In Situ Visualization of Axon Growth and Growth Cone Dynamics in Acute Ex Vivo Embryonic Brain Slice Cultures

Animal experiments must comply with the relevant institutional and federal regulations. Embryonic day 15.5 and 12.5 (E15.5 and E12.5) pregnant female C57BL/6JRj mice were used in this protocol. Experiments were performed in accordance with the Animal Welfare Act of North Rhein-Westphalia State Environmental Agency (Landesamt für Natur, Umwelt und Verbraucherschutz (LANUV)).

1. Preparation of plasmids for injection

1. Isolate DNA using endotoxin-free Maxiprep kit according to manufacturer's protocol (see Table of Materials).
2. Mix selected DNA at the desired concentration (Table 1) and 10% Fast Green solution (see Table of Materials) to visualize the delivery of DNA mixture into brain ventricles.
   NOTE: Specific plasmids were used for sparse labeling of the cortical neurons (Figure 1A), filamentous actin (F-actin) structures in the growth cone (Figure 1B), and dual-labeling of the growth cones within the same cortex (Figure 1C). All plasmids (Table 1) used in this protocol have been deposited in Addgene (see Table of Materials).
3. Prepare glass capillaries using a capillary electrode puller set as per the following program: pressure: 500, heat: 800, pull: 30, and velocity: 40.
4. Load 15 µL of DNA/Fast Green mix into each glass capillary using micro loader pipette tips.
   NOTE: Ensure that no bubbles form.
5. Store DNA-filled capillaries in a 10 cm dish with a piece of modeling clay across the diameter of the dish. Capillaries can be loaded and stored at 4 °C the day before the experiment. Seal the back end of the capillary with flexible film to prevent drying.

2. Preparation of solutions

1. Prepare Hank's Buffered Salt Solution-supplemented with glucose (HBSS-G).
   1. Add 0.5% of 20% glucose stock to a bottle of 1x HBSS. Mix well and store at 4 °C for up to 2 weeks. For the embryo extraction, bubble HBSS-G solution with carbogen (95% O₂ and 5% CO₂) using bubbling stone shortly before embryo collection.
2. Slice media solution
   1. Prepare fresh slice media containing Neurobasal 1x, 5% horse serum, 5% foetal calf serum, B27 supplement 1:50, L-glutamin supplement 1:400, penicillin-streptomycin 1:200, and Neuropan-2 supplement 1:100 (at pH = 7.3), under sterile conditions (see Table of Materials).
   2. Prepare 3 cm dishes with 1 mL of slice media each. Place in the incubator at 35 °C with 5% CO₂ for at least 1 h before the experiment to equilibrate the media's pH through gas exchange.
      NOTE: Media pH equilibration is caused by acidification of the media by the CO₂ from the incubator. Slice media can be stored at 4 °C for up to 1 week.
3. Low melting point agarose solution (3%)
   1. Weigh the desired amount of low melting point agarose powder and dissolve in an appropriate volume of 1x HBSS-G in a glass bottle. Approximately 7 mL of agarose solution per brain is needed.
   2. Place the bottle in a microwave for 2-3 min, with the cap loosely placed, and shake every 10-20 s.
   3. Once the powder has completely dissolved, place the bottle in a water bath or bead bath set to 37 °C at least 1 h before the experiment to allow agarose to cool down.
      NOTE: It is recommended to heat the agarose twice over 15 min to ensure the agarose powder is dissolved. This is crucial for the proper adhesion of agarose to brain tissue. A thermometer should be used to measure the temperature of the agarose solution while embedding brains, making sure it is between 37-40 °C. Brains from different aged animals have different rigidity. It is recommended to test a range of agarose concentrations to find homogeny between tissue and agarose.
   4. Prepare Phosphate Buffered Saline with 0.3% Triton X-100 (PBS-T).
   5. Prepare Phosphate Buffered Saline with 0.2% Sodium Azide (PBS-NaN₃).
      NOTE: Solutions described in steps 2.4-2.5 are for use in the later immunohistochemistry step.

3. Preparation of the surgery station

1. Clean the surgery station with 70%-96% ethanol and place operation underlay on the station surface.
2. Sterilize the surgery instruments by rinsing with 70%-96% ethanol, followed by dry sterilization in a hot bead sterilizer.
3. Clean platinum tweezer electrodes (see Table of Materials) with 70%-96% ethanol before connecting to the pulse generator.
4. Insert a DNA/Fast Green-filled glass capillary into the capillary holder. Immediately before use, gently break off the capillary tip using fine scissors and test solution flow inside a 1.5 mL microcentrifuge tube filled with pre-warmed saline or water.
   NOTE: Figure 2A,B show surgery station set-up and tools used for ex utero electroporation (EUE) and in utero electroporation (IUE).
5. For IUE, warm-up saline solution to 37 °C in a water bath.

