Transcranial Electrical Brain Stimulation in Alert Rodents

For research involving animals, the relevant (country-specific) approvals must be obtained before starting experiments. All animal experiments reported here are performed according to the EU directive 2010/63/EU, the updated German animal protection law ("Tierschutzgesetz") of July 2013, and the updated German animal research regulations of August 2013. Animal protocols have been approved by the local authorities "Commission for Animal Experimentation of the Regional Council of Freiburg" and "Commission for Animal Experimentation of the University Medical Center Freiburg".

1. Preparation of Instrumentation and Material for Surgery

1. Make sure that the items listed in Figure 1 are available and already placed for surgery.
2. Prepare a thin rectangular platinum plate (e.g., 10 mm x 6 mm x 0.15 mm), which will serve as the counter electrode placed subcutaneously on the chest, and punch two small holes in two opposite corners of the plate.
3. Solder an insulated cable with a length of ~10 cm using a lead-free tin-solder to one of the corners (without a hole) of the platinum plate.
4. Apply a small drop of histo-acrylic glue on the soldering joint for isolation.

2. Preparation of the Rodent for Surgery

1. Assign a study number to the rodent and note this on the prepared surgery card.
2. Weigh the rodent and note the weight on the surgery card. Calculate the dose of injection anesthetics (e.g., ketamine 80 mg/kg bodyweight plus xylazine 12 mg/kg bodyweight for rats).
3. Induce anesthesia by intraperitoneal (i.p.) injection of the calculated amount of anesthetics.
   NOTE: When using inhalation anesthesia instead (e.g., isoflurane), place the rodent in an induction chamber with continuous flow of ~4% in 1-2 L/min oxygen.
4. Check depth of anesthesia by the toe pinch reflex starting 5 min post injection. If the toe pinch reflex is still present, reach prolongation and deepening of anesthesia by injection of 30% of the initial dose.
   1. If at any time point in the experiment the toe pinch reflex returns, 30% of the initial dose of anesthesia should be injected.
   2. When using inhalation anesthesia, look for loss of the postural reflex of the rodent in the induction chamber and check the depth of anesthesia by the lack of a toe pinch reflex. If reflexes are still present, extend the duration in the anesthesia chamber. Throughout the whole experiment, adapt the percentage of isoflurane to the depth of anesthesia until reaching a maintenance concentration of ~1-1.5% isoflurane.
   3. When the frequency of breathing decreases and gasping occurs, lower the percentage; when the rodent regains the toe pinch reflex or shows spontaneous movement, increase the percentage of inhalation anesthetic.
5. As soon as reflexes are absent, place the rodent on the lab bench or hold it in hand.
   NOTE: When using inhalation anesthesia, provide continued reduced isoflurane flow (now between 2-3%) by using a nozzle connected to the nebulizer.
6. Remove the hair on the rat's head by shaving the area from ear to ear and from between the rostral eye level to just behind the ears with a clipper. Then remove the hair on the chest by shaving the area between the forelimbs from the xiphoid up to the clavicles.
   NOTE: Keeping the skin under tension eases the shaving.
7. Cover the eyes of the rat with a drop of eye ointment to protect the cornea.
8. Mark the rat's ear according to the assigned study number.
   NOTE: Depending on the length of the study, a tail mark might be sufficient, otherwise standardized earmarking is preferable.

3. Surgical Procedure: Chest Electrode Implantation

NOTE: This step can be skipped when the counter electrode is placed externally on the shaved chest with a vest.

