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><tbody><tr><td>Dental acrylic resin</td><td>Lang Dental Manufacturing</td><td>Ortho-Jet</td><td>to make molded mounter</td></tr><tr><td>Paraffin film</td><td>Bemis</td><td>Parafilm</td><td>to prevent adhesion of acrylic resin to the bar</td></tr><tr><td>Dental adhesive</td><td>Sun Medical</td><td>Super-Bond C&amp;amp;B</td><td>to bond between acrylic resin and skull</td></tr><tr><td>Plexglass or other acrylic plate</td><td></td><td></td><td>to make fixation bar and box platform</td></tr><tr><td>Cyanoacrylate</td><td></td><td></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

Application of Light-cured Dental Adhesive Resin for Mounting Electrodes or Microdialysis Probes in Chronic Experiments

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these acrylics do not bond to the bone, screws have been used as anchors. However, in small experimental animals like finches or mouse, their craniums are very fragile and can not successfully hold the anchors. In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium. Therefore, anchor screws are not required and surgical field can be reduced considerably. Past experiences show that the bonding effect maintains more than 2 months. Conventional resin's window of time when the materials are pliable and workable is a few minutes. However, the window of working time for these dental adhesives is significantly wider and adjustable.

In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium.

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these ...

Biology

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

Behavioral Training Procedures for Head-fixed Virtual Reality in Mice

Virtual reality (VR) combined with head-fixation is increasingly being utilized in behavioral neuroscience studies as it allows complex behavioral assays to be performed in head-fixed mice. This enables precise behavioral recordings while incorporating various neurophysiological techniques that require head-fixation to minimize movement-related signal noise during neural recordings. However, despite the growing use of VR, there is little published data on the detailed methodology of how to implement it. In this study, a training protocol is developed whereby male and female C57B16/J mice are trained to run down a virtual linear corridor, the length of which is increased from 1-3 m over multiple training sessions. Building upon this foundation, this study investigated the feasibility of mice performing complex behaviors within VR using a Y-maze paradigm. The task required navigating to the arm with black walls from the choice point in the Y-maze. After reaching a criterion of two consecutive days equal to or greater than 70% correct, the mice progressed to increasingly difficult sensory discrimination. The findings provide important details on the methodologies useful for the successful training of mice in VR and demonstrate that mice exhibit learning capabilities in navigating the Y-maze. The methodology presented not only offers insights into training duration in VR-based assays but also underscores the potential for probing intricate behaviors in mice, opening avenues for more comprehensive neuroscience investigations.

The article describes the experimental procedures for the commonly used linear track virtual reality (VR) paradigm in mice as well as determining the feasibility of running complex VR tasks by testing a Y-shaped signal discrimination task.

Virtual reality (VR) combined with head-fixation is increasingly being utilized in behavioral neuroscience studies as it allows complex behavioral assays ...

Behavior

Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the identification of the neuronal type in in vivo electrophysiological experiments in broad brain regions. In optogenetics experiments, it is critical to deliver the light to the recording site. However, it is often hard to deliver the stimulation light to the deep brain regions from the brain&#39;s surface. Especially, it is difficult for the stimulation light to reach the deep brain regions when the optical transparency of the brain surface is low, as is often the case with recordings from awake animals. Here, we describe a method to record spike responses to the light from an awake mouse using a custom-made glass optrode. In this method, the light is delivered through the recording glass electrode so that it is possible to reliably stimulate the recorded neuron with light in the deep brain regions. This custom-made optrode system consists of accessible and inexpensive materials and is easy to assemble.

This work introduces a method to perform an optogenetic single-unit recording reliably from an awake mouse using a custom-made glass optrode.

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the ...

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 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 ...

