We thank Shigeru Matsuyama for providing purified bombykol. This work was supported by the Japan Society for the Promotion of Science KAKENHI (grant numbers 22700197 and 24650090) and the Human Frontier Science Program (HFSP).

1. Experimental Animal
<ol>
	<li>Prepare a plastic box to keep the pupae of male silkmoths (B. mori) until their eclosion. Put paper towels at the bottom and pieces of cardboard around the inner wall of the box (Figure 1A). 
	Note: The pieces of cardboard are necessary for the adult moths to hold while extending their wings during eclosion (Figure 1A).</li>
	<li>Put male silkmoth (Bombyx mori) pupae in the box and keep them in an incubator until eclosion under a 16-hr:8-hr light:dark cycle at 25 &#176;C. 
	NOTE: The male and female pupae can be discriminated by the sex markings on the abdomen (Figure 1B).</li>
	<li>Collect adult male moths after eclosion and move them into a new box.</li>
	<li>Keep the adult moths in an incubator under a 16-hr:8-hr light:dark cycle and decrease the temperature to 15 &#176;C to reduce their activity before the experiment.</li>
</ol>
2. Tethering a Silkmoth
<ol>
	<li>Fabrication of an attachment for tethering (Figure 2A) 
	Note: The attachment consists of a copper wire with a strip of a thin plastic sheet at its tip. This ensures the dorsal-ventral movement of the thorax during walking (Figure 2B).
	<ol>
		<li>Prepare a strip of a thin plastic sheet, 2 &#215; 40 mm (thickness: 0.1 mm), and fold it in the middle.</li>
		<li>Attach the folded strip to the tip of a copper wire with an adhesive.</li>
		<li>Bend the tip of the folded strip where the thorax of a silkmoth is attached.</li>
	</ol>
	</li>
	<li>Use adult moths (2-8 days old) during the light period for the experiment. 
	Note: The sensitivity to the pheromone strongly depends on the circadian clock18. Because B. mori is a diurnal moth, the experiment must be performed during the light period.</li>
	<li>Gently remove all scales on the dorsal thorax (mesonotum) using a piece of wet tissue (or cotton swab) and expose the cuticle of the mesonotum (Figure 2C).</li>
	<li>Paste an adhesive on the strip of plastic on the attachment and on the surface of the exposed mesonotum with a small flat-blade screwdriver and wait 5-10 min until the adhesive is no longer sticky. 
	Note: The adhesive should not touch the wing hinge or the forewing tegulae (Figure 2C).</li>
	<li>Bond the mesonotum to the attachment.</li>
	<li>Keep the moth tethered before placing it inside the cockpit of the robot. Hold the attachment on a stand and put a piece of paper under the legs to rest the moth.</li>
</ol>
3. Insect-controlled Robot
<ol>
	<li>Design the hardware of the insect-controlled robot based on previous works16,17,19. 
	Note: The insect-controlled robot consists of an air-supported treadmill with an optical mouse sensor to capture the insect locomotion, custom-built AVR-based microcontroller boards for processing and motor control, and two DC brushless motors (Figures 3 and 4). The robot can run on the basis of the ball rotation with 96% precision or higher, within a time delay of 200 msec. It also ensures the mobility of maximum forward speed (24.8 mm/sec) and angular velocity (96.3&#176;/sec) of the silkmoth during pheromone tracking behavior16. The airflow of the treadmill (Figure 5A) and odor delivery system (Figure 5B) are designed for the onboard moth to walk smoothly on the ball and to acquire an odor by two antennae. The air intake and flow channel of the treadmill is separated from those of the odor delivery system to avoid contamination of the pheromone.</li>
	<li>Design the software for the onboard microcontrollers based on previous works16. 
