This work was supported y a CIRM predoctoral fellowship (EDK), NARSAD Young Investigator Award (DK) and the NIMH BRAINS award (R01MH087495) (DK).

Anesthesia and analgesia: Thirty minutes prior to anesthesia, inject the rat with 0.05 mg/kg subcutaneous buprenorphine for analgesia. Initiate anesthesia with 30-40 mg/kg intraperitoneal sodium pentobarbital. At this point, also inject atropine to prevent respiratory failure (0.4 mg/kg, subcutaneous) and meloxicam as further analgesic (2 mg/kg subcutaneous). If after 5 minutes, the rat is still mobile or responsive to toe pinch, give subsequent doses of sodium pentobarbital at 5 mg/kg (intraperitoneal) until the rat is unresponsive to pain. Before performing the first incision, inject lidocaine (5 mg/kg, intradermal) at the incision site for local anesthesia. Six to eight hours after initial injection, inject the rat with 0.05 mg/kg subcutaneous buprenorphine for analgesia. Buprenorphine can be injected every 6-8 hours thereafter if needed, though this is usually not necessary.
It is important to note that other forms of anesthesia can interfere with excitotoxic lesions. For example, although ketamine is a commonly used form of anesthesia in rodents, it can interfere with lesions induced with NMDA because it is an NMDA receptor antagonist. It is important to select a method of inducing anesthesia that does not reduce lesion size. If gas anesthesia is desired, most stereotaxic devices including those described here can accommodate gas mask adaptors.
Note: Materials are described further in the Table of specific reagents and equipment below.
1. Preparation of the Pump and Stereotax
<ol>
	<li>Fill a 10 &mu;l Hamilton syringe with sterile water and mount it into the caddy of a 6 syringe programmable pump. Secure the end of the syringe plunger into the clamped holder on the pump.</li>
	<li>Prefill gas-sterilized PE20 tubing with sterile water using a needle and 1 ml syringe. Slide the open end of the tubing on to the Hamilton syringe, being careful to avoid creating any air bubbles in the tube.</li>
	<li>Clamp the 30ga, flat-cut, 1 inch infusion needle end of the tubing on to the arm of a stereotaxic device using a barrel-style electrode manipulator. Insert the infusion needle between the clamp and the grooved barrel. Secure the clamp over a piece of the tubing containing metal needle within it to avoid flattening the tubing and restricting fluid flow.</li>
	<li>Adjust the pump rate to &gt; 5 &mu;l/min and turn on the pump. A bead of water should appear at the tip of the infusion needle. Check the length of the tubing and its joints to make sure there are no leaks.</li>
	<li>Wipe away the water bead with a sterile swab and set the pump to withdraw. Withdraw a 2 &mu;l air bubble. The top of the bubble should be visible in the tubing.</li>
	<li>Immerse the infusion needle tip in a 20 mg/ml solution of NMDA in sterile 0.1M PBS and withdraw 4-5 &mu;l. An air bubble should be visible between the sterile water and the NMDA solution in the tip. It is important to note that NMDA is a potent neurotoxin and caution should be used in the preparation and handling of this solution. Gloves and eye protection are advised at all times.</li>
</ol>
2. Mount the Rat in the Stereotaxic Device
<ol>
	<li>Figure 1 depicts the basic parts of the stereotaxic device. After shaving the rat's head from the line of the eyes to the ears, load the rat's front teeth over the bite bar, with the rat's body resting on a heating pad set on medium. The tongue should hang below the bite bar, with the lower jaw. If available, a temperature feedback heating pad would be preferable. Monitoring rectal temperature would also be ideal.</li>
	<li>Holding the rat's neck with thumb and forefinger, move the right horizontal ear canal onto the right ear bar. The bar should feel like it is resting on a solid place, not like it sinks far into the head. Holding the right side of the head steady, slide the left ear bar into the rat's left horizontal ear canal. It should feel like it is resting on a solid place. The pinnae should look symmetrical and should appear to lie flat on the ear bars. Upward flaring of pinnae indicates improper placement of the ear bars. The head should not wobble in response to pressure at the neck.</li>
	<li>Screw down the nose bar. Be gentle. The bones in the nose are delicate and the nose bar does not need to be tight. Check that the rat is still anesthetized before proceeding.</li>
</ol>
3. Preparing the Rat for Surgery
<ol>
	<li>Using a sterile swab, gently place a small amount of veterinary ophthalmic lubricating ointment into each of the rat's eyes to protect them from drying and debris.</li>
	<li>Concentrically from the planned incision site to the edge of the shaved region, wipe the rat's scalp with 2% chlorhexidine solution followed 70% ethanol three times. Chlorhexidine scrub may also be used if more effective disinfectant is desired. Finish by wiping with 10% povidone iodine solution. The prepared scalp is now sterile and should only be touched with sterile materials.</li>
	<li>Don sterile gloves for further steps because you will next deal with the prepared surgical site.</li>
