We thank Patrice Jegouzo for the head devices and headpost development and production. We also thank P. Calvo, A. Mialot, and E. Idoux for their help in the development of previous versions of the device and VVM protocol.
This work was funded by the Centre National des Etudes Spatiales, the CNRS, and the Universit&#233; Paris Descartes. J. C. and M. B. receive support from the French ANR-13-CESA-0005-02. F. F. B. and M. B. receive support from the French ANR-15-CE32-0007.

All animal procedures followed the Paris Descartes University animal regulations.
1. Device assembly
NOTE: The device used in this protocol is a helmet-like structure fixed on mice skulls by means of an implanted headpost.
<ol>
	<li>Using a 3D printer and white opaque poly (lactic acid) (PLA) plastic, print using the design and specification files provided here (see Table of Materials) for both the device and headpost. 
	NOTE: The dimensions of the device are shown in Figure 1 and dimensions of the headpost shown in Figure 2.</li>
	<li>A striped as well as sham device are to be tested (Figure 2A11). To obtain the striped model, using black nail polish, draw 3 mm large vertical stripes on the external surface of the device. The sham condition does not require any modification to the printed device.</li>
</ol>
2. Headpost implantation surgery
All the materials used in this protocol are detailed in the materials list in the supplementary information. Steps 2.7-2.9 use the biomaterials provided in the implantation kit (see Table of Materials). Ensure the use of sterile instruments and arrange surgery and recovery in different zones. Once mastered, the implantation procedure lasts about 30 min.
<ol>
	<li>For analgesia, 30 min before the beginning of the surgery, subcutaneously inject buprenorphine (0.05 mg/kg) and put back the animal in its home cage. 
	NOTE: Buprenorphine&#39;s analgesic effects last approximately 12 h, long after the end of the procedure. In our experience, mice do not show any signs of distress related to this intervention&#160;but a subsequent dose of 0.05mg/kg buprenorphine is recommended 24h after the surgery.&#160;</li>
	<li>Anesthetize the animal in a chamber with 2.5%-3% isoflurane gas. Wait 3 min and check if the mouse is properly anesthetized by observing respiration and lack of movement inside the chamber. Pass the mouse to a nose cone on a surgical table with a heating pad and, by interdigital pinching, verify that there is no withdrawal reflex and lower the isoflurane to 1.5%.</li>
	<li>Shave the head of the mouse using an electric razor. To obtain a sterile environment, rub the shaved area with iodine solution and after with 70% alcohol. Repeat this procedure two more times.</li>
	<li>Inject lidocaine hydrochloride (2%, 2 mg/kg) under the skin of the head for local anesthesia and wait 5 min for the effects to begin. To avoid eye damage due to dryness, cover the mouse&#39;s eyes with topical ophthalmic vet ointment.</li>
	<li>With a pair of blunt forceps, grab the skin at the back of the head, and with a pair of blunt scissors (or scalpel), make a longitudinal incision of about 1.5 cm to expose the skull.</li>
	<li>With the help of a scalpel, scratch the periosteum. Be careful not to scratch too hard, as the fixation of the headpost can be compromised if the skull starts to bleed slightly.</li>
	<li>Apply a drop of the green activator on the middle of the skull. This will improve the fixation of the cement by increasing bone permeability.</li>
	<li>Prepare the cement: mix one spoon (provided in the implantation kit) of polymer with five drops of monomer and one drop of catalyzer. With the help of a brush, apply a generous amount of the cement mix between the lambda and bregma skull landmarks;</li>
	<li>Quickly place the headpost on the cement with a swiping motion going from lambda to bregma. After the headpost has been placed, reapply more cement around the inferior part to ensure that the headpost properly sticks to the skull. To guarantee proper fixation, make sure the cement is applied abundantly and that it dries before continuing to the next step. 
	NOTE: With this fixation procedure, the headpost will not come off and allow for long-term, repeated tests; in our hands, headpost removal is &#60;10%.</li>
	<li>Prepare the resin mix by applying a powder-to-liquid ratio that enables a smooth consistency of the mixture. Apply the resin where the cement was applied as well as around the headpost in order to protect its surface.</li>
	<li>Wait 3 min for the resin to dry and close the skin at the back of the ears with monofilament suture. With a cotton swab, apply diluted (10%-20%) iodine solution to the operated area. 
	NOTE: Make sure the skin does not get stuck to the resin.</li>
	<li>Turn off the anesthesia and place the animal under a red warm light to avoid hypothermia. Place moistened food and&#160;hydrogel or another water source based in gel in the cage&#39;s floor. Do not leave the mouse unattended until it regains consciousness. As soon as the animal fully recovers from the procedure (usually, 30 min to 1 h after), place it in a cage with groups of three or four to stimulate social interactions.</li>
</ol>
3. Device fixation
<ol>
	<li>48 h following the surgery, secure the custom-built head device onto the headpost.
	<ol>
		<li>Using a pair of 1.2 mm screws and a screwdriver (1.3 mm hex), align the holes in the striped device with the holes in the headpost, place the screws and secure them. To fix the sham condition, turn the device upside down and, with the back part (Figure 1A) of the device facing the rostral direction, align the holes in the device with the holes in the headpost. 
		NOTE: It is recommended that this step be done by two operators, one holding the mouse with a one-handed mouse restraint, while the other securing the device to the headpost. If the fixation is done by a single operator, the device can be placed while the mouse is under gas anesthesia.</li>
		<li>Check that the device is well-secured and cannot be removed by the animal and that the device does not apply pressure directly on the mouse&#39;s nose, which could potentially cause pain, difficulty to breath, or skin injury. 
