatiles. The binning of spikes in 100 msec intervals provides enough detail, or signal, about the time-course of the neuronal response to an eluting odorant from the GC.</li>
 <li>To examine the population responses to the different eluting volatiles, integrate the firing rate responses of individual units over a 3 sec sampling window, 1.5 sec before, and 1.5 sec after, the retention time of the volatile (Figure 3). This time period is typical of the duration of an eluting volatile from the GC. We show firing rate responses of the units by color-coding them (red is a high firing rate response; blue is a low response) and arranging them as an activity matrix with each row representing the ensemble response to the GC effluent (columns) (Figure 3).</li>
 </ol>

<ol>
	<li>Hildebrand, J. G., Shepherd, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+olfactory:+converging+evidence+for+common+principles+across+phyla.">Mechanisms of olfactory: converging evidence for common principles across phyla.</a> Annual Review of Neuroscience. 20, 595-631 (1997).</li><li>Reisenman, C. E., Riffell, J. A., Bernays, E. A., 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=Antagonistic+effects+of+floral+scent+in+an+insect-plant+interaction.">Antagonistic effects of floral scent in an insect-plant interaction.</a> Proceedings of the Royal Society B: Biological Sciences. 277, 2371-2379 (2010).</li><li>Reisenman, C. E., Riffell, J. A., 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=.">.</a> International Symposium on Olfaction and Taste. 1170, 462-467 (2009).</li><li>Alarc&#243;n, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Congruence+between+visitation+and+pollen-transport+networks+in+a+California+plant-pollinator+community.">Congruence between visitation and pollen-transport networks in a California plant-pollinator community.</a> Oikos. 119, 35-44 (2010).</li><li>Alarc&#243;n, R., Waser, N. M., Ollerton, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Year-to-year+variation+in+the+topology+of+a+plant-pollinator+interaction+network.">Year-to-year variation in the topology of a plant-pollinator interaction network.</a> Oikos. 117, 1796-1807 (2008).</li><li>Riffell, 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=Behavioral+consequences+of+innate+preferences+and+olfactory+learning+in+hawkmoth-flower+interactions.">Behavioral consequences of innate preferences and olfactory learning in hawkmoth-flower interactions.</a> P. Natl. Acad. Sci. U.S.A. 105, 3404-3409 (2008).</li><li>De Moraes, C. M., Lewis, W. J., Pare, P. W., Alborn, H. T., Tumlinson, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Herbivore-infested+plants+selectively+attract+parasitoids.">Herbivore-infested plants selectively attract parasitoids.</a> Nature. 393, 570 (1998).</li><li>Carey, A. F., Wang, G., Su, C. -. Y., Zwiebel, L. J., 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=Odorant+reception+in+the+malaria+mosquito+Anopheles+gambiae.">Odorant reception in the malaria mosquito Anopheles gambiae.</a> Nature. 464, 66-71 (2010).</li><li>Turner, S. L., 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=Ultra-prolonged+activation+of+CO2-sensing+neurons+disorients+mosquitoes.">Ultra-prolonged activation of CO2-sensing neurons disorients mosquitoes.</a> Nature. 474, 87-91 (2011).</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>Syed, Z., Leal, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiological+measurements+from+a+moth+olfactory+system.">Electrophysiological measurements from a moth olfactory system.</a> J. Vis. Exp. (49), e2489 (2011).</li><li>Roelofs, W. L., Comeau, A., Hill, A., Milicevic, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sex+attractant+of+the+codling+moth:+characterization+with+electroantennogram+technique.">Sex attractant of the codling moth: characterization with electroantennogram technique.</a> Science. 174, 297-299 (1971).</li><li>Riffell, J. A., Lei, H., Christensen, T. A., 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=Characterization+and+coding+of+behaviorally+significant+odor+mixtures.">Characterization and coding of behaviorally significant odor mixtures.</a> Current Biology. 19, 335-340 .</li><li>Riffell, J. 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=Neural+correlates+of+behavior+in+the+moth+Manduca+sexta+in+response+to+complex+odors.">Neural correlates of behavior in the moth Manduca sexta in response to complex odors.