This work is supported in part by American Heart Association Scientist Development Grant and NARSAD Young Investigator Award to Yingwei Mao.

NOTE: All protocols involving animals were followed in accordance with the animal care guidelines of The Pennsylvania State University, IACUC #44057
1. Lentivirus Production
NOTE: The day before transfection, LentiX-293 cells should be at 80% confluency.
<ol>
	<li>Rinse cells with DMEM just before transfection and incubate in 10 ml of DMEM/10% FBS with Penicillin (100 IU/ml) and Streptomycin (100 &#181;g/ml) for 5 min&#160;at RT.</li>
	<li>Dilute DNA and Polyethylenimine (PEI) (1 mg/ml) in 1:3 ratio (1 &#181;g of DNA: 3 &#181;l of PEI) in 1 ml of DMEM and vortex for 1 min. The volume of DNA added is dependent on the concentration of the DNA. Incubate at RT for 10 min.
	<ol>
		<li>For example, use 20 &#181;g of DNA plasmid, comprised of 10 &#181;g of vector plasmid, 5 &#181;g of psPax214 plasmid, 5 &#181;g of vesicular stomatitis virus(VSV) with 60 &#181;l of PEI per dish.</li>
	</ol>
	</li>
	<li>Mix the DNA and PEI solution again by vortexing for 1 min. Incubate at RT for 10 min.</li>
	<li>Add 1/10th volume of total culture medium of DMEM (i.e.&#160;1 ml for 10 ml culture medium in a 100 mm dish). Add 1 ml of DNA-PEI mixture drop by drop into the dish and swirl the dish around until to the culture medium is well-mixed with the DNA.</li>
	<li>Incubate for 6 hr&#160;at RT and remove the supernatant. Add 5 ml of culture medium.</li>
	<li>48 hr&#160;after transfection, collect viral supernatant and spin at 627 x g for 5 min&#160;at 4 &#176;C in a 50 ml tube. Filter 30 ml of viral supernatant through a 0.45 &#181;m syringe filter into an ultracentrifuge tube.</li>
	<li>Add sterile water to balance the tubes, and cover tubes with small piece of parafilm. Spin tubes at 11,249 x g for 120 min&#160;at 4 &#176;C. Remove liquid using a vacuum tip.</li>
	<li>Add 100 &#181;l of cold PBS the tube. Gently rock the tube at 4 &#176;C O/N.</li>
	<li>To re-suspend the virus, pipette the PBS added in step 1.8 over the pellet 10 times, being careful not to touch the pellet with the tip. The pellet will not be re-suspended until this is complete.</li>
	<li>Aliquot the virus at 10-20 &#181;l per tube, flash-freeze in liquid nitrogen and store at -80 &#176;C.</li>
</ol>
2. Stereotactic Injection
2.1) Preparation of Instruments
<ol>
	<li>Place a pair of scissors, blunt-end forceps, a needle holder, scalpel, cotton swabs, and a cloth in a sealed pack with a sterilization indicator, and autoclave prior to surgery. Additionally, obtain 10% povidone iodine, 70% ethyl alcohol, absorbable sutures, a heating pad, artificial tears, a glass bead sterilizer, drill, a drill bit, 2 disposable syringes, an injection syringe, electric razor, analgesic (ketoprofen), anesthetic (avertin), gloves, and a stereotactic apparatus with injection pump.</li>
	<li>Make 1.25% avertin solution fresh that day, filter in a sterile hood using a 0.2 &#181;m sterile syringe filter, and place into a sterile serum vial. Mix 2.5 g of 2,2,2-tribromoethyl alcohol into 5 ml of tert-amyl alcohol, then dissolve in 200 ml water. Keep the pH below 5.15</li>
</ol>
2.2) Preparation of the Mouse
<ol>
	<li>Administer avertin based on the weight of the mouse (375 mg/kg).15 Inject the mouse with the avertin via an intraperitoneal (IP) injection, and then place back into its cage until fully anesthetized.</li>
	<li>Give an analgesic via IP injection (ketaprofen, 5 mg/kg), and place artificial tears on the eyes of the mouse to prevent drying.</li>
	<li>Ensure that the mouse is fully asleep by pinching their foot. If the mouse responds to the foot pinch, then give more anesthetic (in doses of 50 &#181;l). Shave the head of the mouse using an electric razor. Shave the area to be operated on (typically from just behind the ears, to the top of the snout), and the surrounding area.</li>
	<li>Place the mouse into the stereotactic apparatus. Latch the front teeth onto the anterior clamp, and lower the clamp and tighten it so that the jaw is fully secure.</li>
	<li>Insert the ear bars into the ear canal to fully stabilize the head. Be careful not to insert the ear bars in too far, which could damage the inner ear. Once finished, the body of the mouse should be able to be moved slightly without disrupting the position of the head.</li>
	<li>Place a heating pad under the mouse in order to regulate the mouse&#8217;s body temperature throughout the procedure.</li>
	<li>Clean the surgical area thoroughly. Using a cotton swab, rub 10% povidone iodine in a circular motion, starting from the middle of the surgical site and moving outwards. Then, use a cotton swab to rub 70% ethyl alcohol on the surgical site in the same fashion. Repeat two times.</li>
</ol>
2.3) The First Incision
<ol>
	<li>Once the mouse is prepped, open the sterile equipment bag. Change gloves before touching the instruments. Take out the sterile cloth and place the instruments on the cloth. If any instrument touches an un-sterile surface, use the glass bead sterilizer for 15 sec&#160;to sterilize it.</li>
