The authors thank Dr. Peter Gerrits for the drawings used in the video.

Note: Antiseptic techniques should be employed during the whole procedure. All the instruments and materials (cotton-tipped swabs, gauze, etc.) that will be used during the surgery should be sterilized by autoclaving. A surgical mask, hair bonnet and sterile gloves should be worn. The working area and the stereotaxic apparatus should be cleaned thoroughly, and disinfected with a 70% ethanol solution.
1. Pre-surgical procedures
<ol>
 <li>Set up the stereotaxic apparatus and all materials needed. Pre-warm the heating pad.</li>
 <li>Place the cannula in its support and check if it is straight.</li>
 <li>Turn on the gas system - mixture of ambient air and oxygen (30-35% of total flow should be oxygen).</li>
 <li>Weigh the rat and administer the anesthetic. We are using a mixture of ketamine (37.5 mg/kg) and dexmedetomidine (0.25 mg/kg) injected subcutaneously. For different anesthesia protocols, see Flecknell4 and Hellebrekers et al.7.</li>
 <li>After the rat lost consciousness, shave the head area going from the ears to just in between the eyes with an electric razor.</li>
 <li>Place the rat on the heating pad, with its nose in front of the air tubing. Use an oximeter to ensure that the rat has an adequate blood oxygenation level (should not drop &lt;90%). Please follow the manufacturer's instructions for proper use of equipment.</li>
 <li>Apply eye cream (Duratears Z, Alcon) on both corneas to avoid dehydration.</li>
 <li>Check the rat's reflexes (tail reflex or toe-pinch reflex, as demonstrated in Walantus et al.8) to ensure that it is adequately anesthetized. If the rat continues to show strong reflexes, supplementation of anesthesia might be needed.</li>
 <li>If no toe-pinch reflex is shown, place the rat in the stereotaxic apparatus, adjust the ear bars so that it shows equal reading on both sides, and place again the air tubing in front of the animal by fixing it with the nose bar. Check again if the rat shows a blood oxygenation level of 90% or higher. If not, adjust either the tubing, bringing it closer to the nose, or increase the flow of oxygen. Monitor the blood oxygenation level and heart rate throughout the surgery.</li>
 <li>Continuously monitor the rat's temperature with a rectal thermometer (preferentially connected to a heating pad) and record the values at the beginning and end of the surgery. Adjust the heating pad or use a blanket to maintain a body temperature of 37.5 to 38.5&deg;C.</li>
</ol>


