This research was supported by Basic Science Research Program through the NRF funded by the Ministry of Education, Science and Technology (2010-0022362) and (2012R1A2A2A02046812).

Ethics Statement: 
This experiment was approved by the Kyungpook National University Institutional Animal Care and Use Committee, and was performed according to Guide for the Care and Use of Laboratory Animals (National Institute of Health, 1996). After experiment, animals were sacrificed under carbon dioxide euthanasia.
1. Design of the Head-fixation Bar for Rats
NOTE: This step is a general guideline for a fixation bar design. Any modification about the dimension and material is possible depending on animals or implants. Designing a head-fixation bar using CAD software is recommended but not necessary.
<ol>
	<li>Use CAD software to design a fixation bar.
	<ol>
		<li>Design a fixation bar of a rectangular shape (123 &#215; 35 &#215; 6 mm) (Figure 1A). Make a rectangular notch (16 &#215; 20 mm; port) in the middle of the fixation bar. Add slopes or indentations which serve as a guide rail on the borders of the port (Figure 1A-a, Figure 1B-arrows). 
		NOTE: Ensure that the area of the port covers the estimated rectangle of rat&#8217;s skull over the brain to allow vertical accesses using a stereotaxic manipulator to most of the brain areas through a craniotomy.</li>
		<li>Attach plates (50 &#215; 20 &#215; 6 mm) with two elongated anchoring holes on both sides of the fixation bar. Ensure that these holes pass screws of 6 mm diameter (Figure 1A-d). Make a hole of diameter 3 mm in front of the port for a locking hook (Figure 1A-b).</li>
		<li>Make a box having a floor (135 &#215; 305 mm) and side walls (305 &#215; 76 mm). Drill two 6 mm diameter holes as shown in Figure 1A (inset). Add supporting walls on the fixation bar (Figure 1A-e).</li>
	</ol>
	</li>
	<li>Manufacture the designed bar and box and assemble each of them using cyanoacrylate as shown in Figure 1A (inset). Assemble the bar on the box using screws of 6 mm diameter. Apply slope by adjusting the screw position of the bar for ergonomic head fixation (Figure 1A-inset). 
	NOTE: For mice, the dimension of the fixation bar is 60 &#215; 20 &#215; 4 mm (consisting of two layers of 2 mm thick acrylic panels). The port is square of 8 mm width (upper) and 9 x 10 mm (lower) as shown in Figure 1C. Anchoring plates are the same size of rats&#8217;. The box plate and side walls are 100 &#215; 72 mm and 100 &#215; 50 mm, respectively.</li>
</ol>
2. Head Mounter Construction during Implantation Surgery
CAUTION: Be cautious when dealing with dental acrylic resin. Liquid may cause skin irritation. Put on a mask and wear gloves. Work in a well-ventilated area.
<ol>
	<li>Preparation before surgery
	<ol>
		<li>Tightly wrap the border of the port of the fixation bar using a paraffin film to prevent adhesion of applied acrylic resin to the bar (see Figure 1B and Figure 2A). Coat the boarder of the port covered by the paraffin film with a thin layer of acrylic resin.</li>
		<li>Use sterilized surgical instruments. Thoroughly disinfect stereotaxic frame and its surrounding using 70% ethyl alcohol.</li>
	</ol>
	</li>
	<li>Perform stereotaxic implantation surgery
	<ol>
		<li>Anesthetize an animal by intraperitoneal injection of a cocktail mixture (2 ml/kg) of ketamine 10 ml (50 mg/ml), xylazine hydrochloride 1.5 ml (23.32 mg/ml), and saline 2.5 ml. Confirm that the animal fully anesthetized and does not respond to tail and toe pinches. Inject cocktail mixture of 1 ml/kg during surgery when necessary.</li>
		<li>Fix the animal to the stereotaxic apparatus. Apply ophthalmic ointment on eyes to prevent dryness during surgery. Shave and clean the scalp with 70% ethanol. Give subcutaneous injection of 2% lidocaine and incise the scalp along the midline of the skull. Completely scrape off periosteum. Rinse with saline several times.</li>
		<li>Make burr-hole drilling near the rim of the skull. Implant screws 4-5 turns (up to 1 mm depth) from the skull surface to prevent a breach of the dura mater.&#160;&#160;Make sure no leakage of CSF and blood on the skull surface after screw implantation. Rinse with saline and dry the skull surface using air-puff, especially around the screws.</li>
		<li>Coat skull surface with a thin layer of &#8220;dental adhesive (Super-Bond C&#38;B)&#8221;. Do not entirely cover the surface of the skull including the bregma if additional electrode implantation is scheduled. Use the bregma for marking craniotomy coordinates. Implant electrodes and/or microdrive and fix it on the skull by applying acrylic resin around it.</li>
		<li>Apply additional acrylic resin on top of the dental cement. Use appropriate amount of acrylic resin to leave sufficient fill-out space between bar-coated resin and initial mounting resin when fixating the screws and/or electrodes (Figure 2A).</li>
	</ol>
	</li>
	<li>Place head-fixation bar 5 mm or more above the skull surface and align its center (Figure 2A-Left).</li>
	<li>Apply acrylic resin into the gap between the base of the skull and initial mounting resins. (Figure 2A-Center). Wait until the resin become hard.</li>
	<li>Remove ear-bar and pull the molded mounter out of the fixation bar port. (Figure 2A-Right). Carefully apply additional acrylic resin around the edge rails of the molded mounter if necessary.</li>
	<li>Rinse the wound surface of the scalp with sterile saline.&#160;Inject an antibiotic (enrofloxacin, 5 mg/kg s.c.) and an analgesic (butorphanol tartrate, 2 mg/kg s.c) immediately after surgery. House the animal individually in a single cage.</li>
</ol>
3. Habituating Animals to Head-fixation
Note: This step describes habituation schedules based on authors&#8217; empirically successful assessments. For an overview of habituation procedure for head-fixation, see Schwartz&#8217;s10 or Guo&#8217;s11 work.
<ol>
	<li>Give sufficient recovery period (a week or more) to animals. On Day 1, Handle animals frequently on divided short sessions.</li>
	<li>On the next day (Day 2), expose the animal to the fixation environment (box). Do not insert their head mounter into the port of the fixation bar. Leave animals to get accustomed to the environment. Securely grab the head mounter using hand for 1 to 2 sec and release it immediately. Repeat short term fixation by hand-grab several times.</li>
	<li>On Day 3, habituate animals to head-fixation.
	<ol>
		<li>Bring animals from the home cages and settle them in the box directing heads toward the port of the fixation bar. Grab the head mounter and insert it into the port of the fixation bar.</li>
		<li>Bind the mounter (animal&#8217;s head) to the locking hook of the fixation bar using a rubber band. Stand for 5 min. 
		NOTE: Do not touch the trunk or tail of the animal after fixation, which leads the animal to squeak and struggle increasing animal&#8217;s stress. When the animal continues struggling more than 1 min, release it and try later.&#160;From this step, animals show urine and feces excretion whenever exposed to head fixation but this gradually gets reduced as habituation proceeds.</li>
		<li>Retrieve the locking rubber band and eject the mounter out of the port. Put the animals back to the home cage and provide a piece of cereal or a sunflower seed into the home cage as a reward. The next trial begins in 30 min. Repeat 3.4.1 to 3.4.4 five or more within a day.</li>
	</ol>
	</li>
	<li>On Day 4 to 7, expose the animal to head fixation as described in Protocol 3.4. Gradually increase the duration of head fixation (10 min, 20 min, 30 min, and 50 min as habituation proceeds). 
	NOTE: If animals do not feed the food reward, it can be a sign of a high level of stress. Do not increase the fixation time. If animals show repeated struggling behavior, habituate them for a longer period.</li>
	<li>Check animal&#8217;s body weight daily. Consider unsuccessful habituation when body weight is decreasing. Proceed to behavioral tasks with head fixation after successful habituation. 
	NOTE: Successful habituation facilitates subsequent behavioral training (Figure 3E and F).</li>
</ol>

