Name: using single sensillum recording to detect olfactory neuron responses of bed bugs to semiochemicals
Uploaded: 2016-01-18T15:00:00
Description: Watch this Scientific Journal Video about Using Single Sensillum Recording to Detect Olfactory Neuron Responses of Bed Bugs to Semiochemicals at JoVE.com

The project was supported by Award AAES 461Hatch/Multistate Grants ALA08-045 and ALA015-1-10026 to N.L.

1. Preparation of Instruments, Stimuli Solutions, and Bed Bugs
<ol>
	<li>Prepare a 50% KNO2 solution (w/v) in a 20 ml bottle.</li>
	<li>Sharpen two tungsten microelectrodes in KNO2 solution at 5 V by repeatedly dipping the tungsten electrodes in and out of the solution.
	<ol>
		<li>Roughly sharpen the tungsten wire by dipping about 10 mm of the tungsten wire in and out of the KNO2 solution at the speed of 2 dips/sec&#160;for about 5 min, which can greatly consume the front end of the tu...

<ol>
	<li>Bartonicka, T., Gaisler, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Seasonal+dynamics+in+the+numbers+of+parasitic+bugs+(Heteroptera,+Cimicidae):+a+possible+cause+of+roost+switching+in+bats+(Chiroptera,+Vespertilionidae).">Seasonal dynamics in the numbers of parasitic bugs (Heteroptera, Cimicidae): a possible cause of roost switching in bats (Chiroptera, Vespertilionidae).</a> Parasitol Res. 100 (6), 1323-1330 (2007).</li><li>Thomas, I., Kihiczak, G. G., Schwartz, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bed+bug+bites:+a+review.">Bed bug bites: a review.</a> Int J Dermatol. 43 (6), 430-433 (2004).</li><li>Anderson, A. L., Leffler, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bed+bug+infestations+in+the+news:+a+picture+of+an+emerging+public+health+problem+in+the+United+States.">Bed bug infestations in the news: a picture of an emerging public health problem in the United States.</a> J Environ Health. 70 (9), 24-27 (2008).</li><li>Boase, C., Robinnson, W., Bajomi, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bed+bugs+(Hemiptera:+Cimicidae):+an+evidence-based+analysis+of+the+current+situation.">Bed bugs (Hemiptera: Cimicidae): an evidence-based analysis of the current situation.</a> Sixth international conference on urban pests. OOK-Press Kft. , (2008).</li><li>Doggett, S. L., Geary, M. J., Russell, R. 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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biochemical+and+molecular+analysis+of+deltamethrin+resistance+in+the+common+bed+bug+(Hemiptera:+Cimicidae).">Biochemical and molecular analysis of deltamethrin resistance in the common bed bug (Hemiptera: Cimicidae).</a> J Med Entomol. 45 (6), 1092-1101 (2008).</li><li>Wang, L., Xu, Y., Zeng, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resurgence+of+bed+bugs+(Hemiptera:+Cimicidae)+in+mainland+China.">Resurgence of bed bugs (Hemiptera: Cimicidae) in mainland China.</a> Fla Entomol. 96 (1), 131-136 (2013).</li><li>Haynes, K. F., Potter, M. F., Ishaaya, I., Palli, S. R., Horowitz, A. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+progress+in+bed+bug+management.">Recent progress in bed bug management.</a> Advanced technologies for managing insect pests. , 269-278 (2013).</li><li>Carey, A. F., 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=Insect+olfaction+from+model+systems+to+disease+control.">Insect olfaction from model systems to disease control.</a> Proc Natl Acad Sci. 108 (32), 12987-12995 (2011).</li><li>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=Odorant+reception+in+insects:+roles+of+receptors,+binding+proteins,+and+degrading+enzymes.">Odorant reception in insects: roles of receptors, binding proteins, and degrading enzymes.</a> Annu Rev Entomol. 58, 373-391 (2013).</li><li>Den Otter, C. J., Behan, M., Maes, F. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+response+in+female+Pieris+brassicae.+(Lepidoptera:+Pieridae)+to+plant+volatiles+and+conspecific+egg+odours.">Single cell response in female Pieris brassicae. (Lepidoptera: Pieridae) to plant volatiles and conspecific egg odours.</a> J Insect Physiol. 26 (7), 465-472 (1980).</li><li>Levinson, H. Z., Levinson, A. R., Muller, B., Steinbrecht, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+of+sensilla,+olfactory+perception,+and+behavior+of+the+bed+bug,+Cimex+lectularius.,+in+response+to+its+alarm+pheromone.">Structural of sensilla, olfactory perception, and behavior of the bed bug, Cimex lectularius., in response to its alarm pheromone.</a> J Insect Physiol. 20 (7), 1231-1248 (1974).</li><li>Harraca, V., Ignell, R., L&#246;fstedt, C., Ryne, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+antennal+olfactory+system+of+the+bed+bug+(Cimex+lectularius).">Characterization of the antennal olfactory system of the bed bug (Cimex lectularius).</a> Chem Senses. 35 (3), 195-204 (2010).</li><li>Liu, F., Haynes, K. F., Appel, A. G., Liu, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Antennal+olfactory+sensilla+responses+to+insect+chemical+repellents+in+the+common+bed+bug,+Cimex+lectularius.">Antennal olfactory sensilla responses to insect chemical repellents in the common bed bug, Cimex lectularius.</a> J Chem Ecol. 40 (6), 522-533 (2014).</li><li>Olson, J. F., Moon, R. D., Kells, S. A., Mesce, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphology,+ultrastructure+and+functional+role+of+antennal+sensilla+in+off-host+aggregation+by+the+bed+bug,+Cimex+lectularius.">Morphology, ultrastructure and functional role of antennal sensilla in off-host aggregation by the bed bug, Cimex lectularius.</a> Arthropod Struct Dev. 43 (2), 117-122 (2014).</li><li>Bruyne, M., Foster, K., 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=Odor+coding+in+the+Drosophila+antenna.">Odor coding in the Drosophila antenna.</a> Neuron. 30 (2), 537-552 (2001).</li><li>Qiu, Y. T., Loon, J. J. A., Takken, W., Meijerink, J., Smid, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Olfactory+coding+in+antennal+neurons+of+the+malaria+mosquito,+Anopheles+gambiae.">Olfactory coding in antennal neurons of the malaria mosquito, Anopheles gambiae.</a> Chem Senses. 31 (9), 845-863 (2006).</li><li>Ghaninia, M., Ignell, R., 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=Functional+classification+and+central+nervous+projections+of+olfactory+receptor+neurons+housed+in+antennal+trichoid+sensilla+of+female+yellow+fever+mosquito,+Aedes+aegypti.">Functional classification and central nervous projections of olfactory receptor neurons housed in antennal trichoid sensilla of female yellow fever mosquito, Aedes aegypti.</a> Eur J Neurosci. 26 (6), 1611-1623 (2007).</li><li>Hill, S. R., Hanson, B. S., Ignell, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+antennal+trichoid+sensilla+from+female+southern+house+mosquito,+Culex+quinquefasciatus+Say.">Characterization of antennal trichoid sensilla from female southern house mosquito, Culex quinquefasciatus Say.</a> Chem Senses. 34 (3), 231-252 (2009).</li></ol>

The Single Sensillum Recording technique has been extensively used in testing the neural responses of insects such as fruit flies, mosquitoes and bed bugs to different chemical stimuli in the environment. These chemical stimuli are often dissolved and diluted in a common solvent in order to prepare different doses of treatments. However, different solvents can produce quite different release rates for the stimuli. Previous studies on some extensively studied insects such as Drosophila melanogaster, Anopheles...