		NOTE: It is also important to ensure the device is symmetrically inserted on the mouse face, so that eyes are completely covered by the head device. Check that the animal does not show any signs of abnormal pain or distress.</li>
	</ol>
	</li>
	<li>Leave the device on the mouse for 14 days.</li>
</ol>
4. Animal care and surveillance
<ol>
	<li>Once back in their cages, mice will exhibit certain abnormalities in behavior. At first, the animal may stay prostrated and try to remove the device using its forepaws, but this should stop after the first hour. During the next following hours, the animal will usually display difficulties orienting itself inside the cage and reaching for food and water. Therefore, during the 48 h following implantation, monitor the mice and provide easy access to water and food, by placing both directly on the cage floor, for example.</li>
	<li>Keep track of mice&#39;s weights during the duration of the protocol. Weigh the mice right after implantation and again every 24 h. Special attention should be given to animals wearing the striped device, as they normally experience body weight loss (1-2 g) during the first 48 h, but start gaining weight again at a normal pace following that initial period (see Figure 2B11).</li>
	<li>After 2 days, mice are expected to return to their regular faculties. Depending on the system used in animal facilities, the device might be preventing access to the food and water. Ensure the animal is at ease while eating and drinking or adapt the dispensing system accordingly. 
	NOTE: The range of head movements produced by the animals after a few days with the device on is not modified by the device (see Figure 211) (i.e., the range of head movements produced remains similar to natural head movements).</li>
	<li>To further ensure mice&#39;s well-being, ensure daily surveillance and apply the qualitative scale (Table 1) of well-being throughout the duration of the protocol.</li>
	<li>Remove a mouse from the ongoing protocol if one or more of the following criteria apply:
	<ol>
		<li>Mice that have a total score higher than 4 points on the aforementioned qualitative scale must immediately be excluded from the experiment (see Table 1). Regardless of the score, if the mouse does not regain its initial weight after 6 days, the procedure must be stopped.</li>
		<li>The device is not correctly fixed to the headpost if, for instance, the headpost shakes when touched or a part starts to come off. This causes the headpost to come off the mouse&#39;s head and consequently interrupts the learning, which explains why daily surveillances are necessary.</li>
		<li>When a mouse has its headpost ripped off during any part of the protocol. Due to the skull bleeding associated to this detachment, the reimplantation surgery has a low success rate and is not worth attempting.</li>
	</ol>
	</li>
</ol>
5. Removal of the device
<ol>
	<li>After the learning period (in this protocol 14 days), remove the device following the same instructions as for its fixation (section 3). As soon as the device is removed, test the mice with experiments such as video-oculography tests, or, for instance, with in vitro electrophysiology as described previously11. 
	NOTE: As soon as the device is taken off, mice are exposed back to the standard, visually unobstructed environment. Therefore, perform experiments that aim to test the learning effects of this device directly after its removal.</li>
</ol>
6. Video-oculography sessions
NOTE: Video-oculography experiments are performed to record the generated eye movements while the animal is being rotated in the dark (vestibulo-ocular reflex, VOR) or by rotating the animal&#39;s surroundings while the animal is still (optokinetic reflex, OKR). Each mouse was tested for both these reflexes before and after the adaptation protocol. For more details about the video-oculography set-up, see previously published reports12,13.&#160;In order to habituate the mice to the restrained recording conditions, the day before the beginning of the recording, place the animal on the tube at the center of the turntable for 10minutes without performing any test.
<ol>
	<li>Secure the mouse on the turntable by head-fixing it with the help of screws inserted into the headpost. Place a screen dome surrounding the animal and turn off all the lights in the room except for the optokinetic projector. 
	NOTE: Video-oculography recordings require the animal to be still and with its eyes open. Interrupt the recording session and put the animal back on its cage in case the mouse does not voluntarily keep its eyes open, or if the appearance of the eye deteriorates during the recording session. Another attempt can be made following a resting period of at least 12h.</li>
	<li>Start the OKR full-field stimulation (white dot pattern projection) and record at several different velocities in both clockwise and counter clockwise directions. As soon as the recordings are over, remove the dome.</li>
	<li>To be able to record the VOR in pitch dark, apply a drop of 2% pilocarpine to the eyes14. Wait at least 5 min for it to act and gently remove it with a cotton swab. The pilocarpine will keep the pupil constricted with a constant size throughout the measurements, allowing proper quantification of movements in the dark.</li>
	<li>Turn off all the lights in the room and add a box on top of the turntable to keep the animal in pitch dark. Start the horizontal VOR using sinusoidal angular rotations around a vertical axis with different frequencies and/or different velocities.</li>
	<li>Once the recording session is finished, return the mouse to a cage properly illuminated with an infrared lamp. The heat will prevent hypothermia caused by the secondary vasodilator effects of pilocarpine on the body of the mouse. 
	NOTE: Due to the animal being restrained, recording sessions can&#8217;t last more than 90 min. When additional test sessions are needed, let the animal rest for 24h between sessions.</li>
</ol>

