We would like to thank Dr. Kellie Breen Church, Mr. Michael Kreisman, and Ms. Jessica Jang for their contributions to collecting the data shown in the representative results. This work was supported by the National Institutes of Health (NIH) R00 HD104994 (R.B.M.).

All the procedures were approved by the Colorado State University (#3960) and University of California San Diego Institutional Animal Care and Use Committees, where the representative data were collected (S13235, PI Kellie Breen Church). Data from five adult female and two adult male C57/BL6 mice (9-16 weeks old) are depicted in the representative data section. Female mice were ovariectomized 3-4 weeks prior to ICV injection and blood collection as described previously7. Prior to experimentation, these mice were housed with a 12 h light/12 h dark light cycle and had free access to feed and water in accordance with the Guide for the Care and Use of Laboratory Animals.
1. Performing craniotomy
NOTE: Craniotomy can be performed one or more days prior to the actual injection, which makes the injection process quicker on the day of the experiment.
<ol>
	<li>Prepare the materials for the craniotomy as described below.
	<ol>
		<li>Place a clean bench pad or drape material on the work surface. Tape the nose cone of an isoflurane vaporizer to the work surface (close to, but facing away from, the experimenter; see Figure 1A).</li>
		<li>Place silastic tubing on the end of a sterile sharp 18 G needle so that ~1 mm of the needle tip protrudes. Place silastic tubing on the end of a sterile blunt needle so that ~1 mm of the needle tip protrudes. 
		NOTE: Separate and sterile pre-packaged needles (sharp and blunt) should be used for each mouse.</li>
		<li>Gather an electric shaver, iodine scrub, sterile gauze, and alcohol pads for preparing the injection site.</li>
	</ol>
	</li>
	<li>Practice moving a needle 2 mm lateral and 1 mm caudal as described below.
	<ol>
		<li>Use a ruler and a pen to mark a sheet of paper with a starting point (this will be bregma), a mark 2 mm to either the left or right, and another mark 1 mm above the last one (this will be the injection site; see Figure 1B).</li>
		<li>Practice moving the needle from the starting point to mark point 1 to mark point 2 (bregma to injection site) until confident that the movement is repeatable without guides. 
		NOTE: The coordinates described here are effective in adult C57/BL6 mice, but other strains or age groups may require adjustment.</li>
	</ol>
	</li>
	<li>Prepare the mouse for craniotomy as described below.
	<ol>
		<li>Place the mouse in an induction chamber and anesthetize the mouse with 3%-4% isoflurane in medical-grade oxygen. With practice and familiarity with the procedure, reduce the total duration of isoflurane exposure to less than 10 min for the craniotomy procedure. 
		CAUTION: Isoflurane exposure is hazardous to human health; use an approved and inspected isoflurane vaporizer in a well-ventilated area with means of scavenging waste gases.</li>
		<li>Once the toe reflex is absent, shave the head of the animal. Apply eye lubricant.</li>
		<li>Place the mouse on the work surface with the head in the nose cone to maintain anesthesia.</li>
		<li>Administer an analgesic agent, such as buprenorphine (0.6-0.8 mg/kg mouse body weight, subcutaneously), as directed by the supplier.</li>
		<li>Clean the injection site by wiping the head with sterile gauze dipped in an iodine solution (perform three scrubs), and then wipe with an alcohol scrub pad (perform three times). 
		&#8203;NOTE: Remove the fur and clean the skin around the injection site and use sterile instruments and needles, as recommended by the American Veterinary Medical Association to prevent infections.</li>
	</ol>
	</li>
	<li>Firmly hold the head of the mouse with the non-dominant hand. Position the head as flat on the work surface as possible.</li>
	<li>Locate the site of the injection.
	<ol>
		<li>Use the prepared sharp 18 G needle to first identify bregma; for this, drag the needle across the skin of the head along the midline, moving in the rostral-caudal plane. Imagine an equilateral triangle in which the two vertices are the eyes and the third vertex is the approximate location of bregma (see Figure 1C).</li>
		<li>From bregma, move the needle 2 mm lateral and 1 mm caudal to the site of injection.</li>
	</ol>
	</li>
	<li>While holding the needle vertically, firmly push the needle through the skin and bone until the tubing is flush with the skin.</li>