	Note: The onboard microcontroller calculates the robot movements from the insect locomotion measured with an optical sensor (rotational, &#916;x; translational, &#916;y; Figure 6). The travel distance (&#916;L) and turn angle (&#916;&#952;) per unit time of the robot are calculated on the basis of travel distance of each wheel (left, &#916;LL; right, &#916;LR) such as &#916;L = (&#916;LL + &#916;LR) / 2 and &#916;&#952; = (&#916;LL - &#916;LR) / Dwheel, where Dwheel is the distance between the two wheels (120 mm). &#916;LL and &#916;LR are further described as &#916;LL = &#916;Lx,L + &#916;Ly,L and &#916;LR = &#916;Lx,R + &#916;Ly,R, where &#916;Lx,L and &#916;Lx,R are the travel distances of the wheels on the left and right sides controlled by &#916;x, and &#916;Ly,L and &#916;Ly,R are those controlled by &#916;y. Ideally, &#916;Lx,L and &#916;Lx,R are described as &#916;Lx,L = -&#916;Lx,R = G &#916;x (Dwheel / Dball), and &#916;Ly,L and &#916;Ly,R are described as &#916;Ly,L = &#916;Ly,R = G &#916;y, where G is the motor gain and Dball is the diameter of the ball (50 mm). In practice, the motor gain is independently set by each side (left or right wheel) and by each direction (forward or backward rotation) so as to calibrate the robot movement. The independent gains further allow for the setting of asymmetrical motor rotation to generate a turning bias of the robot (see step 6.1).</li>
	<li>Wash the surface of a white expanded polystyrene ball (mass: approximately 2 g; diameter: 50 mm) with water to remove any possible olfactory or visual cues. 
	Note: The surface of a new ball should be roughed with fine-grit sandpaper, such as P400, which ensures the grip of the legs on the ball.</li>
	<li>Turn on the blower fan that supplies air at 9 V to the treadmill and floats the ball (Figure 5A). Observe the ball float approximately 2 mm from the bottom of the cup.</li>
	<li>Using a screw, attach the copper wire of the attachment with the moth (see step 2) to a fixture in the cockpit of the robot (see Figure 3 inset). Make sure that the position of the middle legs is at the center of the ball (Figure 7A).</li>
	<li>Adjust the vertical position of the attachment to enable the moth to walk normally on the ball. Keep the ball at the same height before and after attaching the moth (Figure 7B). 
	Note: A too-low position of the attachment adds pressure on the moth and elicits backward walking to resist the pressure (Figure 7C), whereas a too-high position causes unstable walking and failures of the sensor due to changes in the vertical position of the ball (Figure 7D). To check normal walking behavior, a single-puffed pheromone stimulus is used to trigger walking in the moth (for the pheromone stimulus, see step 4). Note that the test stimulus must be minimal because previous exposure to bombykol habituates silkmoths and decreases their sensitivity (Matsuyama and Kanzaki, unpublished data).</li>
</ol>
4. Odor Source Preparation
Note: Male B. mori are sensitive to the major component of the conspecific female sex pheromone (bombykol: (E,Z)-10,12-hexadecadien-1-ol)20. Any contamination of experimental equipment with bombykol elicits the odor-tracking behavior and affects the responsiveness of the moth.
<ol>
	<li>Drop 10 &#181;l of the bombykol solution dissolved in n-hexane (200 ng/&#181;l) on a piece of filter paper (approximately 10 mm &#215; 10 mm). The amount of bombykol per piece of filter paper is 2,000 ng. 
	Note: To check the normal walking behavior of the moth, prepare a pheromone stimulus cartridge in this step. The cartridge is a glass Pasteur pipette with one piece of filter paper containing 2,000 ng of bombykol. Pushing a bulb puffs the air containing bombykol.</li>
</ol>
5. Odor Source Localization Experiment
<ol>
	<li>Turn on the fan of a pulling-air-type wind tunnel (1,800 &#215; 900 &#215; 300 mm, L &#215; W &#215; H; Figure 8) and set the wind speed to 0.7 m/sec. Ensure that the temperature is more than 20 &#176;C.</li>
	<li>Set the odor source (the piece of filter paper containing bombykol) upstream of the wind tunnel. 
	Note: The plume width should be confirmed prior to the experiment by using TiCl417,19.</li>
	<li>Turn on the microcontroller board of the robot and establish a serial connection to a PC via Bluetooth.</li>
	<li>Launch a custom-made Java program called &#34;BioSignal,&#34; which provides an interface between the PC and the robot. 
	Note: The main window includes buttons for sending commands to the robot, text windows for displaying the input and output of the serial communication, and small boxes to configure parameters. The subsequent commands are sent by clicking corresponding buttons in this program, except for video capturing.</li>
	<li>Click on the &#34;about device&#34; button to confirm the connection by sending a command to the robot via the specified COM port and check that a message is returned by the robot.</li>