2. Surgery
<ol>
 <li>Inject the analgesic. We are using a single peri-operative administration of carprofen (4.0-5.0 mg/kg, subcutaneously). For different analgesic protocols, see Hellebrekers et al.7.</li>
 <li>Clean the shaved area of the skin from the center to the haired perimeter three times with a disinfectant (e.g., chlorhexidine 0.5%) and locally inject a mixture of lidocaine (20 mg/ml) and adrenaline (5 mg/ml) for local anesthesia and vasoconstriction (to prevent excessive bleeding).</li>
 <li>Make an anterior-posterior incision of about 2.5 cm on the midline of the scalp, going from between the eyes until the back of the ears. Use 4-6 bulldog clamps to pinch off the skin and to keep the incision open. Remove any conjunctive tissue with a spatula and/or cotton swabs and clean the area to expose the skull surface.</li>
 <li>Check if the head is level: First, find Lambda and place the guide cannula exactly over this location, touching the skull. Record the dorso-ventral coordinate. Next, place the guide cannula exactly over Bregma, touching the skull, and record its dorso-ventral coordinate. These two coordinates should be identical. If the difference is &gt;0.3 mm, adjust the nose bar to correct it.</li>
 <li>Make two small holes for fixing the skull screws using a sterilized hand drill (one approximately 5 mm anterior to the cannula location in one of the hemispheres and the other 5 mm posterior to the cannula location in the contralateral hemisphere). Place two sterile screws into these holes until they are tightly anchored, without being inserted completely into the skull.</li>
 <li>With the guide cannula placed exactly at Bregma, record the anterior-posterior and lateral coordinates. The correct location of guide cannula placement for each brain region can be calculated by adding or subtracting from Bregma, with the aid of a stereotaxic atlas9-11.</li>
 <li>Position the guide cannula in its correct location, slightly touching the skull. Record the dorso-ventral coordinate. For bilateral cannula placement, find next the cannula location in the other hemisphere, and again record the dorso-ventral coordinate. Both coordinates should be identical (or differ &lt;0.3 mm).</li>
 <li>Mark the cannula locations on the skull with a sterile pencil and, with the hand driller, make the burr holes, checking the size and the correct location with the aid of the guide cannula. Once the holes are made, use a sterile needle to gently punch the meninges to allow for unobstructed insertion of the cannula.</li>
 <li>Place the cannula into the first hole and lower it carefully until it reaches the final ventral coordinate. Prepare the dental cement and generously apply it around the cannula and one or both screws in order to fix the cannula. Wait until the cement has dried completely. Afterwards, carefully remove the cannula support by turning the dorso-ventral bar upwards.</li>
 <li>Place the second cannula into the support and go to the cannula location in the other hemisphere. Place the cannula into the hole and repeat the previous step. Cover the screws and a large surface of the cannulas with the cement, and before the cement is dry, remove any surplus from the skin.</li>
 <li>Inject warm (~37&deg;C) sterile saline (~10 ml/kg, s.c.) to ensure rehydration.</li>
 <li>After the cement has completely dried, remove the cannula support and place a sterile pin into each cannula to prevent obstruction.</li>
 <li>Clean the wound area with sterile saline and suture the front and the back of the wound.</li>
 <li>Remove the animal from the stereotaxic apparatus, replacing the gas tubing in front of its nose. Continue to monitor the oxygen saturation level and body temperature.</li>
 <li>If an injectable anesthetic with dexmedetomidine is used, inject its antagonist atipamezole (0.25 mg/kg, s.c.) and wait until the animal wakes up (approximately 5 minutes).</li>
 <li>Place the rat into a recovery cage. To avoid hypothermia, place the cage in an incubator at 28&deg;C or on a heating pad in a place where you can observe the animal for at least one hour, before returning it to the vivarium room.</li>
</ol>
3. Post-surgical care
<ol>
 <li>During the first 4 days after surgery, monitor the rat's recovery by keeping daily records of weight and other observations concerning the condition of the animal in laboratory logbooks or "animal welfare diaries".</li>
 <li>Animals that show overt signs of sickness, infection of the wound, loss of body weight or other signs of reduced well-being must undergo special care: e.g. an extra dose of analgesics to minimize post-operative pain, a mixture of powder food and water in addition to standard chow to stimulate the rat's appetite, and/or a subcutaneous injection of saline to support rehydration.</li>
 <li>If the rat does not show any improvement after these interventions, or the loss of body weight is &gt;15% (compared to pre-surgery weight), sacrifice the animal with an overdose of anesthetic (humane end-point).</li>
 <li>Rats usually need to recover for at least 7 days before commencement of behavioral experiments.</li>
</ol>
4. Representative Results
To determine whether the various modifications to our surgical procedure, particularly with respect to oxygen supplementation and the continuous monitoring of blood oxygenation levels, heart rate and body temperature, enhanced the animal's survival and improved its general condition after surgery, we compared the non-survival rate of 20 cohorts of animals (consisting of 20 rats each) that underwent surgery after we implemented these modifications with the non-survival rate of 24 cohorts (20 rats each) operated with the standard protocol. As is shown in Figure 1a, the non-survival rate was significantly reduced in the sample of cohorts that was operated with the modified protocol (P &lt; 0.05; Mann-Whitney U test, two tailed). Moreover, as is shown in Figure 1b, post-surgical weight loss of rats operated with the modified protocol was also significantly reduced as compared to that of rats operated with the standard procedure (post-operative day 1: P &lt; 0.05; post-operative day 2: P &lt; 0.01; post-operative day 3: P = 0.17; Student t-tests).
<img alt="Figure 1" src="/files/ftp_upload/3528/3528fig1.jpg" /> 
Figure 1. Effect of surgical modifications on non-survival rate and post-surgical weight loss. (A.) Non-survival rate of rats operated with the modified protocol as compared to that of rats operated with the standard protocol. The non-survival rate (median &plusmn; interquartile ranges) was calculated as the percentage of rats, per cohort of 20 rats, that did not survive surgery. *p &lt; 0.05, Mann-Whitney U test two tailed (n = 20 cohorts for the modified protocol and 24 cohorts for the standard protocol). (B.) Weight loss (mean &plusmn; SEM as percentage of pre-surgery weight) during the first (POD1), second (POD2) and third (POD3) post-operative day. *P &lt; 0.05, **P &lt; 0.01, Student t-test (n = 60 per group).

