We acknowledge Clara Jones, James Stirwalt, Linh Hoang, Young Joon Ha, and Amirsoheil Zareh for their help during training of the rats and preparation of the slings. Funding was provided by the National Institutes of Health (NIH, Grant Number: NS119852) and Leducq Foundation (Grant Number:15CVD02).

All procedures were compliant with National Institute of Health guidelines and approved by the University of California, Irvine Animal Care and Use Committee. Seven males and one female rat (Sprague-Dawley, weight: 185-350 g) were used in this study. After study completion, the rats were sacrificed using carbon dioxide overdose.
1. Design of different components
<ol>
	<li>Design of the head implant:
	<ol>
		<li>Make the head implant using CAD software (Figure 1C) and design it to image the area posterior to the bregma and adjacent to the midline centered on the somatosensory cortex. Ensure that the head implant covers an area of 0.9 mm to 1.9 mm on the skull away from the imaging area.</li>
		<li>Use only three screws to anchor the head implant on the rat&#39;s skull. Design all the screw holes so that they remain on the opposite side of the midline in the contralateral hemisphere of the imaged hemisphere.</li>
		<li>Place a bar, hollowed from the inside, in the upper part of the head implant to allow wires to fix the head cap to the head implant as shown in Figure 1D.</li>
	</ol>
	</li>
	<li>Design of the head cap:
	<ol>
		<li>Ensure that the head cap covers the imaging area completely and protects it from any sort of trauma as shown in Figure 1A, B. Add a curvature to the head cap so that it aligns to the shape of the head without causing difficulty to the animal&#39;s daily activities in the standard enriched cages.</li>
		<li>Cut the inner side of the head cap in a wider rectangular shape so that the upper part of the head implant can fit into it as shown in Figure 1E. Perpendicular to this rectangle, cut two other rectangular regions to anchor the head cap to the head implant.</li>
		<li>Pass one wire through the upper hollowed bar of the head implant for fixation of the head cap on the rat head as shown in Figure 1E-G. Pass the second wire in the same way. 
		NOTE: These wires can be easily removed using pliers or forceps. The 3D printing files are provided (file format: STL) as Supplemental File 1 and Supplemental File 2.</li>
	</ol>
	</li>
	<li>Design of the head frame:
	<ol>
		<li>Design the head frame in a way that one cut part can move through the upper bar of the head implant and is fixed using a clamp.</li>
		<li>Angle the other cut part to provide extra strength for keeping the rat head-fixed to make the contralateral side completely accessible for imaging. For the purpose of this study, cut the steel plate with tin snips to produce the head frame (Figure 1H, I). 
		NOTE: This part can be 3D printed as well.</li>
	</ol>
	</li>
</ol>
2. Initial rat training
<ol>
	<li>Allow rats to acclimate to the vivarium environment in their cages for 2-3 days.</li>
	<li>Start handling the rat in a quiet room. Open the cage and have the experimenter put their hand inside the cage near the rat for 15-20 min to let the rat get habituated.</li>
	<li>Once the rat displays calmness by not getting startled or running away from the experimenter&#39;s hands, gently pick the rat up for handling. Handle the rat for 30-45 min each day before sling training.</li>
</ol>
3. Sling training
<ol>
	<li>Train the rats for at least 2-3 days in the slings before the surgical implantation of the head implant and head cap.</li>
	<li>Arrange the sling setup as shown in Figure 2A. Clean the sling setup using ethanol wipes. 
	NOTE: All the slings are hand-sewn and made of a netting material either on the bottom or on both sides as shown in Figure 2A, B.</li>
