Inducing Cre-lox Recombination in Mouse Cerebral Cortex Through In Utero Electroporation

Methods described here have been approved by the Animal Care and Use Committee (ACUC) of James Madison University, and are in accordance and compliance with all relevant regulatory and institutional guidelines.

1. Mouse Set-up

1. House a young (>P60) male and female homozygous floxed mouse together to set up a breeder pair²⁵.
   NOTE: A good negative control is to set up an additional breeder pair of wildtype mice, in which Cre recombinase expression will not cause a mosaic²⁶.
2. Allow the female to give birth and raise her first litter with the male. Keep the male together with the female to increase the litter's survival. After weaning the litter, e.g. at postnatal day (P) 21, separate the female and male.
3. Murine breeding
   1. Set up a timed mating between the female and male²⁵. Use visual cues to determine the estrus cycle of the female²⁷, and check for vaginal plugs the morning after mating²⁵.
   2. If a plug is found, separate, and weigh the female. Count the day as embryonic day (E) 0.5.
   3. Continue weighing the female every 3-5 days to determine if she is pregnant. Pregnant females will have a 10-20% increase in body weight 7-10 days after conception (E0).
      NOTE: This step confirms pregnancy earlier and more reliably than visual inspection²⁵.
   4. If the female is not pregnant, repeat steps 1.3.1-1.3.3.
4. Once pregnancy is confirmed, choose the date of the in utero electroporation. To target layer II/III cortical pyramidal neuronal progenitors, perform electroporation at E15.5.

2. DNA set-up

1. Choose a single DNA construct that codes for Cre recombinase as well as a fluorescent marker, such as green fluorescent protein (GFP)^28,29. Alternatively, use the "Supernova" system, described in detail in previous publications^23,24. See list of materials for example constructs.
2. Amplify the DNA construct(s) from step 2.1 in E. coli using standard microbiological techniques³⁰.
3. Purify the DNA construct with an endotoxin-free plasmid purification kit following the manufacturer's instructions. Elute the construct with endotoxin-free water.
   NOTE: Choosing an endotoxin-free purification kit over a standard purification kit is critical.
4. Concentrate the DNA eluate to 1-3 mg/mL depending on desired labeling density²⁶.
   NOTE: A titer always needs to be established when working with a new DNA construct, given that they have different lengths and promoters. Consider injecting a construct at several different concentrations and/or amounts to assess expression. For example, injection of 1 µL or 1.5 µL of 2.0 mg/mL GFP.Cre is able to cause sparse (1 µL) or dense (1.5 µL) labeling of layer II/III visual cortical pyramidal neurons²⁶.
5. Add 0.4% trypan blue in 1x phosphate buffered saline (PBS; 137 mM NaCl, 2.7 mM KCl, 10 mM Na₂HPO₄, 1.8 mM KH₂PO₄ in H₂O, adjusted to pH 7.4 with HCl) 1:10 to the concentrated DNA solution.
6. Add 10x PBS 1:10 to the concentrated DNA solution.
7. Store the concentrated DNA solution at 4 °C. The solution may be stored indefinitely.

3. Pipette Set-up

1. Using a glass capillary puller, pull a pipette with a very long (10-15 mm) taper and very fine tip, typical of pipettes for embryonic stem cell injection or nuclear transfer³¹.
   NOTE: To prevent debris from contaminating the pipettes, they should be made within 2 days of the surgery.
   1. Calibrate the glass capillary puller by performing a ramp test, following the manufacturer's instructions. Use the resulting heat value to program a pulling protocol using the following parameters: heat = (ramp test value + 15), pull = 30, vel = 120, time = 100, pressure = 200. Insert the glass capillary, fasten to clamps on puller, and activate the programmed pulling protocol, creating a taper on the glass capillary.
   2. Ensure that the taper length is between 10-15 mm using a compound light microscope (e.g. 10X objective)³¹.
2. Break back the pipette to form a rough-edged tip to 20-25 µm diameter.
   1. Center a single-ply task wipe (see list of materials) over a 50 mL beaker and stretch wipe taut with one hand. With the other hand, hold and push a pipette (taper side down) perpendicularly and fully through the center of the wipe, causing a clean break in the taper³¹.
      NOTE: Pushing the pipette fully through the wipe will produce a 20-25 µm diameter tip about 70% of the time.
   2. Confirm under a compound light microscope (e.g. 20x objective).
      Caution: Use eye protection while breaking the glass.
3. Calibrate a pipette by aspirating 1 µL of 0.4% trypan blue in 1x PBS into a partially filled piptte, then graduating the pipette with 1 µL markings based on the height of 1 µL in the pipette, using a fine-tipped alcohol-resistant marker. Use the calibrated pipette to graduate the other pipettes before discarding. Keep pipettes in a pipette holder free from dust and other particles.