4. Embryo extraction

1. Place pregnant mouse in anesthesia inducer chamber with 5% isoflurane until the mouse is deeply anesthetized. Confirm anesthesia by the absence of pedal withdrawal reflex.
2. Transfer the mouse to an operation underlay and maintain isoflurane at 1.5%-2% through a nose cone.
3. Apply ointment to both eyes to prevent corneal drying.
4. Shave the mouse's abdomen, and then use 70%-96% ethanol-soaked gauze to remove shaved hair. Clean the area with betadine.
5. Using sterile small surgical scissors, make a 2 cm skin incision along the abdominal midline, followed by a 1.5 cm muscle incision.
   NOTE: Incision size depends on the embryo size. Indeed, larger embryos will require a larger incision to accommodate their extraction.
6. Cut a hole in the middle of the gauze sufficiently wide to fit the skin incision (~2 cm in diameter), soak it with warm saline, and place it around the abdominal opening.
7. Pull out both uterine horns using a cotton bud soaked in warm saline or using forceps, carefully grabbing the spaces between embryos to pull them out. Place embryos on wet gauze (Figure 2C).
   NOTE: Even a small damage to blood vessels and capillaries around the uterine horns will likely result in profuse bleeding. Therefore, avoid direct handling of these vascularised areas at all times.
8. Cut open the uterine sack and remove each embryo (Figure 2D).
9. Euthanize each embryo immediately after extraction via a descending diagonal cut, ensuring complete spinal cord transection (Figure 2E).
10. Place the embryos in a 10 cm dish containing HBSS-G on ice.
    NOTE: Beheading the embryo is avoided to prevent DNA/Fast Green mixture leakage from the brain and facilitate easy embryo positioning in the holder (see ex utero electroporation, step 5).
11. Sacrifice the mother immediately after extraction of embryos by performing cervical dislocation.
    ​NOTE: Here, the mother is euthanized under anesthesia to spare it from any further post-procedure pain or suffering in compliance with the protocol approved by the Animal Welfare Act of North Rhein-Westphalia State Environmental Agency (LANUV).

5. Ex utero electroporation (EUE)

1. Pick up an embryo and place it in the holder.
   NOTE: A cut 1 mL pipette tip attached to the end of a cell scraper is used as an embryo holder. It is essential to keep arms of embryos outside the tip during the procedure to prevent them from sliding into the tip (Figure 2F-I). The diameter of the pipette tip is easily adjustable to accommodate embryos of various sizes. Cut a second tip at length where the diameter of the tip matches embryo size and use it as an adapter insert to the holder mentioned above.
2. Carefully insert DNA/Fast Green-filled glass capillary through the embryo's skull into the lateral ventricle and inject 2-3 µL of DNA plasmid mix (Figure 1A,B; Table 1) into each ventricle (Figure 2F).
   NOTE: Use the lambdoidal and sagittal sutures as a guide for the location of DNA injection. The lambdoidal and sagittal sutures are fibrous joints connecting the bone plate of the skull. The former joins the parietal bone with the occipital bone, and the latterjoins the two parietal bones.
3. Hold the embryo's head between platinum tweezer electrodes at the appropriate angle to target the desired brain area (60° angle in this case), with the cathode facing the area where DNA transfer is intended (Figure 2G-H).
4. Apply five pulses at 30 mV with an interval of 1 s and a duration of 50 ms using a square wave pulse generator.
   NOTE: Bear in mind that brains in EUE experience a more effective electric field than those in IUE. Hence, at a given DNA concentration, EUE results in higher efficiency of DNA transfer than IUE, and DNA concentrations need to be adjusted accordingly.
5. If bilateral electroporation is desired, repeat steps 5.3-5.4 with the cathode and anode mirroring the previous position to target the contralateral cortex.
   NOTE: Since both ventricles were injected with DNA, cortices of both hemispheres were targeted.
6. Place the electroporated embryo in a 6 cm dish containing ice-cold HBSS-G. Repeat steps 5.1-5.6 for all embryos required.