	<li>For sling training, anesthetize the rats using 4% isoflurane for induction and 1% for maintenance until there is no hind paw pinch reflex.</li>
	<li>Under isoflurane anesthesia, place the rats on a flexible plastic sheet measuring 20 cm x 8 cm (length x width), where 10 cm x 8 cm of the plastic sheet is fully covered with the softer part of the Velcro. 
	NOTE: Anesthetizing the rats for sling training is an optional step, primarily used to reduce stress and anxiety.</li>
	<li>For the first 2 days of the training, put the rat snuggly into a baby sock (size 0-3 months) with the head out through a small hole incised at the end of the sock.</li>
	<li>Wrap a small piece of absorbent pad around the lower body part to keep the rat dry and collect excrements.</li>
	<li>Wrap the rat in a breathable cotton cloth (size: 25 cm x 25 cm). Place the rat on a plastic sheet that has Velcro strips glued to it.</li>
	<li>Further secure the rat to the plastic sheet using 0.5 cm wide Velcro strips at a distance of 3-6 mm from each other.</li>
	<li>Secure the rat in the sling. Remove the gas anesthesia. Allow the rat to recover from gas anesthesia in the sling.</li>
	<li>When the rat starts whisking, offer a few drops of 10% sucrose solution as a reward every 10-15 min.</li>
	<li>Randomly present the rat with the sensory stimuli that will be used during imaging (here whisker stimulation, every 15-25 min) to make it accustomed to sensory stimuli. Manually stimulate the whiskers at random intervals.</li>
	<li>Train the rat in the sling for 1 h on day 1, 2 h on day 2, and 3 h on day 3 as shown in Figure 2C.</li>
</ol>
4. Presurgical preparation
<ol>
	<li>Print the head implant and head cap using the 3D printer (Figure 1).</li>
	<li>Sterilize all the surgical instruments and headpieces (implants and caps) by immersing the equipment in the Metricide28&#160;germicide for 10 hours. Rinse tools thoroughly with sterile water just before surgery.</li>
	<li>Expose the rat to 4% isoflurane and then maintain at 1%-2% isoflurane until there is no hind paw pinch reflex.&#160;This surgery can be performed under many types of anesthesia, such as isoflurane, sodium pentobarbital, and ketamine-xylazine.</li>
	<li>Inject atropine (0.05 mg/kg) intramuscularly to reduce mucous secretions to help in breathing.</li>
	<li>Shave the head of the rat 5 mm centered around the midline using a hair trimmer starting from between the eyes to the back of the ears.</li>
	<li>Monitor the partial oxygen saturation and heart rate through a pulse oximeter and heart rate monitor probe secured to the hind leg of the rat.</li>
	<li>Wipe the rat&#8217;s head and the surrounding area three times with alternating rounds of betadine and 70% alcohol wipes.</li>
	<li>Fix the rat in a stereotaxic system.</li>
	<li>Insert a petroleum jelly-lubricated rectal probe to measure the rat&#39;s body temperature and maintain it through the heating blanket&#39;s feedback system to avoid hypothermia after anesthetic administration.</li>
	<li>Administer local anesthetic lidocaine hydrochloride at a concentration of 20 mg/ml, 0.07 mg/kg +/-0.2 body weight subcutaneously at the surgical site.</li>
	<li>Apply ophthalmic ointment to both eyes to prevent drying.</li>
	<li>Administer 2% local anesthetic subcutaneously over the surgical site.</li>
	<li>Inject 3 mL of lactated ringer&#39;s solution at room temperature subcutaneously to prevent dehydration and provide nourishment during surgery.</li>
</ol>
5. Surgery
<ol>
	<li>Remove the part of the skin over the surgical site (4 mm diameter centered around the midline and center of the head) using sharp surgical scissors. Dissect and remove part of the skin (~2 mm diameter, over the left somatosensory cortex) between the ear and eye on the temporal part of the head.</li>