The common bed bug Cimex lectularius L (Hemiptera: Cimicidae), as a temporary ectoparasite, is an obligated blood-sucking insect, which means their survival, development, and reproduction require blood sources from hosts, including both humans and animals1,2. Although virus transmission has rarely been reported due to C. lectularius, the biting nuisance generated by an infestation seriously affects hosts both physically and psychologically3. The introduction and widespread use of chemical insecticides, especially DDT, lowered the risk of infestations and by the end of the 1950s infestations were at such a low level that they were no longer a serious public concern. However, a number of possible factors have led to resurgence in bed bug populations worldwide, such as the reduced use of insecticides, a decline of public awareness, increased traveling activity, and the development of resistance to insecticides4-9.
Chemical cues in the environment are detected and recognized by insects through olfactory organs such as antennae and maxillary palps. The olfactory sensilla on the insect antennae play a crucial role in detecting these chemical cues. The chemical molecules enter the antennal cuticle through pores on the cuticle surface. Odorant binding proteins in the antennal lymph bind to these chemical molecules and transport them onto the odorant receptors10. The odorant receptors and their co-receptor from the non-selective cation ion channel on the neural membrane, which will be depolarized once these chemical molecules are recognized by the odorant receptors11.
Single sensillum recording (SSR) was developed to detect the extracellular change in the action potential caused by the application of either chemical or non-chemical stimuli. By inserting a recording electrode into the sensillum lymph and a reference electrode into some other part of the insect body (usually either the compound eyes or the abdomen), the firing rate of the neurons in response to stimuli can be recorded12. Changes in the number of spikes represent the sensitivity of the insect to specific stimuli. Chemical stimuli of different identities and concentration will elicit different neural responses, with different firing rates and temporal structures, and can thus be used to investigate the coding process of the insect to specific chemicals.
For the common bed bug, both sexual forms share the same pattern of olfactory sensilla on the antennae: nine grooved peg C sensilla, 29 hair-like E (E1 and E2) sensilla, and one pair each of D&#945;, D&#946;, D&#947; smooth peg sensilla 13,14. As multiple neurons have been identified in each type of sensillum, it is not easy to distinguish the action potentials from different neurons housed in the same sensillum, so for this experiment the total numbers of action potentials were counted off-line for a 500 msec period before and after stimulation. The number of action potentials after stimulation was then subtracted from the number of action potentials before stimulation and multiplied by two in order to quantify the changes in the firing rate in each individual sensillum in spikes per second15.

<table border="0" cellpadding="0" cellspacing="0" width="1064">
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:299px;">Tungsten wire</td>
			<td style="width:139px;">A-M SYSTEMS</td>
			<td style="width:117px;">#716500</td>
			<td style="width:511px;">Used for preparing the electrode</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">KNO2</td>
			<td>Sigma</td>
			<td>#310484</td>
			<td>Used for sharpening the tungsten wire</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">AC Power Supply</td>
			<td>BK Precision</td>
			<td>1653A</td>
			<td>Providing the voltage in sharpening the tungsten wire</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Leica Z6 APO Microscope</td>
			<td>Leica</td>
			<td>10447424</td>
			<td>Used for observing the sensilla on antennae</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Simulus controller</td>
			<td>Syntech</td>
			<td>CS-55</td>
			<td>Used for controlling the stimulus application</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">4-Channel USB Acquisition Controller</td>
			<td>Syntech</td>
			<td>IDAC-4</td>
			<td>Real-time on screen display of all signals before and during recording</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Light Source</td>
			<td>SCHOTT</td>
			<td>A20500</td>
			<td>Providing light sources for observation</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Micromanupulator</td>
			<td>Leica</td>
			<td>115378</td>
			<td>Used for minor movement of electrode</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Speaker</td>
			<td>Juster</td>
			<td>95a</td>
			<td>Connected with Acquisition Controller IDAC-4 and providing sound for the signal</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Magnetic stand</td>
			<td>Narishige</td>
			<td>GJ-1</td>
			<td>Used to hold the reference electrode, stablized bed bug and stimulus delivery tube</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">TMC Vibration Isolation Table</td>
			<td>TMC</td>
			<td>63-500</td>
			<td>Used for isolating the vibration from the equipments</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Coverslip</td>
			<td>Tedpella</td>
			<td>2225-1</td>
			<td>Used for holding the bed bug</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Double-sided Tape</td>
			<td>3M</td>
			<td>XT6110</td>
			<td>Used for stablizing the bed bug on the coverclip</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dental Wax</td>
			<td>Dentakit</td>
			<td>DK-R012</td>
			<td>Used for supporting the coverclip where bed bug is stablized&#160;</td>
		</tr>
	</tbody>
</table>

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.

