Hypothalamic Kisspeptin Neurons as a Target for Whole-Cell Patch-Clamp Recordings

Josiane do Nascimento Silva; Thais T. Zampieri; Henrique R. Vieira; Renata Frazao

doi:10.3791/64989

JoVE Journal > Neuroscience

Neuroscience

Hypothalamic Kisspeptin Neurons as a Target for Whole-Cell Patch-Clamp Recordings

Published: March 17, 2023

doi:

10.3791/64989

Josiane do Nascimento Silva¹, Thais T. Zampieri¹, Henrique R. Vieira¹, Renata Frazao¹

¹Department of Anatomy, Instituto de Ciencias Biomedicas,Universidade de Sao Paulo

Summary

Here, we present a protocol to perform a whole-cell patch-clamp on brain slices containing kisspeptin neurons, the primary modulator of gonadotrophin-releasing hormone (GnRH) cells. By adding knowledge about kisspeptin neuron activity, this electrophysiological tool has served as the basis for significant advancements in the neuroendocrinology field over the last 20 years.

Abstract

Kisspeptins are essential for the maturation of the hypothalamic-pituitary-gonadal (HPG) axis and fertility. Hypothalamic kisspeptin neurons located in the anteroventral periventricular nucleus and rostral periventricular nucleus, as well as the arcuate nucleus of the hypothalamus, project to gonadotrophin-releasing hormone (GnRH) neurons, among other cells. Previous studies have demonstrated that kisspeptin signaling occurs through the Kiss1 receptor (Kiss1r), ultimately exciting GnRH neuron activity. In humans and experimental animal models, kisspeptins are sufficient for inducing GnRH secretion and, consequently, luteinizing hormone (LH) and follicle stimulant hormone (FSH) release. Since kisspeptins play an essential role in reproductive functions, researchers are working to assess how the intrinsic activity of hypothalamic kisspeptin neurons contributes to reproduction-related actions and identify the primary neurotransmitters/neuromodulators capable of changing these properties. The whole-cell patch-clamp technique has become a valuable tool for investigating kisspeptin neuron activity in rodent cells. This experimental technique allows researchers to record and measure spontaneous excitatory and inhibitory ionic currents, resting membrane potential, action potential firing, and other electrophysiological properties of cell membranes. In the present study, crucial aspects of the whole-cell patch-clamp technique, known as electrophysiological measurements that define hypothalamic kisspeptin neurons, and a discussion of relevant issues about the technique, are reviewed.

Introduction

Hodgkin and Huxley made the first intracellular record of an action potential described in several scientific studies. This recording was performed on the squid axon, which has a large diameter (~500 µm), allowing a microelectrode to be placed inside the axon. This work provided great possibilities for scientific research, later culminating in the creation of the voltage-clamp mode, which was used to study the ionic basis of action potential generation¹^,²^,³^,⁴^,⁵^,⁶^,⁷^,⁸. Over the years, the technique has been improved, and it has become widely applied in scientific research⁶^,⁹. The invention of the patch-clamp technique, which took place in the late 1970s through studies initiated by Erwin Neher and Bert Sakmann, allowed researchers to record single ion channels and intracellular membrane potentials or currents in virtually every type of cell using only a single electrode⁹^,¹⁰^,¹¹^,¹². Patch-clamp recordings can be made on a variety of tissue preparations, such as cultured cells or tissue slices, in either voltage-clamp mode (holding the cell membrane at a set voltage allowing the recording of, for example, voltage-dependent currents and synaptic currents) or current-clamp mode (allowing the recording of, for example, changes in resting membrane potential induced by ion currents, action potentials, and postsynaptic potential frequency).

The use of the patch-clamp technique made several notable discoveries possible. Indeed, the seminal findings on the electrophysiological properties of hypothalamic kisspeptin neurons located at the anteroventral periventricular and rostral periventricular nuclei (AVPV/PeN^Kisspeptin), also known as the rostral periventricular area of the third ventricle (RP3V), and the arcuate nucleus of the hypothalamus (ARH^kisspeptin)¹³^,¹⁴^,¹⁵ are of particular interest. In 2010, Ducret et al. performed the first recordings of AVPV/PeN^Kisspeptinneurons in mice using another electrophysiological tool, the loose-cell patch-clamp technique. These studies provided an electrical description of AVPV/PeN^Kisspeptin neurons and demonstrated that their firing patterns are estrous cycle-dependent¹⁶. In 2011, Qiu et al. used the whole cell patch-clamp technique to demonstrate that ARH^kisspeptin neurons express endogenous pacemaker currents¹⁷. Subsequently, Gottsch et al. showed that kisspeptin neurons exhibit spontaneous activity and express both h-type (pacemaker) and T-type calcium currents, suggesting that ARH^kisspeptin neurons share electrophysiological properties with other central nervous system pacemaker neurons¹⁸. Additionally, it has been demonstrated that ARH^kisspeptin neurons exhibit sexually dimorphic firing rates and that AVPV/PeN^Kisspeptin neurons exhibit a bimodal resting membrane potential (RMP) influenced by ATP-sensitive potassium channels (K_ATP)¹⁹^,²⁰. Furthermore, it was established that gonadal steroids positively affect the spontaneous electrical activity of the kisspeptin neurons in mice¹⁹^,²⁰^,²¹. The first works that study kisspeptin neurons' electrophysiological properties are mentioned¹⁶^,¹⁷^,¹⁸^,¹⁹^,²⁰. Since then, many studies have used the whole-cell patch-clamp technique to demonstrate which factors/neuromodulators are sufficient to modulate the electrical activity of kisspeptin neurons (Figure 1)¹⁷^,²¹^,²²^,²³^,²⁴^,²⁵^,²⁶^,²⁷^,²⁸^,²⁹^,³⁰^,³¹^,³².