</a> Proceedings of the National Academy of Sciences of the U.S.A. 106, 19219-19226 (2009).</li><li>Raguso, R. A., Pellmyr, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+headspace+analysis+of+floral+volatiles:+a+comparison+of+methods.">Dynamic headspace analysis of floral volatiles: a comparison of methods.</a> Oikos. 81, 238-254 (1998).</li><li>Rodriguez-Saona, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Herbivore-induced+blueberry+volatiles+and+intra-plant+signaling.">Herbivore-induced blueberry volatiles and intra-plant signaling.</a> J Vis Exp. , e3440 (2011).</li><li>Nguyen, D. P., 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=Micro-drive+array+for+chronic+in+vivo+recording:+tetrode+assembly.">Micro-drive array for chronic in vivo recording: tetrode assembly.</a> J Vis Exp. , e1098 (2009).</li><li>Schjetnan, A. G. P., Luczak, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recording+large-scale+neuronal+ensembles+with+silicon+probes+in+the+anesthetized+rat.">Recording large-scale neuronal ensembles with silicon probes in the anesthetized rat.</a> J Vis Exp. , e3282 (2011).</li><li>Deisig, N., Giurfa, M., Lachnit, H., Sandoz, J. -. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+representation+of+olfactory+mixtures+in+the+honeybee+antennal+lobe.">Neural representation of olfactory mixtures in the honeybee antennal lobe.</a> European Journal of Neuroscience. 24, 1161-1174 (2006).</li><li>St&#246;kl, 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=A+deceptive+pollination+system+targeting+drosophilids+through+olfactory+mimicry+of+yeast.">A deceptive pollination system targeting drosophilids through olfactory mimicry of yeast.</a> Current Biology. 20, 1846-1852 (2010).</li><li>Schneider, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elektrophysiologische+untersuchungen+von+chemo-+und+mechanorezeptoren+der+antenne+des+seidenspinners+Bombyx+mori+L.">Elektrophysiologische untersuchungen von chemo- und mechanorezeptoren der antenne des seidenspinners Bombyx mori L.</a> Journal of Comparative Physiology A: Neuroethology, Sensory, Neural, and Behavioral Physiology. 40, 8-41 (1957).</li><li>Arn, H., St&#228;dler, E., Rauscher, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+electroantennographic+detector:+a+selective+and+senstitive+tool+in+the+gas+chromatographic+analysis+of+insect+pheromones.">The electroantennographic detector: a selective and senstitive tool in the gas chromatographic analysis of insect pheromones.</a> Zeitschrift f&#252;r Naturforschung. 30c, 722-725 (1975).</li><li>Schneider, D., Boeckh, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rezeptorpotential+und+nervenimpulse+einzelner+olfaktorischer+sensillen+der+insektenantenne.">Rezeptorpotential und nervenimpulse einzelner olfaktorischer sensillen der insektenantenne.</a> Journal of Comparative Physiology A: Neuroethology, Sensory, Neural, and Behavioral Physiology. 45, 405-412 (1962).</li><li>Blight, M. M., Pickett, J. A., Wadhams, L. J., Woodcock, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antennal+perception+of+oilseed+rape+Brassica+napus+(Brassicaceae)+volatiles+by+the+cabbage+seed+weevil+Ceutorhynchus+assimilis+(Coleoptera,+Curculionidae).">Antennal perception of oilseed rape Brassica napus (Brassicaceae) volatiles by the cabbage seed weevil Ceutorhynchus assimilis (Coleoptera, Curculionidae).</a> Journal of Chemical Ecology. 21, 1649-1664 (1995).</li><li>Lin, D. Y., Shea, S. D., Katz, L. C. <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+natural+stimuli+in+the+rodent+main+olfactory+bulb.">Representation of natural stimuli in the rodent main olfactory bulb.</a> Neuron. 50, 937-949 (2006).</li><li>Lei, H., Reisenman, C. E., Wilson, C. H., Gabbur, P., 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=Spiking+patterns+and+their+functional+implications+in+the+antennal+lobe+of+the+tobacco+hornworm+Manduca+sexta.">Spiking patterns and their functional implications in the antennal lobe of the tobacco hornworm Manduca sexta.</a> PLoS ONE. 6, e23382 (2011).</li><li>Syed, Z., Leal, W. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+olfactory+response+of+Culex+mosquitoes+to+a+human-+and+bird-derived+attractant.">Acute olfactory response of Culex mosquitoes to a human- and bird-derived attractant.</a> Proceedings of the National Academy of Sciences. 106, 18803-18808 (2009).</li></ol>