6. In utero electroporation (IUE)

1. Inject pregnant mouse with analgesic; 50 µL of buprenorphine (0.1 mg/kg) (see Table of Materials) subcutaneously, 20 min before the procedure.
2. Perform steps 4.1-4.8 from the embryo extraction section.
   NOTE: Avoid leaving embryos exposed unnecessarily by covering them with sterile gauze soaked in warm saline.
3. Using fingertips, gently rotate the embryo inside the uterus until lambdoidal and sagittal sutures are located (Figure 2J). Carefully insert DNA/Fast Green glass capillary through the uterine wall and embryo's skull into the lateral ventricle and inject 2-3 µL of DNA plasmid mix (Figure 1A,C) into either one or both ventricles as desired (Figure 2K-L).
   NOTE: Excessive finger pressure on uterine horns could lead to amniotic sack collapse.
4. Hold the embryo's head between platinum tweezer electrodes at the appropriate angle to target the desired brain area (60° angle in this case), with the cathode facing the area where the DNA transfer is intended. Avoid squeezing the uterus since it may cause the collapse of the amniotic sack (Figure 2M).
5. Apply five pulses at 35 mV with an interval of 600 ms and a duration of 50 ms using a square wave pulse generator.
6. If both lateral ventricles were injected, repeat steps 6.5-6.6 with the cathode and anode mirroring the previous position to target the contralateral cortex.
7. Repeat steps 6.3-6.6 for all embryos required.
8. Once all the required embryos have been electroporated, use a saline-soaked cotton bud to place uterine horns back inside the abdominal cavity gently.
   NOTE: The addition of saline solution into the peritoneal cavity will help uterine horns slide back into position.
9. Suture muscle and skin incisions using 5-0 suture material. Use suture clips to secure the wound and disinfect the suture wound by spraying it with betadine (Figure 2N-P).
10. Inject the mouse with 200 µL of 5% glucose subcutaneously.
11. Inject the mouse with an antibiotic; 50 µL of Enrofloxacin (5 mg/kg) subcutaneously (see Table of Materials).
12. Place the mouse back in the recovery cage and maintain warmth using a far-infrared warming light or heating pad for at least 20 min post-procedure (Figure 2Q).
13. Monitor the mouse daily, and inject meloxicam after the procedure for pain relief following institutional and federal guidelines.
14. Extract embryos 2 days after the procedure (i.e., E17.5) following step 4.

7. Brain extraction and embedding in agarose

NOTE: It is recommended to carry out the following steps under a dissection microscope for better precision. Avoiding damage to the brain is critical for the success of the procedure.

1. Set up extraction tools in a sterile working space under a dissection hood (Figure 3A).
2. Separate the head of an embryo from the rest of the body using dissection scissors.
3. Fix the head as shown in Figure 3C, and then remove the skin and skull by cutting along the midline, starting from the base of the head toward the nose (Figure 3D).
4. Peel the skin and skull laterally, making a big enough gap (~1 cm) for the brain to be excised.
5. To remove the brain, insert the closed tip of sterile dissection scissors, starting from under the olfactory bulb moving toward the brain stem (Figure 3E).
6. Cut off the brain stem and trim any loose pieces of meninges around the brain (Figure 3F).
   NOTE: Loose meninges often cause slices to remain attached to the agarose block after cutting, leading to detaching of tissue from the agarose during slice collection.
7. Repeat steps 7.1-7.6 for all embryos and keep the brains on ice (ideally no longer than 30 min) until the embedding step.
   NOTE: The following steps 7.7.1-7.7.4 refer to brain extraction of E12.5 brains.
   1. Isolate the top of the head just below the eye, as shown in Figure 3G.
   2. Cut the skin and skull on top of the brainstem, following the dashed line as shown in Figure 3H, without removing the brainstem.
   3. Make a 2 mm skin-skull incision at the back of the head, as shown in Figure 3I (see drawings for clarity).
      NOTE: This incision provides initial gripping points to peel off the layers of skin and skull. Typically, they come off as one layer. The typical incision size is 2 mm, corresponding to the length of the cutting edge of the used micro spring scissors.
   4. Start peeling off skin-skull layers by securing one side of the incision and pulling the other carefully. With the same care, finalize by peeling off the base of the head until freeing the brain (Figure 3J).
      NOTE: This must be done with great care, observing that the brain is not being pulled along the tissue layers. Alternate sides to remove the tissue covering the brain.
8. Pour warm agarose (at 37-40 °C) in a 3 cm dish.
9. Pick up the brain using a perforated spoon and remove excess liquid by dabbing the bottom of the spoon against dry tissue paper. Place the brain in an agarose dish.
   NOTE: It is essential to remove as much liquid from around the brain as possible to allow for better adhesion of agarose to the tissue.
10. Put the dish with liquid agarose on ice. Using a smaller spoon, mix agarose for 10 s for even cooling. Maneuver the brain to the middle of the dish. Place the brain horizontally in the dish with the dorsal side up, ensuring it is completely covered with agarose from all directions (Figure 3K).
    NOTE: Brains will often sink to the bottom of the dish once placed in agarose; lift the brain using a small spoon until a gap of 1-2 mm under the brain is established.
11. Repeat steps 7.8-7.10 for all brains.
12. Once agarose has polymerized, add 500 µL of HBSS-G on top of the agarose block to prevent drying. Then, cover the dish with ice.
    ​NOTE: Keep the sample on ice for 5 min before sectioning to allow the brain temperature to reach 4 °C.