Chronic Implantation of Multiple Flexible Polymer Electrode Arrays

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of scientific and clinical development. The use of flexible polymer electrode arrays can support long-lasting recording, but the same mechanical properties that allow for longevity of recording make multiple insertions and integration into a chronic implant a challenge. Here is a methodology by which multiple polymer electrode arrays can be targeted to a relatively spatially unconstrained set of brain areas.
The method utilizes thin-film polymer devices, selected for their biocompatibility and capability to achieve long-term and stable&#160;electrophysiologic recording interfaces. The resultant implant allows accurate and flexible targeting of anatomically distant regions, physical stability for months, and robustness to electrical noise. The methodology supports up to sixteen serially inserted devices across eight different anatomic targets. As previously demonstrated, the methodology is capable of recording from 1024 channels. Of these, the 512 channels in this demonstration used for single neuron recording yielded 375 single units distributed across six recording sites. Importantly, this method also can record single units for at least 160 days.
This implantation strategy, including temporarily bracing each device with a retractable silicon insertion shuttle, involves tethering of devices at their target depths to a skull-adhered plastic base piece that is custom-designed for each set of recording targets, and stabilization/protection of the devices within a silicone-filled, custom-designed plastic case. Also covered is the preparation of devices for implantation, and design principles that should guide adaptation to different combinations of brain areas or array designs.

Described below is a method for implantation of multiple polymer electrode arrays across anatomically distant brain regions for chronic electrophysiological recording in freely moving rats. Preparation and surgical implantation are described in detail, with emphasis on design principles to guide adaptation of these methods for use in other species.

Simultaneous recordings from large populations of individual neurons across distributed brain regions over months to years will enable new avenues of ...

Successful In vivo Calcium Imaging with a Head-Mount Miniaturized Microscope in the Amygdala of Freely Behaving Mouse

In vivo real-time monitoring of neuronal activities in freely moving animals is one of key approaches to link neuronal activity to behavior. For this purpose, an in vivo imaging technique that detects calcium transients in neurons using genetically encoded calcium indicators (GECIs), a miniaturized fluorescence microscope, and a gradient refractive index (GRIN) lens has been developed and successfully applied to many brain structures1,2,3,4,5,6. This imaging technique is particularly powerful because it enables chronic simultaneous imaging of genetically defined cell populations for a long-term period up to several weeks. Although useful, this imaging technique has not been easily applied to brain structures that locate deep within the brain such as amygdala, an essential brain structure for emotional processing and associative fear memory7. There are several factors that make it difficult to apply the imaging technique to the amygdala. For instance, motion artifacts usually occur more frequently during the imaging conducted in the deeper brain regions because a head-mount microscope implanted deep in the brain is relatively unstable. Another problem is that the lateral ventricle is positioned close to the implanted GRIN lens and its movement during respiration may cause highly irregular motion artifacts that cannot be easily corrected, which makes it difficult to form a stable imaging view. Furthermore, because cells in the amygdala are usually quiet at a resting or anesthetized state, it is hard to find and focus the target cells expressing GECI in the amygdala during baseplating procedure for later imaging. This protocol provides a helpful guideline for how to efficiently target cells expressing GECI in the amygdala with head-mount miniaturized microscope for successful in vivo calcium imaging in such a deeper brain region. It is noted that this protocol is based on a particular system (e.g., Inscopix) but not restricted to it.

In vivo microendoscopic calcium imaging is an invaluable tool that enables real-time monitoring of neuronal activities in freely behaving animals. However, applying this technique to the amygdala has been difficult. This protocol aims to provide a useful guideline for successfully targeting amygdala cells with a miniaturized microscope in mice.

In vivo real-time monitoring of neuronal activities in freely moving animals is one of key approaches to link neuronal activity to behavior. For this ...

Head Implants for the Neuroimaging of Awake, Head-Fixed Rats

Anesthetics, commonly used in preclinical and fundamental scientific research, have a depressive influence on the metabolic, neuronal, and vascular functions of the brain and can adversely influence neurophysiological results. The use of awake animals for research studies is advantageous but poses the major challenge of keeping the animals calm and stationary to minimize motion artifacts throughout data acquisition. Awake imaging in smaller-sized rodents (e.g., mice) is very common but remains scant in rats as rats are bigger, stronger, and have a greater tendency to oppose movement restraints and head fixation over the long durations required for imaging. A new model of neuroimaging of awake, head-fixed rats using customized hand-sewn slings, 3D-printed head implants, head caps, and a headframe is described. The results acquired following a single trial of single-whisker stimulation suggest an increase in the intensity of the evoked functional response. The acquisition of the evoked functional response from awake, head-fixed rats is faster than that from anesthetized rats, reliable, reproducible, and can be used for repeated longitudinal studies.