Single sensillum recording is a powerful investigative technique used in studies of insect chemical ecology and neural physiology. Investigating the neural responses of insects to different volatile compounds, especially those thought to be ecologically related to the survival and development of the insects, not only gives us invaluable insights into the insect olfaction process, but also opens up promising new avenues that could potentially lead to the development of useful new reagents ...

Watch this Scientific Journal Video about Using Single Sensillum Recording to Detect Olfactory Neuron Responses of Bed Bugs to Semiochemicals at JoVE.com

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

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 ...

The overall goal of the single sensillum recording is to investigate neuronal responses of the antennal olfactory sensillum to various odorants. This method can help to answer the key questions in the field of chemical ecology such as if the insect can detect the chemical odorants from their environment and how they process the chemical odorants. The main advantage of this technique is that it allows mirroring the single neuron activity of the function.It provides more precise measurement of the sensitivity of neurons to the chemical odorants. The implication of this technique extended toward the development of attractants or repellants in the insect control. Because this technique can be used large amount of chemicals that are biologically important for the insect.Begin this procedure by anesthetizing bed bugs on ice for two to three minutes. Then fix the insects antennae and body on a microscope cover slip with double sided tape and remove the legs with fine scissors. Use a small pin to gently adjust the antennae on the tape.Next, rest the cover slip against a small ball of dental wax to facilitate manipulation and adjust it to a 90 degree angle for placing the recording electrode. Once secured, place the bed bug under a stereo microscope. Turn on the light source and adjust the intensity of illumination until the antennae is clearly presented.Then focus the microscope on the second flagellum of the bed bug's antennae at a high magnification. In this step, connect the pre-amplifier with a signal acquisition controller, which is connected with the computer for signal recording and visualization. Next, open the software and click record from the menu bar.Then choose wave to start recording the wave signals. Turn on the speaker connected to the pre-amplifier to present the tone of the neuronal responses from the antennal sensillum. Now insert the reference electrode into the abdomen of the stabilized bed bug.After the reference electrode has been connected to the bed bug's abdomen, move the recording electrode towards the posterior end of the bed bug's antennae. When the recording electrode is in contact with the tip of the right antennae, locate the electrode at low magnification. Then adjust the recording electrode while gradually increasing the magnification until both the electrode and the antennal sensillum are in the same plane and clearly visible under the microscope.After that, insert the recording electrode into the shaft of the sensillum using the micro-manipulator. Lower it deeper if the background noise is too high. Once clear action potentials are observed from the recorded sensillum, fill a micro pipette with botanical stimulus.Use the micro pipette to deposit a 10 microliter aliquot of the stimulus onto a filter paper strip inside a glass pastier pipette. Accurately and directly recording electrodes to the sensillum is the key to the success of the experiment. Background noise can be partially reduced after inserting the electrodes a little deeper.Next, connect the loaded pipette to the outlet of the stimulus flow tube and place the pipette tip into the small hole in the tube oriented towards the antennae. When all the connections are stabilized, depress the foot switch of the stimulus controller to deliver a stimulus into the continuous humidified air stream and start recording. The recording process will last for 10 seconds starting one second before the stimulation.Count the action potentials offline for two 500 millisecond periods, one before, and one after the stimulation. To present another stimulus, label a new pipette with 001%eucalyptol to be tested. Apply 10 microliters of the stimulus before placing a small piece of filter paper onto it.Then wait for two to five minutes until the stimulus is completely vaporized in the glass pipette. Attach the pipette onto the outlet of the stimulus flow tube. Subsequently, insert the pipette tip into the small hole of the tube oriented towards the antennae.Depress the foot switch and start the 10 second recording. After that, disconnect the pipette and prepare another pipette with 01%eucalyptol. Test all the rest of the doses of eucalyptol on the antennal sensilla to observe the most dependent responses.In this figure, the signal trace shows the typical neural response of an olfactory sensillum to the solvent which is used as the control in the single sensillum recording. Signal recording started one second before the 0.5 second of puff stimulus. The signal was recorded continuously for 10 seconds after initiating the stimulus puff.This signal trace shows the extremely strong neural response of an olfactory sensillum to a botanical stimulus 10%positive betapynene. After the puff of stimulus is delivered to the olfactory sensillum, the olfactory sensory neurons inside this sensillum fire with high frequency into a long-lasting temporal dynamic. For in this procedure are the message that behavior by an insect can be performed in order to answer additional questions like what are the behavioral responses of insects to certain chemicals with strong neuron responses.After watching this video, you should have a better understanding of how to record the insect olfactory neuron responses to the chemical stimulus.