8. Organotypic slice culture

NOTE: Clean vibratome and surrounding surfaces with 70%-96% ethanol to avoid slice contamination. The set-up of the vibratome workstation (see Table of Materials) is shown in Figure 3B.

1. Fill the vibratome buffer tray with cold HBSS-G and the outer tray with ice to maintain the HBSS-G cold throughout the procedure.
2. Continuously supply HBSS-G in the buffer tray with carbogen using a bubbling stone.
3. Using a fresh blade, make a large cut (~2 x 2 cm) around the brain and remove a block of agarose containing the brain, with enough surrounding agarose to trim the agarose into a small rectangle block.
   NOTE: This step allows adjusting the block's angle so that the sagittal axis of the brain is perpendicular to the vibratome plate and the coronal axis aligns parallel to the blade. Leave around 5 mm of agarose at the dorsal side of the brain for easy handling of the slices.
4. Place a small drop of fast-adhesive solvent-free superglue in the middle of the specimen holder and spread to an area that will cover the bottom of the agarose block.
5. Gently pick up the agarose block and dry the bottom by dabbing against tissue paper. Place the block on the glued area of the specimen holder, with the rostral side of the brain up. Put the specimen holder on ice and allow the glue to dry for 1 min.
6. Once the glue has dried, place the specimen holder in the buffer tray.
7. Cut the brain in coronal slices at an angle of 15°.
   NOTE: The thickness of the slices can vary depending on the application. Here, brains were sliced at a thickness of 150 µm. Set vibratome speed to 1.0-1.5 mm/s for trimming excess agarose on top and trimming olfactory bulbs. Reduce cutting speed to 0.5 mm/s for collecting cortical slices for analysis. Most vibratomes can be paused to collect each slice. If reduced quality of slices or detaching of tissue from agarose are experienced, reducing the cutting speed or replacing the vibratome blade can help.
8. Using clean spatulas, collect brain slices and place them on Polytetrafluoroethylene (PTFE) membrane, immobilized in a 35 mm glass-bottomed dish using paraffin (up to five brain slices/ membrane) (Figure 3L-M).
   NOTE: Fix the PTFE membrane inside a 35 mm glass-bottomed dish using wax. This will stabilize the membrane when adding the slice culture media and also during imaging.
9. Using a 200 µL pipette, remove excess HBSS-G from around the slices on the PTFE membrane, leaving the slices semi-dry.
10. Add 500 µL of slice media (pre-warmed to 35 °C) directly to the space under the PTFE membrane.
    NOTE: No bubbles must form under the membrane when adding the media. This will leave whole or partial slices with no media exchange. Replace 200 µL of media every 2 days in culture or after every imaging session.
11. Incubate the slices at 35 °C with 5% CO₂.

9. Immunohistochemistry

1. Fix slices with 1 mL of 4% paraformaldehyde (PFA)-supplemented with 4% sucrose-per dish. Incubate at RT for 30 min.
   CAUTION: When handling PFA, wear a lab coat and gloves. Perform fixation steps under a chemical hood, and dispose of PFA waste appropriately.
2. Wash the slices twice with 300 µL PBS for 5 min. Transfer the slices to a 24-well plate.
   NOTE: The experiment can be paused at this stage. Add PBS-NaN₃ to the slices and store at 4 °C. NaN₃ is a toxic compound; when handling solutions with it, wear a lab coat and gloves. Steps 9.3-9.10 areto be performed in an orbital shaker.
3. Quench the slices with 300 µL of 0.1 M glycine at 4 °C overnight.
4. Wash out glycine with PBS at RT 3x for 10 min.
5. Permeabilise the slices with 300 µL of PBS-T at RT for 2 h.
6. Block using 10% goat serum in PBS-T at RT for 2 h.
7. Add 300 µL of primary antibody (anti-vimentin antibody at a dilution of 1:200; see Table of Materials) diluted in 10% goat serum in PBS-T solution at 4 °C overnight.
   NOTE: In steps 9.8-9.12, slices were light-protected to prevent loss of fluorescence.
8. Wash primary antibody with PBS at RT 3x for 20 min.
   NOTE: PBS was used instead of PBS-T to wash out Triton X-100.
9. Add 300 µL of secondary antibody (either Alexa Fluor 488 or 647 at a dilution of 1:400; see Table of Materials) in PBS at RT for 2 h.
   NOTE: DAPI is added immediately after removing the secondary antibody at a dilution of 1:10,000 for 5 min.
10. Wash the secondary antibody with PBS at RT 3x for 20 min. Wash with distilled water 2x for 1 min.
11. Transfer the slices to a glass slide using a fine brush, and then dry at 30 °C for 20 min.
12. Mount the slices using an aqueous mounting medium. Keep the slides at RT overnight for the mounting media to curate.

