All animal procedures were approved by the Institute of Biomedical Sciences Animals Ethics Committee at the University of São Paulo and were performed according to the ethical guidelines adopted by the Brazilian College of Animal Experimentation.

1. Preparation of solutions

1. Preparation of internal solution
   NOTE: The internal solution fills the patch-clamp micropipette and will contact the cell's interior (see an example in Figure 2). Internal solutions may vary depending on the type of activity to be measured³³.
   1. Choose the internal solution considering its experimental purpose and the appropriate record type. To record membrane potential in the current-clamp mode, use the internal solution composed of 120 mM K-gluconate, 1 mM NaCl, 5 mM EGTA, 10 mM HEPES, 1 mM MgCl₂, 1 mM CaCl₂, 3 mM KOH, 10 mM KCl, and 4 mM (Mg)-ATP. Weigh all the salts according to the desired final volume, as given in Table 1.
   2. Use enough deionized water to reach 90% of the final volume of the solution. This volume will ensure enough space to adjust the pH and osmolarity.
   3. After adding and mixing all the ingredients thoroughly, adjust the pH to 7.2-7.3 with 5 M KOH using a pH meter. Use an osmometer to check the osmolarity, which should be around 275-280 mOsm (adjust, if necessary, down only).
   4. Prepare the internal solution in advance and store at 8 °C. Add the ATP on the day of the experiment. Store the ATP-containing internal solution at -20 °C (1 mL aliquots) for 3-4 months.
2. Preparation of slicing solution
   1. Prepare slicing solution containing 238 mM sucrose, 2.5 mM KCl, 26 mM NaHCO₃, 1.0 mM NaH₂PO₄, 10 mM glucose, 1.0 mM CaCl_2, and 5.0 mM MgCl₂. Weigh all the salts according to the desired final volume, as shown in Table 2. The exact volume of this solution to be prepared will depend on the cutting chamber's size.
   2. While mixing all the salts in deionized water, constantly saturate with carbogen (95% O₂ and 5% CO₂). Attach polyethylene tubing (1.57 mm outer diameter [OD] x 1.14 mm inner diameter [ID]) to a carbogen cylinder to supply carbogen to the slicing solution. Hold the open end of the tube inside the beaker containing the slicing solution.
   3. After mixing all the ingredients well, adjust the pH to 7.3 with 10% nitric acid using a pH meter. Use an osmometer to check the osmolarity, which should be 290-295 mOsm (adjust, if necessary, down only). Afterward, cool the solution to 0-2 °C.
3. Prepare aCSF solution for recordings
   1. Prepare artificial cerebrospinal spinal fluid (aCSF) solution for recordings containing 124 mM NaCl, 2.8 mM KCl, 26 mM NaHCO₃, 1.25 mM NaH₂PO₄, 1.2 mM MgSO₄, 5 mM glucose, and 2.5 mM CaCl₂. Weigh all the salts according to the desired final volume, given in Table 3.
   2. While mixing all the salts in deionized water, constantly saturate with carbogen (95% O₂ and 5% CO₂), as described in step 1.2.2.
   3. After adding and mixing all the ingredients, adjust the pH to 7.3 (10% nitric acid can be used) using a pH meter. Use an osmometer to check the osmolarity, which should be 290-300 mOsm. Afterward, pour the aCSF in a beaker and maintain in a water bath at 30 °C.

2. Brain dissection and slicing

NOTE: Since different brain structures may require cutting in different planes (coronal, sagittal, or horizontal slices), the exact approach for obtaining the slices depends on the brain region of interest. Typically, to study the Kiss1-expressing cells in the AVPV/PeN and ARH (here denominated as AVPV/PeN^Kisspeptin neurons and ARH^kisspeptin neurons; Figure 2A,B), coronal brain slices (200-300 µm) are usually made^{17,19,20,21,34}. The AVPV/PeN^Kisspeptin neurons are located approximately 0.5 to -0.22 mm from the bregma, whereas ARH^kisspeptin neurons are at -1.22 to -2.70 mm. Nuclei location can be determined by using a stereotaxic mouse brain atlas³⁵ or the Allen Mouse Brain Reference Atlas (http://mouse.brain-map.org/). Adult Kiss1-Cre/GFP female (diestrus-stage) and male mice³⁶ were used in this study.