4. In Utero Electroporation

1. Place Graefe forceps, iris scissors, Hartman mosquito forceps, non-woven gauze sponges, ring-tipped forceps, and Petri dishes on a stainless steel tray, and wrap with aluminum foil. Place tray and 1 L 0.9% saline (0.9% wt/vol NaCl in water) in autoclave. Autoclave at 121 °C and 3.5 kPa for 40 min. Remove from autoclave and place in a disinfected surgical area.
   NOTE: It is critical to maintain sterile conditions during the surgery.
2. Warm the autoclaved 1 L saline bottle in a small water bath to 38 °C at least 2 h before the surgery.
3. Connect the tweezer-type electrodes and the foot pedal to the generator (see list of materials). Following the manufacturer's instructions, set the generator to deliver five 50 ms pulses of 50 V (with a 950 ms interval) when triggered by the foot pedal. Disinfect the tweezer-type electrodes and place on the disinfected surgical area.
4. Make an anesthesia nose cone as previously published³², but use the end of a nitrile or latex glove finger to form a diaphragm that fits securely over the nose cone opening. Cut a hole in the diaphragm large enough to fit over the mouse's snout.
5. Deeply anesthetize pregnant mice at E15.5 in an induction chamber filled with isoflurane (2-4%) in pure oxygen flowing at 1 L/min. After ~1 min, test the righting reflex (gently tilt the induction chamber and ensure that the mouse does not right itself). Weigh mouse to determine dosages of analgesia administered later during the surgery.
6. Transfer the mouse to the surgical area and administer inhaled isoflurane (1-2%) in pure oxygen flowing at 1 L/min through a nose cone to maintain mouse in surgical plane. Use forceps to test the pedal reflex (pinch the paw gently and ensure that the mouse does not return a reflex) to verify anesthetization. Monitor respiratory rate and effort, looking for deep, regular breathing (~70-100 breaths/min), and that mucous membranes are pink in color and moist.
7. Apply enough veterinary ophthalmic ointment over the eyes to cover them completely for protection against drying during the procedure.
8. Flex the metal activator disc of a sealed pouch filled with supersaturated salt solution (see list of materials), activating an exothermic crystallization of the salt. Lay 2 single-ply task wipes on top of the pouch, and place the mouse on the pouch to maintain body temperature.
9. Massage depilatory cream onto the abdominal area with cotton swabs until abdominal fur dissolves (approximately 20-40 s), then wipe off abdominal fur. Carefully clean off any remaining residue and depilatory cream with 70% ethanol and cotton swabs. Position a fenestrated sterile surgical drape over the abdominal area.
10. Using aseptic technique, pull the skin up and away from abdomen with Graefe forceps, and make a ~2 cm ventral midline incision on the skin using iris scissors, ensuring that underlying muscle is not cut.
    NOTE: An acceptable alternative to making an incision with Graefe forceps and iris scissors is to use a scalpel, as published previously¹⁷.
11. Find the linea alba to visualize the midline of the rectus abdominis. Pull muscle up and away from abdomen with Graefe forceps, and make another ~2 cm midline incision there with iris scissors, exposing the abdominal cavity (the uterus is translucent and embryos will be visible beneath). Ensure that underlying uterine and intestinal tissue is not cut. Make the incision only large enough (typically ~2 cm) to pull out and expose the uterus comfortably.
    NOTE: An acceptable alternative to making an incision with Graefe forceps and iris scissors is to use a scalpel, as published previously¹⁷.
12. Using a sterile 10 µL micropipette tip, dispense carprofen (dissolved in sterile 0.9% saline) at 4 mg/kg into the abdominal cavity for analgesia.
13. Clamp one Hartman mosquito forceps on the left edge of the rectus abdominis incision. Rest the forceps on overturned Petri dish to the left of the incision, keeping the left side of the incision open. Clamp another Hartman mosquito forceps on the right edge of the rectus abdominis incision and rest the forceps on an overturned Petri dish to the right of the incision, keeping the right side of the incision open. Place gauze sponge around the area of the incision.