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Research

JoVE Journal

Neuroscience

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Analyzing Responses of Mouse Olfactory Sensory Neurons Using the Air-phase Electroolfactogram Recording

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and other social interactions. The electroolfactogram (EOG) recording is an informative, easy to conduct, and reliable method to assay olfactory function at the level of the olfactory epithelium. Since the 1956 description of the EOG by Ottoson in frogs1, the EOG recording has been applied in many vertebrates including salamanders, rabbits, rats, mice, and humans (reviewed by Scott and Scott-Johnson, 2002, ref. 2). The recent advances in genetic modification in mice have rekindled interest in recording the EOG for physiological characterization of olfactory function in knock-out and knock-in mice. EOG recordings have been successfully applied to demonstrate the central role of olfactory signal transduction components3-8, and more recently to characterize the contribution of certain regulatory mechanisms to OSN responses9-12.
Odorant detection occurs at the surface of the olfactory epithelium on the cilia of OSNs, where a signal transduction cascade leads to opening of ion channels, generating a current that flows into the cilia and depolarizes the membrane13. The EOG is the negative potential recorded extracellularly at the surface of the olfactory epithelium upon odorant stimulation, resulting from a summation of the potential changes caused by individual responsive OSNs in the recording field2. Comparison of the amplitude and kinetics of the EOG thus provide valuable information about how genetic modification and other experimental manipulations influence the molecular signaling underlying the OSN response to odor. 
Here we describe an air-phase EOG recording on a preparation of mouse olfactory turbinates. Briefly, after sacrificing the mouse, the olfactory turbinates are exposed by bisecting the head along the midline and removing the septum. The turbinate preparation is then placed in the recording setup, and a recording electrode is placed at the surface of the olfactory epithelium on one of the medial turbinates. A reference electrode is electrically connected to the tissue through a buffer solution. A continuous stream of humidified air is blown over the surface of the epithelium to keep it moist. The vapor of odorant solutions is puffed into the stream of humidified air to stimulate the epithelium. Responses are recorded and digitized for further analysis.

The electroolfactogram (EOG) recording is an informative, easy-to-conduct, and reliable way of assessing olfactory function at the level of the olfactory epithelium. This protocol describes a recording setup, mouse tissue preparation, data collection, and basic data analysis.

Animals depend on olfaction for many critical behaviors, such as finding food sources, avoiding predators, and identifying conspecifics for mating and ...