	<li>Click on the &#34;memory erase&#34; button to erase previous locomotion data left on the onboard flash memory.</li>
	<li>Click on the &#34;drivemode1&#34; button to send the default motor gains to the robot. 
	Note: The manipulations of the motor gains and the time delay between insect locomotion and robot movement are applied after this step (see steps 6.1 and 6.3, Figure 9).</li>
	<li>Click on the &#34;don&#39;t drive&#34; button to send a command to immobilize the robot until the experiment starts.</li>
	<li>Put the robot at a start position (600 mm downstream from the odor source) and turn on the switch of the motor driver board.</li>
	<li>Push the recording button of the camcorder to start video capture.</li>
	<li>Click on the &#34;rec start&#34; button to send a start command to initiate the robot with a simultaneous recording of the ball rotation on the onboard flash memory. Observe that the robot starts to move and tracks the odor plume.</li>
	<li>Click on the &#34;rec stop&#34; and &#34;don&#39;t drive&#34; buttons to send commands to stop both the robot movement and the recording if the robot localizes the odor source.</li>
	<li>Push the recording button of the camcorder to stop video capture.</li>
	<li>Download recorded locomotion data from the onboard flash memory to the computer via a serial connection. Close the program.</li>
</ol>
6. Manipulation of the Insect-controlled Robot
Note: The timing of each manipulation is indicated in Figure 9.
<ol>
	<li>Manipulation of motor gains 
	Note: This manipulation alters the translational and rotational velocity of the robot. Asymmetrical motor gains generate a turning bias, which can be used to investigate how insects compensate for the bias17.
	<ol>
		<li>Define the rotational gains for forward and backward rotation of the motor on each side17 (Figure 6B) by editing the configuration file named &#34;param2.txt&#34; using a text editor.</li>
		<li>Click on the &#34;set param2&#34; to read the edited configuration file in the software program. Then, click on the &#34;drivemode2&#34; to send the manipulated gains to the robot.</li>
	</ol>
	</li>
	<li>Inversion of the motor output 
	Note: This manipulation provides a condition similar to the inversion of bilateral olfactory input (see step 6.4) and can be used to investigate the significance of bilateral olfaction. However, the inversion of motor output also inverts self-induced visual motion of an onboard moth. The impact of the inverted self-induced visual input can be evaluated by a comparison with the inverted olfactory input19.
	<ol>
		<li>Invert the bilateral motor control by crossing the control cables for each motor.</li>
	</ol>
	</li>
	<li>Manipulation of the time delay between insect locomotion and robot movement. 
	Note: This manipulation allows for the investigation of the acceptable period of time spent on sensory-motor processing for the robotic odor-tracking. The microcontroller stores the locomotion data on a buffer memory and then processes it after the specified time delay. Note that the robot has a maximal internal time delay of 200 msec; therefore, the actual time delay is expected to be the specified time delay plus 200 msec16,17.
	<ol>
		<li>Input a number (from 0-10) in a small box of the main window to specify a time delay from 0-1,000 msec at 100-msec steps.</li>
		<li>Click on the &#34;set delay&#34; button to apply the time delay.</li>
	</ol>
	</li>
	<li>Manipulation of the olfactory input. 
	Note: This manipulation can be used to investigate the significance of bilateral olfactory input. The surge direction of silkmoths is biased on the higher-concentration side22.
	<ol>
		<li>Change the gap between the suction tube tips or invert their positions to alter the difference in odor concentration acquired by each antenna.</li>
	</ol>
	</li>
	<li>Manipulation of visual input 
	Note: This manipulation is to investigate the role of visual input for odor-tracking.
	<ol>
		<li>Cover the canopy with a white paper that occludes 105&#176; and 90&#176; of the horizontal and vertical visual field of the onboard moth, respectively.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