<ol>
	<li>Russell, W. M. S., Burch, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Principles of Humane Experimental Technique. , (1959).</li><li>Richardson, C. A., Flecknell, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anaesthesia+and+post-operative+analgesia+following+experimental+surgery+in+laboratory+rodents:+are+we+making+progress?.">Anaesthesia and post-operative analgesia following experimental surgery in laboratory rodents: are we making progress?.</a> Altern. Lab Anim. 33, 119-127 (2005).</li><li>Stokes, E. L., Flecknell, P. A., Richardson, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reported+analgesic+and+anaesthetic+administration+to+rodents+undergoing+experimental+surgical+procedures.">Reported analgesic and anaesthetic administration to rodents undergoing experimental surgical procedures.</a> Lab Anim. 43, 149-154 (2009).</li><li>Flecknell, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Laboratory Animal Anaesthesia - A Practical Introduction for Research Workers and Technicians. , (2009).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DIRECTIVE+2010/63/EU+OF+THE+EUROPEAN+PARLIAMENT+AND+OF+THE+COUNCIL+of+22+September+2010+on+the+protection+of+animals+used+for+scientific+purposes.">DIRECTIVE 2010/63/EU OF THE EUROPEAN PARLIAMENT AND OF THE COUNCIL of 22 September 2010 on the protection of animals used for scientific purposes.</a> Official Journal of the European Union. , (2010).</li><li>Geiger, B. M., Frank, L. E., Caldera-Siu, A. D., Pothos, E. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survivable+Stereotaxic+Surgery+in+Rodents.">Survivable Stereotaxic Surgery in Rodents.</a> J. Vis. Exp. (20), e880-e880 (2008).</li><li>Hellebrekers, L. J., Booij, L. H. D. J., Flecknell, P. A., Van Zutphen, L. F. M., Baumans, V., Beynen, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anaesthesia,+analgesia+and+euthanasia.">Anaesthesia, analgesia and euthanasia.</a> Principles of Laboratory Animal Science. , 277-311 (2001).</li><li>Walantus, W., Elias, L., Kriegstein, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+utero+intraventricular+injection+and+electroporation+of+E16+rat+embryos.">In utero intraventricular injection and electroporation of E16 rat embryos.</a> J. Vis. Exp. (6), e236-e236 (2007).</li><li>Paxinos, G., Watson, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Rat Brain in Stereotaxic Coordinates. , (1986).</li><li>Paxinos, G., Watson, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Rat Brain in Stereotaxic Coordinates. , (2007).</li><li>Swanson, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Brain Maps: Structure of the Rat Brain. , (1992).</li></ol>

The authors have nothing to disclose.