	<li>Remove, using a scalpel, the underlying skin (pericranium) tissue to expose the skull. Clean the skull using sterilized cotton gauze.</li>
	<li>Retract/resect temporal muscle to expose desired size for imaging area [7.5 mm by 7.5 mm for this study].&#160;</li>
	<li>Expose the skull on the contralateral hemisphere for the head implant. Place the head implant on the skull to ascertain the location of anchoring screws for the implant as shown in Figure 2D-F.</li>
	<li>Mark the skull for drilling the screws using India Ink with drill bit 1. Drill the burr holes for the screws using dental drill bit 3. Screw the head implant in place.</li>
	<li>Dry the skull using sterile gauze. Apply a thin layer of tissue adhesive around and beneath the head implant to glue it to the skull. Apply a layer of dental cement to further support the head implant in place and let the cement dry for 2-3 min. 
	NOTE: The use of tissue adhesive in addition to dental cement ensures a strong hold8.</li>
	<li>Using dental drill bit 3, thin a 7.5 mm x 7.5 mm area on the left side of skull just posterior to the bregma and lateral to the midline. Thin the skull to ~50 &#181;m as shown in Figure 3A.</li>
	<li>Apply topical antibiotic ointment over the surgical site and then cover it with a thin layer of silicone rubber to protect the thinned skull as shown in Figure 3B. Cover the surgical site using the head cap as shown in Figure 3C. Fix it in place with the two small pieces of wires going through both the head implant and head cap as shown in Figure 3D, E. Apply silicone rubber to cover the head cap and skull to stabilize the head cap further on the rat&#39;s head as shown in Figure 3F. 
	NOTE: Silicon rubber provides additional protection to thinned skull.</li>
	<li>Inject the rat with flunixin meglumine (2.5 mg/kg) subcutaneously for pain and inflammation management.&#160;To prevent infection, inject Enrosite antibiotic enrofloxacin (22.7mg/ml, 10mg/kg +/-.01), intraperitoneally.</li>
	<li>Move the rat to the recovery chamber to help maintain its body temperature with a warming blanket and a heat lamp. Monitor the rat continuously until it regains consciousness and can maintain sternal recumbency.</li>
	<li>Return the rat to its separate cage once it fully recovers.</li>
	<li>For the next 3 days, administer flunixin and buprenorphine to alleviate inflammation and pain and enrosite to prevent infection twice daily.</li>
</ol>
6. Awake imaging
<ol>
	<li>Anesthetize the rat with 4% isoflurane for induction and 1% for maintenance when there is no hind paw pinch reflex. Inject acepromazine (0.3-0.5 mg/kg) subcutaneously. 
	NOTE: This concentration of acepromazine is below mild sedation levels and only helps keep the rats calm throughout the imaging process.</li>
	<li>Using customized strips of Velcro, fix the rat on the plastic sheet used during the training procedures. Wrap the lower body part using an absorption pad and place the rat snugly in the sling.</li>
	<li>Remove the silicon rubber. Remove the head cap by removing the fixation wires. Fix the headframe in the head implant as shown in Figure 2G.</li>
	<li>Lock the headframe in clamps as shown in Figure 2H, I.</li>
	<li>Remove gas anesthesia. Flush the imaging area with saline 3x and clean with wet gauze. Dry the imaging area and make a well, using petroleum jelly, around the imaging area. Fill the well with sterilized saline and cover with a glass slide (Figure 2E).</li>
	<li>Refer to the acquisition procedures for intrinsic signal optical imaging, the whisker stimulation protocol, and data analysis and presentation, which have been discussed in detail previously6,7.</li>
	<li>Throughout the experiment, monitor the rats for signs of agitation and restlessness, which can be further reduced by covering the eyes of the rats with a soft cloth or gauze (optional).</li>
</ol>