<ol>
	<li>Mayrhofer, J. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+two-alternative+forced+choice+paradigm+for+bilateral+vibrotactile+whisker+frequency+discrimination+in+head-fixed+mice+and+rats.">Novel two-alternative forced choice paradigm for bilateral vibrotactile whisker frequency discrimination in head-fixed mice and rats.</a> J Neurophysiol. 109, 273-284 (2013).</li><li>Bryant, J. L., Roy, S., Heck, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+technique+for+stereotaxic+recordings+of+neuronal+activity+in+awake,+head-restrained+mice.">A technique for stereotaxic recordings of neuronal activity in awake, head-restrained mice.</a> J Neurosci Methods. 178, 75-79 (2009).</li><li>Ollerenshaw, D. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+tactile+inputs+in+the+rat+vibrissa+pathway.">Detection of tactile inputs in the rat vibrissa pathway.</a> J Neurophysiol. 108, 479-490 (2012).</li><li>Lee, M. G., Manns, I. D., Alonso, A., Jones, B. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sleep-wake+related+discharge+properties+of+basal+forebrain+neurons+recorded+with+micropipettes+in+head-fixed+rats.">Sleep-wake related discharge properties of basal forebrain neurons recorded with micropipettes in head-fixed rats.</a> J Neurophysiol. 92, 1182-1198 (2004).</li><li>Lutcke, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+recording+of+neuronal+activity+with+a+genetically-encoded+calcium+indicator+in+anesthetized+and+freely+moving+mice.">Optical recording of neuronal activity with a genetically-encoded calcium indicator in anesthetized and freely moving mice.</a> Front Neural Circuits. 4, 9 (2010).</li><li>Dombeck, D. A., Khabbaz, A. N., Collman, F., Adelman, T. L., Tank, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+large-scale+neural+activity+with+cellular+resolution+in+awake,+mobile+mice.">Imaging large-scale neural activity with cellular resolution in awake, mobile mice.</a> Neuron. 56, 43-57 (2007).</li><li>Kimura, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reinforcing+operandum:+rapid+and+reliable+learning+of+skilled+forelimb+movements+by+head-fixed+rodents.">Reinforcing operandum: rapid and reliable learning of skilled forelimb movements by head-fixed rodents.</a> Journal of neurophysiology. 108, 1781-1792 (2012).</li><li>Hadlock, T., Kowaleski, J., Mackinnon, S., Heaton, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+of+head+fixation+for+the+study+of+rodent+facial+function.">A novel method of head fixation for the study of rodent facial function.</a> Experimental neurology. 205, 279-282 (2007).</li><li>Roh, M., Jang, I. S., Lee, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Push-in+Head+Restraining+Apparatus+for+Intracranial+Self+Stimulation+Tasks+in+Rats.">Push-in Head Restraining Apparatus for Intracranial Self Stimulation Tasks in Rats.</a> Korean J Physiol Pharmacol. 18, 103-108 (2014).</li><li>Schwarz, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+head-fixed+behaving+rat--procedures+and+pitfalls.">The head-fixed behaving rat--procedures and pitfalls.</a> Somatosensory &amp; motor research. 27, 131-148 (2010).</li><li>Guo, Z. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Procedures+for+Behavioral+Experiments+in+Head-Fixed+Mice.">Procedures for Behavioral Experiments in Head-Fixed Mice.</a> PLoS ONE. 9, (2014).</li><li>Osborne, J. E., Dudman, J. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RIVETS:+a+mechanical+system+for+in+vivo+and+in+vitro+electrophysiology+and+imaging.">RIVETS: a mechanical system for in vivo and in vitro electrophysiology and imaging.</a> PLoS One. 9, e89007 (2014).</li></ol>