	<li>Place a sterile drape so that the fenestration exposes the surgical site.</li>
</ol>
4. Creating the Surgical Window
<ol>
	<li>Using a #10 scalpel blade, make an incision (approximately 2 cm in length) along the longitudinal midline of the scalp. The incision should start posterior to the line of the eyes. With the thumb and forefinger of the non-dominant hand, hold the scalp in tension perpendicular to the direction of the incision.</li>
	<li>Pull the fascia overlying the skull surface to the edges of the surgical site using sterile cotton-tips. It will take some force to clear the skull so it can be viewed.</li>
	<li>Use four curved, sterile hemostats to create a surgical window. Clamp fascia (NOT skin) on each side of the anterior and posterior aspects of the incision and then lay the clamps alongside the animal, thereby pulling the incision open and fully exposing the skull.</li>
</ol>
5. Leveling the Skull and Creating the Pilot Hole
<ol>
	<li>Locate bregma, the anterior point where the three main plates of the skull meet, as depicted in Figure 2. Move the infusion needle so the needle tip just touches the skull at bregma. Note the dorsal-ventral coordinate. Figure 3 demonstrates how to read coordinates on a Stoelting stereotaxic device. Note, at this point, gloves are no longer sterile and should not touch the sterile surgery site directly. Note: After touching the nonsterile stereotaxic device, the surgeon is no longer sterile and should not touch the surgical site nor parts of equipment which will touch the surgical site. </li>
	<li>Locate lambda, the posterior point where the three main plates of the skull meet, as depicted in Figure 2. Raise the infusion needle slightly and move it straight back to lambda. Lower the needle until it just touches the skull at lambda. Read the dorsal-ventral coordinate. If the skull is level, it will be within 0.1 mm of the bregma reading. If it is further off than this, move the needle up (out of the way), loosen the nose bar, either raise or lower the bite bar platform, depending on which end of the head is too high, and then re-measure.</li>
	<li>Move the infusion needle over bregma and record the anterior-posterior and medial-lateral coordinates. Calculate where the needle must be moved based on the coordinates of the brain area of interest according to a brain atlas12. For the BLA, subtract 2.8 mm from the anterior-posterior coordinate and either add or subtract 5.1 mm to/from the medial-lateral coordinate depending on whether you wish to target the left or right. Move the infusion needle to the calculated position.</li>
	<li>Swing the needle out of the way, taking care to keep track of where it was located and then drill a hole approximately 2 mm in diameter in that location using a Dremel moto-tool with a sterile dental drill bit attached. Drill carefully as when the drill breaks through the skull, it will tend to pull down, possibly damaging the dura and brain tissue underneath. Ream the hole to enlarge it.</li>
	<li>Absorb any blood with a sterile cotton swab.</li>
</ol>
6. Infusion of Neurotoxin
<ol>
	<li>Swing the needle back into position. Lower the needle until it touches the surface of the brain. Record the dorsal-ventral coordinate. This is the level of dura. Subtract the appropriate value to reach the brain region of interest. For the basolateral amygdala, subtract 6.7 mm.</li>
	<li>Lower the infusion needle to the calculated level slowly. Then raise it back up, well above the skull. Use a Hamilton syringe cleaning wire to unclog the tip of the infusion needle, being careful not to touch the end of the wire that comes in contact with the infusion needle with non-sterile gloves.</li>
	<li>Run the pump at &gt;5 &mu;l/min and make sure you see a bead of liquid come out of the infusion needle. If not, unclog with the wire and try again.</li>
	<li>Lower the infusion needle to the desired dorsal/ventral level slowly.</li>
	<li>Set the rate on the infusion pump to 0.1 &mu;l/min and start the infusion. Infuse for 2 min.</li>
	<li>Once the infusion has started, mark the two ends of the air bubble in the tubing with a black marker. If both ends do not move, your needle is clogged. Stop the infusion, raise the needle, unclog with the wire, check for flow with a high flow rate, re-lower the needle and try again.</li>
	<li>Once the infusion is complete, stop the pump and wait 5 min for the NMDA to diffuse away from the needle tip.</li>
	<li>Raise the needle 0.5 mm, and infuse again for 1 min. Wait 2.5 min for the NMDA to diffuse away from the needle-tip. This extra infusion helps ensure all of the BLA is covered by the infusion. Whether this is necessary for other brain regions must determined experimentally.</li>
	<li>Raise the needle out of the brain slowly and swing the stereotaxic arm out of the way.</li>
</ol>
7. Closing the Incision
<ol>
	<li>Don new sterile gloves to close the incision.</li>
	<li>Wipe away any excess blood on the skull with sterile cotton-tips and remove the clamps from the fascia.</li>