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The authors have no conflicts of interest to disclose.

The use of awake, head-fixed rat imaging offers many advantages in terms of ease and customization. The custom-designed slings allow the rats to be wrapped through breathable netting material, eliminating the need to enclose animals in closed, plastic restraining chambers for extended periods of time10,11. Rats are kept calm and stress-free throughout the long durations of successive imaging sessions using a very low dose of acepromazine below the levels of mild ...

Most of the basic, preclinical, and translational scientific neuroimaging investigations are acquired from anesthetized animals1,2. Anesthetics ease experimentation but continuously influence the brain&#39;s and body&#39;s metabolism, blood pressure, and heart rate3. The type of anesthetic and the duration and route of administration add confounding variables to data interpretation that could contribute to reproducibility and translational failures4. A major bottleneck of awake, head-fixed rat neuroimaging studies is the requirement to keep the rat stationary and calm throughout the preparation and data acquisition processes. Small movements produce unwarranted motion artifacts, which can adversely affect data analysis and interpretations.
A new model of neuroimaging from awake, head-fixed rats using customized slings, three-dimensional (3D)-printed head implants, head caps, and a headframe has been devised that offers several advantages for easy experimentation. The 3D head implant is light and covers a small portion of the skull needed for transfixing. The 3D-printed head implants and caps are designed using computer-aided design (CAD) software. The protocols of whisker stimulation, data acquisition, data analysis, and results from anesthetized rats have been described in detail in previous work5,6,7.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr >
			<td >Rats</td>
			<td >Charles River</td>
			<td >Sprague Dawley</td>
			<td ></td>
		</tr>
		<tr >
			<td >Isoflurane</td>
			<td >Pivetal</td>
			<td >21295098</td>
			<td>General anesthetic</td>
		</tr>
		<tr >
			<td >Lidocaine HCl 2% injection</td>
			<td >Phoenix</td>
			<td >L-2000-04</td>
			<td>Local anesthetic</td>
		</tr>
		<tr >
			<td >Atropine sulfate injection</td>
			<td >Vedco</td>
			<td >5098907512</td>
			<td>Help in respiration</td>
		</tr>
		<tr >
			<td >Lactated Ringer&#39;s injection solution</td>
			<td >Vedco</td>
			<td >50989088317</td>
			<td></td>
		</tr>
		<tr >
			<td >Flunixin injection</td>
			<td >Vedco</td>
			<td >6064408670</td>
			<td>Pain management</td>
		</tr>
		<tr >
			<td >Enrosite injection (Enrofloxacin 2.27%)</td>
			<td >VetOne</td>
			<td >501084</td>
			<td>Avoid infection</td>
		</tr>
		<tr >
			<td >PromAce injection (Acepromazine maleate)</td>
			<td >Beohringer Ingelheim</td>
			<td >136059</td>
			<td></td>
		</tr>
		<tr >
			<td >Animax ointment</td>
			<td >Dechra Veterinary Products</td>
			<td >122-75</td>
			<td>active ingredients of nystatin 1000units per gram, neomycin sulfate 2.5mg per gram, thiostrepton 2500 units per gram, and triamcinolone acetonide 1mg per gram</td>
		</tr>
		<tr >
			<td >Puralube ophthalmic ointment</td>
			<td >Dechra Veterinary Products</td>
			<td >211-38</td>
			<td></td>
		</tr>
		<tr >
			<td >Povidone-iodine PVP prep pads</td>
			<td >Medline</td>
			<td >MDS093917</td>
			<td>Betadine generic</td>
		</tr>
		<tr >
			<td >Isopropyl alcohol swabs</td>
			<td >BD</td>
			<td >326895</td>
			<td></td>
		</tr>
		<tr >
			<td >Vetbond tissue adhesive</td>
			<td >3M</td>
			<td >1469SB</td>
			<td></td>
		</tr>
		<tr >
			<td >Bur (drill bit), standard operatory carbide</td>
			<td >SS White Burs</td>
			<td >14829</td>
			<td>#3 bur</td>
		</tr>
		<tr >
			<td >Screws, 00-90 x 1/8 flat head stainless steel</td>
			<td >J.I. Morris</td>
			<td >F0090CE125</td>
			<td>Anchor screws</td>
		</tr>
		<tr >
			<td >Stereotaxic system</td>
			<td >Kopf Instruments</td>
			<td >1430</td>
			<td></td>
		</tr>
		<tr >
			<td >Homeothermic heating blanket</td>
			<td >Harvard Apparatus</td>
			<td >50-7220-F</td>
			<td></td>
		</tr>
		<tr >
			<td >Pulse oximeter &#38; heart rate monitor</td>
			<td >Kent Scientific</td>
			<td >MouseStat Jr.</td>
			<td></td>
		</tr>
		<tr >
			<td >Petrolatum</td>
			<td >Fisher Scientific</td>
			<td >P66-1LB</td>
			<td>Vaseline generic</td>
		</tr>
		<tr >
			<td >Wire, bare copper</td>
			<td >Fisher Scientific</td>
			<td >15-545-2C</td>
			<td>20 gauge</td>
		</tr>
		<tr >
			<td >Teets Cold Cure powder</td>
			<td >Pearson Dental</td>
			<td >C73-0054&#160;</td>
			<td>active ingredient: Methyl Methacrylate</td>
		</tr>
		<tr >
			<td >Teets Cold Cure liquid</td>
			<td >Pearson Dental</td>
			<td >C73-0078&#160;</td>
			<td>active ingredient: Methyl Methacrylate</td>
		</tr>
		<tr >
			<td >Silicone mold rubber</td>
			<td >Smooth-On</td>
			<td >Body Double Fast</td>
			<td>silicon polymer</td>
		</tr>
		<tr >
			<td >Metricide 28 (Germicide)</td>
			<td >Metrex</td>
			<td >Oct-05</td>
			<td></td>
		</tr>
		<tr >
			<td >India ink, black</td>
			<td >Pelikan</td>
			<td >301051</td>
			<td></td>
		</tr>
		<tr >
			<td >Dental drill</td>
			<td >NSK Dental</td>
			<td >Ultimate XL-F</td>
			<td></td>
		</tr>
		<tr >
			<td >3D printer</td>
			<td >Prusa Research</td>
			<td >i3 MK3S+</td>
			<td></td>
		</tr>
		<tr >
			<td >Sew on fasteners</td>
			<td >Velcro</td>
			<td >90030</td>
			<td></td>
		</tr>
		<tr >
			<td >Pet screening utility fabric</td>
			<td >Joann</td>
			<td >10173334</td>
			<td>Netting material</td>
		</tr>
		<tr >
			<td >Bur (drill bit), standard operatory carbide</td>
			<td >SS White Burs</td>
			<td >14829</td>
			<td>#1 bur</td>
		</tr>
	</tbody>
</table>