The authors have nothing to disclose.

This protocol demonstrates a simple and convenient head fixation method using materials at hand. For physically successful head fixation, the first critical steps during surgery is to rinse and dry the skull surface clearly before applying the dental adhesive resin cement. Second, this method does not uses screws implanting under temporal muscles and only depends on the screws on the skull, their perpendicular implantation against the surface of skull rim is crucial10. This reduces the possibility of accidenta...

Head fixation or head restraining is a technique of immobilizing the animal&#39;s head by attaching a frame or plate on the skull for rigid clamping. The technique has been adopted for precise sensory stimulation, behavioral sensing with or without neural recording in awake animals1-3 and for juxtacellular labeling during sleep-wake cycle4. It is also used for in vivo brain imaging5,6. Now head fixation technique is a commonly used tool in neuroscience and behavioral research.
Basically, traditional head fixation requires surgical attachment of metal head-posts on top of the skull6-8. The attachments are then clamped to a stationary platform, resulting in immobilization of the head. Neurophysiological research in awake animals such as neural and/or optical recording, brain circuitry intervention, brain imaging is accompanied by implantation surgery using dental acrylics. Therefore, it seems a simple task to implement the head fixation in those laboratories when it is experimentally necessary. The frames should be designed to be light-weight and small enough not to disturb animal&#8217;s natural behavior in their home cages. It also provides mechanical rigidity when clamped to the stationary platform. It further has biocompatibility depending on experimental purposes. Therefore, it is technically difficult to design and fabricate such head-posts in general laboratory equipment.
In this study, a novel head fixation method is described, providing mechanical rigidity through convenient clamping without using metal head-posts. By using the method, head fixation could be done under general laboratory conditions, where implantation surgery is usually performed in rodents for brain study. The metal head-posts are replaced with acrylic resin cement molded partially by a custom-made head fixation bar and the molding process is done during implantation surgery. The fixation bar provides high spatial compatibility with other implants, such as neural recording probes, stimulation electrodes. Also it simplifies steps of clamping the animal&#8217;s head on the fixation platform by removing screw-fastening. We used this method in the previous behavioral study and we concluded that this method is easy to implement and convenient in practical experimentation9.

