Part 1. Prepare the Surgical Area
<ol>
<li>If working with biohazardous reagents (e.g. retroviral vectors), the surgery should be performed inside a Biosafety II cabinet (BSC). The ultrasound backscatter microscope (UBM) should sit next to the BSC, with the transducer inside the BSC, and held in place by a motor controlled support on a specially designed surgical stage. The components needed to construct the stage shown in the video are listed on line. This list, together with images of the stage from several angles (available on request), should permit construction.</li>
<li>The opening and closing of the laparotomy (surgical procedure in which the abdominal cavity is cut open to permit access to the internal organs) is usually performed on an absorbent pad with plastic backing. Prior to surgery all of the necessary tools (e.g. razor, surgical scissors, forceps, loaded autoclip applier, etc), and materials (e.g. PBS, sutures, anesthetic, the mouse holder and overlying plate, etc) should be sterilized and placed in the surgical area. Limiting the amount of time the incision is open is important for maximizing the chances of success with the surgery. All animal handling should be performed with gloves that have been disinfected with MB-10 solution, or the equivalent. Repeated disinfection should be performed as needed to assure that the gloves are sterile.</li>
</ol>
Part 2. Mouse Anesthesia
<ol>
<li>Fill a 1 cc syringe with the anesthetic solution containing 1 part Nembutal (50 mg/ml pentobarbital) to 4 parts filtered sterilized 25 mg/ml magnesium sulfate in PBS. This will result in a solution that is 10 mg/ml pentobarbital and 20 mg/ml magnesium sulfate. Each mouse should be weighted individually, and injected with 90 mg of pentobarbital per kg of bodyweight (e.g., a 28 gm mouse would be injected with 250 &mu;L).</li>
<li>Inject into the abdomen (intraperitoneal, IP) of a pregnant mouse at embryonic day (E) 9.5 (can use other ages as needed) penetrating both the skin and abdominal muscle. Allow 8-12 minutes for the mouse to become non-responsive. Adequate anesthesia should be confirmed by the relaxed posture of the animal, and lack of movement in response to tail and toe pinches. Gently pinching between the finger nails is highly sensitive, and lack of any response to such pinching indicates that the animals is sufficiently anesthetized. Note even animals that are completely unresponsive to tail and toe pinches may some times twitch slightly in response to the initial surgical incision. However, this is a reflexive response and does not indicate inadequate anesthesia. Petroleum-based ophthalmic balm should be applied to the eyes of the mouse to avoid drying during surgery.</li>
<li>Set-up the surgical area and the ultrasound machine while waiting for the mouse to become anesthetized. Since the use of retrovirus or lentivirus requires BSL2 containment, as stated above the viral injections should be performed in a Biosafety II Cabinet. In addition, all solutions and disposable materials that comes into contact with the virus should be decontaminated with a 10% bleach solution. The surgical tools and ultrasound equipment should be treated with a sterilizing solution such as MB-10 (<a href="http://www.quiplabs.com/mb10.htm">http://www.quiplabs.com/mb10.htm</a>), or an equivalent used by the facility in question.</li>
</ol>
Part 3. Exposure of the Embryos
<ol>
<li>If possible (depending upon the constraints of the given animal facility and available space) it is preferred to have the surgical preparation area in a separate location from the surgical area. This will minimize the chances for cross-contamination between animals. To initiate the surgery, place the animal onto its back on a sterile absorbent surface and wash the abdominal area thoroughly with 70% ethanol. More extensive presurgical treatments can be employed (e.g., successive rounds of Betadine and 70% ethanol washes), but in our experience this is not necessary for this type of rodent survival surgery. Using a double edge razor, shave the hair from the abdomen in an area of approximately 1 inch wide and 1.5 inches long. Use short quick motions with the razor at approximately a 45&deg; angle. It is important that the surgical area be suitably treated, but not subjected to excessive wetting, as this can result in needless chilling of the animal, and possibly even hypothermia.</li>
<li>Using fine surgical scissors make a 1" longitudinal incision through the skin first, and then through the abdominal muscle of the animal (cutting through connective tissue holding the skin to the muscle will help with the second incision). Note that throughout the procedure care should be taken to assure that the surgical tools remain sterile, and treatment with disinfectant should be performed as needed (i.e. if the tool tips come in contact with non-sterile material).</li>
<li>Using forceps carefully draw out uterine horns, and record the number of embryos on each side. While leaving two adjacent embryos exposed (selection of which two will depend upon the configuration on each side), place most of the uterine horns back into the animal. In this context, two embryos will be exposed at a given time and bathed in PBS, while the mother remains on her back.</li>
<li>Place the animal on its back into the plexiglass holder where it will be during the surgery (Figure1A)*. Next, lower the 10cm tissue culture dish** (which should be disinfected with MB-10 prior to use) down over the mother, with a pair of forceps placed slightly through the green silastic membrane (made using L RTV silicone rubber base, and curing agent; Dow-Corning product numbers 3142761 and 2156946, respectively). As you lower the plate down over the animal, use the forceps to grip the uterine tissue between the two embryos to be injected, and gently pull them through the membrane. At the same time, the plate is lowered completely down onto the holder, and the thumbtacks are pushed in to hold it in place. Pre-warmed (to 37&deg;C) PBS is then added to the dish to completely cover the embryos.</li>
<li>Place the injection stage containing the animal and exposed embryos under the ultrasound probe (Figure1A) to create real-time imaging of the exposed embryos through the uterine wall (Figure1B). The probe is then lowered (using the motorized controls) into the PBS bath directly over the embryos to be imaged. Images can be obtained when the probe is approximately 2-3 mm away from the embryos. It is very important to make sure that the probe head does not hit the embryos as it oscillates back and forth to collect images.</li>
</ol>
* The holder the mother rests in during surgery can easily be made by a machine shop. It should include a wide plexiglass base, with two plexiglass sides glued on top, and a space in between them (wide enough for the mouse to fit while lying on its back). The sides should have a valley ground out of them, which is then filled with black wax (e.g. from a dissection tray). That wax serves as the material into which thumbtacks will be pressed to hold down the plate of PBS that the embryos are bathed in during injection.

** 10cm bacterial dishes should have a hole (approx. 2cm in diameter) punched out of the center of the bottom (this can be done with a Heavy Duty Portable Punch and a 13/16th inch circular die: <a href="http://roperwhitney.com/punching/2-1011.cfm">http://roperwhitney.com/punching/2-1011.cfm</a>). In addition, two small holes on either side of the center hole (about 1.5cm away) should be burned out with a flame-heated needle (or they could be made with a drill of course). These holes are then covered with vacuum grease and will be where the thumbtacks that hold the plate onto the holder push through into the black wax. The vacuum grease is to prevent the PBS from leaking out.

Part 4. Loading Virus into a Microinjection Needle
<ol>
<li>Fashion a microinjection needle by beveling a 1mm (inner diameter) pulled glass capillary. The sharpness of the needle is critical, and working with an insufficiently sharp needle can seriously compromise penetration of the uterine wall and the amnionic sac, and can lead to experimental failure. The sharpened needle is then placed into the pipette holder, which is connected via tubing to a 25 &mu;L microsyringe mounted in a manual infusion/withdrawal pump (other devices can be used to regulate the loading and discharging of the needle, but this is what is shown in the video). For optimal responsiveness (to avoid the expansion and compression associated with an air filled system), the microsyringe, tubing, and needle should be filled with mineral oil.</li>
<li>Prior to loading the needle make sure there are no air bubbles in the system from the syringe all the way to the tip of the needle. Then draw back a small air bubble (1-2 mm in length) into the needle prior to starting to load virus. This air pocket helps maintain separation between the virus and the mineral oil. The viral stock to be injected should contain polybrene at a final concentration of 0.08 mg/ml (for more information on virus production please refer to references 1, 2). Note that concentrated retroviral stocks generally contain a significant amount of particulate matter, which contains the majority of the infectious material, and so cannot be filtered out without a 5- to 10-fold drop in titer. Also, depending upon the preparation, the viral stock may be viscous from the genomic DNA of lysed packaging cells (especially true if using 293 cells to generate MLV/VSV pseudotyped virus). If the aliquot of virus is viscous, a small amount of DNAse I can and should be added to loosen the stock, or loading the needle will be extremely difficult. A droplet of the viral stock (8-10 &mu;L) should be placed on a sterile piece of parafilm in preparation for needle loading. Then, using the withdrawal pump, carefully load the needle paying attention to avoid clogs. Place the loaded needle into the micromanipulator position in the injection area, being careful not to break the needle on the embryos, or the ultrasound probe. Note that if for some reason the needle is loaded significantly before the injection is to occur it can be helpful to draw a small amount of air into the tip, which can be discharged prior to starting the injections. This air pocket can protect against the viral stock drying at the tip of the needle (an occurrence that will almost always render that loaded needle useless).</li>
</ol>
Part 5. Ultrasound Guided Viral Injection
<ol>
<li>The real-time image created by the ultrasound probe is visualized on the video monitor, and can be used to guide the microinjection needle into the embryo. Using this system virus can be injected into the amniotic sac at E8.5 or either the amniotic sac or the ventricular system from E9.5-E11.5 (Figure1B). This is the most difficult part of the procedure and requires a significant amount of hands on experience to master (on the order of 5-20 hours depending upon the investigator). The needle tip is evident in the ultrasound image as the brightest spot on the screen, and should be seen moving in accordance with the adjustments of the probe position (using the motorized controls in the y and z planes, and the manual control in the x plane). It is critically important that the oscillation direction of the probe be parallel to the length of the needle. This will assure that the needle tip remains in the image plane during movements to position it for injection into the embryo.</li>
<li>The embryo should be positioned such that the target region (in this case the forebrain ventricle) is readily visible and is accessible to the needle with the least amount of uterine material in the injection path. In addition, it is critical not to inject through the placenta. Once the target brain region is lined up with the direction of needle advancement (using the knob on the micromanipulator), the needle should be just touching the uterine wall, and then a short quick jab forward should penetrate the tissue. A similar approach is used to penetrate the amnionic sac and then the brain. In some cases, and experienced investigator can hit the ventricle in one motion, especially if the needle is very sharp. The mechanics of moving the probe, the needle, and the embryos (by moving the mother on the stage), require hands-on practice to master.</li>
<li>Once the desired number of embryos has been injected, close the abdominal wall with 5.0 silk sutures. The mechanics of suturing are not specific to this type of surgery. Finally, use veterinary autoclips to staple the skin together over the stitches in the abdominal muscle. Suturing the skin is not recommended because the mouse will chew out the stitches. Analgesia should be applied to the incision area to reduce the amount of pain experienced by the mouse as it awakens. It is recommended that a 0.25% Bupivacaine solution be injected (0.8 ml/kg, or 2 mg/kg) subcutaneously every few millimeters along the incision site.</li>
<li>While the mouse recovers gently lay the animal on absorbent paper in a clean cage containing a gel pack (to facilitate feeding and hydration). Place the cage onto a slide warmer set at 37&deg;C to maintain body temperature. When awakened, place the mouse and the gel pack into a clean cage and return to the animal rack.</li>
<li>The following day check the mouse for any vaginal bleeding, abdominal contractions, and/or an overall distressed look, all of which indicate abortion. If this is the case, euthanize the mouse immediately. Well groomed hair and an overall healthy appearance indicate successful recovery from surgery. The injected embryos can be sacrificed embryonically or postnatally for further analysis. If the virus expresses a reporter gene, such as human alkaline phosphatase (PLAP) or GFP, a staining will reveal areas of positive viral infection (Figure1C). </li>
</ol>
Representative Results
Injected embryos can be sacrificed 1-2 days post injection all the way through to adulthood. If the virus expresses a reporter gene, such as PLAP or GFP, then reporter expression serves to identify virally infected cells and clones of those cells. For example, shown in Figure 1C is a tissue section of a postnatal brain that was infected in utero at E9.5 with a retroviral vector expressing PLAP. Infected cells are visualized as purple clusters after histochemical staining. In addition to identifying virally infected cells, reporter expression allows one to examine cell morphology and position, as well as gene expression status.
 <img src="/files/ftp_upload/2047/2047fig1.jpg" alt="Figure 1" /> 
Figure 1. Viral injection into the E9.5 mouse forebrain ventricles. A. The anesthetized animal is placed on the injection stage. Two embryos would be exposed and positioned under the ultrasound probe where indicated (arrow). The microinjection needle filled with virus is located near the embryos and lined up with the ultrasound probe. B. Screen capture of an real-time ultrasound image from an E9.5 mouse embryo. V, ventricle; AS, amniotic sac; UW, uterine wall. C. Embryos were injected with a PLAP-expressing virus at E10.5. Histochemical staining for PLAP reveals the location of viral infection events.

<ol>
	<li>Yee, J. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+high-titer+pseudotyped+retroviral+vectors+with+very+broad+host+range.">Generation of high-titer pseudotyped retroviral vectors with very broad host range.</a> Methods Cell Biol. 43, 99-112 (1994).</li><li>Gaiano, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+rapid+gain-of-function+studies+in+the+mouse+embryonic+nervous+system.">A method for rapid gain-of-function studies in the mouse embryonic nervous system.</a> Nat Neurosci. 2, 812-819 (1999).</li></ol>

No conflicts of interest declared.

Viral injections using ultrasound guidance can be conducted from E8.5-E11.5, where only the amniotic sac can be injected at E8.5, while both the amniotic sac and ventricular system can be injected from E9.5-E11.5. When performed correctly, viral particles injected into the ventricular system using ultrasound guidance have access to the cells lining the ventricular system and, therefore, can infect the various structures of the forebrain (neocortex, basal ganglia, diencephalon, eye, etc.), midbrain, and hindbrain (Figure1C). Virus injected into the amniotic sac has access to the cells on the exterior of the embryo. In addition to virus, ultrasound guidance can also be used to inject cells into the developing embryo. Therefore, this technique provides multiple options to choose from (developmental time period, structure, virus/cells) when designing an experiment.
It is critical that the injected virus or cells express a reporter for easy identification of infected/injected cells. Historically, human placental alkaline phosphatase (PLAP) or GFP have been used. From experience, we have found PLAP can be a very useful reporter gene because both PLAP histochemical and immunohistochemical staining yields a sharp and distinct single cell stain that allows one to analyze cell morphology and position, and to conduct a clonal analysis. Since viral GFP expression can be faint, a GFP reporter may not be ideal for staining tissue, but is useful if one wants to obtain single cells by fluorescence activated cell sorting (FACS).

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Micromanipulator </td>
<td>&nbsp;</td>
<td>Narishige</td>
<td>91084361683D</td>
<td>&nbsp;</td>
<tr>
<td>HI-7 pipette holder</td>
<td>&nbsp;</td>
<td>Narishige</td>
<td>9108436141</td>
<td>&nbsp;</td>
<tr>
<td>10 mm support arm</td>
<td>&nbsp;</td>
<td>Carl Zeiss</td>
<td>9108436206</td>
<td>&nbsp;</td>
<tr>
<td>IP iron base plate</td>
<td>&nbsp;</td>
<td>Carl Zeiss</td>
<td>9108436167</td>
<td>&nbsp;</td>
<tr>
<td>CI-1 connector for pipette holder</td>
<td>&nbsp;</td>
<td>Carl Zeiss</td>
<td>9108436257</td>
<td>&nbsp;</td>
<tr>
<td>1mm ID tubing</td>
<td>&nbsp;</td>
<td>Carl Zeiss</td>
<td>911013</td>
<td>&nbsp;</td>
<tr>
<td>3-way luer lock stopcock (pkg 10)</td>
<td>&nbsp;</td>
<td>Carl Zeiss</td>
<td>K420163-4503</td>
<td>&nbsp;</td>
<tr>
<td>Motor controller, 2-4 axes</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>860-c2</td>
<td>&nbsp;</td>
<tr>
<td>Joystick</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>PMC200-J</td>
<td>&nbsp;</td>
<tr>
<td>Micrometer adapter</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>ADAPT-BM17-375	</td>
<td>&nbsp;</td>
<tr>
<td>Motorized drive</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>CMA-25CC</td>
<td>&nbsp;</td>
<tr>
<td>Cable assembly</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>860I-10</td>
<td>&nbsp;</td>
<tr>
<td>25 mm translation stage</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>UMR8-25	</td>
<td>&nbsp;</td>
<tr>
<td>Micrometer</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>BM17-25</td>
<td>&nbsp;</td>
<tr>
<td>Base plate</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>M-PBN8</td>
<td>&nbsp;</td>
<tr>
<td>Inner mounting bracket</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>EQ80-I</td>
<td>&nbsp;</td>
<tr>
<td>Outer mounting bracket</td>
<td>&nbsp;</td>
<td>Newport</td>
<td>EQ80-E</td>
<td>&nbsp;</td>
<tr>
<td>Man. infusion/withdrawl pump</td>
<td>&nbsp;</td>
<td>Stoelting</td>
<td>51222</td>
<td>&nbsp;</td>
<tr>
<td>25 &mu;l microsyringe</td>
<td>&nbsp;</td>
<td>Stoelting</td>
<td>51113</td>
<td>&nbsp;</td>
<tr>
<td>24"x12"x0.5" metric board plate</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT03-640</td>
<td>&nbsp;</td>
<tr>
<td>Set of self-adjusting feet</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT34-841	</td>
<td>&nbsp;</td>
<tr>
<td>Bar-type lens holder	</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT03-669		</td>
<td>&nbsp;</td>
<tr>
<td>Right angle post clamp</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT53-357		</td>
<td>&nbsp;</td>
<tr>
<td>2" post holder</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT03-646		</td>
<td>&nbsp;</td>
<tr>
<td>4&#x201D; steel post	</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT36-499		</td>
<td>&nbsp;</td>
<tr>
<td>6&#x201D; steel post</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT36-503		</td>
<td>&nbsp;</td>
<tr>
<td>12/pk set screws, 1/4-20 x 1/2"</td>
<td>&nbsp;</td>
<td>Edmund</td>
<td>NT03-644		</td>
<td>&nbsp;</td>
<tr>
<td>Additional Materials:</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>PBS, pH7.4</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>10010049</td>
<td>&nbsp;</td>
<tr>
<td>Slide warmer</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>Numerous options</td>
<td>&nbsp;</td>
<tr>
<td>Double Edge Razor Blades</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>72000</td>
<td>&nbsp;</td>
<tr>
<td>Borosilicate Glass O.D: 1.0mm, I.D.: 0.05mm</td>
<td>&nbsp;</td>
<td>Sutter Instruments</td>
<td>B100-50-10</td>
<td>&nbsp;</td>
<tr>
<td>1cc Insulin Syringe U-100 27G5/8</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>329412</td>
<td>&nbsp;</td>
<tr>
<td>Autoclip Applier</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>427630</td>
<td>&nbsp;</td>
<tr>
<td>Autoclips, 9mm (to close skin incision)</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>427631</td>
<td>&nbsp;</td>
<tr>
<td>5.0 Silk sutures</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>S-1173</td>
<td>&nbsp;</td>
<tr>
<td>Sterilube ophthalmic ointment</td>
<td>&nbsp;</td>
<td>Fera pharmaceuticals</td>
<td>48102-012-35</td>
<td>&nbsp;</td>
<tr>
<td>MB-10 or equivalent disinfectant</td>
<td>&nbsp;</td>
<td>Quip labs</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>10% bleach</td>
<td>&nbsp;</td>
<td>Multiple sources</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>
			<div>
			"One of the following ultrasound backscatter microscopes can be used for this procedure. Although in the video the P40 from Paradigm-Medical is used, the P40 is no longer commercially available, and has been replaced by the P60, which is similarly priced and can be used for the same application. The additional materials described for stage construction are for use with Paradigm-Medical device(s). The Vevo 2100 comes with equipment for small animal imaging.
<ol>
<li>P60 Ultrasound BioMicroscope (UBM) from Paradigm-Medical 
(<a href="http://www.paradigm-medical.com/ubms.html">http://www.paradigm-medical.com/ubms.html</a>).</li>
<li>Vevo 2100 from VisualSonics (<a href="http://www.visualsonics.com/">http://www.visualsonics.com/</a>)</li>
</ol>
Anesthetic and Analgesic
The Nembutal solution (50 mg/mL injectable sodium pentobarbital; produced by Abbott laboratories) can be obtained from Bulter Schein Animal Health. Note that this is a schedule II controlled substance and requires a Drug Enforcement Agency (DEA) license to obtain.
The Bupivacaine solution (0.25%; produced by Hospira, Inc, among others) can be obtained from a number of distributors, including Moore Medical.		</div>

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.

Watch this Scientific Journal Video about Ultrasound-Guided Microinjection into the Mouse Forebrain In Utero at E9.5 at JoVE.com

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.

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 ...

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Research

JoVE Journal

Neuroscience

17.9K Views.  Johns Hopkins University School of Medicine. 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.

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

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

Efficient Gene Delivery into Multiple CNS Territories Using In Utero Electroporation

The ability to manipulate gene expression is the cornerstone of modern day experimental embryology, leading to the elucidation of multiple developmental pathways. Several powerful and well established transgenic technologies are available to manipulate gene expression levels in mouse, allowing for the generation of both loss- and gain-of-function models. However, the generation of mouse transgenics is both costly and time consuming. Alternative methods of gene manipulation have therefore been widely sought. In utero electroporation is a method of gene delivery into live mouse embryos1,2 that we have successfully adapted3,4. It is largely based on the success of in ovo electroporation technologies that are commonly used in chick5. Briefly, DNA is injected into the open ventricles of the developing brain and the application of an electrical current causes the formation of transient pores in cell membranes, allowing for the uptake of DNA into the cell. In our hands, embryos can be efficiently electroporated as early as embryonic day (E) 11.5, while the targeting of younger embryos would require an ultrasound-guided microinjection protocol, as previously described6. Conversely, E15.5 is the latest stage we can easily electroporate, due to the onset of parietal and frontal bone differentiation, which hampers microinjection into the brain. In contrast, the retina is accessible through the end of embryogenesis. Embryos can be collected at any time point throughout the embryonic or early postnatal period. Injection of a reporter construct facilitates the identification of transfected cells.
To date, in utero electroporation has been most widely used for the analysis of neocortical development1,2,3,4. More recent studies have targeted the embryonic retina7,8,9 and thalamus10,11,12. Here, we present a modified in utero electroporation protocol that can be easily adapted to target different domains of the embryonic CNS. We provide evidence that by using this technique, we can target the embryonic telencephalon, diencephalon and retina. Representative results are presented, first showing the use of this technique to introduce DNA expression constructs into the lateral ventricles, allowing us to monitor progenitor maturation, differentiation and migration in the embryonic telencephalon. We also show that this technique can be used to target DNA to the diencephalic territories surrounding the 3rd ventricle, allowing the migratory routes of differentiating neurons into diencephalic nuclei to be monitored. Finally, we show that the use of micromanipulators allows us to accurately introduce DNA constructs into small target areas, including the subretinal space, allowing us to analyse the effects of manipulating gene expression on retinal development.

Efficient Gene Delivery into Multiple CNS Territories Using In Utero Electroporation

In utero electroporation allows for rapid gene delivery in a spatially- and temporally-controlled manner in the developing central nervous system (CNS). Here we describe a highly adaptable in utero electroporation protocol that can be used to deliver expression constructs into multiple embryonic CNS domains, including the telencephalon, diencephalon and retina.

The ability to manipulate gene expression is the cornerstone of modern day experimental embryology, leading to the elucidation of multiple developmental ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance molecules, local gene transfer or knock- out is required. Gene targeting knock-in or knock-out into local regions is possible to perform with combination with a specific CRE line, which is laborious, costly, and time consuming. Therefore, a simple transfection method, an in utero electroporation technique, which can be performed with short time, will be handy to test the possible function of candidate genes prior to the generation of transgenic animals 1,2. In addition to this, in utero electroporation targets areas of the brain where no specific CRE line exists, and will limit embryonic lethality 3,4. Here, we present a method of in utero electroporation combining two different types of electrodes for simple and convenient gene transfer into target areas of the developing brain. First, a unique holding method of embryos using an optic fiber optic light cable will make small embryos (from E9.5) visible for targeted DNA solution injection into ventricles and needle type electrodes insertion to the targeted brain area 5,6. The patterning of the brain such as cortical area occur at early embryonic stage, therefore, these early electroporation from E9.5 make a big contribution to understand entire area patterning event. Second, the precise shape of a capillary prevents uterine damage by making holes by insertion of the capillary. Furthermore, the precise shape of the needle electrodes are created with tungsten and platinum wire and sharpened using sand paper and insulated with nail polish 7, a method which is described in great detail in this protocol. This unique technique allows transfection of plasmid DNA into restricted areas of the brain and will enable small embryos to be electroporated. This will help to, open a new window for many scientists who are working on cell differentiation, cell migration, axon guidance in very early embryonic stage. Moreover, this technique will allow scientists to transfect plasmid DNA into deep parts of the developing brain such as thalamus and hypothalamus, where not many region-specific CRE lines exist for gain of function (GOF) or loss of function (LOF) analyses.

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

A gene transfer method into the developing mouse brain is described by using a unique surgical method and special shape of electrodes. This unique technique allows transfection of plasmid DNA temporally and spatially, which will aid many neuroscientists in studying brain development.

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance ...

Time-lapse Imaging of Neuroblast Migration in Acute Slices of the Adult Mouse Forebrain

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells in restricted areas of the adult mammalian brain. Newborn neuroblasts from the subventricular zone (SVZ) migrate along the rostral migratory stream (RMS) to their final destination in the olfactory bulb (OB)1. In the RMS, neuroblasts migrate tangentially in chains ensheathed by astrocytic processes2,3 using blood vessels as a structural support and a source of molecular factors required for migration4,5. In the OB, neuroblasts detach from the chains and migrate radially into the different bulbar layers where they differentiate into interneurons and integrate into the existing network1, 6.
In this manuscript we describe the procedure for monitoring cell migration in acute slices of the rodent brain. The use of acute slices allows the assessment of cell migration in the microenvironment that closely resembling to in vivo conditions and in brain regions that are difficult to access for in vivo imaging. In addition, it avoids long culturing condition as in the case of organotypic and cell cultures that may eventually alter the migration properties of the cells. Neuronal precursors in acute slices can be visualized using DIC optics or fluorescent proteins. Viral labeling of neuronal precursors in the SVZ, grafting neuroblasts from reporter mice into the SVZ of wild-type mice, and using transgenic mice that express fluorescent protein in neuroblasts are all suitable methods for visualizing neuroblasts and following their migration. The later method, however, does not allow individual cells to be tracked for long periods of time because of the high density of labeled cells. We used a wide-field fluorescent upright microscope equipped with a CCD camera to achieve a relatively rapid acquisition interval (one image every 15 or 30 sec) to reliably identify the stationary and migratory phases. A precise identification of the duration of the stationary and migratory phases is crucial for the unambiguous interpretation of results. We also performed multiple z-step acquisitions to monitor neuroblasts migration in 3D. Wide-field fluorescent imaging has been used extensively to visualize neuronal migration7-10. Here, we describe detailed protocol for labeling neuroblasts, performing real-time video-imaging of neuroblast migration in acute slices of the adult mouse forebrain, and analyzing cell migration. While the described protocol exemplified the migration of neuroblasts in the adult RMS, it can also be used to follow cell migration in embryonic and early postnatal brains.

We describe a protocol for real-time videoimaging of neuronal migration in the mouse forebrain. The migration of virally-labeled or grafted neuronal precursors was recorded in acute live slices using wide-field fluorescent imaging with a relatively rapid acquisition interval to study the different phases of cell migration, including the durations of the stationary and migration phases and the speed of migration.

There is a substantial body of evidence indicating that new functional neurons are constitutively generated from an endogenous pool of neural stem cells ...

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 ...

Hippocampal Insulin Microinjection and In vivo Microdialysis During Spatial Memory Testing

Glucose metabolism is a useful marker for local neural activity, forming the basis of methods such as 2-deoxyglucose and functional magnetic resonance imaging. However, use of such methods in animal models requires anesthesia and hence both alters the brain state and prevents behavioral measures. An alternative method is the use of in vivo microdialysis to take continuous measurement of brain extracellular fluid concentrations of glucose, lactate, and related metabolites in awake, unrestrained animals. This technique is especially useful when combined with tasks designed to rely on specific brain regions and/or acute pharmacological manipulation; for example, hippocampal measurements during a spatial working memory task (spontaneous alternation) show a dip in extracellular glucose and rise in lactate that are suggestive of enhanced glycolysis1-3,4-5, and intrahippocampal insulin administration both improves memory and increases hippocampal glycolysis6. Substances such as insulin can be delivered to the hippocampus via the same microdialysis probe used to measure metabolites. The use of spontaneous alternation as a measure of hippocampal function is designed to avoid any confound from stressful motivators (e.g. footshock), restraint, or rewards (e.g. food), all of which can alter both task performance and metabolism; this task also provides a measure of motor activity that permits control for nonspecific effects of treatment. Combined, these methods permit direct measurement of the neurochemical and metabolic variables regulating behavior.

Hippocampal Insulin Microinjection and In vivo Microdialysis During Spatial Memory Testing

Modulation of hippocampally-dependent spatial working memory by direct intrahippocampal microinjection, accompanied and followed by in vivo microdialysis for metabolites in conscious, behaving animals.

Glucose metabolism is a useful marker for local neural activity, forming the basis of methods such as 2-deoxyglucose and functional magnetic resonance ...

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Genetic modification of specific regions of the developing mammalian brain is a very powerful experimental approach. However, generating novel mouse mutants is often frustratingly slow. It has been shown that access to the mouse brain developing in utero with reasonable post-operatory survival is possible. Still, results with this procedure have been reported almost exclusively for the most superficial and easily accessible part of the developing brain, i.e. the cortex. The thalamus, a narrower and more medial region, has proven more difficult to target. Transfection into deeper nuclei, especially those of the hypothalamus, is perhaps the most challenging and therefore very few results have been reported. Here we demonstrate a procedure to target the entire hypothalamic neuroepithelium or part of it (hypothalamic regions) for transfection through electroporation. The keys to our approach are longer narcosis times, injection in the third ventricle, and appropriate kind and positioning of the electrodes. Additionally, we show results of targeting and subsequent histological analysis of the most recessed hypothalamic nucleus, the mammillary body.

Genetic Manipulation of the Mouse Developing Hypothalamus through In utero Electroporation

Despite the functional and medical importance of the hypothalamus, in utero genetic manipulation of its development has rarely been attempted. We show a detailed procedure for in utero electroporation into the mouse hypothalamus and show representative results of total and partial (regional) hypothalamic transfection.

Genetic  modification of specific regions of the developing mammalian brain is a very  powerful experimental approach. However, generating novel mouse ...

Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

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

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

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

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...