All animal experimentation was performed according to the guidelines of the local and institutional Council on Animal Care (University of Bonn, BfArM, LANUV, Germany). In addition, all animal experimentation was carried out in accordance with superior legislation, e.g., the European Communities Council Directive of 24 November 1986 (86/609/EEC), or individual regional or national legislation. Specific effort was made to minimize the number of animals used, as well as their suffering.

1. Animal Housing and EEG Recording Conditions

1. House mice in filter-top cages or use individually-ventilated cages.
2. Transfer mice from the animal facility to ventilated cabinets in special lab rooms that are appropriate for implanted animals and telemetric recordings.
3. Perform all animal experimentation, including EEG electrode implantation and the subsequent recordings, under standardized conditions (22 °C temperature, 12 h/12 h light-dark cycle, 50-60% relative humidity, noise attenuation, etc.)¹⁸.
4. Prior to radiofrequency transmitter implantation, house animals in groups of 3 – 4 in clear type II polycarbonate cages, with food and water ad libitum. Do not isolate individual mice, as this can cause stress and interfere with subsequent experimentation and results.
5. Do not use open housing conditions, but use ventilated cabinets instead during experimentation and recording.

2. Radiotelemetric EEG Electrode Implantation and EEG Recordings

1. Anesthetize the mice using injection narcotics, e.g., ketaminehydrochloride/xylazinehydrochloride (100/10 mg/kg, intraperitoneally, i.p.) or inhalation narcotics, e.g., isoflurane^17,18.
   1. For isoflurane narcosis, position the mouse into an induction chamber with 4-5% isoflurane and 0.8-1% oxygen or carbogen (5% CO₂ and 95% O₂).
   2. Place a silicon facemask on the nose/mouth of the animal to control the desired depth of anesthesia and avoid experimenter exposure to isoflurane using a scavenging system.
   3. Use injectable anesthetics, e.g., esketaminhydrochloride (100 mg/kg, i.p.) and xylazinehydrochloride (10 mg/kg, i.p.), if inhalation anesthesia is not available.
   4. Monitor the depth of anesthesia by checking the tail pinch reflex, foot pinch reflex, and respiration rate. Note that artificial respiration via tracheal intubation is not necessary in mice.
2. Implant the radiofrequency transmitter into a subcutaneous pouch on the back of the animal.
   1. Remove the body hair from the scalp and pretreat the shaved scalp with two disinfectants, i.e., 70% ethanol and an iodine-based scrub.
   2. Using a scalpel, make a midline incision on the scalp from the forehead to the nucheal region.
   3. Starting from the nuchal incision, prepare a subcutaneous pouch on one side of the back of the animal by performing a blunt dissection using surgical scissors or a surgical probe.
   4. Insert the transmitter into the subcutaneous pouch and deposit the excess length of the flexible transmitter leads into the pouch as well. Pay special attention to preventing contamination of the surgical site and transmitter implant. Properly isolate sterile from non-sterile areas using drapes.
3. Place the experimental animal on the stereotaxic frame, e.g., a computerized 3D stereotaxic device. Fix the skull using a nose clamp and ear bars.
4. Pretreat the skull with 0.3% H₂O₂ to remove further tissue from the skull and to illuminate the cranial sutures and craniometrics landmarks, bregma and lambda.
5. Drill holes at the coordinates of choice (see step 2.6) using a high-speed neurosurgical drill in a pressure-free mode at maximum velocity.
   NOTE: Pressure-free drilling avoids the sudden breakthrough of the drill head and damage to the cortex. For a craniotomy, a neurosurgical high-speed drill is recommended. Choose standard drill-head diameters of 0.3 – 0.5 mm, depending on the electrode diameter.
6. Carefully select the electrode type, considering the impedance, diameter, coating, etc.
   NOTE: Parylene-coated tungsten or stainless steel electrodes are the most common. The electrode types should be chosen according to experimental requirements. As a preemptive maneuver, sterilize electrode tips prior to implantation using 70 % ethanol.  Note that the electrode coating does not allow for heat sterilization.
7. For intrahippocampal CA1 EEG recordings, drill a hole stereotaxically at the following coordinates: bregma, -2 mm; mediolateral, 1.5 mm (right hemisphere); dorsoventral, 1.3 mm (target region: cornu ammonis (CA1) pyramidal layer). For the reference electrode, drill a hole above the cerebellar cortex at the following stereotaxic coordinates: bregma, -6.2 mm, mediolateral, 0 mm; dorsoventral, 0 mm.
   NOTE: The cerebellar electrode serves as a pseudoreference electrode, as the cerebellum is a rather silent brain region. The stereotaxic coordinates were derived from a standard mouse brain atlas.
8. Prior to the insertion of the electrodes, shorten them to the required length. Mechanically clip the extracranial part of the electrode to the stainless steel helix of the transmitter.
   NOTE: Soldering should be avoided, as this can introduce substantial noise in the system.
9. Attach the electrode to the vertical arm of the stereotaxic device and insert the electrode according to the aforementioned stereotaxic coordinates.
10. Fix the electrodes using glass ionomer cement and wait until the cement has fully hardened.
11. Close the scalp using over-and-over sutures with non-absorbable 5-0 or 6-0 suture material.
12. For postoperative pain management, administer carprofen (5 mg/kg, subcutaneously, s.c.) once a day for 4 consecutive days post-implantation. Note that carprofen should be injected prior to the initial incision (step 2.2.2).
13. Allow the animals to recover for 10 – 14 days post-implantation before the recordings and/or injection experiments are started.

