This work was supported by National Institute of Neurological Disorders and Stroke grant NS081333 (to CMT), Alzheimer&#8217;s Association grant NIRG-10-171721 and National Institute of Mental Health grant MH096702 (to UH), and National Institute on Aging-funded Alzheimer&#8217;s Disease Research Center at Columbia University pilot grant AG008702 (to YYJ and JB).

Note: For all animal experimentation, Institutional and National guidelines for the care and use of laboratory animals were followed.
1. Prepare Surgical Instruments and Solutions for Surgery
<ol>
	<li>Autoclave all stainless steel surgical instruments.</li>
	<li>Prepare 70% ethanol by diluting 200 proof absolute ethanol with sterile molecular grade deionized distilled water.</li>
	<li>Attach the 29 G needle to the Hamilton syringe. Clean the interior of Hamilton syringe and needle by drawing up and ejecting nanopure water repeatedly for 1 min. Repeat this procedure with 70% ethanol.
	<ol>
		<li>Remove the plunger and needle from the Hamilton syringe. Air dry the parts in a laminar flow hood O/N.</li>
		<li>Irradiate the needle and syringe with ultraviolet light for 30 min before use.</li>
	</ol>
	</li>
	<li>Prepare saline solution by dissolving NaCl in molecular grade water to a final weight to volume concentration of 0.9%. Sterilize the solution by filtering it through 0.2 &#956;m pore filter.</li>
</ol>
2. Prepare Oligomeric A&#946;1-42
<ol>
	<li>Monomerize and resuspend recombinant human A&#946;1-42 in DMSO to 5 mM exactly as described in the publication of Fa&#39; and others13 .</li>
	<li>Dilute 5 mM A&#946;1-42 with sterile (1x) PBS to 100 &#181;M on the day before surgery.</li>
	<li>Mix the solution well by trituration.</li>
	<li>Incubate A&#946;1-42 solution for 12 hr at 4&#730;C. 
	Note: Control solutions such as diluting DMSO in PBS or scrambled/reverse A&#946;1-42 peptide should be prepared the same way as A&#946;1-42.</li>
</ol>
3. Determine Injection Coordinates
<ol>
	<li>Use the adult mouse brain atlas to determine the exact anterior-posterior (AP), medial-lateral (ML), and dorsal-ventral (DV) coordinates for the brain region of interest14. 
	&#8203;Note: To illustrate the technique we picked the dentate gyrus of the hippocampus as the target with the following coordinates from the bregma: AP, -2.00 mm; ML, &#177;1.3 mm; DV, -2.2 mm. The negative sign preceding the AP value indicates that it is 2.00 mm posterior of the bregma. The &#177; sign preceding the ML value indicates left and right direction from the center. Lastly, the negative sign preceding the DV indicates it is moving ventrally from the surface of the brain.</li>
</ol>
4. Stereotaxic Frame Setup
<ol>
	<li>Wear a surgical face mask, clean lab coat, and sterile surgical gloves.</li>
	<li>Wipe down stereotaxic instrument and mouse adaptor with 70% ethanol.</li>
	<li>Lay down sterile surgical drape on the counter.</li>
	<li>Place the stereotaxic instrument with mouse adaptor on top of the sterile surgical drape.</li>
	<li>Wipe the mouse heating pad with 70% ethanol. Position the heating pad over the stereotaxic frame bed. Use general purpose laboratory labeling tape to tape down the heating pad over the mouse adaptor bed.
	<ol>
		<li>Attach the heating pad to the warm water recirculator pump according to manufacturer&#8217;s directions.</li>
		<li>Turn on the warm water recirculator pump and set the temperature to 37 &#176;C, and then wait until it reaches temperature.</li>
	</ol>
	</li>
	<li>Affix the 50 &#181;l Hamilton syringe with a 29 G needle onto the stereotaxic frame by screwing it onto the motorized vertical injecting shaft according to manufacturer&#8217;s directions.</li>
