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

Summary

Cell-autonomous functions of genes in the brain can be studied by inducing loss or gain of function in sparse populations of cells. Here, we describe in utero electroporation to deliver Cre recombinase into sparse populations of developing cortical neurons with floxed genes to cause loss of function in vivo.

Abstract

Cell-autonomous neuronal functions of genes can be revealed by causing loss or gain of function of a gene in a small and sparse population of neurons. To do so requires generating a mosaic in which neurons with loss or gain of function of a gene are surrounded by genetically unperturbed tissue. Here, we combine the Cre-lox recombination system with in utero electroporation in order to generate mosaic brain tissue that can be used to study the cell-autonomous function of genes in neurons. DNA constructs (available through repositories), coding for a fluorescent label and Cre recombinase, are introduced into developing cortical neurons containing genes flanked with loxP sites in the brains of mouse embryos using in utero electroporation. Additionally, we describe various adaptations to the in utero electroporation method that increase survivability and reproducibility. This method also involves establishing a titer for Cre-mediated recombination in a sparse or dense population of neurons. Histological preparations of labeled brain tissue do not require (but can be adapted to) immunohistochemistry. The constructs used guarantee that fluorescently labeled neurons carry the gene for Cre recombinase. Histological preparations allow morphological analysis of neurons through confocal imaging of dendritic and axonal arbors and dendritic spines. Because loss or gain of function is achieved in sparse mosaic tissue, this method permits the study of cell-autonomous necessity and sufficiency of gene products in vivo.

Introduction

Generating a genetic mosaic is a classic experimental paradigm for understanding the function of a gene of interest. To determine if a gene is necessary for a cellular phenotype, the simplest approach is causing a loss of function of the gene throughout the organism (e.g. knockout). However, to determine if a gene is required specifically in a certain cell type, knockout of the gene throughout the organism is not a valid approach. Instead, a method is required that will cause the loss of function of a gene in a given cell while it is surrounded by wildtype (i.e. genetically unperturbed) tissue—in other words, creating mosaic tissue. If the mutant cell shows a mutant phenotype, but surrounding wildtype cells do not, the gene functions in a cell-autonomous manner. Analysis of mosaic tissue, in which mutant cells are surrounded by wildtype tissue, is ideal for understanding cell-autonomous functions of genes, especially in the brain where neurons and glia form a vast interconnected network of tissue.

Several forms of mosaic brain tissue have provided powerful models to investigate cell-autonomous functions of genes. Studies focused on neuronal transplantation¹, female X-linked mosaicism^2,3,4, and endogenous somatic mosaicism^5,6 have drawn their conclusions based on mosaic brain tissue. Conditional deletion of a gene through the Cre-lox recombination system is a method that takes full advantage of the great availability of transgenic mouse lines. In this method, two loxP sites are introduced on either side of a required sequence of a gene (such as an exon), leaving it flanked by loxP sites that both face in the same direction ("floxed"). Cre recombinase excises the sequence between the loxP sites⁷. Cre-mediated recombination can be achieved by crossing floxed mice to another mouse line expressing Cre recombinase along with a fluorescent marker in a subset of cells ("Cre reporter line"). This has been demonstrated in a variety of ways to uncover the functions of a gene in subsets of cells, such as excitatory neurons or astrocytes⁸. Cre reporter lines can express CreER^T2 to allow Cre-mediated recombination to be drug-inducible (single-neuron labeling with inducible Cre-mediated knockout, or SLICK)⁹. In another strategy called mosaic analysis with double markers (MADM)^10,11, Cre-mediated interchromosomal recombination allows a homozygous mutant to be created alongside heterozygous tissue. In these approaches, a new line of mice needs to be produced each time for each candidate gene or cellular subtype that is tested. Alternatively, Cre recombinase can be introduced postnatally through iontophoresis¹² or through viral vectors (e.g. adeno-associated viruses¹³ or lentiviruses¹⁴ carrying cellular subtype-specific promoters). This strategy creates strong and postnatal labeling. To target developing cerebral cortical neurons sparsely and prenatally, an ideal strategy is in utero electroporation of Cre recombinase with a fluorescent marker.

In addition to combining Cre-lox recombination through in utero electroporation to produce mosaic tissue in vivo, we introduce several adaptations to procedures from other published protocols^{15,16,17,18,19,20,21}. We provide information to improve success in breeding timed-pregnant females. We also outline our two strategies to introduce sparse and bright labeling of neurons in cortical tissue: One strategy is to titrate the levels of a single construct coding for Cre recombinase and a fluorescent marker²². Another strategy is to use the "Supernova" system, designed specifically with these parameters in mind^23,24. Additionally, we offer improvements on producing consistent microinjection pipettes and simplifications to the in utero electroporation surgery. Finally, we outline critical steps in a simplified histological preparation that permits the analysis of dendritic spines and dendritic and axonal arbors, without further staining or immunohistochemistry.

Protocol

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.