A detailed new procedure for functional imaging of awake, head-fixed rats is described.

Anesthetics, commonly used in preclinical and fundamental scientific research, have a depressive influence on the metabolic, neuronal, and vascular ...

A Cost-Effective, Versatile Probe Implant System for Electrophysiology in Mice

The DREAM Implant: A Lightweight, Modular, and Cost-Effective Implant System for Chronic Electrophysiology in Head-Fixed and Freely Behaving Mice

Chronic electrophysiological recordings in rodents have significantly improved our understanding of neuronal dynamics and their behavioral relevance. However, current methods for chronically implanting probes present steep trade-offs between cost, ease of use, size, adaptability, and long-term stability.
This protocol introduces a novel chronic probe implant system for mice called the DREAM (Dynamic, Recoverable, Economical, Adaptable, and Modular), designed to overcome the trade-offs associated with currently available options. The system provides a lightweight, modular and cost-effective solution with standardized hardware elements that can be combined and implanted in straightforward steps and explanted safely for recovery and multiple reuse of probes, significantly reducing experimental costs.
The DREAM implant system integrates three hardware modules: (1) a microdrive that can carry all standard silicon probes, allowing experimenters to adjust recording depth across a travel distance of up to 7 mm; (2) a three-dimensional (3D)-printable, open-source design for a wearable Faraday cage covered in copper mesh for electrical shielding, impact protection, and connector placement, and (3) a miniaturized head-fixation system for improved animal welfare and ease of use. The corresponding surgery protocol was optimized for speed (total duration: 2 h), probe safety, and animal welfare.
The implants had minimal impact on animals&#39; behavioral repertoire, were easily applicable in freely moving and head-fixed contexts, and delivered clearly identifiable spike waveforms and healthy neuronal responses for weeks of post-implant data collection. Infections and other surgery complications were extremely rare.
As such, the DREAM implant system is a versatile, cost-effective solution for chronic electrophysiology in mice, enhancing animal well-being, and enabling more ethologically sound experiments. Its design simplifies experimental procedures across various research needs, increasing accessibility of chronic electrophysiology in rodents to a wide range of research labs.

Here, we introduce a lightweight, cost-effective probe implant system for chronic electrophysiology in rodents optimized for ease of use, probe recovery, experimental versatility, and compatibility with behavior.

Chronic electrophysiological recordings in rodents have significantly improved our understanding of neuronal dynamics and their behavioral relevance. ...

Angled Silicon Multielectrode Recordings for Deep Brainstem Structures in Mice

Acute Single-Unit Multi-Electrode Recordings from the Brainstem of Head-Fixed Mice

Silicon multielectrode-based recordings are increasingly popular for studying neuronal activity at the temporal resolution of action potentials in many brain regions. However, recording neuronal activity from deep caudal structures like the brainstem using multi-channel probes remains challenging. A significant concern is finding a trajectory for probe insertion that avoids large blood vessels, such as the superior sagittal venous sinus and the transverse venous sinus. Injuring these large veins can cause extensive bleeding, damage to the underlying brain tissue, and potentially death. This approach describes targeting brainstem structures by coupling anterior coordinates with an angled approach, allowing the recording probe to penetrate the brain below high-risk vascular structures. Compared to a strictly vertical approach, the angled approach maximizes the number of brain regions that can be targeted. Using this strategy, the ventrolateral periaqueductal gray (vlPAG), a brainstem region associated with REM sleep, can be reproducibly and reliably accessed to obtain single-unit, multi-electrode recordings in head-fixed mice before and during sevoflurane anesthesia. The ability to record neuronal activity in the vlPAG and surrounding nuclei with high temporal resolution is a step forward in advancing the understanding of the relationship between REM sleep and anesthesia.

Author Spotlight: Investigating Anesthesia-Induced Sleep Pathways and Neuronal Excitability in Mice

This protocol describes a method to obtain in vivo, high-density single-neuron recordings from the brainstem of head-fixed mice. This approach is deployed to measure the action potential firing of neurons in the ventrolateral periaqueductal gray - a brainstem region inactive during Rapid Eye Movement (REM) sleep - before and during general anesthesia.

Silicon multielectrode-based recordings are increasingly popular for studying neuronal activity at the temporal resolution of action potentials in many ...