	<li>Using London forceps, push the two sides of the incision together. The inner edges of the skin should meet and the outer edge of the skin should not be permitted to curl into the incision. While pressing the incision closed, apply wound clips to close the protrusion firmly. Continue to push the skin together and staple along the length of the incision. This usually takes 3-5 wound clips.</li>
	<li>After placing all the wound clips, use the forceps to pinch the staples again, making sure they are secure. The more secure the clips, the less likely a rat will pull them out before the incision has healed.</li>
	<li>Swab the stapled incision with 10% povidone iodine solution and return the rat to its home cage and monitor until mobility is restored (usually 1-2 hours after anesthesia, total surgery time from induction to closure being 30-45 min). At this point, the rat may be returned to the colony room. After one week, remove staples with a staple remover tool under isoflurane anesthesia.</li>
	<li>Use a hot bead sterilizer to sterilize the tip surfaces of all instruments before use on the next rat. Instruments should be immersed in the beads for no longer than 15 seconds, or they may become too hot to handle. Instruments should be fully cooled before used on another animal.</li>
</ol>
8. Recovery
<ol>
	<li>It is possible that rats will exhibit seizure activity in the days following the lesion. Depending on the brain region damaged, other behaviors such as food consumption or grooming may be affected. It is important that animals be monitored on a daily basis to ensure that they remain healthy during recovery. Supportive care, such as liquid food, high-calorie food, or additional analgesia may be required. Mortality due to anesthesia occurs in a low percentage of rats, usually less than 5%. Mortality due to other causes (infection, bleeding) is extremely rare. Any signs of distress after recovery of anesthesia may be cause for exclusion from future studies, but are extremely rare. Figure 4 depicts the weight loss during the first few days following surgery as it should be expected. Body weight should rebound within 1 week.</li>
</ol>
9. Representative Results
One week or more after NMDA infusion, the lesion can be visualized using a cresyl-violet counterstain and brightfield microscopy. Brains should be perfused with ice cold 4% paraformaldehyde, equilibrated in 30% sucrose in 0.1M PBS and frozen. They should then be sliced at 30-60 &mu;m, in a series of 1:6 to 1:12 and mounted on to gelatin-coated glass slides before undergoing a standard cresyl violet counterstain. The lesion should correspond to a "bald spot" or an area of decreased cresyl violet stain due to cell death following lesion. Figures 5A and B show a representative lesion of the BLA and CeA. This image was obtained using a flatbed scanner with a high resolution scan (1200 dpi). To visualize the lesion, it helps to counterstain a series of sections through much of the tissue surrounding the target area so a misplaced lesion can be located.
<img alt="Figure 1" src="/files/ftp_upload/4079/4079fig1.jpg" /> 
Figure 1. Stereotaxic setup. This figure shows the rat stereotaxic with relevant parts labeled.
<img alt="Figure 2" src="/files/ftp_upload/4079/4079fig2.jpg" /> 
Figure 2. Bregma and lambda on the rat skull. This illustration depicts the locations of bregma and lambda. All coordinates are measured relative to bregma. Lambda is used to ensure that the skull is level.
<img alt="Figure 3" src="/files/ftp_upload/4079/4079fig3.jpg" /> 
Figure 3. Reading stereotaxic coordinates. This drawing depicts a standard stereotaxic reading. To obtain the digits to the left of the decimal place, use the markers on the right side. The labeled digit corresponds to the tens place (i.e. 1 = 10, 2 = 20). The hash marks between the labeled digits correspond to the single units (1-9). The zero line indicates at what integer the stereotax is set. Here, the zero line is between the 10 and 11 markers, indicating that the reading is between 10 and 11. To determine the numbers to the right of the decimal place, use the markers on the left side. The hash marks on the left are numbered 0 to 10, with each unlabeled hash representing one unit difference from those next to it. Whichever of the hash marks on the left lines up best with the hash marks on the right indicates the first decimal place of the coordinate reading. Here, the hash mark on the left corresponding to number 9 lines up best with the right-side hash marks so the decimal place is 9. The final reading is 10.9 mm.
<img alt="Figure 4" src="/files/ftp_upload/4079/4079fig4.jpg" /> 
Figure 4. Weight change after surgery. This graph shows the average (&plusmn; SEM) weight of rats who underwent stereotaxic surgery (n = 17) or did not have surgery (n = 6). Day 0 is the day of surgery. Following surgery, the weight of rats decreased for 2-3 days after surgery and then increased around day 5-6.
<img alt="Figure 5" src="/files/ftp_upload/4079/4079fig5.jpg" /> 
Figure 5. Excitotoxic lesion example. A) This image shows a cresyl violet counterstain on a 30 &mu;m coronal slice with the "bald spot" from NMDA lesion of the basolateral amygdala. The white arrows outline the lesion. It can be compared to the contralateral side of the brain slice which has no lesion. B) A similar lesion of the central nucleus amygdala.