No conflicts of interest declared.

Insect olfactory-mediated behaviors drive many different processes, including reproduction, host-site selection, and the identification of appropriate food resources. The study of these processes requires the ability to identify the volatiles emitted from the source, as well as the ability to identify those compounds that are mediating the behaviors. Complicating matters is that odors are comprised of tens to hundreds of individual compounds that together create a unique scent that is perceived differently than th...

<table border="1">
 <tbody>
 <tr>
 <td>Name of item</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Porapak Type Q 80-100 mesh</td>
 <td>Waters</td>
 <td>WAT027060</td>
 <td></td>
 </tr>
 <tr>
 <td>Reynolds Oven Bags</td>
 <td>Reynolds</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>GC</td>
 <td>Agilent</td>
 <td>7820A</td>
 <td></td>
 </tr>
 <tr>
 <td>GC column</td>
 <td>J&amp;W Scientific, Folsom, CA, USA</td>
 <td>DB-5 (30 m, 0.25 mm, 0.25 &mu;m)</td>
 <td></td>
 </tr>
 <tr>
 <td>Analytical helium carrier gas</td>
 <td>Praxair</td>
 <td>HE K</td>
 <td>1 cc/min</td>
 </tr>
 <tr>
 <td>16-channel silicon electrode</td>
 <td>Neuronexus Technologies</td>
 <td>a4x4-3mm50-177</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine wire NiCr, 0.012 mm diameter) </td>
 <td>Sandvik Kanthal HP Reid</td>
 <td>PX000004 </td>
 <td>For making custom tetrodes and stereotrodes</td>
 </tr>
 <tr>
 <td>Pre-amplifier</td>
 <td>Tucker-Davis System</td>
 <td>PZ-2</td>
 <td></td>
 </tr>
 <tr>
 <td>Amplifier </td>
 <td>Tucker-Davis System</td>
 <td>RZ-2</td>
 <td></td>
 </tr>
 <tr>
 <td>Data acquisition system - OpenEx suite</td>
 <td>Tucker-Davis System</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Online spike-sorting software - SpikePac</td>
 <td>Tucker-Davis System</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Offline spike-sorting software - Mclust Spike-sorting toolbox </td>
 <td>David Redish, Department of Neuroscience, University of Minnesota</td>
 <td>Free download at <a href="http://redishlab.neuroscience.umn.edu/MClust/MClust.html" target="_blank">http://redishlab.neuroscience.umn.edu/MClust/MClust.html</a> </td>
 <td>MATLAB toolbox</td>
 </tr>
 </tbody>
 </table>

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.

In the GCMR assay using the M. lewisii floral scent, we inject 3 &mu;l of the extract into the GC. The total number of volatiles eluting through the GC is typically 60-70 volatiles. The scent of M. lewisii is predominantly composed of monoterpenoids, including &beta;-myrcene (acyclic) and &alpha;-pinene, with the remainder of the scent composed of six-carbon volatiles, such as 2-hexanol, and sesquiterpenoids that comprise &lt; 1% of the headspace.
 GCM...

Watch this Scientific Journal Video about Identification of Olfactory Volatiles using Gas Chromatography-Multi-unit Recordings GCMR in the Insect Antennal Lobe at JoVE.com

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.