Results

The single construct GFP.Cre (see list of materials) was electroporated at E15.5 and visualized at P14. Depending on the concentration of the construct and the volume of injection, a sparse or dense result can be obtained^22,26. For example, injection of 1 µL of 2 mg/mL GFP.Cre results in a sparse distribution of labeled cells, some of which can be bright (Figure 1A), and localized in layer II/III (

Discussion

Here, we introduce the combination of in utero electroporation with Cre recombinase in floxed mice to generate mosaic brain tissue. An advantage of this approach is that a new mouse line does not need to be generated each time a different cellular subtype is to be targeted: in utero electroporation can be used to target excitatory neurons, inhibitory neurons, or glia depending on the time and location of electroporation^15,16,

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
C57BL/6J mice	The Jackson Laboratory	#000664	See "1. Mouse set-up" (step 1.1, "wildtype mice")
GFP.Cre empty vector	AddGene	#20781	See "2. DNA set-up" (step 2.1 "single DNA construct that codes for Cre recombinase as well as a fluorescent marker"). GFP.Cre empty vector was a gift from Tyler Jacks.
pK029.CAG-loxP-stop-loxP-RFP-ires-tTA-WPRE (Supernova)	AddGene	#69138	See "2. DNA set-up" (step 2.1 "Supernova" system) and http://snsupport.webcrow.jp/. pK029.CAG-loxP-stop-loxP-RFP-ires-tTA-WPRE (Supernova) was a gift from Takuji Iwasato.
pK031.TRE-Cre (Supernova)	AddGene	#69136	See "2. DNA set-up" (step 2.1 "Supernova" system) and http://snsupport.webcrow.jp/. pK031.TRE-Cre (Supernova) was a gift from Takuji Iwasato.
pK038.CAG-loxP-stop-loxP-EGFP-ires-tTA-WPRE (Supernova)	AddGene	#85006	See "2. DNA set-up" (step 2.1 "Supernova" system) and http://snsupport.webcrow.jp/. pK038.CAG-loxP-stop-loxP-EGFP-ires-tTA-WPRE (Supernova) was a gift from Takuji Iwasato.
EndoFree Plasmid Maxi Kit (10)	Qiagen	#12362	See "2. DNA set-up" (step 2.3 "endotoxin-free plasmid purification kit")
Trypan Blue powder, BioReagent grade	Sigma	T6146-5G	See "2. DNA set-up" (step 2.5 "trypan blue")
Sodium Chloride, ACS, 2.5 kg	VWR	BDH9286-2.5KG	See "2. DNA set-up" (step 2.5 "NaCl")
Potassium Chloride, ACS, 500 g	VWR	#97061-566	See "2. DNA set-up" (step 2.5 "KCl")
Sodium phosphate dibasic, ReagentPlus, 100 g	Sigma-Aldrich	S0876-100G	See "2. DNA set-up" (step 2.5 "Na2HPO4")
Potassium phosphate monobasic, ReagentPlus, 100 g	Sigma-Aldrich	P5379-100G	See "2. DNA set-up" (step 2.5 "KH2PO4")
Hydrochloric acid, ACS reagent, 500 mL	Fisher Scientific	A144-500	See "2. DNA set-up" (step 2.5 "HCl")
P-97 Micropipette Puller	Sutter Instrument	P-97	See "3. Pipette set-up" (step 3.1 "glass capillary puller")
3.0 mm wide trough filament	Sutter Instrument	FT330B	See "3. Pipette set-up" (step 3.1 "glass capillary puller")
Thin Wall Glass Capillaries, 4", 1 / 0.75 OD/ID	World Precision Instruments	TW100-4	See "3. Pipette set-up" (step 3.1.1 "glass capillary")
Single Ply Soft-Tech Wipes, 4.5"	Phenix	LW-8148	See "3. Pipette set-up" (step 3.2.1 "single-ply task wipe"); other single-ply wipes (e.g. Kimwipes) can be used.
Graefe Forceps, 7 cm, Straight, 0.7 mm 1x2 Teeth	World Precision Instruments	#14140	See "4. In utero electroporation" (step 4.1 "Graefe forceps")
Iris Scissors, 11.5 cm, Straight, 12-pack	World Precision Instruments	#503708-12	See "4. In utero electroporation" (step 4.1 "iris scissors")
Hartman Mosquito Forceps, 9 cm, Straight, 12-pack	World Precision Instruments	#503728-12	See "4. In utero electroporation" (step 4.1 "Hartman mosquito forceps")
General Purpose Non-Woven Sponges, 2" x 2", 4-ply	Medrepexpress	#2204-c	See "4. In utero electroporation" (step 4.1 "non-woven gauze sponges")
Ring Tipped Forceps, 10 cm, Straight, 2.2mm ID	World Precision Instruments	#503203	See "4. In utero electroporation" (step 4.1 "ring-tipped forceps")
Pyrex petri dishes complete, O.D. × H 100 mm × 20 mm	Sigma-Aldrich	CLS3160102-12EA	See "4. In utero electroporation" (step 4.1 "Petri dishes")