	<li>Retract the needle, rotate the needle, and press through the skin and bone again. Repeat the process until a small hole is created in the bone.</li>
	<li>Use the blunt needle to check that a sufficient hole has been produced in the bone. Aim to make an opening in the bone large enough for the blunt needle to pass through.</li>
	<li>Remove the mouse from the nose cone, clean off any blood from the injection site with sterile gauze, and place the mouse in a cage on a cage warmer until awake. Since the opening made by the craniotomy is small (18 G needle), the site does not need to be closed with a suture or wound clips. 
	NOTE: The opening produced here is small (18 G), so many institutions consider this procedure an injection as opposed to a surgery. Moreover, although an opening through skin and bone to the brain is made, holding the skin taut results in the holes not aligning, which likely aids in preventing skin flora or cage bedding from contacting the brain.</li>
</ol>
2. Making the injection
<ol>
	<li>Prepare the materials for the injection as described below.
	<ol>
		<li>Place a clean bench pad or drape material on the work surface. Tape the nose cone of an isoflurane vaporizer to the work surface (close to, but facing away from, the experimenter; see Figure 1A).</li>
		<li>Attach a 10 mm long 27 G needle with a 45&#176; bevel onto a 5 &#181;L glass syringe. Place silastic tubing on the needle so that 3.5 mm of the needle tip protrudes; use laboratory tape to hold the tubing to the syringe body (see Figure 1D).</li>
		<li>Prepare the injection media (drug, virus, or other liquid for injection) in a tube. For the representative results in this study, sterile isotonic saline was injected. 
		NOTE: Select a tube that permits access by the glass syringe and needle, such as a 2 mL microcentrifuge tube.</li>
		<li>Draw 3 &#181;L of injection media into the glass syringe. First, draw more than the desired volume, and eject until 3 &#181;L remains in the syringe.</li>
		<li>Gather iodine scrub, sterile gauze, and alcohol pads for preparing the injection site.</li>
		<li>Start a laboratory timer in count-up mode and place the timer so that it is visible to the experimenter while making injections.</li>
		<li>Practice moving a needle 2 mm lateral and 1 mm caudal as performed on the previous day.</li>
	</ol>
	</li>
	<li>Prepare the mouse for injection as described below.
	<ol>
		<li>Place the mouse in the induction chamber and anesthetize the mouse with isoflurane. Once the toe reflex is absent, apply eye lubricant, and place the mouse on the work surface with the head in the nose cone.</li>
		<li>Clean the injection site with three wipes each of iodine and alcohol. Identify the injection site with an 18 G needle (or blunted needle) as described in step 1.8. The hole created during craniotomy should be detectable. 
		NOTE: If the craniotomy has not been performed in advance, or if the hole in the skull is not detectable, the craniotomy can be performed here.</li>
	</ol>
	</li>
	<li>Perform the injection with a glass syringe as described below.
	<ol>
		<li>Firmly hold the head of the mouse with the non-dominant hand. Position the head as flat on the work surface as possible.</li>
		<li>Place the needle of the glass syringe through the hole in the skull until the tubing placed over the needle prepared in step 2.1.2 rests on the skin of the mouse.</li>
		<li>Hold the syringe as vertically as possible, paying attention to both the coronal and sagittal planes. Slowly inject the media over a period of 1 min.</li>
		<li>Hold the syringe and needle in place for another minute after the completion of the injection to minimize backflow. Slowly retract the needle from the head of the mouse.</li>
		<li>Remove the mouse from the nose cone, clean off any blood from the injection site with sterile gauze (if there is any present), and place the mouse in a cage on a cage warmer until awake.</li>
	</ol>
	</li>
</ol>
3. Confirming the injection location
<ol>
	<li>Deeply anesthetize and perfuse the mouse with saline and 4% paraformaldehyde in phosphate-buffered saline (PBS) as previously described8. Store the brain in 30% sucrose in PBS at 4 &#176;C until the brain sinks (typically 2 days).</li>
	<li>Cut 20-50 &#181;m coronal sections on a cryostat as described previously9. While sectioning, observe the injection tract in the tissue block. Record whether the injection tract clearly intersects the lateral ventricle.</li>
</ol>