The most important points for the successful control of the robot by a silkmoth are letting the moth walk smoothly on the air-supported ball and the stably measuring the ball rotation. Therefore, tethering the silkmoth and mounting it on the ball at the appropriate position are the critical steps in this protocol. Inappropriate adhesion of the moth to the attachment or inappropriate positioning of the moth on the ball will cause unnatural pressure on it, which perturbs its normal walking behavior and/or causes a failure ...

Autonomous robots capable of finding an odor source can be important for the safety and security of society. They can be used for the detection of disaster victims, of drugs or explosive materials at an airport, and of hazardous material spills or leaks in the environment. At present, we rely entirely on well-trained animals (e.g., dogs) for these tasks, and robotic odor source localization has been strongly expected to relieve the workload of these animals. Finding an odor source is a challenging task for robots because odorants are distributed intermittently in an atmosphere1; therefore, continuous sampling of the odor concentration gradient is not always possible. Thus, a search strategy using intermittent odor cues is necessary for the achievement of robotic odor source localization2-4.
Odor source localization is essential for organism survival and includes tasks such as finding food, mating partners, and sites for oviposition. To overcome the difficulty in tracking patchy distributed odorants, organisms have evolved various behavioral strategies consisting of two fundamental behaviors: moving upstream during odor reception and cross-stream during cessation of odor reception5,6. These reactive strategies have been well-documented in insects and further combined with other modalities, such as wind direction and vision5-8. The insect behavioral models have also been useful examples for robotics3,9-11, in which behavioral algorithms or neural circuit models are implemented into mobile robots for the evaluation of odor source localization abilities10,12-15. From biomimetic perspectives, this "bottom-up" approach is certainly a fundamental way to develop biomimetic robots. However, the bottom-up approach is not a shortcut to obtaining a useful search strategy, because biological analyses are still ongoing, and the modeling of the sensory-motor systems behind insect behaviors has not been completed. Therefore, it is still unknown how such a biological system actually works as a controller of a robotic platform.
In this article, we demonstrate the protocol of a straightforward "top-down" approach to develop an odor-tracking mobile robot controlled by a biological system16,17. The robot is controlled by a real insect and can be regarded as a prototype of future insect-mimetic robots. In the robot's cockpit, a tethered adult male silkmoth (Bombyx mori) walked on an air-supported ball in response to the female sex pheromone, which was delivered to each antenna through air suction tubes. The ball rotations caused by the walking of the onboard moth were measured by an optical sensor and were translated into the movement of the two-wheeled robot. The advantage of this "hybrid" approach is that experimenters can investigate how the insect sensory-motor system works on the robotic platform where a pilot insect is in a closed loop between the robot and a real odor circumstance. The manipulation of the robotic hardware alters the closed loop; therefore, the insect-controlled robot is a useful platform for both engineers and biologists. For engineering, the robot represents the first steps of applying a biological model to meet the requirements for robotic tasks. For biology, the robot is an experimental platform for studying sensory-motor control under a closed loop.