1. Brain removal
   1. Prepare the dissection site and the tools needed to extract the brain: decapitation scissors, iris scissors, Castroviejo curved scissors, osteotome, tweezers, spatula, filter paper, Petri dish, razor blade, and cyanoacrylate glue.
   2. Before making the slices, prepare the bench on which the dissection will be performed, as all subsequent procedures must be performed quickly. Make sure to have access to a slicing device such as a vibratome.
   3. Prepare a recording chamber to maintain the brain slices before slicing the tissue. Acquire a recording chamber, bath dimensions of 24 mm x 15 mm x 2 mm (L x W x H), from a commercial brand (Table of Materials) or make one in-house.
      NOTE: An in-house recovery chamber can be fabricated as follows: cut a 24-well plate so that nine wells are available. Glue a nylon screen to the base of the nine wells. With the remainder of the well plate, make a base so that the lower part of the nylon is free. This adapted base can be placed inside a 500 mL beaker containing the aCSF (Supplementary Figure 1).
   4. After preparing everything, anesthetize the animal with inhaled anesthetic using 4%-5% isoflurane. The anesthesia must be in accordance with a protocol approved by the ethics committee. Shortly after the animal is immobile, perform tail and paw pinch tests to ensure that the animal is deeply anesthetized.
   5. Quickly decapitate the mice while the heart is still active to augment cell viability. Harvest the brain quickly after decapitation.
   6. With surgical scissors, make an incision in the skin at the top of the animal's skull, from caudal to rostral, and remove the scalp from the animal's head. Next, cut the interparietal plate along the sagittal suture with the iris scissors and remove the occipital bone. Slide the osteotome under the parietal bone and gently pull it out until the brain is exposed.
   7. After exposing the brain, turn the head upside down, gently lower the brain to visualize the trigeminal nerve on each side, then cut the trigeminal nerve using Castroviejo curved scissors. After visualizing the hypothalamus, identify the optic nerve and cut it gently.
      NOTE: Be careful when cutting the optic nerve, as pulling it will tear the adjacent hypothalamic area containing the AVPV/PeN nucleus.
   8. Cut the most anterior portion of the frontal lobe and remove the brain completely. Immediately immerse the brain in the slicing solution until acquiring the slices.
2. Slicing brain samples
   1. Place the brain on filter paper (supported on a Petri dish) to dry the excess solution. Then, with a sharp cutting razor blade, perform a coronal cut separating the brainstem with the cerebellum from the rest of the tissue.
   2. Next, glue the caudal portion of the brain to the base of the vibratome and fill the chamber of the slicing device with the slicing/aCSF solution cooled to 0-2 °C. During the procedure, pack dry ice around the vibratome chamber to keep the slicing solution cold.
   3. Insert the razor blade into the vibratome and set the device's appropriate cutting parameters: speed = 3, frequency = 9, and feed = 250 μm. Use an acrylic transfer pipette (inverted Pasteur glass pipette attached to a silicone teat) to transfer the slices to the recovery chamber (described in step 2.1.3) during the tissue-slicing procedure. Wait 60 min for tissue recovery after slice acquisition.