Performing Intracerebroventricular Injection in Mice Using a Free-Hand Approach

<ol>
	<li>Roseweir, A. K., 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=Discovery+of+potent+kisspeptin+antagonists+delineate+physiological+mechanisms+of+gonadotropin+regulation.">Discovery of potent kisspeptin antagonists delineate physiological mechanisms of gonadotropin regulation.</a> Journal of Neuroscience. 29 (12), 3920-3929 (2009).</li><li>Kim, J. Y., Grunke, S. D., Levites, Y., Golde, T. E., Jankowsky, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracerebroventricular+viral+injection+of+the+neonatal+mouse+brain+for+persistent+and+widespread+neuronal+transduction.">Intracerebroventricular viral injection of the neonatal mouse brain for persistent and widespread neuronal transduction.</a> Journal of Visualized Experiments. (91), e51863 (2014).</li><li>Taylor, Z. V., Khand, B., Porgador, A., Monsonego, A., Eremenko, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+optimized+intracerebroventricular+injection+of+CD4(+)+T+cells+into+mice.">An optimized intracerebroventricular injection of CD4(+) T cells into mice.</a> STAR Protocols. 2 (3), 100725 (2021).</li><li>Russo, K. A., 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=Circadian+control+of+the+female+reproductive+axis+through+gated+responsiveness+of+the+RFRP-3+system+to+VIP+signaling.">Circadian control of the female reproductive axis through gated responsiveness of the RFRP-3 system to VIP signaling.</a> Endocrinology. 156 (7), 2608-2618 (2015).</li><li>Laursen, S. E., Belknap, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracerebroventricular+injections+in+mice.+Some+methodological+refinements.">Intracerebroventricular injections in mice. Some methodological refinements.</a> Journal of Pharmacological Methods. 16 (4), 355-357 (1986).</li><li>Haley, T. J., McCormick, W. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacological+effects+produced+by+intracerebral+injection+of+drugs+in+the+conscious+mouse.">Pharmacological effects produced by intracerebral injection of drugs in the conscious mouse.</a> British Journal of Pharmacology and Chemotherapy. 12 (1), 12-15 (1957).</li><li>McCosh, R. B., 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=Insulin-induced+hypoglycaemia+suppresses+pulsatile+luteinising+hormone+secretion+and+arcuate+Kiss1+cell+activation+in+female+mice.">Insulin-induced hypoglycaemia suppresses pulsatile luteinising hormone secretion and arcuate Kiss1 cell activation in female mice.</a> Journal of Neuroendocrinology. 31 (12), e12813 (2019).</li><li>Wu, J., 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=Transcardiac+perfusion+of+the+mouse+for+brain+tissue+dissection+and+fixation.">Transcardiac perfusion of the mouse for brain tissue dissection and fixation.</a> Bio-Protocol. 11 (5), e3988 (2021).</li><li>Comba, A., 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=Laser+capture+microdissection+of+glioma+subregions+for+spatial+and+molecular+characterization+of+intratumoral+heterogeneity,+oncostreams,+and+invasion.">Laser capture microdissection of glioma subregions for spatial and molecular characterization of intratumoral heterogeneity, oncostreams, and invasion.</a> Journal of Visual Experiments. (158), e60939 (2020).</li><li>Porteous, 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=Reformulation+of+PULSAR+for+analysis+of+pulsatile+LH+secretion+and+a+revised+model+of+estrogen-negative+feedback+in+mice.">Reformulation of PULSAR for analysis of pulsatile LH secretion and a revised model of estrogen-negative feedback in mice.</a> Endocrinology. 162 (11), (2021).</li><li>Hohmann, J. G., 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=Differential+role+of+melanocortins+in+mediating+leptin's+central+effects+on+feeding+and+reproduction.">Differential role of melanocortins in mediating leptin's central effects on feeding and reproduction.</a> American Journal of Physiology: Regulatory, Integrative, and Comparative Physiology. 278 (1), R50-R59 (2000).</li><li>Gottsch, M. L., 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=A+role+for+kisspeptins+in+the+regulation+of+gonadotropin+secretion+in+the+mouse.">A role for kisspeptins in the regulation of gonadotropin secretion in the mouse.</a> Endocrinology. 145 (9), 4073-4077 (2004).</li><li>Krasnow, S. 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=A+role+for+galanin-like+peptide+in+the+integration+of+feeding,+body+weight+regulation,+and+reproduction+in+the+mouse.">A role for galanin-like peptide in the integration of feeding, body weight regulation, and reproduction in the mouse.</a> Endocrinology. 144 (3), 813-822 (2003).</li></ol>

The authors have nothing to disclose.