10. Imaging acquisition

NOTE: Regardless of the DNA delivery approach (IUE or EUE), slices were analyzed at the same developmental age range (E17.5-E18.5). IUE allows neuronal progenitors to divide and develop for two further days in vivo. EUE, on the other hand, allows for the tracking of early developmental events.

1. Switch on the chamber incubator and set it to 35 °C with 5% CO₂-ideally 4 h before imaging-to allow microscope components to equilibrate at 35 °C.
2. For deep imaging of slices, use water-immersion objectives to reduce the mismatch in refractive index between the tissue and objective.
   NOTE: Here, super-resolution imaging mode was used. Imaging through the PTFE membrane requires an objective with a long working distance (~1 mm). If a long working distance objective is unavailable, slices may be transferred to an 8-well glass-bottomed dish. To transfer slices, add 1 mL of slice media to the top of the membrane, and then use a spatula to lift a slice and transfer it to a well containing 200 µL of media. Remove excess media using a 1 mL pipette tip leaving the slices semi-dry.
3. For imaging axon growth, locate a region of the cortex with low to medium cell density. For imaging growth cone dynamics, locate a growth cone in the cortex's intermediate zone or subventricular zone.
4. Define a z-stack size in the image processing software (see Table of Materials). For axon growth in a large z-stack, set a step size of 2 µm. For growth cones in a smaller z-stack, set a step size of 1 µm.
   NOTE: Always account for potential movement of the growth cone and axon through the x, y, and z planes. Axons grow at a much higher rate in organotypic cultures than in in vitro cultures. Here, a z-stack of around 80 µm to image axon growth was sufficient. For growth cone dynamics, a z-stack of ~6 µm was adequate.
5. For imaging axon growth of neurons in a larger area, define a tile scan.
6. Use the lowest laser power possible to minimize the chances of bleaching growth cones during acquisition.
7. For imaging axon growth, acquire time-lapses for 2 h with an interval of 5 min. For imaging growth cone dynamics, acquire time-lapses for 2-5 min with an interval of 2.5-3 s.

11. Data analysis

1. Measure the speed of axon growth using kymographs
   1. Open the image file in Fiji¹⁴ through File > Open and select the image.
   2. Obtain the maximum intensity projection of the time-lapse through Image > Stacks > Z-Projection > Maximum Intensity Projection.
   3. Go through the time-lapse and locate a growing axon.
   4. Once located, draw a line through the growing axon. Start from the tip of the axon in the first frame, and follow the axon through the entire time-lapse.
   5. Generate a kymograph using the plugin KymoResliceWide.
   6. Set the scale of the kymograph by going to Image > Properties. Set the distance in µm in Pixel Width and set the time in s or min in Pixel Height.
   7. Go to Analyze > Measure.
      NOTE: An angle relative to the x-axis will be given.
   8. Calculate the speed of axon growth by substituting the angle in the following equation: SIN(RADIANS(θ))/COS(RADIANS(θ)) in a spreadsheet.
2. Measure the volume of the growth cone using an image analysis software (see Table of Materials).
   1. Open the image file in the image analysis software through File > Open and select the file of interest.
   2. Select the Add New Surfaces wizard.
      NOTE: A section on the lower-left corner will appear with six steps for manual editing.
   3. In step 1 -under Algorithm Settings- select Segment Only A Region Of Interest. In step 2, crop the frame to fit the entire growth cone in all frames.
   4. Keep thresholding to Absolute Intensity in step 3, and ensure the entire growth cone region is thresholded in step 4.
   5. In step 5, select Number of Voxels lmg = 1 under Filter Type.
      NOTE: In the last step, multiple measurement sets may be created. Here, only one measurement was created for volume.
   6. Select the Execute button to perform all the creation steps and terminate the Add New Surfaces Wizard.
   7. In the Statistics tab at the top of the wizard window, select Specific Values and Volume under the Detailed tab.