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All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. Headpost Implantation Surgery
NOTE: 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 0.05 mg/kg buprenorphine is recommended 24h after the surgery.&#160;</li>
	<li>Anesthetize the animal in a chamber with 2.5%&#8211;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 (see&#160;Table of Materials) 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 (see&#160;Table of Materials) 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%&#8211;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>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >Blunt scissors</td>
			<td >FST</td>
			<td >14079-10</td>
			<td ></td>
		</tr>
		<tr >
			<td >Catalyst V</td>
			<td>Sun Medical, Japan</td>
			<td>LX22</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr >
			<td >Dentalon Plus</td>
			<td>Heraeus</td>
			<td>37041</td>
			<td></td>
		</tr>
		<tr >
			<td >Green activator</td>
			<td>Sun Medical, Japan</td>
			<td>VE-1</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr >
			<td >Monomer</td>
			<td>Sun Medical, Japan</td>
			<td>MF-1</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
		<tr >
			<td >Ocrygel</td>
			<td>TvmLab</td>
			<td>10779</td>
			<td>Ophtalmic vet ointment</td>
		</tr>
		<tr >
			<td >Polymer L-type clear (cement)</td>
			<td>Sun Medical, Japan</td>
			<td>TT12F</td>
			<td>Parkell bio-materials, Kit n&#176;S380</td>
		</tr>
	</tbody>
</table>

mouse headpost implantation: a method for head fixation in rodents

Source: Fran&#231;a de Barros, F., et al. <a href="https://www.jove.com/video/59135/" target="_blank">Long-term Sensory Conflict in Freely Behaving Mice</a>. J. Vis. Exp. (2019).
This video shows a headpost implantation surgery in murine model. The headpost is used not only to head-fix and therefore restrain the mouse during experimentation, but also to securely attach a head device in freely moving animals.

Mouse Headpost Implantation: A Method for Head Fixation in Rodents

Source: Fran&#231;a de Barros, F., et al. Long-term Sensory Conflict in Freely Behaving Mice. J. Vis. Exp. (2019).
This video shows a headpost ...

On an anesthetized mouse whose head is shaved and prepared for surgery, make a length-wise incision to expose the skull. Next, gently remove the periosteum membrane to expose the surface of the skull. This will help the cement adhere better to the bone.
Apply activator to increase bone permeability. Then, in the area between the lambda and bregma landmarks of the skull, apply freshly mixed cement. Slide the headpost from lambda to bregma and apply more cement to ensure it is securely fixed.
Once the cement is completely dry, apply freshly mixed resin on top of the cement and around the headpost. Once the resin is dry, close the cut by suturing the skin.

mouse-headpost-implantation-a-method-for-head-fixation-in-rodents

Research

JoVE Encyclopedia of Experiments

Biology

Rodent Models

Adult

2.0K Views. Source: França de Barros, F., et al. Long-term Sensory Conflict in Freely Behaving Mice. J. Vis. Exp. (2019). This video shows a headpost implantation surgery in murine model. The headpost is used not only to head-fix and therefore restrain the mouse during experimentation, but also to securely attach a head device in freely moving animals. 

Video: Mouse Headpost Implantation: A Method for Head Fixation in Rodents - Concept

A Large Lateral Craniotomy Procedure for Mesoscale Wide-field Optical Imaging of Brain Activity

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, typically 3 mm x 3 mm. This protocol introduces a method for creating a substantially larger 7 mm x 6 mm cranial window exposing most of a cerebral hemisphere over the mouse temporal and parietal cortices (e.g., bregma 2.5 - 4.5 mm, lateral 0 - 6 mm). To perform this surgery, the head must be tilted approximately 30&#176; and much of the temporal muscle must be retracted. Due to the large amount of bone removal, this procedure is intended only for acute experiments with the animal anesthetized throughout the surgery and experiment.
The main advantage of this innovative large lateral cranial window is to provide simultaneous access to both medial and lateral areas of the cortex. This large unilateral cranial window can be used to study the neural dynamics between cells, as well as between different cortical areas by combining multi-electrode electrophysiological recordings, imaging of neuronal activity (e.g., intrinsic or extrinsic imaging), and optogenetic stimulation. Additionally, this large craniotomy also exposes a large area of cortical blood vessels, allowing for direct manipulation of the lateral cortical vasculature.

This protocol presents a method for creating a large unilateral craniotomy over the temporal and parietal regions of the mouse cerebral cortex. This is especially useful for real time imaging over an expansive area of a cortical hemisphere.

The craniotomy is a commonly performed procedure to expose the brain for in vivo experiments. In mouse research, most labs utilize a small craniotomy, ...

JoVE Journal

Neuroscience

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and magnetic resonance imaging (MRI) have enabled in vivo quantification of CSF flow but they are limited by reduced imaging volumes and low spatial resolution, respectively. Recent work has found that CSF enters the brain parenchyma through a network of perivascular spaces surrounding the pial and penetrating arteries of the rodent cortex. This perivascular entry of CSF is a primary driver of the glymphatic system, a pathway implicated in the clearance of toxic metabolic solutes (e.g., amyloid-&#946;). Here, we illustrate a new macroscopic imaging technique that allows real-time, mesoscopic imaging of fluorescent CSF tracers through the intact skull of live mice. This minimally-invasive method facilitates a multitude of experimental designs and enables single or repeated testing of CSF dynamics. Macroscopes have high spatial and temporal resolution and their large gantry and working distance allow for imaging while performing tasks on behavioral devices. This imaging approach has been validated using two-photon imaging and fluorescence measurements obtained from this technique strongly correlate with ex vivo fluorescence and quantification of radio-labeled tracers. In this protocol, we describe how transcranial macroscopic imaging can be used to evaluate glymphatic transport in live mice, offering an accessible alternative to more costly imaging modalities.

Transcranial optical imaging allows wide-field imaging of cerebrospinal fluid transport in the cortex of live mice through an intact skull.

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and ...

Mouse Headpost Implantation: A Method for Head Fixation in Rodents

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