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No conflicts of interest declared.

The stereotaxic method presented here allows for excitotoxic lesion of specific brain areas via infusion of NMDA. The basic stereotaxic methods can be adapted to infuse a variety of pharmacological and biological agents in a site-specific manner. It can also be adapted to target a variety of brain areas, defined by their stereotaxic coordinates in a brain atlas12. Adaptation to other species such as mice can be made with similar equipment built for smaller animals. The present procedures are optimized for 3 mo...

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>N-Methyl-D-aspartic Acid 98%</td>
			<td>Fisher Scientific</td>
			<td>AC32919-0500</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dual Lab Standard Stereotaxic w/45 deg. Ear Bars</td>
			<td>Stoelting</td>
			<td>51653</td>
			<td>Alternative vendor: Kopf *note of caution: assure compatibility of stereotaxic accessories if purchasing from multiple vendors</td>
		</tr>
		<tr>
			<td>10&mu;l SYR SPECIAL (*/*/*)</td>
			<td>Hamilton</td>
			<td>701SN</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Dremel Moto-Tool</td>
			<td>Stoelting</td>
			<td>58600</td>
			<td>Alternative vendor: Kopf</td>
		</tr>
		<tr>
			<td>Carbide burs, handpiece HP, size 2</td>
			<td>Schein Dental</td>
			<td>2284578</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Stoelting 6 Syringe Programmable Pump</td>
			<td>Stoelting</td>
			<td>53140</td>
			<td>Alternative vendor: Kopf</td>
		</tr>
		<tr>
			<td>Stainless Steel 316 Hypodermic Regular Wall Tubing 30 Gauge .0123" OD x .00625" ID x .003" Wall (infusion needle)</td>
			<td>Small Parts</td>
			<td>HTXX-30R-06-05</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Intramedic PE 20 tubing (infusion tubing)</td>
			<td>VWR</td>
			<td>63019-025</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Reflex Clips, 9mm, non-sterile</td>
			<td>Kent Scientific Corp.</td>
			<td>INS500346</td>
			<td>Alternative vendor: Fine Science Tools</td>
		</tr>
		<tr>
			<td>Reflex Clip Applier for 9mm clips</td>
			<td>Kent Scientific Corp.</td>
			<td>12031-09</td>
			<td>Alternative vendor: Fine Science Tools</td>
		</tr>
		<tr>
			<td>Curved Hartman hemostat</td>
			<td>Fine Science Tools</td>
			<td>13003-10</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>London forceps</td>
			<td>Fine Sceince Tools</td>
			<td>11080-02</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>2% chlorhexidine solution</td>
			<td>Allivet</td>
			<td>30159</td>
			<td>Alternative vendor: PetSolutions</td>
		</tr>
		<tr>
			<td>10% povidone iodine solution</td>
			<td>CVS</td>
			<td>SKU #739575</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Hot bead sterilizer</td>
			<td>Harvard Apparatus</td>
			<td>610183</td>
			<td>&nbsp;</td>
		</tr>
	</tbody>
</table>
Table 1. Table of specific reagents and equipment.