1. Place the rodent prone (on the chest) on the operation table.
   NOTE: In case of inhalation anesthesia, keep the rat's snout placed in the anesthesia nozzle, further reducing the isoflurane concentration to 1.5-2%.
2. Disinfect the shaved scalp with a disinfectant spray or with a swab soaked in antiseptic agent (e.g., ethanol 70%) and let air-dry. Repeat two times.
3. Cut the skin with a scalpel in one line from the rostral eye level to the mid ear level.
   NOTE: This allows for tunneling of the connecting cable from the implanted chest electrode toward the top of the head and is also the desired cut for the DCS electrode socket placement.
4. Turn the rat to supine position, so that the chest is exposed.
5. Disinfect the skin of the chest as described in step 3.2.
6. Elevate the lateral skin of the right chest with a tissue forceps and cut a buttonhole with small scissors of about 0.5 cm medial from the right axilla. Then make a straight sagittal cut in cranial orientation with the scissors.
7. Form a subcutaneous pouch by atraumatically disconnecting the skin from the left major pectoral muscle. Do so by repeatedly opening the small scissors (or by a saline soaked cotton swab).
8. Turn the animal on its right side to tunnel the cable path from the left occipital corner of the opened head skin along the neck to exit into the pectoral pouch by penetrating the superficial fascia using homeostatic forceps.
9. Carefully open the homeostatic forceps to grab the end of the electrode cable attached to the platinum electrode without allowing sharp wires to stray. Pull the cable through the tunnel until the electrode enters the pouch, oriented with the soldering point towards the rodents left hindlimb. Turn the rodent back to the prone position.
10. Fix the platinum plate with a sterile synthetic braided non-absorbable suture to the pectoral fascia at the two opposing corner holes (4-5 knots are recommended for stability).
11. Similarly attach the cable to the fascia by a loose knot, forming a slight loop before the entrance of the tissue tunnel.
12. Close the skin with 3-4 cutaneous sutures depending on the size of the cut (the same suture material can be used as for the electrode and cable).

4. Surgical Procedure: Placement of the Epicranial tES Socket

1. Place the animal in a stereotactic frame.
   NOTE: If using inhalation anesthesia, lower the concentration of the anesthetic to a maintenance isoflurane flow of ~1.5-1%, adjusted to the toe pinch reflex and breathing pattern.
2. Disinfect the shaved scalp as described in step 3.2.
3. Cut the skin with a scalpel in one line from the rostral eye level to the mid ear level.
   NOTE: If the chest electrode placement was performed, steps 4.2 and 4.3 have already been performed.
4. Scrape off the periosteum (connective tissue on the skull) to the sides with the scalpel and thoroughly wipe off with cotton swaps. Fixate the connective tissue at the 4 corners of the cut with bulldog clamps and let them hang laterally to keep the surgery field open.
5. Apply 0.9% saline to clean the bone surface and tissue with cotton swabs. Then clean the bone surface with 3% H₂O₂. Avoid contact with the tissue. Hereby the bone is cleaned more thoroughly and minor bleeding from the bone will be stopped. Also, residuals of the periosteum become visible. Remove these residuals with a cotton swab applying moderate pressure.
   NOTE: Removal of the periosteum residuals will increase adhesion and durability of the tES socket glued onto the bone.
   1. In case of unstoppable bleeding, use a bone drill and touch it for 1-3 s with slight pressure on the bone. This mechanical procedure will in most cases stop the bleeding without significant heating. Never use electrocautery on the bone; even brief application will result in brain tissue damage (electrocautery should solely be used for wound tissue bleeding).
6. As fixation screws will improve set-up adherence, choose a drill bit fitting the screw size. Place two burr holes on two different bone plates by pre-drilling with a hand drill and then by slight vertical pressure application with the bone drill. Avoid close proximity to the desired position of the tES socket, as it might hinder screwing in the electrode (e.g., for left primary motor cortical tES, choose right frontal and posterior parietal screw position).
7. In case of an implanted counter electrode, burr a third hole located in the right posterior parietal bone for future fixation of the tunneled cable.
8. Place the plastic screws in the burr holes and screw until the first friction is felt. Then perform three additional 180 ° screw turns. Check with forceps for stability of the screw and add one more turn if not tight enough.
   NOTE: For adult rats this will ensure epidural placement of the screws without damaging the dura or brain (depending on the screw thread design, the turn number might vary) . The use of stainless steel screws should also be feasible, since even at DCS current densities above the neurodegeneration threshold, screw placement did not perturb lesion location or extent below the screws.
9. Turn on the soldering iron and pre-heat for approximately 5 min. Wind the cable exiting the tissue tunnel occipitally around the right parietal screw and then cut it, leaving approximately 1 cm cable behind the winding. Carefully strip the insulation at the end of the cable with a scalpel.
10. Fix the winded cable to the screw and bone with cyanoacrylic glue.
11. Apply a small amount of the lead-free tin-solder to the connector and to the bare wires of the counter electrode cable and connect both by briefly pressing both pre-soldered parts together while touching the soldering tip until the tin-solder melts (about 2-3 s). Remove the soldering tip immediately to avoid excessive metal heating of the cable with subsequent tissue damage.
12. Pick up the custom made tES electrode socket (Figure 1B, in red) with bent, serrated tip forceps and apply a thin layer of cyanoacrylic glue to the bottom rim of the socket. For placement above the motor cortex and using a 4 mm diameter socket, place the mid socket point at 2 mm anterior and 2 mm lateral from the bregma. For this position, the inner medial border of the socket should end directly at the sagittal suture and the caudal border should end at the height of bregma. Press the socket briefly onto the bone (most cyanoacrylic glues harden by pressure).
    NOTE: Placing a light source directly above the socket can ease placing the socket.
13. Ensure that the bone within the area of the socket is free of glue (by checking with light because the glue is reflective). In the case of glue spill, remove the socket, scrape the glue with the scalpel, and repeat step 4.12.
14. After the socket is in place and the future stimulation area is free of glue, first seal the lateral border of the socket to the neighboring tissue with a small drop of cyanoacrylic glue to avoid a fluid bridge that could lead to shunting of current at this location. Do not apply too much glue as it may flow into the stimulation area (if this occurs, return to step 4.12).
    NOTE: Keeping the stimulation area free of glue is crucial as a reduction of the stimulation area might dramatically increase current density (A/m²).
15. Cover all screws with cyanoacrylic glue.
16. Mix the two-component dental acrylic cement in a small silicon tube or glass. As soon as it becomes viscous, apply it with a dental spatula to seal the remaining borders of the socket to the bone. Avoid any flow of dental acrylic cement into the stimulation area.
17. Finally cover the whole skull, screws, counter electrode cable and the socket up to ⅓ of the socket with dental acrylic cement. Ensure that the cement has the correct viscosity: if too fluid, it will flow into the surrounding tissue; if too hard it is difficult to distribute it evenly.
18. When all the bone is covered and the cement is hardened, remove the bulldog clamps; the skin should just touch the built-up cement so that suturing is not needed. (If the initial cut was chosen too long and the connective tissue or muscle is visible, apply a suture as described in step 3.12).
19. Apply one layer of iodine with a cotton swab around the border of the cut skin and subcutaneously inject carprofen (5 mg/kg body weight dissolved in 5-7.5 mL of 0.9% saline for pain treatment and fluid replacement).
    NOTE: If using inhalation anesthesia, turn it off now.
20. Place the rodent in a warming box for recovery from anesthesia until the rodent is awake and postural stability is restored.
    NOTE: Check the animal's weight development, wound state, and general well-being criteria daily according to the institution's recommendation.