Here, a simple and effective means for making ICV injections in mice is described. Since this technique does not require a stereotaxic frame, this approach for the central delivery of drugs and experimental agents is accessible to more researchers. Additionally, this approach is relatively high throughput since the preparation and injection procedure can be performed quickly.
Since this procedure requires the manipulation of needles and a glass syringe by hand using approximate distances, it i...

The delivery of experimental agents, such as drugs1, viruses2, or cells3, to the brain is often necessary for neuroendocrine research. If the agent does not readily cross the blood-brain barrier or the experimental objective is to specifically test the central effects of the agent, it is important to have a reliable method for delivering injections into the brain. Moreover, injection into the intracerebroventricular (ICV) space provides the opportunity to distribute the agent widely in the brain and provides a large target area, thus increasing the likelihood of successful injection2. 
A common method for making ICV injections involves the placement of a permanent indwelling cannula. In this approach, a stereotaxic frame is necessary to position the commercially available or custom-made cannula, as the cannula is glued or cemented in place. Often, upon recovery, a supraphysiological dose of angiotensin II is administered through the cannula, and if drinking behavior is immediately observed, then the cannula is considered correctly placed4. This approach has many advantages, including the ability to perform long-term infusion and the ability to inject the same animal multiple times; additionally, if angiotensin II is employed, correct placement can be confirmed prior to the administration of experimental compounds. However, there are some limitations to placing a permanent cannula, including the requirement for expensive equipment (stereotaxic frame), the possibility of damage to the cannula after placement (e.g., mice can chew on the cannula of a cage mate), and the possibility of infections around the permanent cannula. Single ICV injections can be made with the use of a stereotaxic frame3, which, although effective, requires substantial exposure to anesthesia and, thus, may obscure some acute physiological and behavioral effects of the treatment. Additionally, the placement of mice in a stereotaxic frame requires substantial training to achieve stable placement and prevent the rupturing of the ear canals.
Here, an established method for making free-hand injections in mice is described. This method is based on previous reports5,6. The advantages of this technique are that it is simple, rapid, and does not require specialized equipment such as a stereotaxic frame. As described below, this procedure involves manipulating a glass syringe relative to landmarks on the mouse head to make the injections, which can be done rapidly and, thus, requires only a few minutes of gas anesthesia on the experimental day.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>18-gauge blunt needles</td>
			<td>SAI Infusion</td>
			<td>B18-150</td>
			<td></td>
		</tr>
		<tr>
			<td>18-gauge needles</td>
			<td>BD Medical</td>
			<td>305195</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol pads</td>
			<td>Fisher Scientific</td>
			<td>22-363-750</td>
			<td></td>
		</tr>
		<tr>
			<td>Bench pad</td>
			<td>Fisher Scientific</td>
			<td>14-206-62AC22</td>
			<td></td>
		</tr>
		<tr>
			<td>Betadine solution</td>
			<td>Fisher Scientific</td>
			<td>NC1696484</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td>Patterson Vet Supply</td>
			<td>07-892-5235</td>
			<td>Controlled substance</td>
		</tr>
		<tr>
			<td>Eyelube</td>
			<td>Fisher Scientific</td>
			<td>50-218-8442</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass syringe</td>
			<td>Hamilton</td>
			<td>7634-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Injection needle</td>
			<td>Hamilton</td>
			<td>7803-01</td>
			<td>27 gauge, Small Hub RN needle, point style: 4, Needle length: 10cm, Angle: 45</td>
		</tr>
		<tr>
			<td>Isoflurane&#160;&#160;</td>
			<td>Patterson Vet Supply</td>
			<td>07-893-8441</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer</td>
			<td>Vet Equip</td>
			<td>V-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory Tape</td>
			<td>VWR</td>
			<td>89098-128</td>
			<td></td>
		</tr>
		<tr>
			<td>Medical grade oxygen</td>
			<td>Airgas</td>
			<td>OX USPEA</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Millipore-Sigma</td>
			<td>8.18715.1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline</td>
			<td>Fisher Scientific</td>
			<td>J67802.K2</td>
			<td></td>
		</tr>
		<tr>
			<td>PulsaR Software</td>
			<td>Open source, University of Otago</td>
			<td></td>
			<td>See ref 9</td>
		</tr>
		<tr>
			<td>Ruler</td>
			<td>Fisher Scientific</td>
			<td>12-00-152</td>
			<td></td>
		</tr>
		<tr>
			<td>Silastic tubing (0.040&#34; I.D.)</td>
			<td>DOW</td>
			<td>508-005</td>
			<td></td>
		</tr>
		<tr>
			<td>Silastic tubing (0.078&#34; I.D.)</td>
			<td>DOW</td>
			<td>508-009</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile saline</td>
			<td>VWR</td>
			<td>101320-574</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose&#160;</td>
			<td>Fisher Scientific</td>
			<td>S5-500</td>
			<td></td>
		</tr>
	</tbody>
</table>