Recording Behavioral Responses to Reflection in Crayfish

Social behavior depends on sensory input from the visual, mechanical and olfactory systems. One important issue concerns the relative roles of each sensory modality in guiding behavior. The role of visual inputs has been examined by isolating visual stimuli from mechanical and chemosensory stimuli. In some studies (Bruski &amp; Dunham, 1987: Delgado-Morales et al., 2004) visual inputs have been removed with blindfolds or low light intensity, and effects of remaining sensory modalities have been elucidated. An alternative approach is to study the effects of visual inputs in the absence of any appropriate mechanical and chemosensory cues. This approach aims to identify the exclusive role of visual inputs. 
We have used two methods to provide visual stimuli to crayfish without providing chemical and mechanical cues. In one method, crayfish are videotaped in an aquarium where half of the walls are covered in mirrors to provide a reflective environment, and the other half are covered in a non-reflective (matte finish) plastic. This gives the crayfish a choice between reflective and non-reflective environments. The reflective environment provides visual cues in the form of reflected images of the crayfish as it moves throughout half of the tank; these visual cues are missing from the non-reflective half of the tank. An alternative method is to videotape the behavior of crayfish in an aquarium separated by a smaller chamber at each end, with a crayfish in one small chamber providing visual cues and an inert object in the opposite small chamber providing visual input from a non-moving, non-crayfish source. 
Our published results indicate that responses of crayfish to the reflective environment depend on socialization and dominance rank. Socialized crayfish spent more time in the reflective environment and exhibited certain behaviors more frequently there than in the non-reflective environment; isolated crayfish showed no such differences. Crayfish that were housed in same-sex pairs developed a social rank of either dominant or subordinate. Responses to reflection differed between dominant and subordinate crayfish (May &amp; Mercier, 2006; May &amp; Mercier, 2007). Dominant crayfish spent more time on the reflective side, entered reflective corners more frequently and spent more time in reflective corners compared to the non-reflective side. Subordinate crayfish walked in reverse more often on the reflective side than on the non-reflective side. Preliminary data suggest similar effects from visual cues provided by a crayfish in a small adjoining chamber (May et al., 2008).

We have developed two methods for studying effects of visual cues on behavior in the absence of tactile and chemical cues. One method involves videotaping responses of crayfish to reflective walls in an aquarium; the other examines effects of visual inputs provided by a live crayfish behind a transparent partition.

Social behavior depends on sensory input from the visual, mechanical and olfactory systems.  One important issue concerns the relative roles of each ...

Using a Comparative Species Approach to Investigate the Neurobiology of Paternal Responses

A goal of behavioral neuroscience is to identify underlying neurobiological factors that regulate specific behaviors. Using animal models to accomplish this goal, many methodological strategies require invasive techniques to manipulate the intensity of the behavior of interest (e.g., lesion methods, pharmacological manipulations, microdialysis techniques, genetically-engineered animal models). The utilization of a comparative species approach allows researchers to take advantage of naturally occurring differences in response strategies existing in closely related species. In our lab, we use two species of the Peromyscus genus that differ in paternal responses. The male California deer mouse (Peromyscus californicus) exhibits the same parental responses as the female whereas its cousin, the common deer mouse (Peromyscus maniculatus) exhibits virtually no nurturing/parental responses in the presence of pups. Of specific interest in this article is an exploration of the neurobiological factors associated with the affiliative social responses exhibited by the paternal California deer mouse. Because the behavioral neuroscience approach is multifaceted, the following key components of the study will be briefly addressed: the identification of appropriate species for this type of research; data collection for behavioral analysis; preparation and sectioning of the brains; basic steps involved in immunocytochemistry for the quantification of vasopressin-immunoreactivity; the use of neuroimaging software to quantify the brain tissue; the use of a microsequencing video analysis to score behavior and, finally, the appropriate statistical analyses to provide the most informed interpretations of the research findings.

The comparative species approach allows behavioral neuroscientists to explore various neurobiological factors associated with specific behaviors viewed as characteristic of a specific animal model. Taking advantage of naturally occurring differences in behavior between closely related species, this technique doesn&#x2019;t require invasive techniques to manipulate the expression of the behavior.

A goal of behavioral neuroscience is to identify underlying neurobiological factors that regulate specific behaviors.  Using animal models to accomplish ...

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 ...

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 ...