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 ...

identification-olfactory-volatiles-using-gas-chromatography-multi

Research

JoVE Journal

Neuroscience

14.2K 조회수.  University of Washington. 후각 신호는 곤충의 여러 가지 행동을 중재하고, 휘발성 화합물의 수백 수십 구성 종종 복합 혼합물이다. 곤충 antennal 엽의 멀티 채널 녹음과 가스 크로마토 그래피를 사용하여, 우리는 bioactive 화합물의 식별하는 방법을 설명합니다.

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

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory bulb1-5. Olfactory information in the external world is transmitted to the antennal lobe by olfactory sensory neurons (OSNs), which segregate to distinct regions of neuropil called glomeruli according to the specific olfactory receptor they express. Here, OSN axons synapse with both local interneurons (LNs), whose processes can innervate many different glomeruli, and projection neurons (PNs), which convey olfactory information to higher olfactory brain regions.
Optical imaging of the activity of OSNs, LNs and PNs in the antennal lobe - traditionally using synthetic calcium indicators (e.g. calcium green, FURA-2) or voltage-sensitive dyes (e.g. RH414) - has long been an important technique to understand how olfactory stimuli are represented as spatial and temporal patterns of glomerular activity in many species of insects6-10. Development of genetically-encoded neural activity reporters, such as the fluorescent calcium indicators G-CaMP11,12 and Cameleon13, the bioluminescent calcium indicator GFP-aequorin14,15, or a reporter of synaptic transmission, synapto-pHluorin16 has made the olfactory system of the fruitfly, Drosophila melanogaster, particularly accessible to neurophysiological imaging, complementing its comprehensively-described molecular, electrophysiological and neuroanatomical properties2,4,17. These reporters can be selectively expressed via binary transcriptional control systems (e.g. GAL4/UAS18, LexA/LexAop19,20, Q system21) in defined populations of neurons within the olfactory circuitry to dissect with high spatial and temporal resolution how odor-evoked neural activity is represented, modulated and transformed22-24.
Here we describe the preparation and analysis methods to measure odor-evoked responses in the Drosophila antennal lobe using G-CaMP25-27. The animal preparation is minimally invasive and can be adapted to imaging using wide-field fluorescence, confocal and two-photon microscopes.

Calcium Imaging of Odor-evoked Responses in the Drosophila Antennal Lobe

We describe an established technique to measure and analyze odor-evoked calcium responses in the antennal lobe of living Drosophila melanogaster.

The antennal lobe is the primary olfactory center in the insect brain and represents the anatomical and functional equivalent of the vertebrate olfactory ...

Odorant-induced Responses Recorded from Olfactory Receptor Neurons using the Suction Pipette Technique

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex behaviors such as food choice, predator, conspecific and mate recognition and other socially relevant cues. Olfactory receptor neurons (ORNs) are located in the dorsal part of the nasal cavity embedded in the olfactory epithelium. These bipolar neurons send an axon to the olfactory bulb (see Fig. 1, Reisert &amp; Zhao1, originally published in the Journal of General Physiology) and extend a single dendrite to the epithelial border from where cilia radiate into the mucus that covers the olfactory epithelium. The cilia contain the signal transduction machinery that ultimately leads to excitatory current influx through the ciliary transduction channels, a cyclic nucleotide-gated (CNG) channel and a Ca2+-activated Cl- channel (Fig. 1). The ensuing depolarization triggers action potential generation at the cell body2-4.
In this video we describe the use of the "suction pipette technique" to record odorant-induced responses from ORNs. This method was originally developed to record from rod photoreceptors5 and a variant of this method can be found at jove.com modified to record from mouse cone photoreceptors6. The suction pipette technique was later adapted to also record from ORNs7,8. Briefly, following dissociation of the olfactory epithelium and cell isolation, the entire cell body of an ORN is sucked into the tip of a recording pipette. The dendrite and the cilia remain exposed to the bath solution and thus accessible to solution changes to enable e.g. odorant or pharmacological blocker application. In this configuration, no access to the intracellular environment is gained (no whole-cell voltage clamp) and the intracellular voltage remains free to vary. This allows the simultaneous recording of the slow receptor current that originates at the cilia and fast action potentials fired by the cell body9. The difference in kinetics between these two signals allows them to be separated using different filter settings. This technique can be used on any wild type or knockout mouse or to record selectively from ORNs that also express GFP to label specific subsets of ORNs, e.g. expressing a given odorant receptor or ion channel.