<table border="0" cellpadding="0" cellspacing="0" width="901">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">Dental acrylic resin</td>
			<td style="width:199px;">Lang Dental Manufacturing</td>
			<td style="width:129px;">Ortho-Jet</td>
			<td style="width:343px;">to make molded mounter</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">Paraffin film</td>
			<td style="width:199px;">Bemis</td>
			<td style="width:129px;">Parafilm</td>
			<td>to prevent adhesion of acrylic resin to the bar</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">Dental adhesive</td>
			<td style="width:199px;">Sun Medical</td>
			<td style="width:129px;">Super-Bond C&#38;B</td>
			<td>to bond between acrylic resin and skull</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">Plexglass or other acrylic plate</td>
			<td style="width:199px;"></td>
			<td style="width:129px;"></td>
			<td>to make fixation bar and box platform</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">Cyanoacrylate</td>
			<td style="width:199px;"></td>
			<td style="width:129px;"></td>
			<td>to bond acrylic plates</td>
		</tr>
	</tbody>
</table>

acrylic resin molding based head fixation technique in rodents

Head fixation is a technique of immobilizing animal's head by attaching a head-post on the skull for rigid clamping. Traditional head fixation requires surgical attachment of metallic frames on the skull. The attached frames are then clamped to a stationary platform resulting in immobilization of the head. However, metallic frames for head fixation have been technically difficult to design and implement in general laboratory environment. In this study, we provide a novel head fixation method. Using a custom-made head fixation bar, head mounter is constructed during implantation surgery. After the application of acrylic resin for affixing implants such as electrodes and cannula on the skull, additional resins applied on top of that to build a mold matching to the port of the fixation bar. The molded head mounter serves as a guide rails, investigators conveniently fixate the animal&#8217;s head by inserting the head mounter into the port of the fixation bar. This method could be easily applicable if implantation surgery using dental acrylics is necessary and might be useful for laboratories that cannot easily fabricate CNC machined metal head-posts.

This head fixation method is applicable to both rats and mice. Total eight rats received bilateral implantation of stimulating electrodes into the medial forebrain bundle. For four out of them, customized tungsten wire electrodes (microdrives) were additionally implanted into the M1 forelimb area (Figure 2C). For mice (n = 2), four stainless steel wire electrodes were implanted into frontal/occipital cortical areas. During surgery, head mounters were molded for head fixation in both rats and mice, as des...

Watch this Scientific Journal Video about Acrylic Resin Molding Based Head Fixation Technique in Rodents at JoVE.com

Acrylic Resin Molding Based Head Fixation Technique in Rodents

Traditional head fixation techniques have used metallic frames attached on the skull as an interface for head fixation apparatus. This protocol demonstrates a frameless, acrylic resin molding based head fixation technique for behavioral tasks in rodents.

Head fixation is a technique of immobilizing animal's head by attaching a head-post on the skull for rigid clamping. Traditional head fixation requires ...

acrylic-resin-molding-based-head-fixation-technique-in-rodents

Research

JoVE Journal

Neuroscience

11.7K Views.  Kyungpook National University. Traditional head fixation techniques have used metallic frames attached on the skull as an interface for head fixation apparatus. This protocol demonstrates a frameless, acrylic resin molding based head fixation technique for behavioral tasks in rodents.