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.

The representative optical imaging signals from a single trial of an anesthetized rat and the summed response (of 40 collected trials) of an awake rat are shown (Figure 4). The signal intensity for single-whisker stimulation of an awake rat can be visualized at a higher threshold than for the anesthetized rat, showing a stronger signal from the awake animal. The C2 whiskers of rats are stimulated at 5 Hz for 1 s, and the functional response is displayed as a fractional change compared to the...

Watch this Scientific Journal Video about Head Implants for the Neuroimaging of Awake, Head-Fixed Rats at JoVE.com

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

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 ...

head-implants-for-the-neuroimaging-of-awake-head-fixed-rats

Research

JoVE Journal

Neuroscience

2.3K Views.  University of California Irvine. A detailed new procedure for functional imaging of awake, head-fixed rats is described.

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

Visualizing Protein Kinase A Activity In Head-fixed Behaving Mice Using In Vivo Two-photon Fluorescence Lifetime Imaging Microscopy

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. Despite their importance, technologies for tracking neuromodulatory events with cellular resolution are just beginning to emerge. Neuromodulators, such as dopamine, norepinephrine, acetylcholine, and serotonin, trigger intracellular signaling events via their respective G protein-coupled receptors to modulate neuronal excitability, synaptic communications, and other neuronal functions, thereby regulating information processing in the neuronal network. The above mentioned neuromodulators converge onto the cAMP/protein kinase A (PKA) pathway. Therefore, in vivo PKA imaging with single-cell resolution was developed as a readout for neuromodulatory events in a manner analogous to calcium imaging for neuronal electrical activities. Herein, a method is presented to visualize PKA activity at the level of individual neurons in the cortex of head-fixed behaving mice. To do so, an improved A-kinase activity reporter (AKAR), called tAKAR&#945;, is used, which is based on F&#246;rster resonance energy transfer (FRET). This genetically-encoded PKA sensor is introduced into the motor cortex via in utero electroporation (IUE) of DNA plasmids, or stereotaxic injection of adeno-associated virus (AAV). FRET changes are imaged using two-photon fluorescence lifetime imaging microscopy (2pFLIM), which offers advantages over ratiometric FRET measurements for quantifying FRET signal in light-scattering brain tissue. To study PKA activities during enforced locomotion, tAKAR&#945; is imaged through a chronic cranial window above the cortex of awake, head-fixed mice, which run or rest on a speed-controlled motorized treadmill. This imaging approach will be applicable to many other brain regions to study corresponding behavior-induced PKA activities and to other FLIM-based sensors for in vivo imaging.

A procedure is presented to visualize protein kinase A activities in head-fixed, behaving mice. An improved A-kinase activity reporter, tAKAR&#945;, is expressed in cortical neurons and made accessible for imaging through a cranial window. Two-photon fluorescence lifetime imaging microscopy is used to visualize PKA activities in vivo during enforced locomotion.