3. Spontaneous Recordings of Theta Oscillations and Pharmacological Induction

1. Activate the radiofrequency transmitter using the magnetic switch. Place the animal with its home cage on the receiver plate. Perform long-term hippocampal EEG recordings for at least 24-48 h.
   NOTE: The analysis of EEG amplitude and EEG frequency characteristics of long-term recordings provides detailed insight into the circadian dependency of theta oscillations and their association with specific behavioral and cognitive conditions/tasks. Always combine EEG recordings with video monitoring of the experimental animals.
2. For pharmacological induction of theta oscillations, administer a single dose of urethane (800 mg/kg i.p.) or a single dose of a muscarinic receptor agonists, e.g. pilocarpine (10 mg/kg i.p.), arecoline (0.3 mg/kg i.p.) or oxotremorine (0.03 mg/kg i.p.). Pretreat mice with N-methylscopolamine (0.5 mg/kg i.p.) to avoid peripheral muscarinic reactions. Freshly dissolve all drugs in 0.9% NaCl or Ringer solution. 
   NOTE: Higher dosages of muscarinic receptor agonists might result in seizure induction in experimental animals. Also consider that the dosages given here represent landmarks that require prior dose-effect studies in the mouse line under investigation. Note that urethane is a mutagen and carcinogen that requires appropriate precautions in storage and handling.
3. Inject atropine (50 mg/kg, i.p.) to differentiate atropine-sensitive type II from atropine-insensitive type I theta oscillations.
   NOTE: The atropine dosage is again species- and strain-dependent and requires prior dose-effect evaluation. The optimal time point of atropine injection depends on the pharmacodynamics of muscarinic receptor agonists. For identification of type II theta, atropine injection is recommended 1 h after urethane administration.
4. Try to avoid the subsequent application of several drugs, as systemic drug administration alters global transcriptional and translational patterns, which influence subsequent EEG recordings. Note that short-lasting theta oscillations can also be induced by sensory stimuli, such as tail- or paw-pinching.
5. Extract/export representative EEG data sets of the pre-phase (baseline) and the post-injection phase from the total EEG recording as ASCI or TXT files, considering the pharmacokinetics of the drugs applied and the demands of the individual study protocol.

4. Validation of EEG Electrode Placement

1. Euthanize animals by placing them in an incubation chamber and introduce 100% carbon dioxide. Use a fill rate of 10-30% of the chamber volume per min with carbon dioxide added to the existing air in the incubation chamber; this will result in rapid unconsciousness with minimal stress to the animals.
2. Remove the mouse from the chamber once respiratory arrest occurs and a faded eye color persists for 2-3 min.
3. Cut the stainless steel electrodes and explant the radiofrequency transmitter. Decapitate the mouse using scissors or a guillotine and remove the brain from the neurocranium by gentle manipulation with surgical scissors and forceps.
4. Fix the brains in 4% paraformaldehyde in phosphate buffer saline (PBS) (pH 7.4) overnight. For cryoprotection, transfer the brains to 30% glucose and store them at 4 °C until further processing.
   NOTE: Paraformaldehyde is considered hazardous by the 2012 OSHA Hazard Communication Standard (29 CFR 1910.1200). Take the necessary precautions: use personal protective equipment, ensure appropriate ventilation, and avoid dust formation. In addition, remove sources of ignition and take precautionary measures against static discharges. Paraformaldehyde should not be released into the environment.
5. Mount the extracted brain on the tissue holder of a cryostat using an adhesive and cut the brains into 40 – 75 µm coronal slices. Mount the slices onto glass slides and stain them with Nissl blue using standard histological procedure; this procedure will visualize the branch channel that reflects the previous electrode position. Note that it is also possible to cut coronal slices from the native brain using a vibroslicer
6. Incorporate only those animals that meet correct EEG electrode placement criteria; for the CA1 region, the tip of the deep electrode should be localized inside the CA1 pyramidal layer.