Video: Acrylic Resin Molding Based Head Fixation Technique in Rodents

Preparing Undercut Model of Posttraumatic Epileptogenesis in Rodents

Partially isolated cortex ("undercut") is an animal model of posttraumatic epileptogenesis. The surgical procedure involves cutting through the sensorimotor cortex and the underneath white matter (undercut) so that a specific region of the cerebral cortex is largely isolated from the neighboring cortex and subcortical regions1-3. After a latency of two or more weeks following the surgery, epileptiform discharges can be recorded in brain slices from rodents1; and electrical or behavior seizures can be observed in vivo from other species such as cat and monkey4-6. This well established animal model is efficient to generate and mimics several important characteristics of traumatic brain injury. However, it is technically challenging attempting to make precise cortical lesions in the small rodent brain with a free hand. Based on the procedure initially established in Dr. David Prince's lab at the Stanford University1, here we present an improved technique to perform a surgery for the preparation of this model in mice and rats. We demonstrate how to make a simple surgical device and use it to gain a better control of cutting depth and angle to generate more precise and consistent results. The device is easy to make, and the procedure is quick to learn. The generation of this animal model provides an efficient system for study on the mechanisms of posttraumatic epileptogenesis.

Partially isolated cortex (&#x201C;undercut&#x201D;) is an efficient animal model of posttraumatic epileptogenesis. Here we demonstrate how to make a novel surgical device and use it to make more precise and consistent lesions to generate this model.

Partially isolated cortex ("undercut") is an animal model of posttraumatic epileptogenesis. The surgical procedure involves cutting through the ...

Whole Animal Perfusion Fixation for Rodents

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small pieces of tissue, larger specimens like the intact brain pose a problem for immersion fixation because the fixative does not reach all regions of the tissue at the same rate 5,7. Often, changes in response to hypoxia begin before the tissue can be preserved 12. The advantage of directly perfusing fixative through the circulatory system is that the chemical can quickly reach every corner of the organism using the natural vascular network. In order to utilize the circulatory system most effectively, care must be taken to match physiological pressures 3. It is important to note that physiological pressures are dependent on the species used. Techniques for perfusion fixation vary depending on the tissue to be fixed and how the tissue will be processed following fixation. In this video, we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain for immunohistochemistry. The main advantage of this technique (vs. gravity-fed systems) is that the circulatory system is utilized most effectively.

Here we describe a low-cost, rapid, controlled and uniform fixation procedure using 4% paraformaldehyde perfused via the vascular system: through the heart of the rat to obtain the best possible preservation of the brain.

The goal of fixation is to rapidly and uniformly preserve tissue in a life-like state. While placing tissue directly in fixative works well for small ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

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

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

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

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

Transplantation of Olfactory Ensheathing Cells to Evaluate Functional Recovery after Peripheral Nerve Injury

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory system is characterized by its ability to give rise to new neurons even in adult animals. This particular ability is partly due to the presence of OECs which create a favorable microenvironment for neurogenesis. This property of OECs has been used for cellular transplantation such as in spinal cord injury models. Although the peripheral nervous system has a greater capacity to regenerate after nerve injury than the central nervous system, complete sections induce misrouting during axonal regrowth in particular after facial of laryngeal nerve transection. Specifically, full sectioning of the recurrent laryngeal nerve (RLN) induces aberrant axonal regrowth resulting in synkinesis of the vocal cords. In this specific model, we showed that OECs transplantation efficiently increases axonal regrowth.
OECs are constituted of several subpopulations present in both the olfactory mucosa (OM-OECs) and the olfactory bulbs (OB-OECs). We present here a model of cellular transplantation based on the use of these different subpopulations of OECs in a RLN injury model. Using this paradigm, primary cultures of OB-OECs and OM-OECs were transplanted in Matrigel after section and anastomosis of the RLN. Two months after surgery, we evaluated transplanted animals by complementary analyses based on videolaryngoscopy, electromyography (EMG), and histological studies. First, videolaryngoscopy allowed us to evaluate laryngeal functions, in particular muscular cocontractions phenomena. Then, EMG analyses demonstrated richness and synchronization of muscular activities. Finally, histological studies based on toluidine blue staining allowed the quantification of the number and profile of myelinated fibers.
All together, we describe here how to isolate, culture, identify and transplant OECs from OM and OB after RLN section-anastomosis and how to evaluate and analyze the efficiency of these transplanted cells on axonal regrowth and laryngeal functions.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth of the primary olfactory neurons. This specific property can be used for cellular transplantation. We present here a model of cellular transplantation based on the use of OECs in a laryngeal nerve injury model.

Olfactory ensheathing cells (OECs) are neural crest cells which allow growth and regrowth of the primary olfactory neurons. Indeed, the primary olfactory ...