free-hand intracerebroventricular injections in mice

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. An intracerebroventricular (ICV) injection allows the widespread delivery of the experimental agent throughout the brain (particularly in the structures near the ventricles). Here, methods for making free-hand ICV injections in adult mice are described. By using visual and tactile landmarks on the heads of mice, injections into the lateral ventricles can be made rapidly and reliably. The injections are made with a glass syringe held in the experimenter&#39;s hand and placed at approximate distances from the landmarks. Thus, this technique does not require a stereotaxic frame. Furthermore, this technique requires only brief isoflurane anesthesia, which permits the subsequent assessment of mouse behavior and/or physiology in awake, freely behaving mice. Free-hand ICV injection is a powerful tool for the efficient delivery of experimental agents into the brains of living mice and can be combined with other techniques such as frequent blood sampling, neural circuit manipulation, or in vivo recording to investigate neuroendocrine processes.

When performed successfully, this technique allows the rapid delivery of an experimental agent into the ventricular system. A luteinizing hormone (LH) pulse profile from an ovariectomized mouse that received an ICV injection of 3 &#181;L of sterile isotonic saline, the vehicle for many pharmacological compounds, is shown in Figure 2A. This example demonstrates that brief exposure to gas anesthesia and the injection of 3 &#181;L of fluid into the ventricular system alone did not alter the pul...

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Author Spotlight: Insights, Innovations, and Future Avenues in Reproductive Neurocontrol 

Here, a simple and rapid approach for performing intracerebroventricular injections in mice using a free-hand approach (that is, without a stereotaxic device) is described.

Free-Hand Intracerebroventricular Injections in Mice

The investigation of neuroendocrine systems often requires the delivery of drugs, viruses, or other experimental agents directly into the brains of mice. ...

free-hand-intracerebroventricular-injections-in-mice

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10.0K Views.  Colorado State University. Here, a simple and rapid approach for performing intracerebroventricular injections in mice using a free-hand approach (that is, without a stereotaxic device) is described.

Video: Author Spotlight: Insights, Innovations, and Future Avenues in Reproductive Neurocontrol 

Measuring Motor Coordination in Mice

Mice are increasingly being used in behavioral neuroscience, largely replacing rats as the behaviorist's animal of choice. Before aspects of behavior such as emotionality or cognition can be assessed, however, it is vital to determine whether the motor capabilities of e.g. a mutant or lesioned mouse allow such an assessment. Performance on a maze task requiring strength and coordination, such as the Morris water maze, might well be impaired in a mouse by motor, rather than cognitive, impairments, so it is essential to selectively dissect the latter from the former. For example, sensorimotor impairments caused by NMDA antagonists have been shown to impair water maze performance2. 
Motor coordination has traditionally been assessed in mice and rats by the rotarod test, in which the animal is placed on a horizontal rod that rotates about its long axis; the animal must walk forwards to remain upright and not fall off. Both set speed and accelerating versions of the rotarod are available.
The other three tests described in this article (horizontal bar, static rods and parallel bars) all measure coordination on static apparatus. The horizontal bar also requires strength for adequate performance, particularly of the forelimbs as the mouse initially grips the bar just with the front paws. 
Adult rats do not perform well on tests such as the static rods and parallel bars (personal observations); they appear less well coordinated than mice. I have only tested male rats, however, and male mice seem generally less well coordinated than females. Mice appear to have a higher strength:weight ratio than rats; the Latin name, Mus musculus, seems entirely appropriate. The rotarod, the variations of the foot fault test12 or the Catwalk (Noldus)15 apparatus are generally used to assess motor coordination in rats.

Protocols are presented for two established motor coordination tasks, the accelerating rotarod and horizontal bar, also two tests developed in Oxford recently, the static rods and parallel bars. These tests can detect motor impairments potentially of interest in their own right, as well as being possible variables in tests of other areas of behavior.

Mice are increasingly being used in behavioral neuroscience, largely replacing rats as the behaviorist's animal of choice. Before aspects of behavior such ...