<table border="0" cellpadding="0" cellspacing="0" width="961">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="43">
			<td height="43" style="height:43px;width:267px;">Male adult silkmoth 
			&#160;(Bombyx mori)</td>
			<td style="width:159px;"></td>
			<td style="width:171px;"></td>
			<td style="width:365px;">Rear from eggs, or purchase as pupae.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Incubator</td>
			<td style="width:159px;">Panasonic</td>
			<td style="width:171px;">MIR-254</td>
			<td style="width:365px;">Store pupae or adult silkmoths at a constant temperature, 238 L.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Plastic box</td>
			<td style="width:159px;">Sunplatec</td>
			<td style="width:171px;">O-3</td>
			<td style="width:365px;">Store pupae or adult silkmoths, 299 &#215; 224 &#215; 62 mm L &#215; W &#215; H.</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:267px;">Copper wire</td>
			<td style="width:159px;"></td>
			<td style="width:171px;"></td>
			<td style="width:365px;">2-mm diameter for the attachment. Any rigid bar can be used as an alternative for making the attachment to tether a silkmoth.&#160;</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Plastic sheet</td>
			<td style="width:159px;">Kokuyo</td>
			<td style="width:171px;">VF-1420N</td>
			<td style="width:365px;">Sold as overhead projector film with thickness of 0.1 mm. Use at the tip of the attachment.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Forceps</td>
			<td style="width:159px;">As one</td>
			<td style="width:171px;">5SA</td>
			<td style="width:365px;">Remove scales on the thorax.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Adhesive</td>
			<td style="width:159px;">Konishi</td>
			<td style="width:171px;">G17</td>
			<td style="width:365px;">Bond a silkmoth to the attachment.</td>
		</tr>
		<tr height="93">
			<td height="93" style="height:93px;width:267px;">Insect-controlled robot</td>
			<td style="width:159px;">Custom</td>
			<td style="width:171px;"></td>
			<td style="width:365px;">Bearing an air-supported treadmill, an optical sensor, custom-built AVR-based microcontroller boards, and two DC brushless motors. It is powered by 8 &#215; AA and 3 &#215; 006P batteries.</td>
		</tr>
		<tr height="28">
			<td height="28" style="height:28px;width:267px;">Microcontroller</td>
			<td style="width:159px;">Atmel</td>
			<td style="width:171px;">ATMEGA8</td>
			<td style="width:365px;">A component of the insect-controlled robot.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">DC blower</td>
			<td style="width:159px;">Nidec</td>
			<td style="width:171px;">A34342-55</td>
			<td style="width:365px;">A component of the insect-controlled robot for floating a ball in an air-supported treadmill.&#160;</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:267px;">DC fan</td>
			<td style="width:159px;">Minebea</td>
			<td style="width:171px;">1606KL-04W-B50</td>
			<td style="width:365px;">A component of the insect-controlled robot for suctioning air containing an odor.</td>
		</tr>
		<tr height="65">
			<td height="65" style="height:65px;width:267px;">Optical mouse sensor</td>
			<td style="width:159px;">Agilent technologies</td>
			<td style="width:171px;">HDNS-2000</td>
			<td style="width:365px;">A component of the insect-controlled robot, obtained from an optical mouse (M-GUWSRSV, Elecom, Japan).</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Brushless motor</td>
			<td style="width:159px;">Maxon</td>
			<td style="width:171px;">EC-45</td>
			<td style="width:365px;">A component of the insect-controlled robot for driving a wheel.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">White polystyrene ball</td>
			<td style="width:159px;"></td>
			<td style="width:171px;"></td>
			<td style="width:365px;">A component of the insect-controlled robot. Diameter 50 mm, mass approximately 2 g.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Bombykol: 
			&#160;(E,Z)-10,12-hexadecadien-1-ol</td>
			<td style="width:159px;">Shin-Etsu chemical</td>
			<td style="width:171px;"></td>
			<td style="width:365px;">Custom synthesis.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">n-hexane</td>
			<td style="width:159px;">Wako</td>
			<td style="width:171px;">085-00416</td>
			<td style="width:365px;">Solvent for bombykol.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:267px;">Wind tunnel</td>
			<td style="width:159px;">Custom</td>
			<td style="width:171px;"></td>
			<td style="width:365px;">Pulling-air type, sized 1800 &#215; 900 &#215; 300 mm L &#215; W &#215; H.</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:267px;">BioSignal program</td>
			<td style="width:159px;">Custom</td>
			<td style="width:171px;"></td>
			<td style="width:365px;">A program to establish serial communication between the insect-controlled robot and a PC via Bluetooth. Used for sending commands to start/stop the robot or configuring its motor properties.&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:267px;">Camcorder</td>
			<td style="width:159px;">Sony</td>
			<td style="width:171px;">HDR-XR520V</td>
			<td style="width:365px;">Capture robot movements.</td>
		</tr>
	</tbody>
</table>

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.

We present here the basic characteristics of the insect-controlled robot required for the successful localization of an odor source. The comparison between the robot and silkmoths, the effectiveness of the odor delivery system, and the significance of accurate bilateral olfactory and visual inputs are examined.
The comparison of odor-tracking behaviors between freely-walking moths and the insect-controlled robot is shown in <str...

Watch this Scientific Journal Video about Insect-controlled Robot: A Mobile Robot Platform to Evaluate the Odor-tracking Capability of an Insect at JoVE.com

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

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 ...

insect-controlled-robot-mobile-robot-platform-to-evaluate-odor

Research

JoVE Journal

Neuroscience

14.6K Views.  The University of Tokyo. 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.

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

An Experimental Platform to Study the Closed-loop Performance of Brain-machine Interfaces

The non-stationary nature and variability of neuronal signals is a fundamental problem in brain-machine interfacing. We developed a brain-machine interface to assess the robustness of different control-laws applied to a closed-loop image stabilization task. Taking advantage of the well-characterized fly visuomotor pathway we record the electrical activity from an identified, motion-sensitive neuron, H1, to control the yaw rotation of a two-wheeled robot. The robot is equipped with 2 high-speed video cameras providing visual motion input to a fly placed in front of 2 CRT computer monitors. The activity of the H1 neuron indicates the direction and relative speed of the robot's rotation. The neural activity is filtered and fed back into the steering system of the robot by means of proportional and proportional/adaptive control. Our goal is to test and optimize the performance of various control laws under closed-loop conditions for a broader application also in other brain machine interfaces.

We use a closed-loop fly-machine interface to investigate general principles in neuronal control.

The non-stationary nature and variability of neuronal signals is a fundamental problem in brain-machine interfacing. We developed a brain-machine ...

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

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

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

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

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

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 ...