Deep Brain Stimulation with Simultaneous fMRI in Rodents

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for using blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI) to image rodents with simultaneous DBS. DBS fMRI presents a number of technical challenges, including accuracy of electrode implantation, MR artifacts created by the electrode, choice of anesthesia and paralytic to minimize any neuronal effects while simultaneously eliminating animal motion, and maintenance of physiological parameters, deviation from which can confound the BOLD signal.&#160;Our laboratory has developed a set of procedures that are capable of overcoming most of these possible issues. For electrical stimulation, a homemade tungsten bipolar microelectrode is used, inserted stereotactically at the stimulation site in the anesthetized subject. In preparation for imaging, rodents are fixed on a plastic headpiece and transferred to the magnet bore. For sedation and paralysis during scanning, a cocktail of dexmedetomidine and pancuronium is continuously infused, along with a minimal dose of isoflurane; this preparation minimizes the BOLD ceiling effect of volatile anesthetics. In this example experiment, stimulation of the subthalamic nucleus (STN) produces BOLD responses which are observed primarily in ipsilateral cortical regions, centered in motor cortex. Simultaneous DBS and fMRI allows the unambiguous modulation of neural circuits dependent on stimulation location and stimulation parameters, and permits observation of neuronal modulations free of regional bias. This technique may be used to explore the downstream effects of modulating neural circuitry at nearly any brain region, with implications for both experimental and clinical DBS.

This protocol describes a standard method for simultaneous functional magnetic resonance imaging and deep brain stimulation in the rodent. The combined use of these experimental tools allows for the exploration of global downstream activity in response to electrical stimulation at virtually any brain target.

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for ...

Transcranial Electrical Brain Stimulation in Alert Rodents

Transcranial electrical brain stimulation can modulate cortical excitability and plasticity in humans and rodents. The most common form of stimulation in humans is transcranial direct current stimulation (tDCS). Less frequently, transcranial alternating current stimulation (tACS) or transcranial random noise stimulation (tRNS), a specific form of tACS using an electrical current applied randomly within a pre-defined frequency range, is used. The increase of noninvasive electrical brain stimulation research in humans, both for experimental and clinical purposes, has yielded an increased need for basic, mechanistic, safety studies in animals. This article describes a model for transcranial electrical brain stimulation (tES) through the intact skull targeting the motor system in alert rodents. The protocol provides step-by-step instructions for the surgical set-up of a permanent epicranial electrode socket combined with an implanted counter electrode on the chest. By placing a stimulation electrode into the epicranial socket, different electrical stimulation types, comparable to tDCS, tACS, and tRNS in humans, can be delivered. Moreover, the practical steps for tES in alert rodents are introduced. The applied current density, stimulation duration, and stimulation type may be chosen depending on the experimental needs. The caveats, advantages, and disadvantages of this set-up are discussed, as well as safety and tolerability aspects.

This protocol describes a surgical set-up for a permanent epicranial electrode socket and an implanted chest electrode in rodents. By placing a second electrode into the socket, different types of transcranial electrical brain stimulation can be delivered to the motor system in alert animals through the intact skull.

Transcranial electrical brain stimulation can modulate cortical excitability and plasticity in humans and rodents. The most common form of stimulation in ...

Intracranial Pharmacotherapy and Pain Assays in Rodents

Pain is a salient sensory experience with affective and cognitive dimensions. However, central mechanisms for pain remain poorly understood, hindering the development of effective therapeutics. Intracranial pharmacology presents an important tool for understanding the molecular and cellular mechanisms of pain in the brain, as well as for novel treatments. Here we present a protocol that integrates intracranial pharmacology with pain behavior testing. Specifically, we show how to infuse analgesic drugs into a select brain region, which may be responsible for pain modulation. Furthermore, to determine the effect of the candidate drug in the central nerve system, pain assays are performed after intracranial treatment. Our results demonstrate that intracranial administration of analgesic drugs in a targeted region can provide relief of pain in rodents. Thus, our protocol successfully demonstrates that intracranial pharmacology, combined with pain behavior testing, can be a powerful tool for the study of pain mechanisms in the brain.

Here we present a protocol to perform intracranial pharmacological experiments followed by pain behavior assays in rodents. This protocol allows researchers to deliver molecular and cellular targets in the brain, for pharmacologic agents in the treatment of pain.

Pain is a salient sensory experience with affective and cognitive dimensions. However, central mechanisms for pain remain poorly understood, hindering the ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in ...

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a&#160;good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) ...