Video-oculography in Mice

Eye movements are very important in order to track an object or to stabilize an image on the retina during movement. Animals without a fovea, such as the mouse, have a limited capacity to lock their eyes onto a target. In contrast to these target directed eye movements, compensatory ocular eye movements are easily elicited in afoveate animals1,2,3,4. Compensatory ocular movements are generated by processing vestibular and optokinetic information into a command signal that will drive the eye muscles. The processing of the vestibular and optokinetic information can be investigated separately and together, allowing the specification of a deficit in the oculomotor system. The oculomotor system can be tested by evoking an optokinetic reflex (OKR), vestibulo-ocular reflex (VOR) or a visually-enhanced vestibulo-ocular reflex (VVOR). The OKR is a reflex movement that compensates for "full-field" image movements on the retina, whereas the VOR is a reflex eye movement that compensates head movements. The VVOR is a reflex eye movement that uses both vestibular as well as optokinetic information to make the appropriate compensation. The cerebellum monitors and is able to adjust these compensatory eye movements. Therefore, oculography is a very powerful tool to investigate brain-behavior relationship under normal as well as under pathological conditions (f.e. of vestibular, ocular and/or cerebellar origin).
Testing the oculomotor system, as a behavioral paradigm, is interesting for several reasons. First, the oculomotor system is a well understood neural system5. Second, the oculomotor system is relative simple6; the amount of possible eye movement is limited by its ball-in-socket architecture ("single joint") and the three pairs of extra-ocular muscles7. Third, the behavioral output and sensory input can easily be measured, which makes this a highly accessible system for quantitative analysis8. Many behavioral tests lack this high level of quantitative power. And finally, both performance as well as plasticity of the oculomotor system can be tested, allowing research on learning and memory processes9.
Genetically modified mice are nowadays widely available and they form an important source for the exploration of brain functions at various levels10. In addition, they can be used as models to mimic human diseases. Applying oculography on normal, pharmacologically-treated or genetically modified mice is a powerful research tool to explore the underlying physiology of motor behaviors under normal and pathological conditions. Here, we describe how to measure video-oculography in mice8.

Video-oculography is a very quantitative method to investigate ocular motor performance as well as motor learning. Here, we describe how to measure video-oculography in mice. Applying this technique on normal, pharmacologically-treated or genetically modified mice is a powerful research tool to explore the underlying physiology of motor behaviors.

Eye movements are very important in order to track an object or to stabilize an image on the retina during movement. Animals without a fovea, such as the ...

Intracerebroventricular Viral Injection of the Neonatal Mouse Brain for Persistent and Widespread Neuronal Transduction

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene expression in the mouse brain. Here we describe a technique first introduced by Passini and Wolfe for direct intracranial delivery of virally-encoded transgenes into the neonatal mouse brain. In its most basic form, the procedure requires only an ice bucket and a microliter syringe. However, the protocol can also be adapted for use with stereotaxic frames to improve consistency for researchers new to the technique. The method relies on the ability of adeno-associated virus (AAV) to move freely from the cerebral ventricles into the brain parenchyma while the ependymal lining is still immature during the first 12-24 hr after birth. Intraventricular injection of AAV at this age results in widespread transduction of neurons throughout the brain. Expression begins within days of injection and persists for the lifetime of the animal. Viral titer can be adjusted to control the density of transduced neurons, while co-expression of a fluorescent protein provides a vital label of transduced cells. With the rising availability of viral core facilities to provide both off-the-shelf, pre-packaged reagents and custom viral preparation, this approach offers a timely method for manipulating gene expression in the mouse brain that is fast, easy, and far less expensive than traditional germline engineering.

Here we demonstrate a technique for widespread neuronal transduction by intraventricular injection of adeno-associated virus into the neonatal mouse brain. This method provides a rapid and easy way to attain lifelong expression of virally-delivered transgenes.

With the pace of scientific advancement accelerating rapidly, new methods are needed for experimental neuroscience to quickly and easily manipulate gene ...