5. Data Acquisition

1. Record CA1 intrahippocampal EEGs with an appropriate sampling rate with no a priori filter cut-off.
   NOTE: The sampling rate, which is transmitter-specific, determines the upper frequency limit of the EEG analysis.
2. Process the recorded data with an analysis software. Program time-frequency analyses and calculations with custom-made procedures for the adequate control of analysis methods (Figure 5)²⁰.
3. Cut the recorded EEG into sections with a length of 1 h each. Use fast computer processors, since computation time is high. Additionally, make use of software that can parallelize computing on multiple kernels²⁰.

6. EEG Data Analysis

1. Analyze data segments using a complex Morlet wavelet to calculate both the frequency and the amplitude of oscillations.
   NOTE: This wavelet (e.g.,Ψ(x)=(πb)^(−1/2) exp(2iπcx) exp(−x²/b), where b is the bandwidth parameter, c the center frequency, and i the imaginary unit) has often been applied in literature to study EEG data, as it guarantees optimal resolution in both frequency and time^23,24.
2. Use a bandwidth parameter and center frequency setting that particularly weights the frequency resolution to distinguish the frequency differences on the 0.1-Hz level while still not neglecting a sufficient time resolution.
   NOTE: Neuronal processes in the gamma band are short-lasting²⁵, and this can also hold true for theta rhythms. Thus, the analytical approach must consider adequate temporal resolution.
3. Analyze the EEG data in the frequency range of 0.2 – 12 Hz, with a step-size of 0.1 Hz, thus including the typical delta, theta, and alpha frequency ranges.
4. Set up an automated, complex analytical tool for the elaboration of the theta frequency architecture, which can replace a standard visual inspection of theta oscillations; this procedure is called the theta detection method (TDM).
5. Compute the quotient of the maximum amplitude in the theta frequency range (3.5-8.5 Hz) and the maximum amplitude in the upper delta frequency range (2-3.4 Hz) for time windows of 2.5 s each.
   NOTE: The value of this quotient serves as a measure to decide if a theta oscillation occurred. The definition of the theta frequency range can vary depending on the functional state and neuroanatomical circuitry/system to be analyzed.
6. Classify a segment as a "theta oscillatory epoch," if the computed ratio during this segment is above 1.5.
   NOTE: This guarantees that the maximum theta amplitude is at least 50% higher than the amplitude in the upper delta band during the related 2.5-s EEG segment. Note that the ratio might need adaptation depending on the line and/or species used. An interval of 2.5 s represents a minimal duration for a theta oscillation, prevents false-positive detections of certain noisy epochs, and lies within the range definitions of other publications^19,26. The upper delta frequency range serves as a control frequency band because physiologically-relevant delta activity appears during non-theta epochs, for example, during slow-wave sleep, which is highly dampened during theta activity.
7. Repeat this procedure for every 1 h section; therefore, every section consists of 1,440 EEG segments with lengths of 2.5 s each.
8. Statistically evaluate the data of the identified theta oscillation epochs.
9. Compute the statistics of the total duration times of detected theta epochs; distinct or predefined groups; cycles, such as light/dark; and others parameters.
   NOTE: Statistics may include t-test, ANOVA, or MANOVA, depending on the variable, the number of groups, the conditions, etc.
10. Calculate the statistics of the amplitude of the detected theta epochs, but only in the theta frequency range (3.5 – 8.5 Hz).
11. Evaluate the statistics of the frequency of the detected theta epochs, but only in the theta frequency range (3.5 – 8.5 Hz).
    NOTE: The theta frequency of a theta epoch is defined as the frequency belonging to the maximum theta amplitude of a theta epoch.