Placement of Extracranial Stimulating Electrodes and Measurement of Cerebral Blood Flow and Intracranial Electrical Fields in Anesthetized Mice

All the animal procedures were approved by the Institutional Animal Care and Use Committee at Duke University or the equivalent local authority regulating research involving animals. See the Table of Materials for details about all the materials, instruments, and equipment used in this protocol.

1. Instrument preparation

1. Make sure that all the required items and surgical instruments are in place (Figure 2): scalp cleaning solution (alcohol pads), tape, forceps, scissors, and a drill for placing the small (0.5 mm) burr holes.
2. Prepare the extracranial surface electrodes for skull application, and make sure any surgical glue has been cleaned from them if they have been used previously.
3. Verify the impedance of these tACS electrodes directly before applying them to the skull. For this, use the built-in measurement function of the tACS stimulator with both electrodes placed in a saline bath.
   NOTE: The preferred impedance is <5 KΩ per electrode pair to allow sufficient current to be passed across the skull. The stimulator device checks the impedance prior to delivering constant-current pulses and gives the value directly.

Figure 2: A photograph of the required instrumentation, including dissecting instruments and scissors, for preparing the extracranial stimulation. 1. Micro dissecting scissors, 11.5 cm; 2. Forceps, 11.5 cm, slight curve, serrated; 3. Dumont #7 forceps, curved; 4. Dumont #5 forceps; 5. Micro curette, 13 cm; 6. Q-tips; 7. Surgical tape; 8. Alcohol pads. Please click here to view a larger version of this figure.

2. Preparation of the animal for surgery

NOTE: For these experiments, we used 14 C57BL/6 control mice between 12 weeks and 33 weeks of age, of which five were male and nine were female.

1. Anesthetize the animal in an induction chamber with isoflurane in 30% O₂ at ~1.5 L/min, with ~4% initially to induce and ~1.25%-1.5% to maintain at a level of anesthesia with spontaneous breathing and sufficient to eliminate the tail pinch response.
2. Transfer the animal to the stereotaxic frame after induction, and then secure the head into the nose cone and ear bars for the subsequent electrode application and burr hole procedure (Figure 1 and Figure 3).
3. Connect the nose cone of the stereotaxic frame to the vaporizer via an inlet and to an outlet to remove any isoflurane residual through a scavenger system (e.g., charcoal or a vacuum). Make sure there are no air leaks from the nose cone, both to maintain the anesthesia level with the isofluorane and to prevent accidental leakage into the room air (Figure 3).
4. Check the position of the mouse in the stereotaxic frame, including the position of the nose cone, to allow spontaneous respiration without intubation, as well as appropriate anesthesia recovery and scavenging to protect the research personnel (Figure 3).
5. Place the probes for measuring the pulse, pulse oxygen saturation (pulse OX), blood pressure, and temperature on the animal; ensure that the minimum pulse oxygenation is 90% and the pulse is >450/min (the alarm lower limit is shown as 380 pulses/min). Record these parameters during the procedure at regular intervals or continuously, depending on the recording system (Figure 3).
6. Prior to starting the procedure, check the level of sedation of the animal using (for example) a toe pinch to check the reflexes. If there is no reflex, then the level of sedation is optimal, as long as the animal maintains spontaneous respiration and adequate pulse oxygenation. If there is a reflex, increase the delivery of isoflurane to deepen the level of the anesthesia, and then recheck the reflex. Continuously observe and monitor the respiratory frequency of the animal, and adjust the isoflurane delivery accordingly.
7. Shave the scalp hair or remove the hair with depilatory cream (clean the residual cream with alcohol pad passages).
8. Apply eye ointment, and then aseptically clean the scalp with three passages of iodine and alcohol prior to excision using scissors.

Figure 3: An image of the animal in the stereotactic frame, with the skull exposed and only the tACS stimulator electrodes in place (prior to the burr hole placement). Note the blood pressure device around the tail and the pulse oximeter on the paw, with the reading on the left. There are scavenging tubes for the isoflurane around the nose cone. Abbreviation: tACS = transcranial alternating current stimulation. Please click here to view a larger version of this figure.

3. Surgical procedure: Applying the stimulating electrodes and making the burr holes

1. For a terminal study, remove the scalp using surgical scissors, and expose the skull ~3 mm from the lambdoid suture caudally and ~3 mm frontal to the bregma to expose part of the posterior frontal suture. Excise the scalp parietally to expose the initial part of the temporal muscle on both sides (Figure 3).
2. Remove any residual subcutaneous connective tissues so that the skull is clean and dry for the application of the stimulating electrodes.
3. Apply conductive gel or paste to the side of the electrodes that will be in contact with the skull, and secure the electrodes with surgical superglue around the edge at intermittent spots.
   NOTE: Do not allow the conducting gel to interfere with the surgical superglue in order to allow a better bond to the skull surface. The outer surface of the electrodes can also be insulated (from the scalp if this is closed during a survival surgery) using surgical superglue.
4. Use either commercial flat electrodes, or create in-house electrodes using insulated wire with a diameter of 100 µm (soldered to the plate) and a 1 mm x 3 mm flexible, insulated (on one surface) copper plate cut according to the size of the skull.
5. Apply lidocaine paste on the temporal muscle and scalp on both sides without disturbing the electrodes to reduce the muscle and peripheral nerve activation.
6. Once the extracranial stimulating electrodes are in place 4 mm laterally to each side of the skull (between the bregma and lambda), drill two 0.5 mm burr holes for the glass electrodes 2 mm on each side of the midline, 4 mm apart from each other, orthogonal to the sagittal suture (Figure 1). Fill these burr holes with sterile mineral oil to prevent current ingress into the skull from the extracranial electrodes.
7. If desired for a particular experiment to induce spreading depression (i.e., potassium-induced spreading depression [K+-SD]), add, on the right side of the skull, a third 0.5 mm burr hole ~1.5 mm rostral to the coronal suture and ~1 mm lateral to the posterior frontal suture. Fill this burr hole with saline for later 1 M KCl application to induce K+-SD.
8. Test the impedance of the extracranial stimulating electrodes both prior to the burr hole placement (and compared to the same electrodes placed in a saline bath) and after the burr hole placement to verify that the burr holes do not interfere with the current flow into the brain (i.e., ensure that the resistance is unchanged).
   NOTE: The impedance measurement is provided directly by the stimulating device. Generally, we have found the overall system impedance (i.e., from the extracranial electrodes across the skull/brain pathway, typically ~3 KΩ) to be relatively constant regardless of the burr holes and glass microelectrodes, indicating that there is minimal current leakage directly into the brain through the burr holes.
9. Place the chronic transcranial stimulation electrodes for chronic stimulation in a similar fashion. In this case, insulate the outer surface of the electrodes, close the scalp, and either tunnel the insulated wires out through the scalp or route them into a fixed head stage mounted to the skull.