14. Using ring forceps (without an attached limit screw), pull on the uterus between any two neighboring embryos without crushing or injuring any tissue. Begin pulling out all embryos through the incision, laying them on top of the gauze sponge. When pulling out the last embryos from the abdominal cavity, make sure not to pull on the ovaries or cervix. Once exposed, it is critical to keep the uterus moist with warm saline. The uterus typically contains between 5 and 8 embryos.
    NOTE: It is possible to add penicillin and streptomycin to the saline¹⁷.
15. Connect a calibrated pipette to the aspirator tube assembly (see list of materials), and inject exactly 1 µL of the DNA solution prepared in step 2 through the uterine wall into a lateral ventricle of each embryo. Lateral ventricles appear as two patches on the dorsal telencephalon of the embryo (patches are darker than surrounding tissue of telencephalon). Use the thumb and forefinger during microinjection and electroporation to manipulate the embryos, revealing the lateral ventricles and allowing the embryos to be pushed gently against the uterine wall during injection. Confirm successful injection by observing trypan blue filling the lateral ventricle.
16. Place the cathode of tweezer-type electrodes (see list of materials) on the uterus, directly over the medial and caudal cortex to target the visual cortex (targeting alternative areas of the brain will require different electrode placement )^16,26. Place the anode on the uterus, just inferior and anterior to the embryo's head.
17. Confirm that the anode and cathode are touching the uterus surrounding the embryo at the appropriate locations. With a foot pedal, trigger the delivery of five 50 ms pulses of 50 V (with a 950 ms interval) across the electrodes.
    NOTE: The injected, negatively-charged DNA will move toward the cathode and will be incorporated into ventricular zone cells closest to the cathode.
18. Return the uterus to the abdominal cavity in the same orientation it was found. Use saline to lubricate the uterus while guiding it manually and extremely gently, taking care not to displace embryos from their position in the uterus.
19. Close the abdominal muscles with absorbable sutures (see list of materials) using a simple interrupted stitch, tying the ends with a surgeon's knot. Do not clip the ends too close to the knot or the knot will become unfastened.
    NOTE: It is critical to suture the abdominal muscles such that the edges of the wound are completely closed shut, but without causing blanching of the muscle. Knots should be tight so they cannot be unfastened.
20. Close the skin layer with absorbable sutures (simple interrupted stitch, surgeon's knot, as in step 4.19). Apply a small amount of tissue adhesive to seal the wound. Apply tissue adhesive on the knots to prevent unfastening. Use a micropipette to remove any tissue adhesive surrounding the wound that can be aspirated by the micropipette.
21. Lower isoflurane levels to 0.5-1.5%. Once tissue adhesive is dry (test by touching glove to adhesive), remove from isoflurane and allow female to recover alone in a warm cage. Administer buprenorphine intraperitoneally at 0.03 mg/kg using a 1 mL syringe with a 26G, ½" needle.
22. Continue to monitor the mouse until the mouse is fully recovered and behaving normally (i.e. grooming, exploring cage, eating, or drinking). Once recovered, place female back in cage with male.
    NOTE: If all steps are followed, the mouse will typically regain consciousness within 2 min and immediately begin moving about the cage, showing no signs of discomfort.
23. If the mouse exhibits signs of discomfort (e.g. hunched posture, porphyrin secretions)³³, administer carprofen at 0.1 mg/mL in water supply. If several signs of discomfort are present, or if discomfort persists for more than 4 h despite carprofen administration, immediately euthanize the mouse by administering an intraperitoneal lethal injection of ketamine (240 mg/kg)-xylazine (48 mg/kg)-acepromazine (1.85 mg/kg) cocktail using a 1 mL syringe with a 26 G, ½" needle, and follow with an approved secondary form of euthanasia³³.
24. To increase the survival of electroporated embryos after birth, keep the cage in a quiet holding room, free from noises and vibrations. Enrich the environment with plastic igloos, nesting materials, and dietary supplements such as sunflower seeds (see list of materials). Do not move or disturb the cage, particularly during the first 3 days after parturition.