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The authors declare no conflicts of interest.

The long-term sensory perturbation described here consists of a visuo-vestibular mismatch produced in freely-behaving mice. To implant the device that mice wear for 14 days, a simple and short surgery using a commercially available surgical kit is performed. Mice recover in less than 1 h from this headpost implantation procedure and show no associated signs of distress from it. Subsequently, in the given example of application of this protocol, VOR and OKR are measured using the video-oculography technique. Nonetheless, this device-induced long-term learning protocol could be used in a variety of experiments such as in vitro electrophysiology1, neuronal imaging, and various behavioral assays. The rationale behind the development of this technique was inspired by the prism-based methodology used in humans and monkeys. This technique, however, differs because it impairs rather than modifies vision. Thus, it constitutes (in its current form) an extreme case of visuo-vestibular mismatch. The authors believe that the provided technical information may be useful for designing a prism-like version of the device or further developing specific feature-restricting devices16.
Made of a light (0.9 g) poly (lactic acid) plastic, the head device was designed to fit the head of a young adult mouse, allowing protection of the snout and leaving enough space laterally to let the animal groom. The front part of this device exposes the end of the snout to permit feeding and grooming behaviors. The device is slightly opaque, so that the animal is deprived of precise vision of the surrounding but still receives luminance stimulation. The striped and sham implantations are tested to ensure that the measured effects are due primarily to the visuo-vestibular mismatch caused by the high-contrast visual signal during self-generated movements of the striped device and not by proprioceptive modification (i.e, the weight of the device applied in the mouse&#180;s head and neck).
Experimentally, mice that wore the striped device showed a significant VOR gain decrease of 50% after the learning period; still, there can be an inter-individual variability for absolute gain values. Sham mice showed no significant VOR gain alterations, thus demonstrating that the VOR reduction is caused by the sensory conflict and not by motor impairment. Furthermore, young mice (&#60;P26) showed VOR and OKR gain values lower than older animals17. For that reason, animal age has to be taken into account while planning the experiment. Finally, the aforementioned mice exclusion criteria (section 4.5) are a crucial step that should be followed to ensure well-being as well as establish reliable results.
One of the advantages of this protocol is the time that it saves experimenters during the learning period, compared to other types of VOR/OKR adaptation protocols. So far, VOR adaptation in mice has been studied by head-fixing and training the animal on a rotating turntable6,8,18,19, which is time-consuming, especially when a lot of animals must be trained. The presented protocol allows the training of several animals at once and saves time. In addition, in these classical experiments the trainings are typically limited to 1 h per day, leaving long periods of putative unlearning that cause adaptation to be an iterated alternation of learning/unlearning with different dynamics20. Here, the head-fixation of the device allows for uninterrupted learning. Another advantage is that since the learning period is generated in a freely behaving head-free situation, mice are able to learn through a range of natural head movements that are actively generated. In the classical protocols, the animal is head-fixed while being passively rotated on the turntable so that the learning occurs at a determined stimulation (one frequency, one velocity)21 that does not reflect the natural range of head movements. It is important to note that the vestibular system encodes movements differently when they are actively generated by the subject or when externally applied10; thus, the cellular mechanisms triggered in both situations may also differ.
Overall, the described methodology is suitable for combined in vivo/in vitro studies on long-term sensory adaptations occurring after a visual conflict and/or visuo-vestibular mismatch in freely behaving mice. Sensory conflicts are a recognized cause of motion sickness, which is a field that has recently attracted use of mice22,23. It was recently demonstrated that the gain adaptation caused by the use of this device offers protection against motion sickness when mice are exposed to a provocative stimulus15. Hence, this protocol could be used to identify the cellular mechanisms underlying adaptation to a sensory conflict as well as to develop anti-motion sickness treatments.