The main purpose of this video presentation is to familiarize behavioral neuroscientists with the basic principles of stereotaxic surgery. Researchers who are already performing stereotaxic surgery might also benefit from this video and consider some of the procedural refinements for use in their own laboratory. An ever-increasing knowledge about optimizing surgical techniques1-3, the development of new anesthetics and analgesics for use in human and veterinary medicine4,12, public awareness concern...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 </tr>
 <tr>
 <td>Alcohol 70%</td>
 <td>VWR</td>
 </tr>
 <tr>
 <td>Antisedan (atipamezole)</td>
 <td>Orion</td>
 </tr>
 <tr>
 <td>Atropine sulfate 0,5</td>
 <td>Pharmachemie BV</td>
 </tr>
 <tr>
 <td>Bulldog haemostatic clamp</td>
 <td>Aesculap</td>
 </tr>
 <tr>
 <td>Cannulas</td>
 <td>Component Supply Co.</td>
 </tr>
 <tr>
 <td>Chlorhexidine 0.5%</td>
 <td>AppepPharma</td>
 </tr>
 <tr>
 <td>Cleaning powder</td>
 <td>Alconox</td>
 </tr>
 <tr>
 <td>Cotton sticks</td>
 <td>Hartmann BV</td>
 </tr>
 <tr>
 <td>Dexdomitor (dexmedetomidine)</td>
 <td>Orion</td>
 </tr>
 <tr>
 <td>Drill</td>
 <td>Dremel 8000</td>
 </tr>
 <tr>
 <td>Duratears Z</td>
 <td>Alcon</td>
 </tr>
 <tr>
 <td>Durogrip Naaldvoerder converse 130mm</td>
 <td>Aesculap</td>
 </tr>
 <tr>
 <td>Durotip Fijne schaar ret.sp/st 110 mm</td>
 <td>Aesculap</td>
 </tr>
 <tr>
 <td>Gauze</td>
 <td>Medicomp (5x5)</td>
 </tr>
 <tr>
 <td>Heating pad</td>
 <td>Harvard Apparatus</td>
 </tr>
 <tr>
 <td>Insect pins (stylets)</td>
 <td>Entosphinx (Czech Republik)</td>
 </tr>
 <tr>
 <td>Ketamine 10% (ketamine)</td>
 <td>Alfasan</td>
 </tr>
 <tr>
 <td>Micro-halsted straight tip</td>
 <td>Aesculap-vet</td>
 </tr>
 <tr>
 <td>Molinea</td>
 <td>Hartmann BV</td>
 </tr>
 <tr>
 <td>NaCl 0,9%</td>
 <td>Baxter</td>
 </tr>
 <tr>
 <td>Needles (25G)</td>
 <td>Terumo</td>
 </tr>
 <tr>
 <td>Oximeter</td>
 <td>Edan Instruments, Inc.</td>
 </tr>
 <tr>
 <td>Pentobarbital</td>
 <td>Pharmacy of the UMCG</td>
 </tr>
 <tr>
 <td>Rimadyl</td>
 <td>Pfizer</td>
 </tr>
 <tr>
 <td>Scalpel blade No. 23</td>
 <td>Swann Morton</td>
 </tr>
 <tr>
 <td>Scalpelholder NR. 4 133 mm</td>
 <td>Aesculap</td>
 </tr>
 <tr>
 <td>Screw driver</td>
 <td>conventional Hardware Store</td>
 </tr>
 <tr>
 <td>Simplex Rapid (dental cement)</td>
 <td>Kemdent</td>
 </tr>
 <tr>
 <td>Skull screws</td>
 <td>Component Supply Co.</td>
 </tr>
 <tr>
 <td>Spatula</td>
 <td>VWR</td>
 </tr>
 <tr>
 <td>Spongestan special</td>
 <td>Johnson &amp; Johnson</td>
 </tr>
 <tr>
 <td>Stereotacts</td>
 <td>Kopf Instruments</td>
 </tr>
 <tr>
 <td>Surgical forceps 100mm</td>
 <td>Aesculap</td>
 </tr>
 <tr>
 <td>Suture material Safil 5/0</td>
 <td>Aesculap</td>
 </tr>
 <tr>
 <td>Syringe 10 ml</td>
 <td>Omnifix</td>
 </tr>
 <tr>
 <td>Syringe 1ml</td>
 <td>Terumo</td>
 </tr>
 <tr>
 <td>Syringe 5ml</td>
 <td>Omnifix</td>
 </tr>
 <tr>
 <td>Xylocaine (lidocaine/adrenaline)</td>
 <td>Astra Zeneca</td>
 </tr>
 </tbody>
</table>

rodent stereotaxic surgery and animal welfare outcome improvements for behavioral neuroscience

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to investigate the effects of locally manipulated neurotransmitter and signaling pathways in awake, behaving animals. Moreover, the stereotaxic implantation of electrodes for electrophysiological stimulation and recording studies has been instrumental to our current understanding of neuroplasticity and brain networks in behaving animals. Ever-increasing knowledge about optimizing surgical techniques in rodents1-4, public awareness concerning animal welfare issues and stringent legislation (e.g., the 2010 European Union Directive on the use of laboratory animals5) prompted us to refine these surgical procedures, particularly with respect to implementing new procedures for oxygen supplementation and the continuous monitoring of blood oxygenation and heart rate levels during the surgery as well as introducing a standardized protocol for post-surgical care. Our observations indicate that these modifications resulted in an increased survival rate and an improvement in the general condition of the animals after surgery (e.g. less weight loss and a more active animal). This video presentation will show the general procedures involved in this type of stereotaxic surgery with special attention to our several modifications. We will illustrate these surgical procedures in rats, but it is also possible to perform this type of surgery in mice or other small laboratory animals by using special adaptors for the stereotaxic apparatus6.