5. Transcranial Electrical Stimulation Procedure

NOTE: As anesthesia affects tES effects, performing the stimulation in alert rodents whenever possible is recommended. Allow the rodent to recover for at least 5 days (healing of the head and chest wound) before starting experiments. Experiments can be performed at earlier time points after surgery when using an external counter electrode fixed with a vest, as the chest wound is most irritable; but animals need to be habituated to the electrode vest for several days and interference with behavioral tasks might occur.

1. Fill the tES electrode socket half with 0.9% saline and remove air bubbles.
2. Before cathodal tDCS sessions, always check chlorination, and if needed (such as a shiny silver surface), re-chlorinate the Ag/AgCl electrode. Before anodal tDCS sessions, remove possible excess AgCl deposits from previous stimulations with sandpaper to allow for good conductivity during stimulation. Screw in the tES electrode screw cap (Figure 1B, grey piece).
   CAUTION: Failure to re-chlorinate the electrode between cathodal tDCS sessions will lead to exhaustion of chlorination during stimulation and to toxic build-up by electrochemical reaction. This will induce tissue damage. Re-chlorination is not needed within a single session if stimulation duration is shorter than 20 min.
3. Connect the cables to the two connectors on the head (for anodal stimulation, the anodal cable is connected to the connector on the screw cap, for cathodal stimulation, it is opposite).
   NOTE: When using an externally placed counter electrode, cover the counter electrode with conductive gel and place on the rodent's chest. This is easiest if the electrode is pre-fixed in a small rodent vest, which the rodent can wear during stimulation.
4. Place the rodent into the experimental cage, with the cables connected to a swivel above the cage that allows for free movement.
5. Turn on the stimulator and adjust the stimulation parameters (stimulation intensity, duration, ramp up and down time).
6. When not using a commercially available stimulation device with a safety shut down and disconnection alarm, include a meter in the circuit to check the constant current flow.
   NOTE: With this set-up, stimulation can be applied during performance or training of behavioral tasks.
7. Check for signs of stress or discomfort of the rodent during stimulation.
8. After the end of the stimulation, disconnect the cables, unscrew the electrode cap on the head, and clean and dry the socket with a cotton swab. Return the rodent to the home environment or proceed with a behavioral procedure if desired.