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.

Watch this Scientific Journal Video about Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat at JoVE.com

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

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 ...

stereotaxic-surgery-for-excitotoxic-lesion-specific-brain-areas-adult

Research

JoVE Journal

Neuroscience

35.3K visualizações.  University of California Berkeley. Ablação alvo de região específica do cérebro (s) por infusão de uma excitotoxina utilizando coordenadas estereotáxico é descrito. Esta técnica pode também ser adaptado para infusão de outros produtos químicos para o cérebro de rato.

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

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.

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 ...

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for tissue formation during embryonic development and tissue homeostasis during adulthood 1. Currently, great efforts are invested towards controlling the switch of somatic stem cells from expansion to differentiation because this is thought to be fundamental for developing novel strategies for regenerative medicine 1,2. However, a major challenge in the study and use of somatic stem cell is that their expansion has been proven very difficult to control.
Here we describe a system that allows the control of neural stem/progenitor cell (altogether referred to as NSC) expansion in the mouse embryonic cortex or the adult hippocampus by manipulating the expression of the cdk4/cyclinD1 complex, a major regulator of the G1 phase of the cell cycle and somatic stem cell differentiation 3,4. Specifically, two different approaches are described by which the cdk4/cyclinD1 complex is overexpressed in NSC in vivo. By the first approach, overexpression of the cell cycle regulators is obtained by injecting plasmids encoding for cdk4/cyclinD1 in the lumen of the mouse telencephalon followed by in utero electroporation to deliver them to NSC of the lateral cortex, thus, triggering episomal expression of the transgenes 5-8. By the second approach, highly concentrated HIV-derived viruses are stereotaxically injected in the dentate gyrus of the adult mouse hippocampus, thus, triggering constitutive expression of the cell cycle regulators after integration of the viral construct in the genome of infected cells 9. Both approaches, whose basic principles were already described by other video protocols 10-14, were here optimized to i) reduce tissue damage, ii) target wide portions of very specific brain regions, iii) obtain high numbers of manipulated cells within each region, and iv) trigger high expression levels of the transgenes within each cell. Transient overexpression of the transgenes using the two approaches is obtained by different means i.e. by natural dilution of the electroporated plasmids due to cell division or tamoxifen administration in Cre-expressing NSC infected with viruses carrying cdk4/cyclinD1 flanked by loxP sites, respectively 9,15.
These methods provide a very powerful platform to acutely and tissue-specifically manipulate the expression of any gene in the mouse brain. In particular, by manipulating the expression of the cdk4/cyclinD1 complex, our system allows the temporal control of NSC expansion and their switch to differentiation, thus, ultimately increasing the number of neurons generated in the mammalian brain. Our approach may be critically important for basic research and using somatic stem cells for therapy of the mammalian central nervous system while providing a better understanding of i) stem cell contribution to tissue formation during development, ii) tissue homeostasis during adulthood, iii) the role of adult neurogenesis in cognitive functions, and perhaps, iv) better using somatic stem cells in models of neurodegenerative diseases.

Expansion of Embryonic and Adult Neural Stem Cells by In Utero Electroporation or Viral Stereotaxic Injection

Controlling the expansion of somatic stem cells is a major factor hampering their study and use in therapy. Here we describe a system to temporally control neural stem cells expansion during development and adulthood, which can be used to increase the number of neurons generated in the mouse brain.

Somatic stem cells can divide to generate additional stem cells (expansion) or more differentiated cell types (differentiation), which is fundamental for ...

Micromanipulation of Gene Expression in the Adult Zebrafish Brain Using Cerebroventricular Microinjection of Morpholino Oligonucleotides

Manipulation of gene expression in tissues is required to perform functional studies. In this paper, we demonstrate the cerebroventricular microinjection (CVMI) technique as a means to modulate gene expression in the adult zebrafish brain. By using CVMI, substances can be administered into the cerebroventricular fluid and be thoroughly distributed along the rostrocaudal axis of the brain. We particularly focus on the use of antisense morpholino oligonucleotides, which are potent tools for knocking down gene expression in vivo. In our method, when applied, morpholino molecules are taken up by the cells lining the ventricular surface. These cells include the radial glial cells, which act as neurogenic progenitors. Therefore, knocking down gene expression in the radial glial cells is of utmost importance to analyze the widespread neurogenesis response in zebrafish, and also would provide insight into how vertebrates could sustain adult neurogenesis response. Such an understanding would also help the efforts for clinical applications in human neurodegenerative disorders and central nervous system regeneration. Thus, we present the cerebroventricular microinjection method as a quick and efficient way to alter gene expression and neurogenesis response in the adult zebrafish forebrain. We also provide troubleshooting tips and other useful information on how to carry out the CVMI procedure.