Watch this Scientific Journal Video about Rodent Stereotaxic Surgery and Animal Welfare Outcome Improvements for Behavioral Neuroscience at JoVE.com

Rodent Stereotaxic Surgery and Animal Welfare Outcome Improvements for Behavioral Neuroscience

Stereotaxic surgery on rodents allows for targeted administration of drugs or electrical stimulation and recordings in awake, behaving animals. In this video presentation we will demonstrate recent procedural refinements to this long-standing procedure that successfully improved survival rate and reduced post-surgical weight loss.

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to ...

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38.9K Views.  University of Groningen. Stereotaxic surgery on rodents allows for targeted administration of drugs or electrical stimulation and recordings in awake, behaving animals. In this video presentation we will demonstrate recent procedural refinements to this long-standing procedure that successfully improved survival rate and reduced post-surgical weight loss.

Video: Rodent Stereotaxic Surgery and Animal Welfare Outcome Improvements for Behavioral Neuroscience

Systemic and Local Drug Delivery for Treating Diseases of the Central Nervous System in Rodent Models

Thorough preclinical testing of central nervous system (CNS) therapeutics includes a consideration of routes of administration and agent biodistribution in assessing therapeutic efficacy. Between the two major classifications of administration, local vs. systemic, systemic delivery approaches are often preferred due to ease of administration. However, systemic delivery may result in suboptimal drug concentration being achieved in the CNS, and lead to erroneous conclusions regarding agent efficacy. Local drug delivery methods are more invasive, but may be necessary to achieve therapeutic CNS drug levels. Here, we demonstrate proper technique for three routes of systemic drug delivery: intravenous injection, intraperitoneal injection, and oral gavage. In addition, we show a method for local delivery to the brain: convection-enhanced delivery (CED). The use of fluorescently-labeled compounds is included for in vivo imaging and verification of proper drug administration. The methods are presented using murine models, but can easily be adapted for use in rats.

Thorough preclinical testing of drugs that act in the central nervous system often involves assessing and comparing drug biodistribution in association with specific routes of administration. Here, three commonly used methods of systemic delivery (intravenous, intraperitoneal, and oral) as well as a method for local delivery (convection-enhanced delivery) are demonstrated in mice.

Thorough preclinical testing of central nervous system (CNS) therapeutics includes a consideration of routes of administration and agent biodistribution ...

Whole Animal Perfusion Fixation for Rodents

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

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

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

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the function of various brain regions for behavior or other experimental outcomes is to implement a localized ablation of function. In humans, patient populations with localized brain lesions are often studied for deficits, in hopes of revealing the underlying function of the damaged area. In rodents, one can experimentally induce lesions of specific brain regions.
Lesion can be accomplished in several ways. Electrolytic lesions can cause localized damage but will damage a variety of cell types as well as traversing fibers from other brain regions that happen to be near the lesion site. Inducible genetic techniques using cell-type specific promoters may also enable site-specific targeting. These techniques are complex and not always practical depending on the target brain area. Excitotoxic lesion using stereotaxic surgery, by contrast, is one of the most reliable and practical methods of lesioning excitatory neurons without damaging local glial cells or traversing fibers.
Here, we present a protocol for stereotaxic infusion of the excitotoxin, N-methyl-D-aspartate (NMDA), into the basolateral amygdala complex. Using anatomical indications, we apply stereotaxic coordinates to determine the location of our target brain region and lower an injection needle in place just above the target. We then infuse our excitotoxin into the brain, resulting in excitotoxic death of nearby neurons. While our experimental subject of choice is a rat, the same methods can be applied to other mammals, with the appropriate adjustments in equipment and coordinates.
This method can be used on a variety of brain regions, including the basolateral amygdala1-6, other amygdala nuclei6, 7, hippocampus8, entorhinal cortex9 and prefrontal cortex10. It can also be used to infuse biological compounds such as viral vectors1, 11. The basic stereotaxic technique could also be adapted for implantation of more permanent osmotic pumps, allowing more prolonged exposure to a compound of interest.

Targeted ablation of specific brain region(s) by infusion of an excitotoxin using stereotaxic coordinates is described. This technique could also be adapted for infusion of other chemicals into the rat brain.

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the ...