Neuromodulation exerts powerful control over brain function. Dysfunction of neuromodulatory systems results in neurological and psychiatric disorders. ...

Injections of AAV Vectors for Optogenetics in Anesthetized and Awake Behaving Non-Human Primate Brain

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of optogenetics, however, requires that safety and efficacy be demonstrated in animal models, ideally in non-human primates (NHPs), because of their neurological similarity to humans. The number of candidate vectors that are potentially useful for neuroscience and medicine is vast, and no high-throughput means to test these vectors yet exists. Thus, there is a need for techniques to make multiple spatially and volumetrically accurate injections of viral vectors into NHP brain that can be identified unambiguously through postmortem histology. Described herein is such a method. Injection cannulas are constructed from coupled polytetrafluoroethylene and stainless-steel tubes. These cannulas are autoclavable, disposable, and have low minimal-loading volumes, making them ideal for the injection of expensive, highly concentrated viral vector solutions. An inert, red-dyed mineral oil fills the dead space and forms a visible meniscus with the vector solution, allowing instantaneous and accurate measurement of injection rates and volumes. The oil is loaded into the rear of the cannula, reducing the risk of co-injection with the vector. Cannulas can be loaded in 10 min, and injections can be made in 20 min. This procedure is well suited for injections into awake or anesthetized animals. When used to deliver high-quality viral vectors, this procedure can produce robust expression of optogenetic proteins, allowing optical control of neural activity and behavior in NHPs.

As currently implemented, optogenetics in non-human primates requires injection of viral vectors into the brain. An optimal injection method should be reliable and, for many applications, capable of targeting individual sites of arbitrary depth that are readily and unambiguously identified in postmortem histology. An injection method with these properties is presented.

Optogenetic techniques have revolutionized neuroscience research and are poised to do the same for neurological gene therapy. The clinical use of ...

Abdominal VNS for Treating Inflammatory Conditions in Rat Models

Implantation Surgery for Abdominal Vagus Nerve Stimulation and Recording Studies in Awake Rats

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does not have any respiratory and cardiac off-target effects commonly associated with cervical VNS. The lack of respiratory and cardiac off-target effects means that the intensity of stimulation does not need to be lowered to reduce side effects commonly experienced during cervical VNS. Few recent studies demonstrate the anti-inflammatory effects of abdominal VNS in rat models of inflammatory bowel disease, rheumatoid arthritis, and glycemia reduction in a rat model of type 2 diabetes. Rat is a great model to explore the potential of this technology because of the well-established anatomy of the vagus nerve, the large size of the nerve that allows easy handling, and the availability of many disease models. Here, we describe the methods for cleaning and sterilizing the abdominal VNS electrode array and surgical protocol in rats. We also describe the technology required for confirmation of suprathreshold stimulation by recording evoked compound action potentials. Abdominal VNS has the potential to offer selective, effective treatment for a variety of conditions, including inflammatory diseases, and the application is expected to expand similarly to cervical VNS.

Author Spotlight: Exploring Abdominal VNS for Inflammatory Conditions

The present protocol describes the surgical technique for implanting an electrode array onto the abdominal vagus nerve in rats, along with methods for chronic electrophysiology testing and stimulation using the implanted device.

Abdominal vagus nerve stimulation (VNS) can be applied to the subdiaphragmatic branch of the vagus nerve of rats. Due to its anatomical location, it does ...

Multilayer Imaging with Microprism in Head-Fixed and Freely Moving Animals

Long-Term Imaging of Identified Neural Populations using Microprisms in Freely Moving and Head-Fixed Animals

With the advancement of multi-photon microscopy and molecular technologies, fluorescence imaging is rapidly growing to become a powerful approach for studying the structure, function, and plasticity of living brain tissues. In comparison to conventional electrophysiology, fluorescence microscopy can capture the neural activity as well as the morphology of the cells, enabling long-term recordings of the identified neuron populations at single-cell or subcellular resolution. However, high-resolution imaging typically requires a stable, head-fixed setup that restricts the movement of the animal, and the preparation of a flat surface of transparent glass allows visualization of neurons at one or more horizontal planes but is limited in studying the vertical processes running across different depths. Here, we describe a procedure to combine a head plate fixation and a microprism that gives multilayer and multimodal imaging. This surgical preparation not only gives access to the entire column of the mouse visual cortex but allows two-photon imaging in a head-fixed position and one-photon imaging in a freely moving paradigm. Using this approach, one can sample identified cell populations across different cortical layers, register their responses under head-fixed and freely moving states, and track the long-term changes over months. Thus, this method provides a comprehensive assay of the microcircuits, enabling direct comparison of neural activities evoked by well-controlled stimuli and under a natural behavioral paradigm.