3. Cell sealing for recording

1. Ensure that all the pieces of equipment (microscope, amplifier, digitizer, micromanipulator, and others) are turned on before starting the recording.
2. Fill the recording chamber, from a commercial brand (Table of Materials), attached to the microscope with the aCSF solution for recordings. Use a perfusion pump to constantly perfuse the aCSF at a rate of 2 mL/min.
3. Transfer a brain slice of interest (one at a time) to the recording chamber. Use an acrylic transfer pipette (inverted Pasteur glass pipette attached to a silicone teat) to transfer the hypothalamic slices to the chamber. Use a slice anchor (Table of Materials) to hold the slice so it does not move during the aCSF perfusion.
4. Place a slice at the center of the recording chamber attached to the microscope. The slice position is critical to allow a good view of the desired region under the microscope and for a perfect reach of the recording micropipette.
5. Use an immersion microscope's low-power objective lens (10x or 20x) to assist in positioning the slice and locating the region of interest.
6. After locating the region of interest, switch the objective lens to the high-power lens (63x) and focus on the tissue level, observing the endogenous fluorescent protein and shapes of the cells in the target region to locate the kisspeptin cells on the surface of the brain slice²⁰.
7. When a possible target cell is located, mark it on the computer screen with the mouse cursor or by drawing a format, like a square, over the area of interest. The computer screen mark helps guide the recording micropipette's position to the cell.
8. After determining the exact location of the target cell, lift the objective and introduce the recording micropipette filled with the internal solution. When placing the micropipette in the electrode holder, ensure that the internal solution is in contact with the silver electrode.
   NOTE: For micropipette preparation, placement, and positioning on the electrode holder, please refer to³³.
9. Apply positive pressure before submerging the micropipette in the aCSF solution, to prevent debris from entering the micropipette, using a 1-3 mL air-filled syringe connected to the micropipette holder through a polyethylene tubing (≈130 cm longer); apply nearly 100-200 μL of air.
10. Using the micromanipulator, guide the micropipette below the center of the objective. Move the buttons on the micromanipulator to guide the micropipette on the X-Y-Z axis toward the cell of interest.
11. Adjust the focus to see the tip of the micropipette and bring the focus closer, but not too close, to the slice. Reduce the speed of the micromanipulator and slowly lower the micropipette to the plane of focus. Ensure that the micropipette tip does not abruptly penetrate the slice, but rather slowly descends until it touches the surface of the cell/target region.
12. Apply light positive pressure (≈100 μL) with the 1-3 mL air-filled syringe attached to the micropipette holder to clear any debris from the approach path.
13. Focus on the target cell and slowly move the micromanipulator on the X-Y-Z axis to bring the micropipette closer to the target cell. When touching the micropipette to the cell, a dimple caused by the pressure applied through the micropipette tip will be observed (Figure 2C).
14. After forming the dimple, due to the micropipette's proximity to the cell, apply weak, brief suction by mouth (1-2 s) through the tube connected to the micropipette holder to generate the seal between the micropipette to the cell (gigaohm seal or gigaseal >1 GΩ; Figure 2D). To form the seal, use the voltage-clamp mode on the software. For seal formation details, please refer to³³.
15. If the seal remains stable (the gigaohm seal should be mechanically stable and without noise interference, determined by observation for about 1 min), set the holding voltage at the closest physiological resting potential of the cell of interest. For kisspeptin hypothalamic neurons, -50 mV is recommended.
16. Apply brief suction by mouth (negative pressure) with the micropipette sealed to the cell to break the plasma membrane (Figure 2E). Adequate whole-cell configuration is achieved when suction is performed with sufficient force so that the ruptured membrane does not clog the micropipette and does not attract a sizable portion of the membrane or even the cell.
17. Check the system settings manual used. Use the software (see Table of Materials) to digitally check and calculate the series resistance (SR) and the whole-cell capacitance (wcc).
18. On voltage-clamp mode, after breaking the cell membrane, enable the whole cell option, and click on the Auto command referring to the whole cell tab. The cell's SR and wcc will be automatically calculated and instantly displayed by the software. These parameters can also be checked by performing the membrane test with the amplifier mentioned in the Table of Materials³³.
19. Make sure to check cell viability parameters. For kisspeptin neurons, check that the electrophysiological measurements are: SR < 25 mΩ, input resistance > 0.3 GΩ, and holding current absolute value < 30 pA (personal observations and reference²¹). The mean value of the wcc of AVPV/PeN^Kisspeptin or ARH^kisspeptin neurons is ≈ 10-12 pF in gonad-intact mice²⁰.
20. Monitor the SR and the cell steady-state capacitance during the experiments. Ensure the SR does not change more than 20% during a recording and that the membrane capacitance is stable.
21. Check the software settings. Create specific protocols for the recordings according to the type of experiment. To record membrane potential in the current-clamp mode, with equipment mentioned in the Table of Materials, low-pass filter the electrophysiological signals at 2-4 kHz and analyze results offline in a software (see the Table of Materials for software information).
22. Once the whole-cell configuration is properly achieved, measure synaptic currents in voltage-clamp mode (Figure 2G). Record changes in resting membrane potential (RMP) and induced RMP variations in the current-clamp mode (Figure 2H). Changes in RMP, such as depolarization of the cell membrane, can be induced by administering a known drug/neurotransmitter to the bath (described in step 3.2), as illustrated in Figure 1.
    NOTE: In current-clamp mode, positive or negative current can be injected to hold the membrane voltage at a desired voltage. For kisspeptin neurons, we usually set zero current injection (I = 0) to record the spontaneous variation of the membrane potential.