	<li>Turn on the bead sterilizer and wait until it reaches the manufacturer&#8217;s preset temperature. 
	Note: This is used for sterilizing stainless steel instruments between animals. It takes instruments 20 sec of contact with the beads for sterilization.</li>
	<li>Turn on hot plate and set it to 42 &#176;C and place a clean empty cage atop.</li>
	<li>While the Hamilton syringe is fixed on the stereotaxic frame use the motorized stereotaxic injector to draw up the A&#946;1-42 solution. 
	&#8203;Note: Draw up more than 4 &#956;l of the solution into the syringe and then use the stereotaxic injector to set the injecting volume. Operate the injector according to manufacturer&#8217;s instructions.</li>
</ol>
5. Animal Preparation
<ol>
	<li>Determine the weight of the mouse using the weigh scale.</li>
	<li>Anesthetize mouse with ketamine/xylazine cocktail at 100 mg/kg ketamine and 10 mg/kg xylazine by injecting intraperitoneally (IP). Monitor the depth of anesthesia by the loss of toe pinch reflex.</li>
	<li>After the mouse is sedated take the hair clipper and shave its head to expose the skin over the skull.</li>
	<li>Place the mouse on top of the heating pad on top of the stereotaxic bed.</li>
	<li>Use the spatula to open the mouth and place the incisor teeth inside the teeth guard. Secure the nosepiece over the face of the mouse. Clamp it down gently and not too tight.</li>
	<li>Position the bilateral ear crossbars into auditory meatus to secure the head. Move each crossbar in until it hits the skull, and then turn the screw to lock it.
	<ol>
		<li>To make sure the head is secured use your index finger to gently push down on the head. If the head is properly secured it will not give out or move when pressed.</li>
	</ol>
	</li>
	<li>Apply a drop of eye cream or eye drop to the eyes to keep them moist. Make sure the eyes are moist throughout the surgical procedure.</li>
	<li>To disinfect the surgical site use sterile cotton swabs to apply betadine solution to the skin over the skull, followed by 70% ethanol. Perform the alternating betadine and 70% ethanol cleaning 2 more times.</li>
</ol>
6. Surgery and Infusion
<ol>
	<li>Use a scalpel to make a 2-3 mm incision in the midline of the scalp, and then use a straight fine scissors to extend the incision line to 1.0-1.5 cm to expose the sagittal suture, bregma, and lambda of the skull, landmarks which the stereotaxic coordinates are based on. Use micro clamps to keep the skin apart. 
	Note: The bregma and lambda positions are explained in the mouse brain atlas &#8220;The Mouse Brain in Stereotaxic Coordinates&#8221;14.</li>
	<li>Take a cotton swab, soak it in sterile saline, and use it to clean the skull. Then, use a clean dry cotton swab to dry the skull.</li>
	<li>Use a sterile fine point pen to mark a dot on the bregma. Use the stereotaxic micromanipulator to position the Hamilton syringe so that the tip of the needle just touches the dot on the bregma.
	<ol>
		<li>Record the DV coordinate.</li>
		<li>To check if the AP axis of the skull is level move the Hamilton syringe posteriorly so that the tip of the needle touches the lambda point.</li>
		<li>Record the DV position. Make sure the DV coordinates for bregma and lambda are within 0.5 mm. 
		Note: The goal is to minimize the DV difference between bregma and lambda.</li>
	</ol>
	</li>
	<li>Use the micromanipulator to return the Hamilton syringe needle tip back to the bregma dot.</li>
	<li>Record the starting AP position.</li>
	<li>Calculate the ending AP position by subtracting 2.00 mm from the starting AP coordinate.</li>
	<li>Use the micromanipulator to reposition the Hamilton syringe needle to the final AP coordinate.</li>
	<li>Record the current ML coordinate.</li>