Olfactory receptor neurons (ORNs) convert odor signals first into a receptor current that in turn triggers action potentials that are conveyed to second order neurons in the olfactory bulb. Here we describe the suction pipette technique to record simultaneously the odorant-induced receptor current and action potentials from mouse ORNs.

Animals sample the odorous environment around them through the chemosensory systems located in the nasal cavity. Chemosensory signals affect complex ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

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.

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.

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

Using Insect Electroantennogram Sensors on Autonomous Robots for Olfactory Searches

Robots designed to track chemical leaks in hazardous industrial facilities1 or explosive traces in landmine fields2 face the same problem as insects foraging for food or searching for mates3: the olfactory search is constrained by the physics of turbulent transport4. The concentration landscape of wind borne odors is discontinuous and consists of sporadically located patches. A pre-requisite to olfactory search is that intermittent odor patches are detected. Because of its high speed and sensitivity5-6, the olfactory organ of insects provides a unique opportunity for detection. Insect antennae have been used in the past to detect not only sex pheromones7 but also chemicals that are relevant to humans, e.g., volatile compounds emanating from cancer cells8 or toxic and illicit substances9-11. We describe here a protocol for using insect antennae on autonomous robots and present a proof of concept for tracking odor plumes to their source. The global response of olfactory neurons is recorded in situ in the form of electroantennograms (EAGs). Our experimental design, based on a whole insect preparation, allows stable recordings within a working day. In comparison, EAGs on excised antennae have a lifetime of 2 hr. A custom hardware/software interface was developed between the EAG electrodes and a robot. The measurement system resolves individual odor patches up to 10 Hz, which exceeds the time scale of artificial chemical sensors12. The efficiency of EAG sensors for olfactory searches is further demonstrated in driving the robot toward a source of pheromone. By using identical olfactory stimuli and sensors as in real animals, our robotic platform provides a direct means for testing biological hypotheses about olfactory coding and search strategies13. It may also prove beneficial for detecting other odorants of interests by combining EAGs from different insect species in a bioelectronic nose configuration14 or using nanostructured gas sensors that mimic insect antennae15.

We describe a protocol for using insect antennae in the form of electroantennograms (EAGs) on autonomous robots. Our experimental design allows stable recordings within a day and resolves individual odor patches up to 10 Hz. The efficiency of EAG sensors for olfactory searches is demonstrated in driving a robot toward an odor source.

Robots designed to track chemical leaks in hazardous industrial facilities1 or explosive traces in landmine fields2 face the same problem as insects ...

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first neural circuit in the AOS pathway, called the accessory olfactory bulb (AOB), plays an important role in establishing sex-typical behaviors such as territorial aggression and mating. This small (&#60;1 mm3) circuit possesses the capacity to distinguish unique behavioral states, such as sex, strain, and stress from chemosensory cues in the secretions and excretions of conspecifics. While the compact organization of this system presents unique opportunities for recording from large portions of the circuit simultaneously, investigation of sensory processing in the AOB remains challenging, largely due to its experimentally disadvantageous location in the brain. Here, we demonstrate a multi-stage dissection that removes the intact AOB inside a single hemisphere of the anterior mouse skull, leaving connections to both the peripheral vomeronasal sensory neurons (VSNs) and local neuronal circuitry intact. The procedure exposes the AOB surface to direct visual inspection, facilitating electrophysiological and optical recordings from AOB circuit elements in the absence of anesthetics. Upon inserting a thin cannula into the vomeronasal organ (VNO), which houses the VSNs, one can directly expose the periphery to social odors and pheromones while recording downstream activity in the AOB. This procedure enables controlled inquiries into AOS information processing, which can shed light on mechanisms linking pheromone exposure to changes in behavior.