In this article, we demonstrate a method for manipulation of gene expression in the ventricular cells of the adult zebrafish telencephalon using antisense morpholino oligonucleotides. We present this method as an efficient and quick protocol that can be used for functional studies in the adult vertebrate brain.

Manipulation of gene expression in tissues is  required to perform functional studies. In this paper, we demonstrate the  cerebroventricular ...

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion channel, and transporter cell surface presentation on a minutes time scale.&#160;There is a broad diversity of mechanisms that control endocytic trafficking of individual proteins. Studies investigating the molecular underpinnings of trafficking have primarily relied upon surface biotinylation to quantitatively measure changes in membrane protein surface expression in response to exogenous stimuli and gene manipulation.&#160;However, this approach has been mainly limited to cultured cells, which may not faithfully reflect the physiologically relevant mechanisms at play in adult neurons.&#160;Moreover, cultured cell approaches may underestimate region-specific differences in trafficking mechanisms.&#160;Here, we describe an approach that extends cell surface biotinylation to the acute brain slice preparation.&#160;We demonstrate that this method provides a high-fidelity approach to measure rapid changes in membrane protein surface levels in adult neurons.&#160;This approach is likely to have broad utility in the field of neuronal endocytic trafficking.

Brain Slice Biotinylation: An Ex Vivo Approach to Measure Region-specific Plasma Membrane Protein Trafficking in Adult Neurons

Neuronal membrane trafficking dynamically controls plasma membrane protein availability and significantly impacts neurotransmission.&#160;To date, it has been challenging to measure neuronal endocytic trafficking in adult neurons.&#160;Here, we describe a highly effective, quantitative method to measure rapid changes in surface protein expression ex vivo in acute brain slices.

Regulated endocytic trafficking is the central mechanism facilitating a variety of neuromodulatory events, by dynamically controlling receptor, ion ...

Using Fluorescence Activated Cell Sorting to Examine Cell-Type-Specific Gene Expression in Rat Brain Tissue

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant cell type in the brain, neurons are the most widely studied of these cell types given their direct role in impacting behaviors. Other cell types in the brain also impact neuronal function and behavior via the signaling molecules they produce. Neuroscientists must understand the interactions between the cell types in the brain to better understand how these interactions impact neural function and disease. To date, the most common method of analyzing protein or gene expression utilizes the homogenization of whole tissue samples, usually with blood, and without regard for cell type. This approach is an informative approach for examining general changes in gene or protein expression that may influence neural function and behavior; however, this method of analysis does not lend itself to a greater understanding of cell-type-specific gene expression and the effect of cell-to-cell communication on neural function. Analysis of behavioral epigenetics has been an area of growing focus which examines how modifications of the deoxyribonucleic acid (DNA) structure impact long-term gene expression and behavior; however, this information may only be relevant if analyzed in a cell-type-specific manner given the differential lineage and thus epigenetic markers that may be present on certain genes of individual neural cell types. The Fluorescence Activated Cell Sorting (FACS) technique described below provides a simple and effective way to isolate individual neural cells for the subsequent analysis of gene expression, protein expression, or epigenetic modifications of DNA. This technique can also be modified to isolate more specific neural cell types in the brain for subsequent cell-type-specific analysis.

The goal of this protocol is to use the fluorescence activated cell sorting (FACS) technique to sort specific types of neural cells for subsequent analysis of cell-type-specific gene expression, epigenetic markers, and or protein expression.

The brain is comprised of four primary cell types including neurons, astrocytes, microglia and oligodendrocytes. Though they are not the most abundant ...

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 ...

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, ...