Author Spotlight: Comparative Imaging of Neural Activity in Awake and Freely Moving States

When integrated with a head-plate and an optical design compatible with both single- and two-photon microscopes, the microprism lens presents a significant advantage in measuring neural responses in a vertical column under diverse conditions, including well-controlled experiments in head-fixed states or natural behavioral tasks in freely moving animals.

With the advancement of multi-photon microscopy and molecular technologies, fluorescence imaging is rapidly growing to become a powerful approach for ...

Fabrication and Surgical Implantation of Electromyographic Electrodes in Mice

Recording Forelimb Muscle Activity in Head&#45;Fixed Mice with Chronically Implanted EMG Electrodes

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with unprecedented precision in head-fixed mice performing a variety of tasks. The small size of the mouse makes the measurement of motor output difficult, as the traditional method of electromyographic (EMG) recording of muscle activity was designed for larger animals like cats and primates. Pending commercially available EMG electrodes for mice, the current gold-standard method for recording muscle activity in mice is to make electrode sets in-house. This article describes a refinement of established procedures for hand fabrication of an electrode set, implantation of electrodes in the same surgery as headplate implantation, fixation of a connector on the headplate, and post-operative recovery care. Following recovery, millisecond-resolution EMG recordings can be obtained during head-fixed behavior for several weeks without noticeable changes in signal quality. These recordings enable precise measurement of forelimb muscle activity alongside in vivo neural recording and/or perturbation to probe mechanisms of motor control in mice.

Author Spotlight: Investigating Mouse Motor Cortex Interactions from Muscle Activity to Neural Dynamics

This protocol describes the hand fabrication and surgical implantation of electromyographic (EMG) electrodes in the forelimb muscles of mice to record muscle activity during head-fixed behavior experiments.

Powerful genetic and molecular tools available in mouse systems neuroscience research have enabled researchers to interrogate motor system function with ...

Open&#45;Source Implant for Multi&#45;Region Electrophysiological Monitoring in Rats

The TD Drive: A Parametric, Open-Source Implant for Multi&#45;Area Electrophysiological Recordings in Behaving and Sleeping Rats

Intricate interactions between multiple brain areas underlie most functions attributed to the brain. The process of learning, as well as the formation and consolidation of memories, are two examples that rely heavily on functional connectivity across the brain. In addition, investigating hemispheric similarities and/or differences goes hand in hand with these multi-area interactions. Electrophysiological studies trying to further elucidate these complex processes thus depend on recording brain activity at multiple locations simultaneously and often in a bilateral fashion. Presented here is a 3D-printable implant for rats, named TD Drive, capable of symmetric, bilateral wire electrode recordings, currently in up to ten distributed brain areas simultaneously. The open-source design was created employing parametric design principles, allowing prospective users to easily adapt the drive design to their needs by simply adjusting high-level parameters, such as anterior-posterior and mediolateral coordinates of the recording electrode locations. The implant design was validated in n = 20 Lister Hooded rats that performed different tasks. The implant was compatible with tethered sleep recordings and open field recordings (Object Exploration) as well as wireless recording in a large maze using two different commercial recording systems and headstages. Thus, presented here is the adaptable design and assembly of a new electrophysiological implant, facilitating fast preparation and implantation.

Author Spotlight: Deciphering Memory and Learning Through Neural Implants for Multi&#45;Region Brain Studies

Here, we present a unique, 3D-printable implant for rats, named TD Drive, capable of symmetric, bilateral wire electrode recordings, currently in up to ten distributed brain areas simultaneously.

Intricate interactions between multiple brain areas underlie most functions attributed to the brain. The process of learning, as well as the formation and ...

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. ...

Fabrication of TD Drive Implants for Electrophysiological Recordings of Sleeping and Live Rats

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 ...