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 ...

Fabrication of High Contact-Density, Flat-Interface Nerve Electrodes for Recording and Stimulation Applications

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic applications. This protocol describes a fabrication technique of a modified cylindrical nerve cuff electrode to meet these criteria. Minimum computer-aided design and manufacturing (CAD and CAM) skills are necessary to consistently produce cuffs with high precision (contact placement 0.51 &#177; 0.04 mm) and various cuff sizes. The precision in spatially distributing the contacts and the ability to retain a predefined geometry accomplished with this design are two criteria essential to optimize the cuff&#39;s interface for selective recording and stimulation. The presented design also maximizes the flexibility in the longitudinal direction while maintaining sufficient rigidity in the transverse direction to reshape the nerve by using materials with different elasticities. The expansion of the cuff&#39;s cross sectional area as a result of increasing the pressure inside the cuff was observed to be 25% at 67 mm Hg. This test demonstrates the flexibility of the cuff and its response to nerve swelling post-implant. The stability of the contacts&#39; interface and recording quality were also examined with contacts&#39; impedance and signal-to-noise ratio metrics from a chronically implanted cuff (7.5 months), and observed to be 2.55 &#177; 0.25 k&#8486; and 5.10 &#177; 0.81 dB respectively.

This article provides a detailed description on the fabrication process of a high contact-density flat interface nerve electrode (FINE). This electrode is optimized for recording and stimulating neural activity selectively within peripheral nerves.

Many attempts have been made to manufacture multi-contact nerve cuff electrodes that are safe, robust and reliable for long term neuroprosthetic ...

An Improved Method for Collection of Cerebrospinal Fluid from Anesthetized Mice

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central nervous system and thus, can be used to analyze the diseased state of the brain or spinal cord without directly accessing these tissues. However, in mice it is difficult to obtain from the cisterna magna due to its closeness to blood vessels, which often contaminate samples. The area for CSF collection in mice is also difficult to dissect to and often only small samples are obtained (maximum of 5-7 &#181;L or less). This protocol describes in detail a technique that improves on current methods of collection to minimize contamination from blood and allow for the abundant collection of CSF (on average 10-15 &#181;L can be collected). This technique can be used with other dissection methods for tissue collection from mice, as it does not impact any tissues during CSF extraction. Thus, the brain and spinal cord are not affected with this technique and remain intact. With greater CSF sample collection and purity, more analyses can be used with this fluid to further aid neuroscience research and better understand diseases affecting the brain and spinal cord.

This protocol describes an improved technique for the abundant collection of cerebrospinal fluid (CSF) with no contamination from blood. With greater sample collection and purity, more analyses can be performed using CSF to further our understanding of diseases that affect the brain and spinal cord.

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central ...

A Rat Model of Central Fatigue Using a Modified Multiple Platform Method

In this article, we introduced a rat model of central fatigue using the modified multiple platform method (MMPM). The Multiple Platform box was designed as a water tank with narrow platforms on the bottom. The model rats were put into the tank and stood on the platforms for 14 h (18:00 - 8:00) per day for a consecutive 21 days, with a blank control group set for contrast. At the end of modeling, rats in the model group showed an obvious fatigued appearance. To assess the model, we performed several behavioral tests, including the open field test (OFT), the elevated plus maze (EPM) test, and the exhaustive swimming (ES) test. The results showed that anxiety, spatial cognition impairment, poor muscle performance, and declined voluntary activity presented in model rats confirm the diagnosis of&#160;central fatigue. Changes of the central neurotransmitters also verified the result. In conclusion, the model successfully simulated central fatigue, and future study with the model may help reveal the pathological mechanism of the disease.

Here, we present a protocol to introduce a rat model of central fatigue using the modified multiple platform method (MMPM).

In this article, we introduced a rat model of central fatigue using the modified multiple platform method (MMPM). The Multiple Platform box was designed ...