Assessment of Long-term Depression Induction in Adult Cerebellar Slices

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from parallel fibers (PF) to Purkinje cells (PC) is considered the basis for motor learning, and deficiencies of both LTD and motor learning are observed in various gene-manipulated animals. Common motor learning sets, such as adaptation of the optokinetic reflex (OKR), the vestibular-ocular reflex (VOR), and rotarod test were used for evaluation of motor learning ability. However, results obtained from the GluA2-carboxy terminus modified knock-in mice demonstrated normal adaptation of the VOR and the OKR, despite lacking PF-LTD. In that report, induction of LTD was only attempted using one type of stimulation protocol at room temperature. Thus, conditions to induce cerebellar LTD were explored in the same knock-in mutants using various protocols at near physiological temperature. Finally, we found stimulation protocols, by which LTD could be induced in these gene-manipulated mice. In this study, a set of protocols are proposed to evaluate LTD-induction, which will more accurately allow examination of the causal relationship between LTD and motor learning. In conclusion, experimental conditions are crucial when evaluating LTD in gene-manipulated mice.

In some gene-manipulated animals, using a single protocol may fail to induce LTD in cerebellar Purkinje cells, and there may be a discrepancy between LTD and motor learning. Multiple protocols are necessary to assess LTD-induction in gene-manipulated animals. Standard protocols are shown.

Synaptic plasticity provides a mechanism for learning and memory. For cerebellar motor learning, long-term depression (LTD) of synaptic transmissions from ...

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset (Callithrix jacchus)

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the neurosciences. Phylogenetically, these animals are much closer to humans than rodents. They also display complex behaviors, including a wide range of vocalizations and social interactions. Here, an effective stereotaxic neurosurgical procedure for implantation of recording electrode arrays in the common marmoset is described. This protocol also details the pre- and postoperative steps of animal care that are required to successfully perform such a surgery. Finally, this protocol shows an example of local field potential and spike activity recordings in a freely behaving marmoset 1 week after the surgical procedure. Overall, this method provides an opportunity to study the brain function in awake and freely behaving marmosets. The same protocol can be readily used by researchers working with other small primates. In addition, it can be easily modified to allow other studies requiring implants, such as stimulating electrodes, microinjections, implantation of optrodes or guide cannulas, or ablation of discrete tissue regions.

Stereotaxic Surgery for Implantation of Microelectrode Arrays in the Common Marmoset Callithrix jacchus

This work presents a protocol to perform a stereotaxic, neurosurgical implantation of&#160;microelectrode arrays in the common marmoset. This method specifically enables electrophysiological recordings in freely behaving animals but can be easily adapted to any other similar neurosurgical intervention in this species (e.g., cannula for drug administration or electrodes for brain stimulation).

Marmosets (Callithrix jacchus) are small non-human primates that are gaining popularity in biomedical and preclinical research, including the ...

Unraveling the Role of Discrete Areas of the Rat Brain in the Regulation of Ovulation through Reversible Inactivation by Tetrodotoxin Microinjections

Many experimental approaches have been used for studying the role of the brain in the regulation of ovulation. Examples include the lesion and deafferentation of neuronal groups, which are both invasive methods that permanently impair the integrity of the target area. These methods are accompanied by collateral effects that can affect the analysis of acute and temporal regulatory mechanisms. The stereotaxic implantation of guide cannulas aimed at specific brain regions, followed by a recovery period, allows researchers to microinject different drugs after the disappearance of the undesired effects of the surgery. Tetrodotoxin has been used to determine the roles of several brain areas in diverse physiological processes because it transiently inhibits the sodium-dependent action potentials, thus blocking all neural activity in the target region. This protocol combines this method with strategies for the assessment of the estrous cycle and ovulation to reveal the role of discrete brain regions in the regulation of ovulation at particular times of any given stage of the estrous cycle. Awake and unrestrained rats (Rattus norvegicus) were used to avoid the blocking effects that anesthetics and stress hormones exert on ovulation. This protocol can be easily adapted to other species, brain targets and pharmacological agents to study different physiological processes. Future improvements to this method include the design of a microinjection system using glass capillaries of small diameter instead of guide cannulas. This will reduce the amount of tissue damaged during the implantation and decrease the spread of the infused drugs outside the target area.

This protocol describes the construction of a low-cost microinjection system, its stereotaxic implantation into deep-brain structures and the procedure for timed microinjections of tetrodotoxin in awake and unrestrained rats. The goal is to reveal the participation of hypothalamic structures in the regulation of ovulation by inhibiting their neural activity.