Sensory conflicts, such as visual ones, are present in daily life, for instance, when one wears glasses or during an entire lifespan (developmental growth, changes in sensory acuity, etc.). Due to a well-described circuit anatomy, easily controlled sensory inputs, quantifiable motor outputs, and precise quantification methods1, gaze stabilization reflexes have been used as models of motor learning in many species. In humans and monkeys, the vestibulo-ocular reflex (VOR) adaptation is studied through the use of prisms that the subject wears for several days2,3,4,5. Since the rodent model allows the combination of behavioral and cellular experiments, we developed a new method to create long-term sensory conflict in freely behaving mice with a helmet-like device. Inspired by the methodology used in humans and monkeys, the protocol generates a mismatch between the vestibular and visual inputs (i.e., visuo-vestibular mismatch, VVM) that leads to a decrease in VOR gain.
Classical protocols triggering a VOR gain-down adaptation in rodents consist of rotating the head-fixed animal on a turntable while rotating the visual field in phase. This paradigm creates a visuo-vestibular conflict, which makes the VOR counter-productive. Long-term adaptation protocols consist of an iteration of this procedure over the course of several consecutive days6,7,8. As a result, when a large group of animals needs to be tested, classical methodology requires a great amount of time. In addition, because the animal is head-fixed, the learning is mostly limited to a discrete frequency/velocity and consist of discontinuous trainings interrupted by intertrial intervals of variable duration6. Finally, classical protocols use passive learning, as the vestibular stimulation is not actively generated by the animal&#39;s voluntary movements, a situation that greatly shapes vestibular processing9,10.
The aforementioned experimental constraints are surpassed by the presented innovative methodology. The required surgical approach is straightforward, and the materials used are readily available commercially. The sole part that relies on more expensive material is the quantification of the behavior; nonetheless, the fundamentals of the protocol may be used for any experiment, from in vitro investigations to other behavioral studies of learning. Overall, by generating a temporary visual impairment and a visuo-vestibular conflict over several days, this methodology can easily be transposed to any study concerned with sensory perturbation or motor learning.

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	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:216px;">3D printer</td>
			<td style="width:123px;">Ulimaker, USA</td>
			<td style="width:188px;">S5</td>
			<td style="width:223px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Blunt scissors</td>
			<td>FST</td>
			<td>14079-10</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Catalyst V</td>
			<td>Sun Medical, Japan</td>
			<td>LX22</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Dentalon Plus</td>
			<td>Heraeus</td>
			<td>37041</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Eyetracking system and software</td>
			<td>Iscan</td>
			<td>ETN200</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Green activator</td>
			<td>Sun Medical, Japan</td>
			<td>VE-1</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Monomer</td>
			<td>Sun Medical, Japan</td>
			<td>MF-1</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Ocrygel</td>
			<td>TvmLab</td>
			<td>10779</td>
			<td>Ophtalmic vet ointment</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Polymer L-type clear (cement)</td>
			<td>Sun Medical, Japan</td>
			<td>TT12F</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sketchup</td>
			<td>Trimble</td>
			<td></td>
			<td>3D modeling software used for the device&#39;s ready-to-print design file</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Turntable</td>
			<td></td>
			<td></td>
			<td>Not commercially available</td>
		</tr>
	</tbody>
</table>