	<li>Calculate the left and right ending ML coordinates by adding or subtracting 1.3 mm from the starting ML coordinate.</li>
	<li>Use the micromanipulator to move the Hamilton syringe needle to either the ending ML coordinate.
	<ol>
		<li>Touch the needle tip to the surface of the skull on both ending ML coordinates and record the DV coordinates and make sure the values are within 0.5 mm. 
		Note: The goal is to minimize the DV difference between the two ending ML coordinates.</li>
		<li>While at an ending coordinate pull the needle up 0.5-1 cm over the skull. Take a sterile fine point pen to mark a dot on the surface of the skull. Repeat this step for the contralateral side of the skull.</li>
	</ol>
	</li>
	<li>Swing the Hamilton syringe clear out of the way.</li>
	<li>Attach 0.8 mm drill head to the drill. Take the drill with both hands, set elbows on the surface of the table for stability, and position the drill head slightly above the sharpie dot on either the left or right ML coordinate. Activate the drill, lower the drill tip onto the skull surface to introduce a hole in the skull. Repeat this step for the contralateral side. 
	Note: Some bleeding may occur from the newly introduced holes. If there is bleeding then use a clean dry cotton swab to dab the blood.</li>
	<li>Move the Hamilton syringe back into position over either of the newly introduced ML holes. Lower the Hamilton needle just past the skull. At this point be careful not to puncture the brain. To make sure the needle has passed the skull take your index finger and gently push the needle against the skull to make sure the needle doesn&#8217;t exit the hole.</li>
	<li>Record the starting DV coordinate.</li>
	<li>Calculate the ending DV coordinate by subtracting 2.2 mm from the starting DV coordinate.</li>
	<li>Lower the needle down to the ending DV coordinate.</li>
	<li>Activate the stereotaxic injector pump to pump 4 &#181;l of A&#946;1-42 into the dentate gyrus at a rate of 0.5 &#181;l/min.</li>
	<li>When the infusion is complete let the needle remain in place for an additional 1 min to minimize backflow of solution out of the injection site.</li>
	<li>Move the needle to the other side of the brain and repeat steps 6.13-6.18.</li>
</ol>
7. Animal Removal and Postoperative Care
<ol>
	<li>Unscrew the ear bars and face/nose guard. Remove the mouse from the apparatus.</li>
	<li>Use a student standard pattern forceps to pull close the scalp and then seal the wound with suture.</li>
	<li>Inject 1 ml of sterile saline into the mouse via IP for hydration.</li>
	<li>Inject buprenorphine (0.1 mg/kg) subcutaneously to relieve pain.</li>
	<li>Maintain the mouse in the clean empty cage set on top of 42 &#176;C hot plate until it wakes, then return it to its housing cage with ample food and water.</li>
	<li>Administer buprenorphine via subcutaneous injection every 12 hr for 3 days for pain relief.</li>
</ol>
8. Suture Removal
<ol>
	<li>Anesthetize mouse with ketamine/xylazine cocktail at 100 mg/kg ketamine and 10 mg/kg xlyazine by IP. Note: This is done 7-10 days after surgery.</li>
	<li>Use a student standard pattern forceps and straight fine scissors to remove the suture.</li>
	<li>Inject 1 ml of sterile saline into the mouse via IP for hydration.</li>
	<li>Maintain the mouse in a clean empty cage set on top of 42 &#176;C hot plate until it wakes, and then return it to its housing cage with ample food and water.</li>
</ol>
9. Animal Sacrifice and Tissue Processing
<ol>
	<li>Anesthetize the mouse and perfuse it with 4% paraformaldehyde15, remove the brain, and process it for cryosectioning16. 
	Note: The timing of animal sacrifice depends on the experimenter. However, for our studies we chose 7 and 14 days post surgery.</li>
</ol>