Simultaneous Electrophysiological Recording and Micro-injections of Inhibitory Agents in the Rodent Brain

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing system was developed that allows for the injection of a drug while recording electrophysiological signals from the affected neuronal population. This method provides a simple and economical alternative to commercial solutions. A glass pipette was modified by combining it with a hypodermic needle and a silver filament. The injectrode is attached to commercial microsyringe pump for drug delivery. This results in a technique that provides real-time pharmacodynamics feedback through multi-unit extracellular signals originating from the site of drug delivery. As a proof of concept, we recorded neuronal activity from the superior colliculus elicited by flashes of light in rats, concomitantly with delivery of drugs through the injectrode. The injectrode recording capacity permits the functional characterization of the injection site favoring precise control over the localization of drug delivery. Application of this method also extends far beyond what is demonstrated here, as the choice of chemical substance loaded into the injectrode is vast, including tracing markers for anatomic experiments.

Here, we craft a glass pipette with dual functions: inhibition of deep brain structures by microinjections of drugs and real-time monitoring of their effects through simultaneous electrophysiological recordings.

Here we describe a method for the construction of a single-use &#8220;injectrode&#8221; using commercially accessible and affordable parts. A probing ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Intracerebroventricular Injection of Amyloid-&#946; Peptides in Normal Mice to Acutely Induce Alzheimer-like Cognitive Deficits

Amyloid-&#946; (A&#946;) is a major pathological mediator of both familial and sporadic Alzheimer's disease (AD). In the brains of AD patients, progressive accumulation of A&#946; oligomers and plaques is observed. Such A&#946; abnormalities are believed to block long-term potentiation, impair synaptic function, and induce cognitive deficits. Clinical and experimental evidences have revealed that the acute increase of A&#946; levels in the brain allows development of Alzheimer-like phenotypes. Hence, a detailed protocol describing how to acutely generate an AD mouse model via the intracerebroventricular (ICV) injection of A&#946; is necessary in many cases. In this protocol, the steps of the experiment with an A&#946;-injected mouse are included, from the preparation of peptides to the testing of behavioral abnormalities. The process of preparing the tools and animal subjects before the injection, of injecting the A&#946; into the mouse brain via ICV injection, and of assessing the degree of cognitive impairment are easily explained throughout the protocol, with an emphasis on tips for effective ICV injection of A&#946;. By mimicking certain aspects of AD with a designated injection of A&#946;, researchers can bypass the aging process and focus on the downstream pathology of A&#946; abnormalities.

The amyloid-&#946; (A&#946;)-injected animal model enables the administration of a defined quantity and species of A&#946; fragments and reduces individual differences within each study group. This protocol describes the intracerebroventricular (ICV) injection of A&#946; without stereotactic instruments, enabling the production of Alzheimer-like behavioral abnormalities in normal mice.

Amyloid-&#946; (A&#946;) is a major pathological mediator of both familial and sporadic Alzheimer's disease (AD). In the brains of AD patients, ...

Intracerebroventricular and Intravascular Injection of Viral Particles and Fluorescent Microbeads into the Neonatal Brain

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the hematogenous route, which involves infection of the endothelial cells and pericytes of the brain. The second is the intracerebroventricular (ICV) route. Once within the central nervous system (CNS), viruses may spread to the subarachnoid space, meninges, and choroid plexus via the cerebrospinal fluid. In experimental models, the earliest stages of CNS viral distribution are not well characterized, and it is unclear whether only certain cells are initially infected. Here, we have analyzed the distribution of cytomegalovirus (CMV) particles during the acute phase of infection, termed primary viremia, following ICV or intravascular (IV) injection into the neonatal mouse brain. In the ICV injection model, 5 &#181;l of murine CMV (MCMV) or fluorescent microbeads were injected into the lateral ventricle at the midpoint between the ear and eye using a 10-&#181;l syringe with a 27 G needle. In the IV injection model, a 1-ml syringe with a 35 G needle was used. A transilluminator was used to visualize the superficial temporal (facial) vein of the neonatal mouse. We infused 50 &#181;l of MCMV or fluorescent microbeads into the superficial temporal vein. Brains were harvested at different time points post-injection. MCMV genomes were detected using the in situ hybridization method. Fluorescent microbeads or green fluorescent protein expressing recombinant MCMV particles were observed by fluorescent microscopy. These techniques can be applied to many other pathogens to investigate the pathogenesis of encephalitis.

Here, we describe a simple method of intracerebroventricular and intravascular injection of viral particles or fluorescent microbeads into the neonatal mouse brain. The localization pattern of the virus and nanoparticles could be detected by microscopic evaluation or by in situ hybridization.

In the study on the pathogenesis of viral encephalitis, the infection method is critical. The first of the two main infectious routes to the brain is the ...