4. Physiological procedure

1. Commence with the physiological aspects of the experiment, once the animal is fully prepared for the non-survival, physiological experiment. Maintain the anesthesia level sufficient for both spontaneous respiration and adequate pulse Ox, respiratory, and pulse levels.
2. Measure the CBF resulting from the extracranial stimulation by one of the following two methods.
   1. Place the mouse under a laser speckle imaging device with or without intracranial recording electrodes to measure the intracranial electrical field during stimulation episodes (Figure 3).
   2. Transfer the animal to a physiological preparation for the placement of bilateral laser Doppler probes and intracranial electrodes to measure the intracranial electrical field during stimulation episodes (Figure 1).

5. Placement of bilateral laser Doppler and glass electrodes

1. Transfer the animal to a microscope stage for the application of bilateral laser Doppler probes. Place the probes on the top of the skull surface between the bilateral burr holes and the coronal suture (Figure 1).
2. Fill pulled glass microelectrodes (~0.1 µM, 2-6 MΩ impedance) with 0.2 M NaCl, and place them using a micromanipulator into the two burr holes placed laterally to the sagittal suture^3,14 (Figure 1).
   NOTE: These burr holes are between the two symmetrical extracranial stimulation electrodes (Figure 1).
3. Once inserted into the brain, ensure these glass microelectrodes are ~1 mm within the cerebral cortex. Perform depth profiles at various symmetrical depths. Refill the burr holes with sterile mineral oil to insulate this pathway for current flow.

6. Stimulation procedure and measurement of the intensity of the transcranial alternating current stimulation (tACS) or transcranial direct current stimulation (tDCS)

1. Record continuous data from the dual laser Doppler probes on the skull and the two intracranial microelectrode outputs (recorded using a DC amplifier with headstages) using a digitizing system and software with at least four channels (at a sampling rate of 1 KHz). Once all the values have been recorded over a sufficiently stable baseline duration (i.e., >10 min), test the extracranial stimulation.
   NOTE: Figure 4 shows an example of the four channels with the two intracranial recording electrodes in the upper channels and the CBF response in the lower channels.
2. Apply brief periods of on/off stimulation at various amplitudes (i.e., 20-30 s, 0.5-2.0 mA, in the tolerable range) to obtain a clear baseline before and after stimulation (Figure 4). Apply the stimulation between the two skull tACS electrodes on either side (Figure 1) using a commercial, human-compatible stimulating device that delivers a constant current.
3. Closely observe the mouse for muscle twitches or other responses to the tACS, such as a change in the pulse or respiration, to create an upper limit of tolerability (generally ~2 mA).
4. Continue to monitor the impedance across the electrodes with stimulation epochs to ensure this is constant.
5. Add a small amount (2-3 µL) of 1 M KCl to the anterior burr hole¹⁴ to induce spontaneous K+-SD events. These generate a large CBF response and interactions between the K+-SD-induced CBF response and the CBF response. Estimate the tACS CBF response, applying the tACS stimulation both before and after the occurrence of the SD.
6. At the end of the experiment, perform euthanasia through an overdose of isoflurane (5%) and then decapitate once the respirations and heartbeats have ceased.

Figure 4: Data showing four channels of raw data in response to low-intensity tACS. The data are arranged with the upper two rows as the intracranial, direct DC electrical recordings (labeled as Input 1 [IN0] and input 2 [IN1]) and the lower two rows as the bilateral laser Doppler recordings of the cerebral blood flow. Note that the responses are asymmetrical between the right (upper) and left (lower) electrical and cerebral blood flow traces. (A) A small response (16% increase in blood flow) in response to a 1.2 mV/mm 20 s stimulus (0.75 mA). (B) A larger response (21% increase in blood flow) in response to a 1.4 mV/mm stimulus (1.0 mA). Abbreviation: tACS = transcranial alternating current stimulation. Please click here to view a larger version of this figure.

7. Calculation of the electrical field

1. Measure the difference in the output from the two intracranial electrodes using the difference in the half-wave (one cycle) of the two sine waves recorded (the upper two traces in Figure 4). Divide this difference (mV) by the distance between the two electrodes (mm, here ~4 mm but directly measured in each case) to arrive at the field strength (mV/mm)^3,6.