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The authors declare that they have no competing financial interests.

To achieve a successful A&#946;1-42 injection the experimenter or surgeon must: 1) use aseptic technique; 2) correctly identify the brain region of interest with accurate coordinates; 3) be able to properly secure the mouse in the stereotaxic frame with the brain leveled in the AP and ML axis; 4) have the ability to operate the micromanipulator with precision; 5) ensure proper post-operative care. If these key steps are followed the mouse should survive the surgery with no observable infection.
<p class="j...

Amyloid plaque deposits, which are composed of amyloid-beta (A&#946;1-42), are a key feature of the pathology of Alzheimer&#8217;s disease (AD). Numerous studies have shown that high or toxic levels of recombinant oligomeric A&#946;1-42 elicit neuronal death, synaptic dystrophy, loss and dysfunction; as well as learning and memory deficits1-4. Brain regions affected include the hippocampus, the cortex, and subcortical structures such as the basal forebrain and the amygdala5,6. To date, there are multiple transgenic mouse models that attempt to simulate the A&#946;1-42 pathology of AD. Depending on the strain these animals prove to be useful in examining select pathological features of AD. Unfortunately, with the exception of 2 transgenic lines, APP23 and 5XFAD, these mice never fully replicate neuronal loss, a key aspect of AD. Even with the neuronal loss observed in APP23 and 5XFAD, the neuronal death observed was subtle, age dependent, and isolated to a few select regions7,8.
The direct infusion of oligomeric A&#946;1-42 into the wild-type mouse brain provides an excellent in vivo model which replicates the neuronal death aspect of amyloidopathy1,9,10. Unlike the commonly utilized transgenic mouse models the oligomeric A&#946;1-42 infusion model is ideal for acutely elevating A&#946;1-42 levels in a spatial and temporal manner. The advantage of using wild-type mice for this model obviates potential compensation or side effects from the mutations introduced in transgenic mouse lines. Past studies have shown that infusing toxic levels of A&#946;1-42 into the hippocampus elicits neuron death in the vicinity of the injection site within 1 week1. Moreover, consistent with the observation that A&#946;1-42 is toxic for cholinergic neurons11 the basal forebrain cholinergic neuron (BFCN) population which projects to the hippocampus is decreased 20-50% within 7-14 days following beta-amyloid infusion1,10 in mice, effectively allowing for the examinations of isolated neuronal circuitry in the brain. Since BFCN project ipsilaterally to the dentate gyrus of the hippocampus12, for the most part control/vehicle and oligomeric A&#946;1-42 solutions can be injected on either side of the brain allowing comparisons to be made between the left and right hemispheres1.
In this report we will provide a detailed surgical and injection methodology for adult wild-type C57BL/6J mice. This mouse strain is chosen because of its wide use in research. Technically, any brain region can be targeted for infusion, however here we will use the dentate gyrus of the hippocampus as the target to illustrate the technique.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Ketamine HCl (100mg/ml)</td>
			<td>Henry Schein Medical</td>
			<td>1049007</td>
			<td>100 mg ketamine per 1 kg animal</td>
		</tr>
		<tr>
			<td>Xylazine (20mg/ml)</td>
			<td>Henry Schein Medical</td>
			<td>not available</td>
			<td>10 mg xylazine per 1 kg animal</td>
		</tr>
		<tr>
			<td>Buprenex (0.3mg/ml)</td>
			<td>Henry Schein Medical</td>
			<td>1217793</td>
			<td>0.1 mg buprenex per 1 kg animal</td>
		</tr>
		<tr>
			<td>A&#946;1-42</td>
			<td>David Teplow/UCLA</td>
			<td>not available</td>
			<td>100 &#956;M; This amyloid was used in the paper</td>
		</tr>
		<tr>
			<td>A&#946;1-42</td>
			<td>Bachem</td>
			<td>H-1368</td>
			<td>Can be used in place of amyloid from the Teplow lab</td>
		</tr>
		<tr>
			<td>A&#946;1-42</td>
			<td>American Peptide</td>
			<td>62-0-80B</td>
			<td>Can be used in place of amyloid from the Teplow lab</td>
		</tr>
		<tr>
			<td>Scrambled A&#946;1-42</td>
			<td>American Peptide</td>
			<td>62-0-46B</td>
			<td>Can be used as control peptide for comparing A&#946;1-42</td>
		</tr>
		<tr>
			<td>NU4 Antibody (Oligomeric Amyloid Antibody)</td>
			<td>Gift from William Klein/Northwestern U.</td>
			<td>not available</td>
			<td>1:2000 dilution</td>
		</tr>
		<tr>
			<td>Anti-Amyloid Oligomeric Antibody&#160; (Polyclonal Rabbit)</td>
			<td>EMD Millipore</td>
			<td>AB9234</td>
			<td>May be used in place of Nu4; needs to&#160; be tested by the end user</td>
		</tr>
		<tr>
			<td>6E10 Antibody (Monoclonal Mouse) (Amyloid Antibody)</td>
			<td>Biolegend</td>
			<td>sig-39320</td>
			<td>1:1000 dilution</td>
		</tr>
		<tr>
			<td>ChAT Antibody (Polyclonal Goat)</td>
			<td>Millipore</td>
			<td>AB144P</td>
			<td>1:100 dilution</td>
		</tr>
		<tr>
			<td>DeadEnd Fluorometric TUNEL system</td>
			<td>Promega</td>
			<td>G3250</td>
			<td>Follow manufacturer&#39;s directions for use</td>
		</tr>
		<tr>
			<td>Prolong Gold Antifade Reagent with DAPI</td>
			<td>Invitrogen</td>
			<td>P36935</td>
			<td>Use when coverslipping slides</td>
		</tr>
		<tr>
			<td>Fluorogold</td>
			<td>Fluorochrome, LLC</td>
			<td>not available</td>
			<td>2% solution</td>
		</tr>
		<tr>
			<td>Absolute Ethanol (200 proof)</td>
			<td>Fisher Scientific</td>
			<td>BP2818-4</td>
			<td>For making 70% ethanol for sanitizing and disinfecting</td>
		</tr>
		<tr>
			<td>Novex 10-20% Tricine gel</td>
			<td>Life Technologies</td>
			<td>EC6625BOX</td>
			<td>For separating A&#946;1-42</td>
		</tr>
		<tr>
			<td>Novex Tricine SDS Running Buffer (10X)</td>
			<td>Life Technologies</td>
			<td>LC1675</td>
			<td>For running 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>Novex Tris-Glycine Transfer Buffer (25X)</td>
			<td>Life Technologies</td>
			<td>LC3675</td>
			<td>For transferring 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>SuperSignal Western Blot Enhancer</td>
			<td>Thermo Scientific</td>
			<td>46640</td>
			<td>For enhancing A&#946;1-42 signal; follow manufacturer&#39;s protocol</td>
		</tr>
		<tr>
			<td>Protran BA79 Nitrocellulose Blotting Membrane, 0.1 &#956;m</td>
			<td>GE Healthcare Life Sciences</td>
			<td>10402088</td>
			<td>For transferring 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>Xcell SureLock Mini-Cell</td>
			<td>Life Technologies</td>
			<td>EI0001</td>
			<td>Electrophoresis aparatus for running 10-20% Tricine gels</td>
		</tr>
		<tr>
			<td>GenTeal Lubricant Eye Gel</td>
			<td>Novartis</td>
			<td>not available</td>
			<td>For keeping the mouse eyes moist during surgery; can be found in local pharmacy stores</td>
		</tr>
		<tr>
			<td>Refresh Optive Lubricant Eye Drops</td>
			<td>Allergan</td>
			<td>not available</td>
			<td>For keeping the mouse eyes moist during surgery; can be found in local pharmacy stores; Can be used in place of GenTeal</td>
		</tr>
		<tr>
			<td>Betadine</td>
			<td>Stoelting</td>
			<td>50998</td>
			<td>For sanitizing and disinfecting</td>
		</tr>
		<tr>
			<td>Round/Tapered Spatula&#160;</td>
			<td>VWR</td>
			<td>82027-490</td>
			<td>For opening animal mouth</td>
		</tr>
		<tr>
			<td>Bulldog Serrefines Clamps (Jaw Dims. 9X1.6mm; Length 28mm)</td>
			<td>Fine Science Tools</td>
			<td>18050-28</td>
			<td>Optional; For keeping scalp skin apart during injection</td>
		</tr>
		<tr>
			<td>Straight Fine Scissors (Cutting edge 25mm; Length 11.5cm)</td>
			<td>Fine Science Tools</td>
			<td>14060-11</td>
			<td>For cutting scalp</td>
		</tr>
		<tr>
			<td>#3 Scalpel Handle</td>
			<td>Fine Science Tools</td>
			<td>10003-12</td>
			<td></td>
		</tr>
		<tr>
			<td>#11 Surgical Blade</td>
			<td>Fine Science Tools</td>
			<td>10011-00</td>
			<td>For making scalp incision</td>
		</tr>
		<tr>
			<td>Student Standard Pattern Forcep (Tip Dims. 2.5x1.5mm; Length 11.5cm)</td>
			<td>Fine Science Tools</td>
			<td>91100-12</td>
			<td>For holding scalp closed during suturing</td>
		</tr>
		<tr>
			<td>Trimmer Combo Kit</td>
			<td>Kent Scientific</td>
			<td>CL9990-1201</td>
			<td>For shaving hair</td>
		</tr>
		<tr>
			<td>T/Pump Warm Water Recirculator&#160;</td>
			<td>Kent Scientific&#160;</td>
			<td>TP-700</td>
			<td>For warming animal during surgery</td>
		</tr>
		<tr>
			<td>Resusable Warmining Pad (5&#34; x 10&#34;)</td>
			<td>Kent Scientific&#160;</td>
			<td>TPZ-0510FEA</td>
			<td>For attaching it to the T/Pump warm water recirculator to warm the animal during surgery</td>
		</tr>
		<tr>
			<td>Cordless Micro Drill</td>
			<td>Stoelting</td>
			<td>58610</td>
			<td>Use 0.8mm steel burrs to drill holes in the skull</td>
		</tr>
		<tr>
			<td>Lab Standard Stereotaxic Instrument with Mouse &#38; Neonatal Rat Adaptor</td>
			<td>Stoelting</td>
			<td>51615</td>
			<td></td>
		</tr>
		<tr>
			<td>Just for Mouse Stereotaxic Instrument</td>
			<td>Stoelting</td>
			<td>51730</td>
			<td>Can use this in place of Stoelting Cat. #51615</td>
		</tr>
		<tr>
			<td>Quintessential Stereotaxic Injector</td>
			<td>Stoelting</td>
			<td>53311</td>
			<td></td>
		</tr>
		<tr>
			<td>Dry Glass Bead Sterilizer</td>
			<td>Stoelting</td>
			<td>50287</td>
			<td>For sterilizing stainless steel instruments</td>
		</tr>
		<tr>
			<td>Sterile Surgical Drape (18&#34; x 26&#34;)</td>
			<td>Stoelting</td>
			<td>50981</td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton Syringe 50 ml, Model 705 RN SYR, NDL</td>
			<td>Hamilton Company</td>
			<td>7637-01</td>
			<td>For brain injection; use different syringes for different solutions</td>
		</tr>
		<tr>
			<td>29 Gauge Needle, Small Hub RN NDL</td>
			<td>Hamilton Company</td>
			<td>7803-06</td>
			<td>For attaching to the Hamilton syringe for brain injection</td>
		</tr>
		<tr>
			<td>1 ml BD Tuberculin Syringes</td>
			<td>VWR</td>
			<td>BD309659</td>
			<td>For administering anesthesia and saline</td>
		</tr>
		<tr>
			<td>30 Gauge Needle (0.5&#34;)</td>
			<td>VWR</td>
			<td>BD305106</td>
			<td>For administering anesthesia and saline</td>
		</tr>
		<tr>
			<td>Portable Electronic CS Series Scale (Ohaus)</td>
			<td>VWR</td>
			<td>65500-202</td>
			<td>For weighing animals to determine anesthesia dose</td>
		</tr>
		<tr>
			<td>Hot plate (Top Plate Dims. 7.25x7.25in)</td>
			<td>VWR</td>
			<td>47751-148</td>
			<td>For warming animals post-surgery</td>
		</tr>
		<tr>
			<td>Sofsilk Silk Suture C-1 Cutting 3/8, 12 mm</td>
			<td>Covidien</td>
			<td>S1173</td>
			<td>For closing wound</td>
		</tr>
		<tr>
			<td>Vetbond Tissue Adhesive (3M)</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-361931</td>
			<td>Optional: for aiding in wound closure; Use with suture.</td>
		</tr>
		<tr>
			<td>Cotton-Tipped Wooden-Shaft Sterile Applicators</td>
			<td>Fisher scientific</td>
			<td>22-029-488</td>
			<td>For cleaning and drying surgical wound</td>
		</tr>
		<tr>
			<td>Fisherbrand Superfrost Plus Microscope Slides</td>
			<td>Fisher Scientific&#160;</td>
			<td>12-550-15</td>
			<td>For collecting brain sections</td>
		</tr>
		<tr>
			<td>VWR Micro Cover Glass 24 X 50 mm</td>
			<td>VWR</td>
			<td>48393241</td>
			<td>For mounting microscope slides</td>
		</tr>
		<tr>
			<td>Thermo Scientific Nalgene Syringe Filter 0.2 &#956;m</td>
			<td>Fisher Scientific</td>
			<td>194-2520</td>
			<td>For sterilizing saline solution</td>
		</tr>
		<tr>
			<td>Sterile dual tip skin markers by Viscot Medical</td>
			<td>Medline</td>
			<td>VIS1422SRL91</td>
			<td>For marking coordinates on the skull</td>
		</tr>
	</tbody>
</table>

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.

The present method of preparing human recombinant oligomeric A&#946;1-42 yields soluble oligomeric species consisting of monomers, dimers, trimers, and tetramers (Figure 1A). These low molecular weight A&#946;1-42 species, but not the fibrils and plaques, have been shown in numerous settings to be most toxic to neurons1,4,9,17-19. To determine whether or not oligomeric A&#946;1-42 induces neuron death in the mouse brain A&#946;1-42 (4 &#181;l of 100 ...

Watch this Scientific Journal Video about Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus at JoVE.com

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

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-infusion-oligomeric-amyloid-beta-into-mouse

Research

JoVE Journal

Neuroscience

18.1K Views.  Columbia University Medical Center. 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.

Video: Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development. However, because the uterine wall is opaque during early embryogenesis, the ability to target specific parts of the embryo for microinjection is greatly limited. Fortunately, high-frequency ultrasound imaging permits the generation of images that can be used in real time to guide a microinjection needle into the embryonic region of interest. Here we describe the use of such imaging to guide the injection of retroviral vectors into the ventricular system of the mouse forebrain at embryonic day (E) 9.5. This method uses a laparotomy to permit access to the uterine horns, and a specially designed plate that permits host embryos to be bathed in saline while they are imaged and injected. Successful surgeries often result in most or all of the injected embryos surviving to any subsequent time point of interest (embryonically or postnatally). The principles described here can be used with slight modifications to perform injections into the amnionic fluid of E8.5 embryos (thereby permitting infection along the anterior posterior extent of the neural tube, which has not yet closed), or into the ventricular system of the brain at E10.5/11.5. Furthermore, at mid-neurogenic ages (~E13.5), ultrasound imaging can be used direct injection into specific brain regions for viral infection or cell transplantation. The use of ultrasound imaging to guide in utero injections in mice is a very powerful technique that permits the molecular and cellular manipulation of mouse embryos in ways that would otherwise be exceptionally difficult if not impossible.

Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5

In utero survival surgery in mice permits the molecular manipulation of gene expression during development. Here we describe the use of high-frequency ultrasound imaging to guide the injection of retroviral vectors into the mouse brain at embryonic day (E) 9.5.

In utero survival surgery in mice permits the molecular manipulation of gene expression during development.  However, because the uterine wall is opaque ...

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 ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central nervous system. We demonstrate a stereotaxic injection technique that allows targeted gene expression or silencing in the dorsal horn of the mouse spinal cord. The surgical procedure is brief. It requires laminectomy of a single vertebra, providing for quick recovery of the animal and unimpaired motility of the spine. Controlled injection of a small vector suspension volume at low speed and use of a microsyringe with beveled glass cannula minimize the tissue lesion. The local immune response to the vector depends on the intrinsic properties of the virus employed; in our experience, it is minor and short-lived when a recombinant adeno-associated virus is used. A reporter gene such as enhanced green fluorescent protein facilitates monitoring spatial distribution of the vector, and the efficacy and cellular specificity of the transfection.

Viral vectors allow for targeted gene manipulation. We demonstrate a method for conditional gene expression or ablation in the mouse spinal cord, using stereotaxic injection of a viral vector into the dorsal horn, a prominent site of synaptic contact between primary somatosensory afferents and neurons of the central nervous system.

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central ...