	<li>Take the scalpel in the dominant hand and blunt-end forceps in the non-dominant hand. Use the blunt-end forceps to gently grip the skin of the mouse, and make an incision using the scalpel. Begin the incision about 1.5 cm above the ears (toward the nose) and extend to about 0.5 cm below the ears. Extend the incision vertically down the middle of the mouse&#8217;s head to expose bregma (Figure 1).</li>
	<li>Once bregma is visualized, take a sterile cotton tip to gently remove any blood covering the surface of the skull. Use two cotton tips to push the skin toward the side of the head.</li>
</ol>
2.4) Equipment Setup
<ol>
	<li>After any blood is removed from the surface of the skull, and bregma is clearly visualized, prepare the syringe (concentration of 108 TU/ml, volume of 1 &#181;l). In the fume hood, fill the syringe with the virus to be injected. Ensure that no bubbles are inside. Place the syringe in the stereotactic apparatus, making sure that it is fully secured.</li>
	<li>Slowly lower the syringe until it is right above the surface of the skull, so that the tip of the sterile syringe needle is set to the intersection of bregma and the interaural line. Set this point as zero, and determine coordinates from this point.</li>
	<li>Depending on the brain area of interest, vary the stereotactic coordinates. Determine these coordinates by utilizing a brain atlas4. Once the coordinates are determined, move the needle of the syringe to match those coordinates. Target the dentate gyrus using the coordinates: M/L= +/- 1 mm, A/P= -1.82 mm. Lower the needle to right above the skull to visualize where the hole needs to be drilled.</li>
	<li>Raise the syringe slightly, take the drill and place the drill bit right above the target drill site at about a 45&#176; angle to the skull. Begin drilling, and keep drilling until the skull gives way. When the skull gives way, a drop in resistance can be detected. Be careful not to drill into the brain, so as to prevent cortical damage.</li>
	<li>Take a sterile cotton tip and wipe away any blood from the hole.</li>
	<li>Lower the syringe so that the tip sits right on the surface of the brain (not the skull). Set the D/V coordinate (depth) to 0. Lower the syringe to the desired depth, based on the brain atlas coordinates. To reach the dentate gyrus, set D/V to -1.79 mm.</li>
	<li>Start the injection at a rate of 0.2 &#181;l/min. Watch to ensure that the syringe does not slip lower than the desired depth. If this occurs, gently raise the syringe to the desired depth again.</li>
	<li>After the injection is finished, wait about 2 min&#160;to ensure that any residual virus has been absorbed. Slowly raise the syringe and use a cotton tip to remove any liquid from the injection site.</li>
	<li>Repeat for the other hemisphere.</li>
</ol>
2.5) Suturing
<ol>
	<li>Remove the sterile suture and needle from its packaging and grip the needle using the needle holders. Face the pointed edge of the needle away from the needle holder. Hold the blunt-end forceps in the non-dominant hand, and use it to grip the skin on the mouse. Start with the dominant hand.</li>
	<li>Gently push the suture through the skin and use the blunt-end forceps to grab the skin on the other side of the incision. Pull the suture material through until about 0.75 inch remains.</li>
	<li>Let go of the needle, and use the blunt-end forceps to wrap the suture around the needle holder one time, then, grab the 0.75 inch remaining suture with the needle holder, and pull the suture through, so that the needle and needle holder end up on the side of the head opposite to that which they were on originally. This should form a knot.</li>
	<li>Repeat this step three more times, alternating the side of the head the needle holder ends up on each time.</li>
	<li>Cut the suture and repeat steps 2.5.1-2.5.3 until the incision is closed.</li>
</ol>
2.6) Post-operative Care
<ol>
	<li>Gently remove the ear bars from the mouse, and remove its jaw from the anterior clamp.</li>
	<li>Keep the mouse on the heating pad until it wakes up and moves around on its own. Monitor the mouse daily, and keep and eye out for signs of pain, such as not grooming, eating or drinking. Give additional analgesic when deemed necessary.</li>
</ol>
3. Open Field Test
3.1) Set Up
<ol>
	<li>Obtain an empty square, plastic arena with dimensions 50 cm x 50 cm, with 50 cm high walls (but can be slightly larger). Clean the arena with 70% ethyl alcohol prior to the beginning of the experiment. Make sure the ethyl alcohol has fully dried before placing a mouse into the arena.</li>
	<li>Place the arena on the floor, and try to ensure the lighting is set to minimize shadows and glare.</li>
	<li>If using tracking software, position the camera directly overhead of the open field, approximately 1.5 ft above the open field (far enough away to full encompass the whole arena, but close enough to have a clear view of the mouse).</li>
	<li>Set up a zone in the center of the box (approximately an area of 30 cm x 30 cm).16 To set a zone, click on the zone 1 tab in the side panel.
	<ol>
		<li>Select a shape outline in the upper panel, and outline the perimeter of the center zone. Select add zone, and click on the inside of the center zone. Using the program prompts, name the zone (this will be referred to as &#8216;center&#8217; for this procedure). If the behavioral test will be scored by hand, make sure the bottom of the field is divided into 10 cm x 10 cm squares, marked using tape.</li>
	</ol>
	</li>
	<li>Set up the software so that the settings are correct, and the mouse is easily visible on the screen. Accomplish this by adjusting the detection settings on the machine to be compatible with the testing arena (lighting) and mouse color. 
	NOTE: When the settings are correct, the system should track only the mouse, and not any shadows or urine/fecal matter. The specific software settings depend on the lighting of the arena, and the color of the mouse.</li>
</ol>
3.2) Acclimation and Test
<ol>
	<li>Move experimental mice into the behavioral room 1 hr&#160;before the test in order to acclimate them to their surroundings. Leave the mice in their cage for the acclimation.</li>
	<li>Turn on the camera to tape the behavioral task and place the mouse toward the middle front of the arena so that the mouse is facing the wall.</li>
	<li>Set the timer for 5 min, and step away from the arena. If possible, leave the room so as to not accidentally influence the mouse.</li>
	<li>Once 5 min&#160;are up, remove the mouse and place them in their cage.</li>
	<li>Use 70% ethyl alcohol to thoroughly clean out the field before proceeding to the next mouse. Make sure the ethyl alcohol has fully dried.</li>
</ol>
3.3) Scoring
<ol>
	<li>Consider the mouse as being in the center of the field when all four paws are inside the middle 30 cm x 30 cm.</li>
	<li>Quantify the amount of time the mouse spends in the center. Alternatively, determine this time using tracking software (such as Noldus ethovison).
	<ol>
		<li>To do this, go to the analysis tab, and click on &#8220;movement&#8221; under analysis profiles. Delete the categories currently set, and then select &#8220;in zone&#8221; under the location tab. Select the center zone. Next, select &#8220;analysis output&#8221; (under the results tab) to view results. A table listing the time spent immobile for each trial should appear.</li>
	</ol>
	</li>
	<li>Otherwise, score the OFT by hand. The open field should have 25 10 cm x 10 cm squares outlined on the bottom of the open field (as outlined in step 3.1.3). Quantify the amount of time the mouse spends in the center, in addition to the number of entries into the center of the arena.16 
	NOTE: The center 30 cm x 30 cm area is considered the center of the open field. A mouse is regarded as inhabiting the center region if all four paws are inside the 30 x 30 cm area.</li>
</ol>
4. Forced Swim Test
NOTE: Allow a minimum of five days between behavioral tasks.
4.1) Set Up
<ol>
	<li>Obtain a 2 L clear bucket, and fill with 22 &#176;C water. Place the bucket on a table, and try to ensure the lighting is set to minimize shadows and glare.</li>
	<li>If using tracking software, position the camera directly overhead of the bucket.</li>
	<li>Set up software so that the mouse is easily visible on the screen.</li>
</ol>
4.2) Acclimation and Test
<ol>
	<li>Move experimental mice into the behavioral room 1 hr before the test to acclimate the surroundings. Leave the mice in their cage for the acclimation.</li>
	<li>Turn on the camera to tape the behavioral task. Gently, place the mouse in the middle of the bucket of water. Make sure not to drop the mouse in. Slowly lower the mouse, so its front feet touch the water first. Take care to prevent their head from submerging.</li>
	<li>Set the timer for 6 min, and step away from the behavioral area.</li>
	<li>Once 6 min&#160;are up, remove the mouse from the bucket, and wipe off excess water before placing the mouse back in their cage.</li>
</ol>
4.3) Scoring
<ol>
	<li>If using tracking software, set percent immobility to 11%, as suggested by Noldus (this should be the default setting), to quantify time floating vs. swimming.</li>
	<li>To quantify using the tracking system, go to the analysis tab, and click on movement under analysis profiles. Delete the categories currently set, and then select mobility state, on the left side panel (under individual behavior). Set immobility to 11%.</li>
	<li>Select track visualization, under the results tab. Select analysis output (under the results tab) to view results. A table listing the time spent immobile for each trial should appear.</li>
	<li>If hand scoring, measure the time the mouse spends swimming or climbing, and amount of time the mouse spends immobile. In addition, measure the latency to the first time immobile. See Figure 2 in Results Section.</li>
</ol>

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The authors have nothing to disclose.

Successful stereotactic injections rely on three factors: keeping the mouse alive, setting the correct zero point for coordinates (tip of the needle on the intersection of bregma and the interaural line), and setting the right depth to the zero point (tip of the needle just touching the exterior of the brain tissue). The viability of mice is important. Surgery survival can be aided by making sure that the mouse is properly anesthetized and receives adequate analgesic. Pain is known to be a major cause of poor recovery af...

The mouse has been widely used in neurobiology because it is easy to manipulate genetically. Gene knockout techniques allow researchers to investigate how each genetic factor shapes mouse behaviors. Moreover, the cre-loxp system provides a valuable tool for tissue- and cell type- specific gene knockout in mice, which enables researchers to study gene function in different tissues.1 However, in practice, expression patterns of cre promoters are difficult to control, and so far many established cre drivers have not achieved region-specific expression.2,3
Alternatively, stereotactic injection is a method that targets specific brain regions of the adult mouse. By injecting genetically engineered viruses expressing cDNA or shRNA, region-specific modulation of gene expression can be achieved. Although the brain size of each mouse varies, the location of specific brain regions can be determined using stereotactic coordinates set from landmarks on the skull of the mouse brain. The most commonly used landmarks are bregma, lambda, and the interaural line. Using coordinates obtained from a brain atlas,4 the exact location of each brain area can be identified by the antero-posterior (A/P), medial-lateral (M/L), and dorsal-ventral (D/V) axes from bregma/interaural line intersection. Typically viruses injected into the brains of mice are tagged with either red, or green fluorescent protein (RFP or GFP), so that injections can be confirmed by fluorescent microscopy.
Behavioral assessments of mice are especially necessary for basic research of psychiatric disorders. Symptoms of psychiatric disorders in patients typically involve abnormal behaviors. Some of these human behaviors are evolutionarily conserved and can be directly mimicked and observed in the mouse. For example, depression can be modeled in the mouse by measuring behavioral despair. People with depression often feel as though nothing they do will ever help, a symptom that can eventually lead to suicide. In rodents, this can be modeled using the Forced Swim Test (FST), which measures the amount of time a mouse swimming versus floating in a pool of water (viewed as giving up). This paradigm is validated by rescuing the phenotype with anti-depressants.7,8,9 Mice which have received anti-depressants will spend significantly less time immobile compared to untreated controls. Another behavioral test, the Open Field Test (OFT) is designed to assess locomotion in mice, and additionally can be used to analyze the anxious phenotype in mice.5,6 This test is based on the premise that mice feel safer when they are close to the wall in a novel open field. Wild type mice will eventually explore the novel environment, as they are curious animals. However, spending less time in the center of the field indicates anxiety in the mouse, as the mouse will not be able to overcome the initial fear brought on by a novel environment. The anxiety of the mouse, as quantified by the amount of time spent in the center of an open field, can be compared to clinical anxiety in humans, which is present in many psychiatric disorders.
The combination of stereotactic injections with behavioral paradigms is a novel way to alter the expression of a specific gene in a targeted brain area. The effect of modulated gene expression on mouse behaviors can then be determined. In contrast to whole brain knockout, this method is particularly useful as it only targets specific brain areas. In addition, stereotactic injections are typically performed in the adult wild type mouse, therefore, endogenous gene expression has been maintained throughout developmental stages. This method will avoid the confound effect if the gene is required for survival during the embryonic or postnatal stage of development. One major limitation is that the experimental mice need to go through an invasive surgery, in which the skulls of mice have to be opened. Moreover, the degree of gene modulation is determined by the titer and efficiency of the virus. The virus needs to be injected into the correct region using stereotactic coordinates, which requires special instruments. Verification of the correct injection site can only be completed post mortem.
This method has been previously used to test the involvement of a specific gene in various neurological diseases. For example, viral mediated RNAi targeting the Th gene (which allows dopamine to be synthesized) was injected into the substantia nigra compacta, and locomotor behavior analysis was conducted.10 Another study used stereotactic injection of a lentivirus silencing DISC1 to assess mouse behavior in relation to schizophrenia. Knockdown of DISC1 led to increased locomotion in response to novelty (parallels positive symptoms in schizophrenia), and greater immobility in the FST.11 Similarly, an additional study found that 5-HT1B overexpression led to increased exploratory behavior in the OFT, consistent with an anti-anxiety phenotype using this method.12 Stereotactic injections can deliver cre virus to induce recombination in cre-loxp mice. This method was used to selectively delete the Y2 receptor in the amygdala and the bed nucleus of the stria terminalis. Upon behavioral analysis, these mice were found to an anti-depressive phenotype when the gene was deleted in the central amygdala, but no phenotype when the gene was deleted in the basolateral amygdala or the bed nucleus of the stria terminalis.13 Thus, this technique provides a unique tool to study the genetic effect on animal behaviors.

<table><tbody><tr><td>Stereotactic Apparatus item 51725</td><td>Stoelting co.</td><td>51725</td><td></td></tr><tr><td>Quintessential stereotaxic pump&amp;nbsp;</td><td>Stoelting co.</td><td>53311</td><td></td></tr><tr><td>Injection Styringe, 65 RN</td><td>Hamilton</td><td>7633-01</td><td></td></tr><tr><td>DMEM</td><td>Sigma-Aldrich</td><td>D5796</td><td></td></tr><tr><td>PEI&amp;nbsp;</td><td>Polysciences Inc.</td><td>23966</td><td></td></tr><tr><td>Scissors</td><td>Fine Surgical Tools</td><td>14084-08</td><td></td></tr><tr><td>Blunt end forceps</td><td>Fine Surgical Tools</td><td>11002-12</td><td></td></tr><tr><td>Needle holder&amp;nbsp;</td><td>Fine Surgical Tools</td><td>12001-13</td><td></td></tr><tr><td>Drill</td><td>Ram Products Inc</td><td>Microtorque control box, Tech2000 handpiece</td><td>with pedal</td></tr><tr><td>Glass bead sterilizer</td><td>Inotech&amp;nbsp;</td><td>IS-400</td><td></td></tr><tr><td>Absorbable sutures&amp;nbsp;</td><td>Unify</td><td>M-K518r19</td><td>Absorbable, reverse cutting</td></tr><tr><td>Cotton swabs</td><td>VWR</td><td>89031-270</td><td></td></tr><tr><td>Heating pad</td><td>Gaymar</td><td>T-pump TP-500 PN11184-000</td><td></td></tr><tr><td>Artificial tears</td><td>Rugby</td><td>NDC 0536-6550-91</td><td></td></tr><tr><td>Disposable syringe</td><td>BD Syringe</td><td>309623</td><td></td></tr><tr><td>Cloth</td><td></td><td></td><td></td></tr><tr><td>Gloves&amp;nbsp;</td><td>Ansell</td><td>Senseitouch #7823</td><td></td></tr><tr><td>Avertin or other anesthetic</td><td>see recipe citation</td><td></td><td></td></tr><tr><td>Ketoprofen or other analgesic</td><td>see veternarian&amp;nbsp;</td><td></td><td></td></tr><tr><td>Tracking software</td><td>Noldus</td><td>Ethovision XT</td><td></td></tr><tr><td>Tracking Camera</td><td>Noldus</td><td>Media Recorder</td><td></td></tr></tbody></table>

applying stereotactic injection technique to study genetic effects on animal behaviors

Stereotactic injection is a useful technique to deliver high titer lentiviruses to targeted brain areas in mice. Lentiviruses can either overexpress or knockdown gene expression in a relatively focused region without significant damage to the brain tissue. After recovery, the injected mouse can be tested on various behavioral tasks such as the Open Field Test (OFT) and the Forced Swim Test (FST). The OFT is designed to assess locomotion and the anxious phenotype in mice by measuring the amount of time that a mouse spends in the center of a novel open field. A more anxious mouse will spend significantly less time in the center of the novel field compared to controls. The FST assesses the anti-depressive phenotype by quantifying the amount of time that mice spend immobile when placed into a bucket of water. A mouse with an anti-depressive phenotype will spend significantly less time immobile compared to control animals. The goal of this protocol is to use the stereotactic injection of a lentivirus in conjunction with behavioral tests to assess how genetic factors modulate animal behaviors.

Accurate stereotactic injection relies heavily on setting the correct coordinates. The tip of the needle used to inject the virus should be set directly on the intersection of bregma and the interaural line (Figure 1). It is helpful to use a stereomicroscope to ascertain whether or not the needle is placed correctly. When looking through the microscope, the needle should be positioned so that if the virus were injected, it would land directly on the intersection of bregma and the interaural line. That is...

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Applying Stereotactic Injection Technique to Study Genetic Effects on Animal Behaviors

Stereotactic injection of lentiviruses expressing cDNAs or shRNAs can modulate gene expression in specific brain areas of mice. Here, we present a protocol to combine stereotactic injections with behavioral tasks, such as the Open Field Test (OFT) and the Forced Swim Test (FST).

Stereotactic injection is a useful technique to deliver high titer lentiviruses to targeted brain areas in mice. Lentiviruses can either overexpress or ...

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Research

JoVE Journal

Behavior

14.1K Views.  The Pennsylvania State University. Stereotactic injection of lentiviruses expressing cDNAs or shRNAs can modulate gene expression in specific brain areas of mice. Here, we present a protocol to combine stereotactic injections with behavioral tasks, such as the Open Field Test (OFT) and the Forced Swim Test (FST).

Video: Applying Stereotactic Injection Technique to Study Genetic Effects on Animal Behaviors

Stereotactic Injection of MicroRNA-expressing Lentiviruses to the Mouse Hippocampus CA1 Region and Assessment of the Behavioral Outcome

MicroRNAs (miRNAs) are small regulatory single-stranded RNA molecules around 22 nucleotides long that may each target numerous mRNA transcripts and dim an entire gene expression pathway by inducing destruction and/or inhibiting translation of these targets. Several miRNAs play key roles in maintaining neuronal structure and function and in higher-level brain functions, and methods are sought for manipulating their levels for exploring these functions. Here, we present a direct in vivo method for examining the cognitive consequences of enforced miRNAs excess in mice by stereotactic injection of miRNA-encoding virus particles. Specifically, the current protocol involves injection into the hippocampal CA1 region, which contributes to mammalian memory consolidation, learning, and stress responses, and offers a convenient injection site. The coordinates are measured according to the mouse bregma and virus perfusion is digitally controlled and kept very slow. After injection, the surgery wound is sealed and the animals recover. Lentiviruses encoding silencers of the corresponding mRNA targets serve to implicate the specific miRNA/target interaction responsible for the observed effect, with na&iuml;ve mice, mice injected with saline and mice injected with "empty" lentivirus vectors as controls. One month post-injection, the animals are examined in the Morris Water Maze (MWM) for assessing their navigation learning and memory abilities. The MWM is a round tank filled with colored water with a small platform submerged 1 cm below the water surface. Steady visual cues around the tank allow for spatial navigation (sound and the earth's magnetic field may also assist the animals in navigating). Video camera monitoring enables measuring the route of swim and the time to find and amount the platform. The mouse is first taught that mounting the hidden platform offers an escape from the enforced swimming; it is then tested for using this escape and finally, the platform is removed and probe tests examine if the mouse remembers its previous location. Repeated tests over several consecutive days highlight improved performance of tested mice at shorter latencies to find and mount the platform, and as more direct routes to reach the platform or its location. Failure to show such improvement represents impaired learning and memory and/or anxiety, which may then be tested specifically (e.g. in the elevated plus maze). This approach enables validation of specific miRNAs and target transcripts in the studied cognitive and/or stress-related processes.

MicroRNAs have significant roles in brain structure and function. Here we describe a method to enforce hippocampal miRNA over-expression using stereotactic injection of an engineered miRNA-expressing lentivirus. This approach can serve as a relatively rapid way to assess the in vivo effects of over-expressed miRNAs in specific brain regions.

MicroRNAs (miRNAs) are small regulatory single-stranded RNA molecules around 22 nucleotides long that may each target numerous mRNA transcripts and dim an ...

Stereotaxic Microinjection of Viral Vectors Expressing Cre Recombinase to Study the Role of Target Genes in Cocaine Conditioned Place Preference

Microinjecting recombinant adenoassociated viral (rAAV) vectors expressing Cre recombinase into distinct mouse brain regions to selectively knockout genes of interest allows for enhanced temporally- and regionally-specific control of gene deletion, compared to existing methods. While conditional deletion can also be achieved by mating mice that express Cre recombinase under the control of specific gene promoters with mice carrying a floxed gene, stereotaxic microinjection allows for targeting of discrete brain areas at experimenter-determined time points of interest. In the context of cocaine conditioned place preference, and other cocaine behavioral paradigms such as self-administration or psychomotor sensitization that can involve withdrawal, extinction and/or reinstatement phases, this technique is particularly useful in exploring the unique contribution of target genes to these distinct phases of behavioral models of cocaine-induced plasticity. Specifically, this technique allows for selective ablation of target genes during discrete phases of a behavior to test their contribution to the behavior across time. Ultimately, this understanding allows for more targeted therapeutics that are best able to address the most potent risk factors that present themselves during each phase of addictive behavior.

This article describes how to microinject viral vectors into mouse brain and then test in a conditioned place preference paradigm that includes an acquisition, extinction and reinstatement phase.

Microinjecting recombinant adenoassociated viral (rAAV) vectors expressing Cre recombinase into distinct mouse brain regions to selectively knockout genes ...

Stable and Efficient Genetic Modification of Cells in the Adult Mouse V-SVZ for the Analysis of Neural Stem Cell Autonomous and Non-autonomous Effects

Relatively quiescent somatic stem cells support life-long cell renewal in most adult tissues. Neural stem cells in the adult mammalian brain are restricted to two specific neurogenic niches: the subgranular zone of the dentate gyrus in the hippocampus and the ventricular-subventricular zone (V-SVZ; also called subependymal zone or SEZ) in the walls of the lateral ventricles. The development of in vivo gene transfer strategies for adult stem cell populations (i.e. those of the mammalian brain) resulting in long-term expression of desired transgenes in the stem cells and their derived progeny is a crucial tool in current biomedical and biotechnological research. Here, a direct in vivo method is presented for the stable genetic modification of adult mouse V-SVZ cells that takes advantage of the cell cycle-independent infection by LVs and the highly specialized cytoarchitecture of the V-SVZ niche. Specifically, the current protocol involves the injection of empty LVs (control) or LVs encoding specific transgene expression cassettes into either the V-SVZ itself, for the in vivo targeting of all types of cells in the niche, or into the lateral ventricle lumen, for the targeting of ependymal cells only. Expression cassettes are then integrated into the genome of the transduced cells and fluorescent proteins, also encoded by the LVs, allow the detection of the transduced cells for the analysis of cell autonomous and non-autonomous, niche-dependent effects in the labeled cells and their progeny.

Here we describe a procedure based on the use of lentiviral particles for the long-term genetic modification of neural stem cells and/or their adjacent ependymal cells in the adult ventricular-subventricular neurogenic niche which allows the separate analysis of cell autonomous and non-autonomous, niche-dependent effects on neural stem cells.

Relatively quiescent somatic stem cells support life-long cell renewal in most adult tissues. Neural stem cells in the adult mammalian brain are ...

Developmental Biology

Rodent Brain Microinjection to Study Molecular Substrates of Motivated Behavior

Brain microinjection can aid elucidation of the molecular substrates of complex behaviors, such as motivation. For this purpose rodents can serve as appropriate models, partly because the response to behaviorally relevant stimuli and the circuitry parsing stimulus-action outcomes is astonishingly similar between humans and rodents. In studying molecular substrates of complex behaviors, the microinjection of reagents that modify, augment, or silence specific systems is an invaluable technique. However, it is crucial that the microinjection site is precisely targeted in order to aid interpretation of the results. We present a method for the manufacture of surgical implements and microinjection needles that enables accurate microinjection and unlimited customizability with minimal cost. Importantly, this technique can be successfully completed in awake rodents if conducted in conjunction with other JoVE articles that covered requisite surgical procedures. Additionally, there are many behavioral paradigms that are well suited for measuring motivation. The progressive ratio is a commonly used method that quantifies the efficacy of a reinforcer to maintain responding despite an (often exponentially) increasing work requirement. This assay is sensitive to reinforcer magnitude and pharmacological manipulations, which allows reinforcing efficacy and/ or motivation to be determined. We also present a straightforward approach to program operant software to accommodate a progressive ratio reinforcement schedule.

Rodents are an appropriate model to investigate the molecular substrates of behavior and complex psychiatric disorders. Brain microinjection in awake rodents can be used to elucidate disease substrates. An efficient and customizable brain microinjection method as well as the execution of an operant paradigm that quantifies motivation is presented.

Brain microinjection can aid elucidation of the molecular substrates of complex behaviors, such as motivation. For this purpose rodents can serve as ...

Modeling Brain Metastases Through Intracranial Injection and Magnetic Resonance Imaging

Metastatic spread of cancer is an unfortunate consequence of disease progression, aggressive cancer subtypes, and/or late diagnosis. Brain metastases are particularly devastating, difficult to treat, and confer a poor prognosis. While the precise incidence of brain metastases in the United States remains hard to estimate, it is likely to increase as extracranial therapies continue to become more efficacious in treating cancer. Thus, it is necessary to identify and develop novel therapeutic approaches to treat metastasis at this site. To this end, intracranial injection of cancer cells has become a well-established method in which to model brain metastasis. Previously, the inability to directly measure tumor growth has been a technical hindrance to this model; however, increasing availability and quality of small animal imaging modalities, such as magnetic resonance imaging (MRI), are vastly improving the ability to monitor tumor growth over time and infer changes within the brain during the experimental period. Herein, intracranial injection of murine mammary tumor cells into immunocompetent mice followed by MRI is demonstrated. The presented injection approach utilizes isoflurane anesthesia and a stereotactic setup with a digitally controlled, automated drill and needle injection to enhance precision, and reduce technical error. MRI is measured over time using a 9.4 Tesla instrument in The Ohio State University James Comprehensive Cancer Center Small Animal Imaging Shared Resource. Tumor volume measurements are demonstrated at each time point through use of ImageJ. Overall, this intracranial injection approach allows for precise injection, day-to-day monitoring, and accurate tumor volume measurements, which combined greatly enhance the utility of this model system to test novel hypotheses on the drivers of brain metastases.

Intracranial brain metastasis modeling is complicated by an inability to monitor tumor size and response to treatment with precise and timely methods. The presented methodology couples intracranial tumor injection with magnetic resonance imaging analysis, which when combined, cultivates precise and consistent injections, enhanced animal monitoring, and accurate tumor volume measurements.

Metastatic spread of cancer is an unfortunate consequence of disease progression, aggressive cancer subtypes, and/or late diagnosis. Brain metastases are ...

Cancer Research

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, ...

Neuroscience

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

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

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

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

Adaptable Angled Stereotactic Approach for Versatile Neuroscience Techniques

Stereotactic surgery is an essential tool in the modern neuroscience lab. However, the ability to precisely and accurately target difficult-to-reach brain regions still presents a challenge, particularly when targeting brain structures along the midline. These challenges include avoiding of the superior sagittal sinus and third ventricle and the ability to consistently target selective and discrete brain nuclei. In addition, more advanced neuroscience techniques (e.g., optogenetics, fiber photometry, and two-photon imaging) rely on targeted implantation of significant hardware to the brain, and spatial limitations are a common hindrance. Presented here is a modifiable protocol for stereotactic targeting of rodent brain structures using an angled coronal approach. It can be adapted to 1) mouse or rat models, 2) various neuroscience techniques, and 3) multiple brain regions. As a representative example, it includes the calculation of stereotactic coordinates for targeting of the mouse hypothalamic ventromedial nucleus (VMN) for an optogenetic inhibition experiment. This procedure begins with the bilateral microinjection of an adeno-associated virus (AAV) encoding a light-sensitive chloride channel (SwiChR++) to a Cre-dependent mouse model, followed by the angled bilateral implantation of fiberoptic cannulae. Using this approach, findings show that activation of a subset of VMN neurons is required for intact glucose counterregulatory responses to insulin-induced hypoglycemia.

Described here is a stereotactic procedure that can target challenging and difficult-to-reach brain regions (due to spatial limitations) using an angled coronal approach. This protocol is adaptable to both mouse and rat models and can be applied to diverse neuroscientific applications, including cannula implantation and microinjections of viral constructs.

Stereotactic surgery is an essential tool in the modern neuroscience lab. However, the ability to precisely and accurately target difficult-to-reach brain ...

Stereotaxic Intracranial Delivery of Chemicals, Proteins or Viral Vectors to Study Parkinson's Disease

Parkinson's disease (PD) is a progressive disorder traditionally defined by resting tremor and akinesia, primarily due to loss of dopaminergic neurons in the substantia nigra. Affected brain areas display intraneuronal fibrillar inclusions consisting mainly of alpha-synuclein (asyn) proteins. No animal model thus far has recapitulated all characteristics of this disease. Here, we describe the use of stereotaxic injection to deliver chemicals, proteins, or viral vectors intracranially in order to mimic various aspects of PD. These methods are well-established and widely used throughout the PD field. Stereotaxic injections are incredibly flexible; they can be adjusted in concentration, age of animal used for injection, brain area targeted and in animal species used. Combinations of substances allow for rapid variations to assess treatments or alter severity of the pathology or behavioral deficits. By injecting toxins into the brain, we can mimic inflammation and/or a severe loss of dopaminergic neurons resulting in substantial motor phenotypes. Viral vectors can be used to transduce cells to mimic genetic or mechanistic aspects. Preformed fibrillar asyn injections best recapitulate the progressive phenotype over an extended period of time. Once these methods are established, it can be economical to generate a new model compared to creating a new transgenic line. However, this method is labor intensive as it requires 30 minutes to four hours per animal depending on the model used. Each animal will have a slightly different targeting and therefore create a diverse cohort which on one hand can be challenging to interpret results from; on the other hand, help mimic a more realistic diversity found in patients. Mistargeted animals can be identified using behavioral or imaging readouts, or only after being sacrificed leading to smallercohort size after the study has already been concluded. Overall, this method is a rudimentary but effective way to assess a diverse set of PD aspects.

We describe how to successfully inject solutions into specific brain areas of rodents using a stereotaxic frame. This survival surgery is a well-established method used to mimic various aspects of Parkinson's disease.

Parkinson's disease (PD) is a progressive disorder traditionally defined by resting tremor and akinesia, primarily due to loss of dopaminergic neurons in ...

Optimizing Stereotaxic Microinjection Accuracy in Mice

Preliminary Validation of Stereotaxic Injection Coordinates via Cryosectioning

Stereotaxic injection of a specific brain region constitutes a fundamental experimental technique in basic neuroscience. Researchers commonly base their choice of stereotaxic injection parameters on mouse brain atlases or published materials that employed various populations/ages of mice and different stereotaxic equipment, necessitating further validation of the stereotaxic coordinate parameters. The efficacy of calcium imaging, chemogenetic, and optogenetic manipulations relies on the precise expression of reporter genes within the region of interest, often requiring several weeks of effort. Thus, it is a time-consuming task if the coordinates of the target brain region are not verified in advance. Using an appropriate dye instead of a virus and implementing cryosectioning, researchers can observe the injection site immediately following dye administration. This facilitates timely adjustments to coordinate parameters in cases where discrepancies exist between the actual injection site and the theoretical position. Such adjustments significantly enhance the accuracy of viral expression within the target region in subsequent experiments.

Author Spotlight: Accelerating Cognitive Impairment Research in Mice Through Stereotaxic Injection and Cost-Effective Dyes

The present protocol describes a practical strategy to expedite the verification step of stereotaxic injection coordinates before conducting viral tracing using dyes and frozen sections.

Stereotaxic injection of a specific brain region constitutes a fundamental experimental technique in basic neuroscience. Researchers commonly base their ...