Ex Vivo Preparations of the Intact Vomeronasal Organ and Accessory Olfactory Bulb

The mouse accessory olfactory bulb (AOB) has been difficult to study in the context of sensory coding. Here, we demonstrate a dissection that produces an ex vivo preparation in which AOB neurons remain functionally connected to their peripheral inputs, facilitating research into information processing of mouse pheromones and kairomones.

The mouse accessory olfactory system (AOS) is a specialized sensory pathway for detecting nonvolatile social odors, pheromones, and kairomones. The first ...

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular recording has historically been a common technique used to study sensitivities of individual photoreceptor cells to different light stimuli that is still being used today. However, there remains a dearth of detailed methodology in the literature for researchers wishing to replicate intracellular recording experiments in the eye. Here we present the insect as a model for examining eye physiology more generally. Insect photoreceptor cells are located near the surface of the eye and are therefore easy to reach, and many of the mechanisms involved in vision are conserved across animal phyla. We describe the basic procedure for in vivo intracellular recording of photoreceptor cells in the eye of a butterfly, with the goal of making this technique more accessible to researchers with little prior experience in electrophysiology. We introduce the basic equipment needed, how to prepare a live butterfly for recording, how to insert a glass microelectrode into a single cell, and finally the recording procedure itself. We also explain the basic analysis of raw response data for determining spectral sensitivity of individual cell types. Although our protocol focuses on determining spectral sensitivity, other stimuli (e.g., polarized light) and variations of the method are applicable to this setup.

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular ...

Insect-controlled Robot: A Mobile Robot Platform to Evaluate the Odor-tracking Capability of an Insect

Robotic odor source localization has been a challenging area and one to which biological knowledge has been expected to contribute, as finding odor sources is an essential task for organism survival. Insects are well-studied organisms with regard to odor tracking, and their behavioral strategies have been applied to mobile robots for evaluation. This "bottom-up" approach is a fundamental way to develop biomimetic robots; however, the biological analyses and the modeling of behavioral mechanisms are still ongoing. Therefore, it is still unknown how such a biological system actually works as the controller of a robotic platform. To answer this question, we have developed an insect-controlled robot in which a male adult silkmoth (Bombyx mori) drives a robot car in response to odor stimuli; this can be regarded as a prototype of a future insect-mimetic robot. In the cockpit of the robot, a tethered silkmoth walked on an air-supported ball and an optical sensor measured the ball rotations. These rotations were translated into the movement of the two-wheeled robot. The advantage of this "hybrid" approach is that experimenters can manipulate any parameter of the robot, which enables the evaluation of the odor-tracking capability of insects and provides useful suggestions for robotic odor-tracking. Furthermore, these manipulations are non-invasive ways to alter the sensory-motor relationship of a pilot insect and will be a useful technique for understanding adaptive behaviors.

The capability to localize an odor source is necessary for insect survival and is expected to be applicable to artificial odor-tracking. The insect-controlled robot is driven by an actual silkmoth and enables us to evaluate the odor-tracking capability of insects through a robotic platform.

Robotic odor source localization has been a challenging area and one to which biological knowledge has been expected to contribute, as finding odor ...

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 ...

Investigating Object Representations in the Macaque Dorsal Visual Stream Using Single-unit Recordings

Previous studies have shown that neurons in parieto-frontal areas of the macaque brain can be highly selective for real-world objects, disparity-defined curved surfaces, and images of real-world objects (with and without disparity) in a similar manner as described in the ventral visual stream. In addition, parieto-frontal areas are believed to convert visual object information into appropriate motor outputs, such as the pre-shaping of the hand during grasping. To better characterize object selectivity in the cortical network involved in visuomotor transformations, we provide a battery of tests intended to analyze the visual object selectivity of neurons in parieto-frontal regions.

A detailed protocol to analyze object selectivity of parieto-frontal neurons involved in visuomotor transformations is presented.

Previous studies have shown that neurons in parieto-frontal areas of the macaque brain can be highly selective for real-world objects, disparity-defined ...