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral cortex. We here share our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex using a method developed by Lima and colleagues1. Recordings are performed in mice expressing Channelrhodopsin-2 (ChR2) in specific neuronal subpopulations. Members of the population are identified by their response to a brief flash of blue light. This technique &#8211; termed &#8220;PINP&#8221;, or Photostimulation-assisted Identification of Neuronal Populations &#8211; can be implemented with standard extracellular recording equipment. It can serve as an inexpensive and accessible alternative to calcium imaging or visually-guided patching, for the purpose of targeting extracellular recordings to genetically-identified cells. Here we provide a set of guidelines for optimizing the method in everyday practice. We refined our strategy specifically for targeting parvalbumin-positive (PV+) cells, but have found that it works for other interneuron types as well, such as somatostatin-expressing (SOM+) and calretinin-expressing (CR+) interneurons.

A Guide to In vivo Single-unit Recording from Optogenetically Identified Cortical Inhibitory Interneurons

Here we describe our strategy for obtaining stable, well-isolated single-unit recordings from identified inhibitory interneurons in the anesthetized mouse cortex. Neurons expressing ChR2 are identified by their response to blue light. The method uses standard extracellular recording equipment, and serves as an inexpensive alternative to calcium imaging or visually-guided patching.

A major challenge in neurophysiology has been to characterize the response properties and function of the numerous inhibitory cell types in the cerebral ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a considerable amount of information has been generated regarding ORN responses to odorants, the role of specific ORNs in driving behavioral responses remains poorly understood. Complications in behavior analyses arise due to different volatilities of odorants that activate individual ORNs, multiple ORNs activated by single odorants, and the difficulty in replicating naturally observed temporal variations in olfactory stimuli using conventional odor-delivery methods in the laboratory. Here, we describe a protocol that analyzes Drosophila larval behavior in response to simultaneous optogenetic stimulation of its ORNs. The optogenetic technology used here allows for specificity of ORN activation and precise control of temporal patterns of ORN activation. Corresponding larval movement is tracked, digitally recorded, and analyzed using custom written software. By replacing odor stimuli with light stimuli, this method allows for a more precise control of individual ORN activation in order to study its impact on larval behavior. Our method could be further extended to study the impact of second-order projection neurons (PNs) as well as local neurons (LNs) on larval behavior. This method will thus enable a comprehensive dissection of olfactory circuit function and complement studies on how olfactory neuron activities translate in to behavior responses.

Tracking Drosophila Larval Behavior in Response to Optogenetic Stimulation of Olfactory Neurons

This protocol analyzes navigational behavior of Drosophila larva in response to simultaneous optogenetic stimulation of its olfactory neurons. Light of 630 nm wavelength is used to activate individual olfactory neurons expressing a red-shifted channel rhodopsin. Larval movement is simultaneously tracked, digitally recorded, and analyzed using custom-written software.

The ability of insects to navigate toward odor sources is based on the activities of their first-order olfactory receptor neurons (ORNs). While a ...

Single Sensillum Recordings for Locust Palp Sensilla Basiconica

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially for the detection of non-volatile chemical cues through sensilla chaetica (previously named terminal sensilla or crested sensilla). There is now increasing evidence that these palps also have an olfactory function. An odorant receptor (LmigOR2) and an odorant-binding protein (LmigOBP1) have been localized in the neurons and accessory cells, respectively, in the sensilla basiconica of the palps. Single sensillum recording (SSR) is used for recording the responses of odorant receptor neurons, which is an effective method for screening active ligands on specific odorant receptors. SSR is used in functional studies of odorant receptors in palp sensilla. The structure of the sensilla basiconica located on the dome of the palps differs somewhat from the structure of those on the antennae. Therefore, when performing an SSR elicited by odorants, some specific advice may be helpful for obtaining optimum results. In this paper, a detailed and highly effective protocol for an SSR from insect palp sensilla basiconica is introduced.

This paper describes a detailed and highly effective protocol for single sensillum recordings from the sensilla basiconica on the palps of insect mouthparts.

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially ...