Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

Jane transferred to the Developing Mouse Inner Ear by in Vivo, electroporation Ling Wang, Han Jang, and John Burgandy, Oregon Hearing Research Center, Oregon Health and Science University.Okay. My name is John Burgandy. My lab is at the Oregon Hearing Research Center at the Oregon Health and Science University.

My name is Ang. I'm a poster in Johns lab. I'm Ang also in the Johns lab as a research assistant.

Our presentation is broken down into four parts. The first is ventral laparotomy. We demonstrate sodium pen, barbital based anesthesia, how to test for the adequacy of anesthesia, corneal protection, preparation of the surgical site, disinfection mouse survival surgery data sheet, the surgical suite ventral midline incision through the skin and the abdominal wall and externalization of the uterine horns.

Part two is trans uterine microinjection. We demonstrate trans illumination of the embryo, embryo, reorientation, microinjection, pipette, fabrication, and microinjection into the Otis at embryonic day 11.5 and embryonic day 12.5. Part three is in vivo electroporation.

We demonstrate paddle style electrode placement, and electroporation of the embryonic day 11.5 mouse O assist. Part four is representative results. We show the expression of GFP in late embryonic and early postnatal cochleas.

That resulted from in vivo electroporation of the Otis with expression plasmid at embryonic day 11.5 part one ventral laparotomy. The dam is held securely by grasping the skin behind the neck, the tail, and the left hind limb. An intraperitoneal injection of sodium pentobarbital based anesthetic is administered after swabbing the abdomen with 70%ethanol.

The mouse is placed inside its home cage for four to five minutes to allow the anesthetic to act. Foot and tail pinches are used to assess the adequacy of anesthesia. This dam shutters in response to the tail pinch and needs two more minutes before becoming unresponsive to noxious stimuli.

Sterile ophthalmic ointment is applied to the eyes to protect the corneas from desiccation. We prepare the surgical site in a chemical fume hood to minimize the spread of fur and dander. The fur is shaved with a fine blade to expose the skin covering the abdominal wall.

Complete fur removal facilitates healing of the incision postoperatively. Disinfection of the abdominal skin is achieved by alternating passes of 70%ethanol Betadine, and 70%ethanol. Always swipe from rostral to coddle and use a fresh surgical cotton ball or cotton tipped applicator.

For each pass, we assess the depth and quality of respirations during disinfection to ensure that the dam is unresponsive After, After the second 70%ethanol pass, we immediately placed the damn ventral surface down on a sterile drape and set her on a heated pad for two to three minutes to warm her. This is a convenient time to record preoperative data on the mouse survival surgery log sheet. We have a appended our mouse survival surgery data sheet as supplementary information.

Consider including the data sheet and your animal care protocol so your institutional animal care and use committee can see how you'll monitor preoperative operative and postoperative data. We complete one log sheet for each surgical subject. We prefer to conduct surgery in a sterile environment, though this is typically not required for rodents.

The work surface is separate from the blower motor and plenum housing, so no vibration is transmitted to the user. During microinjection, we have appended as supplementary information, a schematic of the modifications we made to our laminar flow hood to facilitate In utero gene transfer, the essential changes are low wat challenge and track lighting cutouts and hidden receptacles for instrument power cords, and a routing path for accessory electrical circuits. The surgical suite consists of the electric lactated ringers IV bag, warm bath, XY, Z manipulator, stereo fluorescence, dissecting microscope, fiber optic light source, and surgical instruments.

Foot pedals are used to trigger the micro injector, electro perter enter focus. The scope surgical instruments are sterilized in an autoclave. Instruments must be re-sterilized between mice using a glass bead sterilizer or by re autoclaving the needle driver.

Balli scissors ring forceps and vascular forceps are the essential tools. Ventral midline incision, grasp the skin with vascular forceps and excise the integument with the ball tipped scissors, extend the incision 10 to 14 Millimeters. Identify the white avascular band of connected tissue called the linea alba and the ventral midline of the abdomen incise the linear alba with the ball tipped scissors.

Being careful not to nick the gut or the inguinal fat pads. Immediately irrigate the abdomen with sterile Prewarm lactated ringer solution. Assess the incision sites in the skin and abdominal wall for bleeding.

Direct pressure with blunt forceps will arrest minor bleeding if present, externalize the right and left uterine horns with ring tipped forceps. Gently catch the right uterine horn with one ringed end of the forcep and draw the uterus toward the midline incision. Use your free hand to gently manipulate the lateral side of the dam to facilitate hooking the uterus.

The forceps are never used to compress the uterus and pull it out. Since this could damage the uterine vasculature or the embryos, Repeat the externalization procedure for the left uterine horn. Immediately Irrigate the newly externalized uterine horns with prewarm lactated ringer solution gently reinsert the inal fat pads or the intestines should they externalize with the uterus.

This is not encountered often. Part two uterine micro injection. The dam is arranged beneath the long working distance objective and a soft cable fiber optic light is set against the uterine wall.

The uterus is gently pressed with a finger of the free hand. To facilitate identification of the embryo's orientation. Light from the fiber optic cable should be at the lowest intensity required to accurately assess embryo orientation.

Intense light can transmit heat to the uterus that may cause complications. We hold the fiber optic light and the uterus in one hand and use a finger on our free hand to reposition the embryo by Gentle palpation. We demonstrate in the following Three video by microscopy EC clips and the common embryo orientations observed by of the uterus.

The viewer will see the microscopic view that we see during trans illumination, though in black and white trans illumination of the uterus permits detection of the embryo's orientation in vivo. In this example, the embryo is in a headstand position with its ventral surface facing the viewer. By toggling the uterus back and forth, we can detect the right and left hind limbs, the tail and the left eye cephalon.

In this Example, the embryo is again in the headstand position, but rotated about 90 degrees with respect to the anterior posterior axis. By toggling the uterus gently back and forth, we can see the paddle shapes that define the left hind limb and the left for limb. The hind brain Is also evident.

The embryo is in an upright Position with its left side facing the viewer. The nascent fourth ventricle is detectable as a bright patch in the hind brain. The left eye cephalon, the left four limb and the left hind limbs are also evident.

Gentle pressure on the uterus reorients the embryos slightly to enable detection of the main trunk of the primary head vein and its anterior and posterior branches, which are marked with white and black asterisk respectively. As we will see, the Otis is located midway between the anterior and posterior branches of the primary head vein. The Otis itself cannot be seen through the uterus by transillumination.

We demonstrate in two successive video microscopy clips how to reorient the embryo for microinjection into the Otis assist. The goal is to identify the embryonic landmarks that permit interpolation of the otis's location in the lateral head. Mesen con, our goal is to reorient the embryo from the dorsal to lateral position so we can identify the primary head vein.

The mesencephalon fourth ventricle and left for limb are evident in the dorsal position. Gentle pressure on the uterus shifts the embryo's orientation from dorsal to later. The left eye cephalon, mesencephalon, and fourth ventricle are readily apparent.

The primary head vein and its posterior branch indicated by the asterisk are also obvious. The anterior branch of the primary head vein is not clearly detected. Our goal, again is to orient the embryo so it can identify the primary head vein, but under higher magnification appropriate for uterine microinjection.

The uterine vessels decidual band and left eye are seen in this lateral perspective. Gentle pressure on the uterus shifts the position of the embryo, so we can detect the fourth ventricle left eye. The primary head vein and its posterior branch marked by the asterisk, the primary head vein and its branches make the shape of American football uprights.

The otic vesicle cannot be seen through the uterus, but it is located In the center of the uprights. Lynn has initiated an Injection sequence for one embryo. She trans illuminates the uterus, reorients the embryo for a micro injection, and brings the micro injection pipette filled with fast green into the field.

The micro injection pipette is attached to a pipette holder that is held by an X, Y, z microm manipulator. The manipulator magnetic stand is attached to a steel plate with a Teflon base that permits effortless gross positioning of the microinjection pipette. The needle is advanced using the forward micrometer control knob on the manipulator appropriately.

Fabricated pipettes are essential for a traumatic microinjection into the early Otis stage mouse embryo. We pull thick walled borosil glass capillary tubing with a horizontal pipette puller. The resulting pipette shown in A is broken manually with forceps at about the 14 to 16 micron outer diameter position indicated by the arrowhead in B.The rough break of the pipette shown in C is jagged and fragile.

We bevel the pipette at about 20 degrees to sharpen the pipette tip as seen in Panel D parameters for pulling and beveling microinjection pipettes are provided in the accompanying text. We demonstrate in the next two video microscopy clips microinjection into the mouse otic assist at embryonic days 11.5 and 12.5. Our goal is to demonstrate uterine microinjection into the embryonic day, 11.5 mouse otics.

First, we inject the otics without the uterus present to show unambiguously how a correctly targeted injection appears. Then we demonstrate authentic trans uterine micro injection. For this first injection, the uterus was incised fally to allow the visceral sac to extrude the decidual band overlying the visceral yolk sac was removed.

The placenta remains attached to the uterus and the embryo is alive. With the uterus focally removed, we see the fourth ventricle, the primary head vein, and its anterior and posterior branches, the pipette and the approximate location of the cyst in white. At the start of this injection sequence, blood flow is detected in the visceral yolk sac vasculature.

The pipette is advanced through the visceral yolk sac and dai is injected into the osis note, the endo lymphatic duct dorsally and the position of the otics in the center of the uprights. The otics is midway between the anterior and posterior branches of the primary head vein, uterine micro injection into the embryonic day. 11.5 mouse otics requires detection of the primary head vein and at least one of its branches to estimate the location of the otic assist.