Building An Open-source Robotic Stereotaxic Instrument

This protocol includes the designs and software necessary to upgrade an existing stereotaxic instrument to a robotic (CNC) stereotaxic instrument for around $1,000 (excluding a drill), using industry standard stepper motors and CNC controlling software. Each axis has variable speed control and may be operated simultaneously or independently. The robot&#39;s flexibility and open coding system (g-code) make it capable of performing custom tasks that are not supported by commercial systems. Its applications include, but are not limited to, drilling holes, sharp edge craniotomies, skull thinning, and lowering electrodes or cannula. In order to expedite the writing of g-coding for simple surgeries, we have developed custom scripts that allow individuals to design a surgery with no knowledge of programming. However, for users to get the most out of the motorized stereotax, it would be beneficial to be knowledgeable in mathematical programming and G-Coding (simple programming for CNC machining).
The recommended drill speed is greater than 40,000 rpm. The stepper motor resolution is 1.8&#176;/Step, geared to 0.346&#176;/Step. A standard stereotax has a resolution of 2.88 &#956;m/step. The maximum recommended cutting speed is 500 &#956;m/sec. The maximum recommended jogging speed is 3,500 &#956;m/sec. The maximum recommended drill bit size is HP 2.

This protocol includes the designs and software necessary to upgrade an existing stereotaxic instrument to a robotic (computer numeric controlled; CNC) stereotaxic instrument for around $1,000 (excluding a drill).

This protocol includes the designs and software necessary to upgrade an existing stereotaxic instrument to a robotic (CNC) stereotaxic instrument for ...

Mouse Microsurgery Infusion Technique for Targeted Substance Delivery into the CNS via the Internal Carotid Artery

Animal models of central nervous system (CNS) diseases and, consequently, blood-brain barrier disruption diseases, require the delivery of exogenous substances into the brain. These exogenous substances may induce injurious impact or constitute therapeutic strategy. The most common delivery methods of exogenous substances into the brain are based on systemic deliveries, such as subcutaneous or intravenous routes. Although commonly used, these approaches have several limitations, including low delivery efficacy into the brain. In contrast, surgical methods that locally deliver substances into the CNS are more specific and prevent the uptake of the exogenous substances by other organs. Several surgical methods for CNS delivery are available; however, they tend to be very traumatic. Here, we describe a mouse infusion microsurgery technique, which effectively delivers substances into the brain via the internal carotid artery, with minimal trauma and no interference with normal CNS functionality.

Mouse Microsurgery Infusion Technique for Targeted Substance Delivery into the CNS via the Internal Carotid Artery

The present protocol describes a mouse microsurgery infusion technique, which effectively delivers substances directly into the brain via the internal carotid artery.

Animal models of central nervous system (CNS) diseases and, consequently, blood-brain barrier disruption diseases, require the delivery of exogenous ...

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, ...

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Transplantation of Chemogenetically Engineered Cortical Interneuron Progenitors into Early Postnatal Mouse Brains

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is a complicated task, which is particularly difficult in regards to the development of &#947;-aminobutyric acid (GABA)ergic cortical interneurons (CIs). CIs are the main inhibitory neurons in the cerebral cortex, and they play key roles in neuronal networks, by regulating both the activity of individual pyramidal neurons, as well as the oscillatory behavior of neuronal ensembles. They are generated in transient embryonic structures (medial and caudal ganglionic eminences -&#160;MGE and CGE) that are very difficult to efficiently target using in utero electroporation approaches. Interneuron progenitors migrate long distances during normal embryonic development, before they integrate in the cortical circuit. This remarkable ability to disperse and integrate into a developing network can be hijacked by transplanting embryonic interneuron precursors into early post-natal host cortices. Here, we present a protocol that allows genetic modification of embryonic interneuron progenitors using focal ex vivo electroporation. These engineered interneuron precursors are then transplanted into early post-natal host cortices, where they will mature into easily identifiable CIs. This protocol allows the use of multiple genetically encoded tools, or the ability to regulate the expression of specific genes in interneuron progenitors, in order to investigate the impact of either genetic or environmental variables on the maturation and integration of CIs.

Here we present a protocol, designed to use chemogenetic tools to manipulate the activity of cortical interneuron progenitors transplanted into the cortex of early postnatal mice.

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is ...

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).

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

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

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

Transplantation of Chemogenetically Engineered Cortical Interneuron Progenitors into Early Postnatal Mouse Brains

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