long-term sensory conflict in freely behaving mice

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning in mice. By permanently wearing a device fixed on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages. Therefore, this protocol readily enables the study of the visual system and multisensory interactions over an extended timeframe that would not be accessible otherwise. In addition to lowering the experimental costs of long-term sensory learning in naturally behaving mice, this approach accommodates the combination of in vivo and in vitro experiments. In the reported example, video-oculography is performed to quantify the vestibulo-ocular reflex (VOR) and optokinetic reflex (OKR) before and after learning. Mice exposed to this long-term sensory conflict between visual and vestibular inputs presented a strong VOR gain decrease but exhibited few OKR changes. Detailed steps of device assembly, animal care, and reflex measurements are hereby reported.

The following figures illustrate the results obtained with mice that underwent the 2 week adaptation protocol wearing either a striped or sham device. Figure 3 shows an example of raw traces seen during recording sessions. As shown by comparing the traces, the VOR response decreases after the VVM protocol (Figure 3A, before vs. after striped). The VOR of sham mice remained unaltered after the adaptation (Figure 3A, before vs. after sham). The OKR of mice wearing the striped device (Figure 3B) is comparable to the period prior to the VVM protocol and to sham mice. Figure 4 shows a quantification example of the mean VOR gains at a fixed frequency of 0.5 Hz and at 40 degrees per second, before and after the VVM protocol, for both striped and sham devices. There is a strong gain decrease after mice wore the striped device, while the sham mice did not have significant gain changes. Effects of VOR decrease tested at different velocities/frequencies have been reported by Carcaud et al.11 and Idoux et al.15.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59135/59135fig1.jpg" /> 
Figure 1: Head device depicted with dimensions, in millimeters. Views: (A) back, (B) side, (C) bottom, and (D) aerial. <a href="https://www.jove.com/files/ftp_upload/59135/59135fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59135/59135fig2.jpg" /> 
Figure 2: Headpost depicted with dimensions, in millimeters. Fixed in the implantation surgery, this light (0.2 g) poly (lactic acid) plastic headpost allows the locking of the adaptation device to the mouse and head-fixing of the animal on the turntable during the video-oculography sessions. <a href="https://www.jove.com/files/ftp_upload/59135/59135fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59135/59135fig3v2.jpg" /> 
Figure 3: Example raw traces of eye movements during VOR and OKR stimulations. (A, left) Left: VOR performed at 0.5 Hz at 40 &#176;/s and (B, right) optokinetic stimulation at a constant velocity of 10 &#176;/s (black line), in a clockwise direction, before (green lines) and after (yellow) wearing the striped or sham (purple) device. <a href="https://www.jove.com/files/ftp_upload/59135/59135fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59135/59135fig4v2.jpg" /> 
Figure 4: Example mean VOR and OKR gain values after adaptation to either striped or sham device. Gains were plotted according to time (days) for the striped (n = 10) and sham (n = 6) devices at stimulations of 40&#176;/s and 0.5 Hz for the VOR (left), and 10 &#176;/s clockwise direction for the OKR (right). On the timescale, &#34;before&#34; day represents the day immediately prior to the adaptation and &#34;day 0&#34; represents the day when the device is removed. Error bars represent the standard deviation, ***p &#60; 0.001, not significant. <a href="https://www.jove.com/files/ftp_upload/59135/59135fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Points</td>
			<td>Body weight alterations</td>
			<td>Physical appearance</td>
			<td>Behavior</td>
		</tr>
		<tr>
			<td>0</td>
			<td>none or weight gain</td>
			<td>standard</td>
			<td>no signs of distress and normal locomotion</td>
		</tr>
		<tr>
			<td>1</td>
			<td>weight loss &#60;10%</td>
			<td>no body grooming</td>
			<td>impaired locomotion or cage orientation</td>
		</tr>
		<tr>
			<td>2</td>
			<td>weight loss between 10%-20%</td>
			<td>dehydration</td>
			<td>--</td>
		</tr>
		<tr>
			<td>3</td>
			<td>weight loss &#62;20%</td>
			<td>wounds</td>
			<td>nervous ticks (e.g. scratching, biting)</td>
		</tr>
	</tbody>
</table>
Table 1: Qualitative scale for the well-being assessment. Listed are the qualitative parameters that must be assessed during the duration of the protocol. The sum weight alterations, physical appearance, and behavior scores should not be greater than four points.
Supplemental File 1. Device.stl.&#160;<a href="https://www.jove.com/files/ftp_upload/59135/Device.stl" target="_blank">Please click here to download this file.</a>
Supplemental File 2. Headpost.stl.&#160;<a href="https://www.jove.com/files/ftp_upload/59135/Headpost.stl" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Long-term Sensory Conflict in Freely Behaving Mice at JoVE.com