A Neurosphere Assay to Evaluate Endogenous Neural Stem Cell Activation in a Mouse Model of Minimal Spinal Cord Injury

Neural stem cells (NSCs) in the adult mammalian spinal cord are a relatively mitotically quiescent population of periventricular cells that can be studied in vitro using the neurosphere assay. This colony-forming assay is a powerful tool to study the response of NSCs to exogenous factors in a dish; however, this can also be used to study the effect of in vivo manipulations with the proper understanding of the strengths and limitations of the assay. One manipulation of the clinical interest is the effect of injury on endogenous NSC activation. Current models of spinal cord injury provide a challenge to study this as the severity of common contusion, compression, and transection models cause the destruction of the NSC niche at the site of the injury where the stem cells reside. Here, we describe a minimal injury model that causes localized damage at the superficial dorsolateral surface of the lower thoracic level (T7/8) of the adult mouse spinal cord. This injury model spares the central canal at the level of injury and permits analysis of the NSCs that reside at the level of the lesion at various time points following injury. Here, we show how the neurosphere assay can be utilized to study the activation of the two distinct, lineally-related, populations of NSCs that reside in the spinal cord periventricular region - primitive and definitive NSCs (pNSCs and dNSCs, respectively). We demonstrate how to isolate and culture these NSCs from the periventricular region at the level of injury and the white matter injury site. Our post-surgical spinal cord dissections show increased numbers of pNSC and dNSC-derived neurospheres from the periventricular region of injured cords compared to controls, speaking to their activation via injury. Furthermore, following injury, dNSC-derived neurospheres can be isolated from the injury site &#8212; demonstrating the ability of NSCs to migrate from their periventricular niche to sites of injury.

Here, we demonstrate the performance of a minimal spinal cord injury model in an adult mouse that spares the central canal niche housing endogenous neural stem cells (NSCs). We show how the neurosphere assay can be used to quantify activation and migration of definitive and primitive NSCs following injury.

Neural stem cells (NSCs) in the adult mammalian spinal cord are a relatively mitotically quiescent population of periventricular cells that can be studied ...

A Volumetric Method for Quantification of Cerebral Vasospasm in a Murine Model of Subarachnoid Hemorrhage

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor determining patient outcome and is therefore frequently taken as a study endpoint. However, in small animal studies on SAH, quantification of cerebral vasospasm is a major challenge. Here, an ex vivo method is presented that allows quantification of volumes of entire vessel segments, which can be used as an objective measure to quantify cerebral vasospasm. In a first step, endovascular casting of the cerebral vasculature is performed using a radiopaque casting agent. Then, cross-sectional imaging data are acquired by micro computed tomography. The final step involves 3-dimensional reconstruction of the virtual vascular tree, followed by an algorithm to calculate center lines and volumes of the selected vessel segments. The method resulted in a highly accurate virtual reconstruction of the cerebrovascular tree shown by a diameter-based comparison of anatomical samples with their virtual reconstructions. Compared with vessel diameters alone, the vessel volumes highlight the differences between vasospastic and non-vasospastic vessels shown in a series of SAH and sham-operated mice.

The goal of this article is to present a method that allows a 3-dimensional reconstruction of the cerebrovascular tree in mice after micro computed tomography and determination of volumes of entire vessel segments that can be used to quantify cerebral vasospasm in murine models of subarachnoid hemorrhage.

Subarachnoid hemorrhage (SAH) is a subtype of hemorrhagic stroke. Cerebral vasospasm that occurs in the aftermath of the bleeding is an important factor ...

Functional MRI in Conjunction with a Novel MRI-compatible Hand-induced Robotic Device to Evaluate Rehabilitation of Individuals Recovering from Hand Grip Deficits

Functional magnetic resonance imaging (fMRI) is a non-invasive magnetic resonance imaging technique that images brain activation in vivo, using endogenous deoxyhemoglobin as an endogenous contrast agent to detect changes in blood-level-dependent oxygenation (BOLD effect). We combined fMRI with a novel robotic device (MR-compatible hand-induced robotic device [MR_CHIROD]) so that a person in the scanner can execute a controlled motor task, hand-squeezing, which is a very important hand movement to study in neurological motor disease. We employed parallel imaging (generalized auto-calibrating partially parallel acquisitions [GRAPPA]), which allowed higher spatial resolution resulting in increased sensitivity to BOLD. The combination of fMRI with the hand-induced robotic device allowed precise control and monitoring of the task that was executed while a participant was in the scanner; this may prove to be of utility in rehabilitation of hand motor function in patients recovering from neurological deficits (e.g., stroke). Here we outline the protocol for using the current prototype of the MR_CHIROD during an fMRI scan.

We performed functional MRI using a novel MRI-compatible hand-induced robotic device to evaluate its utility for monitoring hand motor function in individuals recovering from neurological deficits.

Functional magnetic resonance imaging (fMRI) is a non-invasive magnetic resonance imaging technique that images brain activation in vivo, using endogenous ...