Our target in this diagrammatic example, both the anterior and posterior branches of the primary head vein are detected with the OUC territory marked in white. A more realistic presentation of the embryo is one where the main trunk of the primary head vein is detected along with only one of its branches and the example shown only the posterior branch of the primary head vein is seen, but that is enough to interpretate the location of the otics. The PT is advanced through the uterus and dye is injected into the otics.

We wait 30 seconds to allow dye to clear from the amniotic cavity, at which point we can see the endo lymphatic duct dorsally and the vestibule. We conclude with a low magnification view of the embryo whose cyst we just injected to provide anatomical perspective in this view, the fourth ventricle left eye and uterine vasculature are seen. We demonstrate trans and microinjection into the embryonic day 12 mouse otic cyst and microinjection into the nascent fourth ventricle of the hind brain.

Transillumination allows us to detect the primary head vein in this left lateral view as well as the eye. The pipette is advanced through the uterus into the otic vesicle and dies injected to fill its lumen. To inject the fourth ventricle, we rotate the embryo 90 degrees for a dorsal view of the hind brain.

The injected left cyst is now on the left side. The pipette is advanced through the uterus and fast green mixed with lor. Conjugated dextran is injected into the fourth ventricle.

Note that the volume of dye injected does not enter the midbrain in vivo. Fluorescence is assessed after the injection to validate dextran localization in the fourth ventricle. The embryo is then reoriented from dorsal to lateral position.

The endo lymphatic duct is just ventral to the fourth ventricle in this lateral view. In vivo fluorescence validates localization of the dextran in the fourth ventricle. At birth, we screen pups for green fluorescence in the fourth ventricle using an epi fluorescence.

Dissecting microscope pups with green fluorescence in their hind brains possess an inner ear that was manipulated during embryogenesis part three in vivo electroporation. After filling the cyst with expression plasmid, the uterus is freshly irrigated with prewarm lactated ringer solution. Transillumination facilitates centering the ODU cyst in the electrode field.

The metal head of the fiber optic light is retracted from the uterine surface. Before initiating the square wave pulse train, we demonstrate in the next video microscopy clip a complete electroporation cycle in vivo. Electroporation of the embryonic day, 11.5 mouse otic cyst requires gentle placement of the paddle style electrodes on the uterus.

In this example, the left otic cyst has been filled with expression plasmid and the uterus has been freshly irrigated with lactated ringers. The cathode is positioned laterally with respect to the filled otic cyst and the and ode is positioned medially at the start of the ation sequence, the otic cyst is centered in the paddle field. The uterus is gently compressed and the square wave pulse train is initiated with a foot pedal switch.

Ideally 60 to a hundred milliamps of current is delivered per pulse. The uterus is then released and immediately irrigated. The freshly irrigated uterus is returned to the abdominal cavity by gentle pressure with the ring forceps.

Once the uterus is internalized, the abdominal cavity is flushed with about four mils of lactated ringers leave about one mil of lactated ringers in the abdominal cavity to help maintain hydration of the dam who will drink less water than usual immediately after abdominal surgery. The abdominal wall and then the skin are closed with resorbable suture using a needle driver. Our needle driver has a built-in scissor that simplifies closing.

We prefer a running stitch for both the abdominal wall and the skin. We lock every other stitch to provide additional support at the incision site. We administer a nonsteroidal anti-inflammatory medication subcutaneously before placing the dam in the recovery cage.

Consult with your veterinary staff regarding which prophylactic analgesics are preferred by your institutional animal care and use committee. The dam is nestled in a sterile drape and placed on the floor of the heated recovery cage. The recovery cage contains nesting material, a red tube for cover, lab chow, and a water bottle.

The cage is reassembled with a Filter bonnet on top the morning after surgery. The Dam is beautifully groomed, ambulatory, and able to balance on her hind limbs to inquire about the open cage top. The suture line in the abdomen is clean, dry and shows no evidence of separation, redness or swelling.

We have appended a document that discusses animal care protocol development for in utero gene transfer. The document contains language that may be useful as you design an institution specific protocol in consultation with your animal care and use committee. Part four, representative results.

Representative results panels A and B show a representative cochlea from a postnatal day six pup whose Otis was injected with an an enhanced green fluorescent protein expression plasmid at embryonic day 11.5 and then electro. The cochlear lateral wall was removed from the midterm and apex. Only GFP expression is present from the base to the midterm of the cochlea and follows the distribution of the hair cell marker myosin seven A shown in panel B.Panel C shows a low magnification confocal projection of the embryonic day.

18.5 mouse organ of corde immuno stained with a myosin seven A antibody transfected cells are distributed throughout the organ of corde. Panel D shows a high magnification confocal projection of the embryonic day. 18.5 mouse organ of corde stained with cin.Seven.

A GFP expression is detected in inner and outer hair cells, interphalangeal cells, pillar cells, and TER cells. Electroporation mediated gene transfer to the Otis at embryonic day 11.5 Transfect progenitors that give rise to all of the component cell types within the organ of corde.