Long-term Sensory Conflict in Freely Behaving Mice

The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning. By permanently wearing a fixed device on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages.

Long-term sensory conflict protocols are a valuable means of studying motor learning. The presented protocol produces a persistent sensory conflict for ...

long-term-sensory-conflict-in-freely-behaving-mice

Research

JoVE Journal

Neuroscience

6.5K Views.  CNRS. The presented protocol produces a persistent sensory conflict for experiments aimed at studying long-term learning. By permanently wearing a fixed device on their heads, mice are continuously exposed to a sensory mismatch between visual and vestibular inputs while freely moving in home cages.

Video: Long-term Sensory Conflict in Freely Behaving Mice

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

Long-term Behavioral Tracking of Freely Swimming Weakly Electric Fish

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and social interactions can be best studied by observing unrestrained, freely behaving animals. Weakly electric fish (WEF) display readily observable exploratory and social behaviors by emitting electric organ discharge (EOD). Here, we describe three effective techniques to synchronously measure the EOD, body position, and posture of a free-swimming WEF for an extended period of time. First, we describe the construction of an experimental tank inside of an isolation chamber designed to block external sources of sensory stimuli such as light, sound, and vibration. The aquarium was partitioned to accommodate four test specimens, and automated gates remotely control the animals&#39; access to the central arena. Second, we describe a precise and reliable real-time EOD timing measurement method from freely swimming WEF. Signal distortions caused by the animal&#39;s body movements are corrected by spatial averaging and temporal processing stages. Third, we describe an underwater near-infrared imaging setup to observe unperturbed nocturnal animal behaviors. Infrared light pulses were used to synchronize the timing between the video and the physiological signal over a long recording duration. Our automated tracking software measures the animal&#39;s body position and posture reliably in an aquatic scene. In combination, these techniques enable long term observation of spontaneous behavior of freely swimming weakly electric fish in a reliable and precise manner. We believe our method can be similarly applied to the study of other aquatic animals by relating their physiological signals with exploratory or social behaviors.

We describe a set of techniques for studying spontaneous behavior of freely swimming weakly electric fish over an extended period of time, by synchronously measuring the animal&#39;s electric organ discharge timing, body position and posture both accurately and reliably in a specially designed aquarium tank inside a sensory isolation chamber.

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and ...

Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent and divergent anatomical connections including feed forward and feedback wiring. Furthermore, information of the same origin is partially sent via parallel pathways to different and sometimes into the same brain areas. To understand the evolutionary benefits as well as the computational advantages of these wiring strategies and especially their temporal dependencies on each other, it is necessary to have simultaneous access to single neurons of different tracts or neuropiles in the same preparation at high temporal resolution. Here we concentrate on honeybees by demonstrating a unique extracellular long term access to record multi unit activity at two subsequent neuropiles1, the antennal lobe (AL), the first olfactory processing stage and the mushroom body (MB), a higher order integration center involved in learning and memory formation, or two parallel neuronal tracts2 connecting the AL with the MB.&#160;The latter was chosen as an example and will be described in full. In the supporting video the construction and permanent insertion of flexible multi&#160;channel wire electrodes is demonstrated. Pairwise differential amplification of the micro&#160;wire electrode channels drastically reduces the noise and verifies that the source of the signal is closely related to the position of the electrode tip. The mechanical flexibility of the used wire electrodes allows stable invasive long&#160;term recordings over many hours up to days, which is a clear advantage compared to conventional extra&#160;and intracellular in vivo recording techniques.

Simultaneous extracellular long term recordings from two different brain neuropiles or two different anatomical tracts were established in honeybees. These recordings allow the investigation of temporal aspects of neuronal processing across different brain areas at the single neuron as well as at the ensemble level in a behaving animal.