5. Histological Preparation for Fluorescence Microscopy

NOTE: This histology protocol is optimized for preparing tissue from animals older than postnatal day (P) 13 that were electroporated in utero. To prepare tissue from younger postnatal animals (P0-P13), it is recommended to follow all steps (including transcardial perfusion), though the brain should be embedded in agar prior to preparing sections (step 5.9). Tissue may even be prepared from embryos within 1-2 days after electroporation, using methods described previously^18,20.

1. Prepare paraformaldehyde (PFA) solution.
   NOTE: It is imperative to prepare fresh PFA, within one day of the experiment.
   Caution: Paraformaldehyde is toxic and should be handled in a fume hood with appropriate personal protective equipment.
   1. To make 50 mL of 4% PFA, heat 40 mL water to 60 °C. Add 2 g PFA while stirring with glass rod or stirring bar. Add 10 µL of 10 M NaOH every 2 min until PFA dissolves.
   2. Add 5 mL of 10x PBS and bring solution to a final volume of 50 mL with water.
   3. After bringing solution to 50 mL, adjust pH to 7.4 as needed with 10 M HCl. Allow solution to cool and store at 4 °C.
2. Euthanize mouse with an intraperitoneal injection of ketamine (240 mg/kg)-xylazine (48 mg/kg)-acepromazine (1.85 mg/kg) cocktail using a 1 mL syringe with a 26G, ½" needle. Transcardially perfuse 10 or 20 mL ice-cold 1x PBS, followed by 25 or 40 mL freshly-made, ice-cold 4% PFA¹⁷. Use lower volumes for young postnatal (P0-P13) mice and higher volumes for older mice (P14 and above).
3. Immediately after perfusion, decapitate mouse carcass between occipital bone and C1 vertebra with large scissors and peel away and discard most of the skin surrounding skull using iris scissors, particularly skin surrounding frontal, parietal, and occipital bones. Keep the skull and brain intact.
4. Place the skull and brain in 10 mL 4% PFA in a 50 mL conical tube for 24 h at 4 °C to post-fix tissue. Then add 30 mL 1x PBS to conical tube to dilute the solution to 1% PFA for storage at 4 °C.
   NOTE: Use a separate conical tube for each skull and brain to be post-fixed. Do not incubate the brain in 1% PFA for longer than 2 weeks, or fluorescence will begin to fade.
5. Place the brain and skull on a flat surface with single-ply task wipes moistened with 1x PBS. Use a pair of tweezers (see list of materials) to remove any remaining skin or membrane surrounding the skull.
6. Using tweezers, first remove the occipital bone, then carefully remove the parietal bones, moving the tweezers out and away from the surface of the brain. Carefully remove any meninges to prevent damage to the cortex.
7. Wedge the back of the tweezers under the brain along the skull to sever any cranial nerves, and remove the brain.
8. For young postnatal (P0-P13) brains
   1. Make 25 mL of 4% agar by heating 25 mL 1x PBS to 80 °C. Stir and dissolve 1 g agar with a glass rod or stirring bar).
   2. Cool solution to 35 °C while continuing to stir. Place brain in a small container (e.g. the cap of a 50 mL conical tube), and pour agar solution over brain. Allow agar to harden.
9. Make a coronal cut manually using a single-edge razor blade through the brain, approximately 0.5 mm rostral to bregma. Make another coronal cut through the brain approximately 0.5 mm caudal to visual cortex. Place a pool of cyanoacrylate glue with the same diameter as the rostral end of the brain on the center of the vibrating microtome specimen disc. Place the rostral end of the brain on top of the pool of glue, ensuring that the entire rostral surface of the tissue is bound to the specimen disc with glue.
10. Mount buffer tray onto vibrating microtome and secure with built-in clamping lever. Insert blade into knife holder, and mount knife holder onto vibrating microtome with built-in clamping screw. Secure specimen disc with brain onto buffer tray using built-in clamping device. Set speed setting to 5.70 (= 0.285 mm/s), and frequency setting to 5.33 (53.3 Hz).
11. Fill the chamber above the level of the brain with 1x PBS and lower the blade into the PBS. Begin taking 100 µm coronal sections through the tissue.
12. If not requiring further processing, such as 4',6-diamidino-2-phenylindole (DAPI) nuclear staining or immunohistochemistry.
    1. Use a fine tipped paintbrush to transfer sections directly from the vibrating microtome onto a microscope slide, fitting 4-6 sections onto each slide.
    2. Allow the sections to dry by wicking away solution with a fine tipped paintbrush, such that the brain slices no longer drift on the slides.
    3. Then, place one drop of mountant (see list of materials) on each section on the slide.
    4. Gently lay a 24 x 50 mm coverslip on top, ensuring that 1) no air bubbles form on the sections, 2) the sections do not move, and 3) the mountant fully covers the area between the slide and coverslip. Allow the mountant to cure for at least 24-48 h.
13. To stain nuclei with DAPI for histological examination of cortical laminae
    1. Use a fine tipped paintbrush to transfer each section into a solution containing 10 µg/mL DAPI (see list of materials) diluted in 1x PBS from stock solution (20 mg/mL DAPI in water).
    2. Incubate in DAPI solution for 5 min at room temperature, then use a fine tipped paintbrush to transfer section from DAPI solution to 1x PBS and incubate in 1x PBS for 5 min at room temperature. Transfer section two more times to fresh 1x PBS for 5 min each. Then, using a fine tipped paintbrush, transfer the section onto a microscope slide, follow steps 5.11.2-5.11.4, and proceed with step 5.13.
14. Heat a glass Petri dish containing Valap (equal parts Vaseline, lanolin, and paraffin) on a hot plate at a low heat setting until fully melted. Seal exposed edges of slides by brushing on melted Valap.
15. To inspect fluorescence in tissue, use a light microscope or confocal microscope (see list of materials) and a filter set adequate for viewing the fluorophore (e.g. DAPI: excitation 325-375 nm, emission 435-485 nm; GFP: excitation 450-490 nm, emission 500-550 nm)¹⁷. Confocal images may be taken with low- (4X, numerical aperture, NA = 0.20), medium- (20X, NA = 0.75), and high-power (60X, NA = 1.40) objectives.