Intracerebroventricular Treatment with Resiniferatoxin and Pain Tests in Mice

The transient receptor potential vanilloid type 1 (TRPV1), a thermosensitive cation channel, is known to trigger pain in the peripheral nerves. In addition to its peripheral function, its involvement in brain functions has also been suggested. Resiniferatoxin (RTX), an ultrapotent TRPV1 agonist, has been known to induce long-term desensitization of TRPV1, and this desensitization has been an alternative approach for investigating the physiological relevance of TRPV1-expressing cells. Here we describe a protocol for intracerebroventricular (i.c.v.) treatment with RTX in mice. Procedures are described for testing nociception to peripheral TRPV1 stimulation (RTX test) and mechanical stimulation (tail pressure test) then follow. Although the nociceptive responses of mice that had been administered RTX i.c.v. were comparable to those of the control groups, RTX-i.c.v.-administered mice were insensitive to the analgesic effect of acetaminophen, suggesting that i.c.v. RTX treatment can induce supraspinal-selective TRPV1 desensitization. This mouse model can be used as a convenient experimental system for studying the role of TRPV1 in brain/supraspinal function. These techniques can also be applied to studies of the central actions of other drugs.

The transient receptor potential vanilloid type 1 (TRPV1) in the supraspinal region has been suggested to play some roles in the brain function. Described here is a protocol for intracerebroventricular injection of resiniferatoxin for supraspinal TRPV1 desensitization in mice. Procedures for some pain tests are also presented.

The transient receptor potential vanilloid type 1 (TRPV1), a thermosensitive cation channel, is known to trigger pain in the peripheral nerves. In ...

In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices

Knowledge of cell-type specific synaptic connectivity is a crucial prerequisite for understanding brain-wide neuronal circuits. The functional investigation of long-range connections requires targeted recordings of single neurons combined with the specific stimulation of identified distant inputs. This is often difficult to achieve with conventional and electrical stimulation techniques, because axons from converging upstream brain areas may intermingle in the target region. The stereotaxic targeting of a specific brain region for virus-mediated expression of light-sensitive ion channels allows selective stimulation of axons originating from that region with light. Intracerebral stereotaxic injections can be used in well-delimited structures, such as the anterior thalamic nuclei, in addition to other subcortical or cortical areas throughout the brain.
Described here is a set of techniques for precise stereotaxic injection of viral vectors expressing channelrhodopsin in the mouse brain, followed by photostimulation of axon terminals in the brain slice preparation. These protocols are simple and widely applicable. In combination with whole-cell patch clamp recording from a postsynaptically connected neuron, photostimulation of axons allows the detection of functional synaptic connections, pharmacological characterization, and evaluation of their strength. In addition, biocytin filling of the recorded neuron can be used for post-hoc morphological identification of the postsynaptic neuron.

This protocol describes a set of methods to identify the cell-type specific functional connectivity of long-range inputs from distant brain regions using optogenetic stimulations in ex vivo brain slices.

Knowledge of cell-type specific synaptic connectivity is a crucial prerequisite for understanding brain-wide neuronal circuits. The functional ...

Quantitative Measurement of Intrathecally Synthesized Proteins in Mice

Cerebrospinal fluid (CSF), a fluid found in the brain and the spinal cord, is of great importance to both basic and clinical science. The analysis of the CSF protein composition delivers crucial information in basic neuroscience research as well as neurological diseases. One caveat is that proteins measured in CSF may derive from both intrathecal synthesis and transudation from serum, and protein analysis of CSF can only determine the sum of these two components. To discriminate between protein transudation from the blood and intrathecally produced proteins in animal models as well as in humans, CSF protein profiling measurements using conventional protein analysis tools must include the calculation of the albumin CSF/serum quotient (Qalbumin), a marker of the integrity of the blood-brain interface (BBI), and the protein index (Qprotein/Qalbumin), an estimate of intrathecal protein synthesis. This protocol illustrates the entire procedure, from CSF and blood collection to quotients and indices calculations, for the quantitative measurement of intrathecal protein synthesis and BBI impairment in mouse models of neurological disorders.

Elevated spinal fluid protein levels can either be the result of diffusion of plasma protein across an altered blood-brain barrier or intrathecal synthesis. An optimized testing protocol is presented in this article that helps to discriminate both cases and provides quantitative measurements of intrathecally synthesized proteins.

Cerebrospinal fluid (CSF), a fluid found in the brain and the spinal cord, is of great importance to both basic and clinical science. The analysis of the ...