In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure the activity of neurons during various behaviors. To correlate neural activity with behavior, the animal should not be immobilized but should be able to move. Many behavioral changes occur during long time scales and require recording over many hours of behavior. This also makes it necessary to culture the worms in the presence of food. How can worms be cultured and their neural activity imaged over long time scales? Agarose Microchamber Imaging (AMI) was previously developed to culture and observe small larvae and has now been adapted to study all life stages from early L1 until the adult stage of C. elegans. AMI can be performed on various life stages of C. elegans. Long-term calcium imaging is achieved without immobilizing the animals by using short externally triggered exposures combined with an electron multiplying charge-coupled device (EMCCD) camera recording. Zooming out or scanning can scale up this method to image up to 40 worms in parallel. Thus, a method is described to image behavior and neural activity over long time scales in all life stages of C. elegans.

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Imaging behavior and neural activity over long time scales without immobilization of the animal is a prerequisite to understand behavior. Agarose microfluidic chambers imaging (AMI) can be used to image neural activity and behavior for all life stages of Caenorhabditis elegans.

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure ...

Generation and Long-term Maintenance of Nerve-free Hydra

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The interstitial cells can be eliminated through two consecutive treatments with colchicine, a plant-derived toxin that kills dividing cells, thus erasing the potential for renewal of the differentiated cells that are derived from the interstitial stem cells. This allows for the generation of Hydra that lack nerve cells. A nerve-free polyp cannot open its mouth to feed, egest, or regulate osmotic pressure. Such animals, however, can survive and be cultured indefinitely in the laboratory if regularly force-fed and burped. The lack of nerve cells allows for studies of the role of the nervous system in regulating animal behavior and regeneration. Previously published protocols for nerve-free Hydra maintenance involve outdated techniques such as mouth-pipetting with hand-pulled micropipette tips to feed and clean the Hydra. Here, an improved protocol for maintenance of nerve-free Hydra is introduced. Fine-tipped forceps are used to force open the mouth and insert freshly killed Artemia. Following force-feeding, the body cavity of the animal is flushed with fresh medium using a syringe and hypodermic needle to remove undigested material, referred to here as "burping". This new method of force-feeding and burping nerve-free Hydra through the use of forceps and syringes eliminates the need for mouth-pipetting using hand-pulled micropipette tips. It thus makes the process safer and significantly more time efficient. To ensure that the nerve cells in the hypostome have been eliminated, immunohistochemistry using anti-tyrosine-tubulin is conducted.

Generation and Long-term Maintenance of Nerve-free Hydra

Through a double treatment with colchicine, a plant-derived toxin that kills dividing cells, nerve-free Hydra vulgaris can be generated. These Hydra cannot feed or egest on their own. This paper describes an improved method for long-term maintenance of nerve free Hydra vulgaris in the laboratory.

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...

Mechanical Conflict-Avoidance Assay to Measure Pain Behavior in Mice

Pain comprises of both sensory (nociceptive) and affective (unpleasant) dimensions. In preclinical models, pain has traditionally been assessed using reflexive tests that allow inferences regarding pain's nociceptive component but provide little information about the affective or motivational component of pain. Developing tests that capture these components of pain are therefore translationally important. Hence, researchers need to use non-reflexive behavioral assays to study pain perception at that level. Mechanical conflict-avoidance (MCA) is an established voluntary non-reflexive behavior assay, for studying motivational responses to a noxious mechanical stimulus in a 3 chamber paradigm. A change in a mouse's location preference, when faced with competing noxious stimuli, is used to infer the perceived unpleasantness of bright light versus tactile stimulation of the paws. This protocol outlines a modified version of the MCA assay which pain researchers can use to understand affective-motivational responses in a variety of mouse pain models. Though not specifically described here, our example MCA data use the intraplantar complete Freund's adjuvant (CFA), spared nerve injury (SNI), and a fracture/casting model as pain models to illustrate the MCA procedure.

The mechanical conflict-avoidance assay is used as a non-reflexive readout of pain sensitivity in mice which can be used to better understand affective-motivational responses in a variety of mouse pain models.

Pain comprises of both sensory (nociceptive) and affective (unpleasant) dimensions. In preclinical models, pain has traditionally been assessed using ...