A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and sensorimotor plasticity1. Consistent sensory perturbations result in compensatory changes in motor output, reflecting changes in feedforward motor control that reduce the experienced feedback error. By quantifying how different sensory feedback errors affect human behavior, prior studies have explored how visual signals are used to recalibrate arm movements2,3 and auditory feedback is used to modify speech production4-7. The strength of this approach rests on the ability to mimic naturalistic errors in behavior, allowing the experimenter to observe how experienced errors in production are used to recalibrate motor output.
Songbirds provide an excellent animal model for investigating the neural basis of sensorimotor control and plasticity8,9. The songbird brain provides a well-defined circuit in which the areas necessary for song learning are spatially separated from those required for song production, and neural recording and lesion studies have made significant advances in understanding how different brain areas contribute to vocal behavior9-12. However, the lack of a naturalistic error-correction paradigm - in which a known acoustic parameter is perturbed by the experimenter and then corrected by the songbird - has made it difficult to understand the computations underlying vocal learning or how different elements of the neural circuit contribute to the correction of vocal errors13.
The technique described here gives the experimenter precise control over auditory feedback errors in singing birds, allowing the introduction of arbitrary sensory errors that can be used to drive vocal learning. Online sound-processing equipment is used to introduce a known perturbation to the acoustics of song, and a miniaturized headphones apparatus is used to replace a songbird's natural auditory feedback with the perturbed signal in real time. We have used this paradigm to perturb the fundamental frequency (pitch) of auditory feedback in adult songbirds, providing the first demonstration that adult birds maintain vocal performance using error correction14. The present protocol can be used to implement a wide range of sensory feedback perturbations (including but not limited to pitch shifts) to investigate the computational and neurophysiological basis of vocal learning.

We describe the design and assembly of miniaturized headphones suitable for replacing a songbird&#x2019;s natural auditory feedback with a manipulated acoustic signal. Online sound processing hardware is used to manipulate song output, introduce real-time errors in auditory feedback via the headphones, and drive vocal motor learning.

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the over-production of amyloid-beta and the deposition of amyloid-beta plaques in brain regions of the afflicted individuals. Throughout the years scientists have generated numerous Alzheimer&#8217;s disease mouse models that attempt to replicate the amyloid-beta pathology. Unfortunately, the mouse models only selectively mimic the disease features. Neuronal death, a prominent effect in the brains of Alzheimer&#8217;s disease patients, is noticeably lacking in these mice. Hence, we and others have employed a method of directly infusing soluble oligomeric species of amyloid-beta - forms of amyloid-beta that have been proven to be most toxic to neurons - stereotaxically into the brain. In this report we utilize male C57BL/6J mice to document this surgical technique of increasing amyloid-beta levels in a select brain region. The infusion target is the dentate gyrus of the hippocampus because this brain structure, along with the basal forebrain that is connected by the cholinergic circuit, represents one of the areas of degeneration in the disease. The results of elevating amyloid-beta in the dentate gyrus via stereotaxic infusion reveal increases in neuron loss in the dentate gyrus within 1 week, while there is a concomitant increase in cell death and cholinergic neuron loss in the vertical limb of the diagonal band of Broca of the basal forebrain. These effects are observed up to 2 weeks. Our data suggests that the current amyloid-beta infusion model provides an alternative mouse model to address region specific neuron death in a short-term basis. The advantage of this model is that amyloid-beta can be elevated in a spatial and temporal manner.

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Alzheimer&#8217;s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the ...

Non-restraining EEG Radiotelemetry: Epidural and Deep Intracerebral Stereotaxic EEG Electrode Placement

Implantable EEG radiotelemetry is of central relevance in the neurological characterization of transgenic mouse models of neuropsychiatric and neurodegenerative diseases as well as epilepsies. This powerful technique does not only provide valuable insights into the underlying pathophysiological mechanisms, i.e., the etiopathogenesis of CNS related diseases, it also facilitates the development of new translational, i.e., therapeutic approaches. Whereas competing techniques that make use of recorder systems used in jackets or tethered systems suffer from their unphysiological restraining to semi-restraining character, radiotelemetric EEG recordings overcome these disadvantages. Technically, implantable EEG radiotelemetry allows for precise and highly sensitive measurement of epidural and deep, intracerebral EEGs under various physiological and pathophysiological conditions. First, we present a detailed protocol of a straight forward, successful, quick and efficient technique for epidural (surface) EEG recordings resulting in high-quality electrocorticograms. Second, we demonstrate how to implant deep, intracerebral EEG electrodes, e.g., in the hippocampus (electrohippocampogram). For both approaches, a computerized 3D stereotaxic electrode implantation system is used. The radiofrequency transmitter itself is implanted into a subcutaneous pouch in both mice and rats. Special attention also has to be paid to pre-, peri- and postoperative treatment of the experimental animals. Preoperative preparation of mice and rats, suitable anesthesia as well as postoperative treatment and pain management are described in detail.