Flat Type Instrument Tray, Stainless Steel, 13-5/8" x 9-3/4" x 5/8"	Amazon	B007SHGAHA	See "4. In utero electroporation" (step 4.1 "stainless steel tray")
Platinum Tweezertrode, 5 mm	BTX	#45-0489	See "4. In utero electroporation" (step 4.3 and 4.16 "tweezer-type electrodes")
ECM 830 Foot Pedal	BTX	#45-0211	See "4. In utero electroporation" (step 4.3 and 4.17 "foot pedal")
ECM 830 Generator	BTX	#45-0052	See "4. In utero electroporation" (step 4.3 "generator")
Single Animal Isoflurane Anesthesia System with Small Induction Box	Harvard Apparatus	#72-6468	See "4. In utero electroporation" (step 4.4 and 4.6 "nose cone", step 4.4 "induction chamber")
Ophthalmic ointment	Hanna Pharmaceutical Supply Co	#0536108691	See "4. In utero electroporation" (step 4.7 "veterinary ophthalmic ointment")
Space Gel (AIMS)	VWR	#95059-640	See "4. In utero electroporation" (step 4.8 "sealed pouch filled with supersaturated salt solution")
Hair Remover Gel Cream, Sensitive Formula	Veet	#062200809951	See "4. In utero electroporation" (step 4.9 "depilatory cream")
10ul Low Retention Tip Starter (960 tips/pk)	Phenix Research Products	TSP-10LKIT	See "4. In utero electroporation" (step 4.12 "sterile 10 µL micropipette tip")
Aspirator tube assemblies for calibrated microcapillary pipettes	Sigma-Aldrich	A5177	See "4. In utero electroporation" (step 4.15 "aspirator tube assembly")
Braided Absorbable Suture, 4-0, Needle NFS-2(FS-2), 27"	Medrepexpress	MV-J397	See "4. In utero electroporation" (step 4.19 "absorbable sutures")
“LiquiVet Rapid” Tissue Adhesive	Medrepexpress	VG3	See "4. In utero electroporation" (step 4.20 "tissue adhesive")
Hypodermic syringes, polypropylene, Luer lock tip, capacity 1.0 mL	Sigma-Aldrich	Z551546-100EA	See "4. In utero electroporation" (step 4.21 "1 mL syringe")
BD Precisionglide syringe needles gauge 26, L 1/2 in.	Sigma-Aldrich	Z192392-100EA	See "4. In utero electroporation" (step 4.21 "26G, ½” needle")
Nestlets Nesting Material	Ancare	NES3600	See "4. In utero electroporation" (step 4.24 "nesting materials")
Sunflower Seeds, Black Oil, Sterile	Bio-Serv	S5137	See "4. In utero electroporation" (step 4.24 "sunflower seeds")
Paraformaldehyde, 97%	Alfa Aesar	A11313	See "5. Histology" (step 5.1.1 "PFA")
Economy Tweezers #3, 11 cm, 0.2 x 0.4 mm tips	World Precision Instruments	#501976	See "5. Histology" (step 5.5 "tweezers")
Agar powder	Alfa Aesar	#10752	See "5. Histology" (step 5.8.1 "agar")
Single-edge razor blades, #9 blade	Stanley Tools	#11-515	See "5. Histology" (step 5.9 "single-edge razor blade")
Specimen disc S D 50 mm	Leica	#14046327404	See "5. Histology" (step 5.9 "vibrating microtome specimen disc")
Buffer tray S assembly	Leica	#1404630132	See "5. Histology" (step 5.10 "buffer tray")
VT1000 S Vibratome	Leica	#14047235612	See "5. Histology" (step 5.10 "vibrating microtome")
Double Edge Razor Blades	Personna	BP9020	See "5. Histology" (step 5.10 "blade")
Knife Holder S	Leica	#14046230131	See "5. Histology" (step 5.10 "knife holder")
Studio Elements Golden Taklon Short Handle Round Brush Set	Amazon	B0089KU6XE	See "5. Histology" (step 5.12.1 "fine tipped paintbrush")
Superfrost Plus Slides	Electron Microscopy Services	#71869-11	See "5. Histology" (step 5.12.1 "microscope slide")
ProLong Diamond Antifade Mountant, 10 ml	Thermofisher	P36970	See "5. Histology" (step 5.12.3-5.12.4 "mountant")
Cover Glass, 24 x 50 mm, No. 1	Phenix Research Products	MS1415-10	See "5. Histology" (step 5.12.4 "coverslip")
4′,6-Diamidino-2-phenylindole dihydrochloride (DAPI)	Sigma-Aldrich	D9542	See "5. Histology" (step 5.13.1 "DAPI")
Fixed Stage Upright Microscope	Olympus	BX51WI	See "5. Histology" (step 5.15 "light microscope")
Laser Scanning Confocal Microscope	Nikon	TE2000/C2si	See "5. Histology" (step 5.15 "confocal microscope")
4x objective, NA = 0.20	Nikon	CFI Plan Apo Lambda 4X	See "5. Histology" (step 5.15 "low-power objective")
20x objective, NA = 0.75	Nikon	CFI Plan Apo Lambda 20X	See "5. Histology" (step 5.15 "medium-power objective")
60x objective, NA = 1.40	Nikon	CFI Plan Apo VC 60X Oil	See "5. Histology" (step 5.15 "high-power objective")