Non-restraining EEG radiotelemetry is a valuable methodological approach to record in vivo long-term electroencephalograms from freely moving rodents. This detailed protocol describes stereotaxic epidural and deep intracerebral electrode placement in different brain regions in order to obtain reliable recordings of CNS rhythmicity and CNS related behavioral stages.

Implantable EEG radiotelemetry is of central relevance in the neurological characterization of transgenic mouse models of neuropsychiatric and ...

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, they may function to regulate the developmental process and/or physiological function in neonatal brains. To investigate neurobiological functions of specific genes in the brain, it is essential to inactivate genes in the brain. Here, we describe a simple stereotaxic method to inactivate gene expression in the striatum of transgenic mice at neonatal time windows. AAV-eGFP-Cre viruses were microinjected into the striatum of Ai14 reporter gene mice at postnatal day (P) 2 by stereotaxic brain surgery. The tdTomato reporter gene expression was detected in P14 striatum, suggesting a successful Cre-loxP mediated DNA recombination in AAV-transduced striatal cells. We further validated this technique by microinjecting AAV-eGFP-Cre viruses into P2Foxp2fl/fl mice. Double labeling of GFP and Foxp2 showed that GFP-positive cells lacked Foxp2 immunoreactivity in P9 striatum, suggesting the loss of Foxp2 protein in AAV-eGFP-Cre transduced striatal cells. Taken together, these results demonstrate an effective genetic deletion by stereotaxically microinjected AAV-eGFP-Cre viruses in specific neuronal populations in the neonatal brains of floxed transgenic mice. In conclusion, our stereotaxic technique provides an easy and simple platform for genetic manipulation in neonatal mouse brains. The technique can not only be used to delete genes in specific regions of neonatal brains, but it also can be used to inject pharmacological drugs, neuronal tracers, genetically modified optogenetics and chemogenetics proteins, neuronal activity indicators and other reagents into the striatum of neonatal mouse brains.

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, ...

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in regulating neuronal growth and development. However, the mechanisms underlying Drd1&#945; receptor abnormalities mediating behavioral responses and modulating working memory function are still unclear. Using a novel RNA in situ hybridization assay, the current study identified dopamine Drd1&#945; receptor and tyrosine hydroxylase (TH) RNA expression from DA-related circuitry in the nucleus accumbens (NAc) area and substantia nigra region (SNR), respectively. Drd1&#945; expression in the NAc shows a &#34;discrete dot&#34; staining pattern. Clear sex differences in Drd1&#945; expression were observed. In contrast, TH shows a &#34;clustered&#34; staining pattern. Regarding TH expression, female rats displayed a higher signal expression per cell relative to male animals. The methods presented here provide a novel in situ hybridization technique for investigating changes in dopamine system dysfunction during the progression of central nervous system diseases.

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

Identification of dopamine D1-alpha receptor in the nucleus accumbens is critical for clarifying D1 receptor dysfunction during a central nervous system disease. We performed a novel RNA in situ hybridization assay to visualize single RNA molecules in a specific brain area.

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in ...

Noninvasive EEG Recordings from Freely Moving Piglets

The method allows the recording of high-quality electroencephalograms (EEGs) from freely moving piglets directly in the pigpen. We use a one-channel telemetric electroencephalography system in combination with standard self-adhesive hydrogel electrodes. The piglets are calmed down without the use of sedatives. After their release into the pigpen, the piglets behave normally&#8212;they drink and sleep in the same cycle as their siblings. Their sleep phases are used for the EEG recordings.

Here, we present a protocol to record telemetric electroencephalograms (EEGs) from freely moving piglets directly in the pigpen without the use of a sedative, making it possible to record typical EEG patterns during non-REM sleep, like spindle bursts.

The method allows the recording of high-quality electroencephalograms (EEGs) from freely moving piglets directly in the pigpen. We use a one-channel ...