Given the importance of this technique for the study of neurons that are required for reproduction, among other cell types not covered here, this article describes the basic steps for the development of the whole-cell patch-clamp technique, such as preparing the solutions, dissecting and slicing the brain, and performing the seal of the cell membrane for recordings. Moreover, relevant issues about the technique are discussed, such as its advantages, technical limitations, and important variables that must be controlled for optimal experimental performance.

Protocol

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

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.
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.
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 made¹⁷^,¹⁹^,²⁰^,²¹^,³⁴. The AVPV/PeN^Kisspeptinneurons 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.

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.
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

Ensure that all the pieces of equipment (microscope, amplifier, digitizer, micromanipulator, and others) are turned on before starting the recording.
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.
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.
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.
Use an immersion microscope's low-power objective lens (10x or 20x) to assist in positioning the slice and locating the region of interest.
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.
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.
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.
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.
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.
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.
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.
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).
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.
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.
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.
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).
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.
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.
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.
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).
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.

Representative Results

To study the possible effects of human recombinant growth hormone (hGH) on the activity of hypothalamic kisspeptin neurons, we performed whole-cell patch-clamp recordings in brain slices and assessed whether this hormone causes acute changes in the activity of AVPV/PeN^Kisspeptin and ARH^kisspeptin neurons. Adult Kiss1-Cre/GFP female (diestrus-stage) and male mice³⁶ were used in this study. Gonad-intact animals were selected for the experiments, since the properties of their hypothalamic kisspeptin neurons may vary depending on sex steroid levels¹⁹^,²⁰. While it was beyond the scope of the present study to evaluate differences between sexes, we refer the reader to Croft et al. and Frazão et al.¹⁹^,²⁰ for more information. Genetically modified animals, such as mice and rats, represent exciting tools for this type of experiment, since cells expressing a specific gene or a selective-induced ablation can be identified by a fluorescent protein such as GFP, among others²³^,²⁶^,³⁶. The use of genetically modified mice represents a breakthrough in understanding kisspeptin neuron activity.

Recorded neurons (26 cells out of 12 animals) were determined according to neuroanatomical features³⁵ and the expression of the endogenous GFP²⁰. In current-clamp mode, neurons were recorded under I = 0 in whole-cell patch-clamp configuration. The AVPV/PeN^Kisspeptin neurons (12 cells from nine animals) exhibited an average RMP of -59.0 mV ± 3.0 mV (range: -75 mV to -46 mV), input resistance of 0.9 ± 0.1 GΩ, wcc of 12.3 pF ± 1.6 pF, and SR of 19.4 ± 1.9 mΩ. Among the recorded neurons, three out of 12 AVPV/PeN^Kisspeptin cells showed spontaneous discharges of action potentials (APs) at rest (0.1 Hz ± 0.06 Hz; average RMP of the spontaneously active cells was -50.7 mV ± 2.7 mV). The average RMP of ARH^kisspeptin neurons (14 cells from 12 animals) was -50.0 mV ± 1.5 mV (range: -62 mV to -39 mV), the average input resistance was 1.7 ± 0.1 GΩ, wcc was 9.2 pF ± 0.7 pF, and SR of 16.9 ± 1.7 mΩ. Most ARH^kisspeptin neurons were quiescent, whereas four out of 14 cells showed spontaneous APs at rest (0.9 Hz ± 0.5 Hz; average RMP of the spontaneously active cells was -52.7 mV ± 1.4 mV).

The administration of hGH (20 µg/g) to the bath induced a significant hyperpolarization of the RMP of many of the recorded neurons, five out of 12 AVPV/PeN^Kisspeptin neurons (≈55% of cells from 9 mice), and nine out of 14 recorded ARH^kisspeptin neurons (≈65% of cells from 12 mice, p = 0.0006, Fisher's exact test; Figure 3). The AVPV/PeN^Kisspeptinand ARH^kisspeptin hyperpolarized neurons significantly changed the RMP compared to the unresponsive cells (Figure 3B,E; Mann-Whitney test). The effects on RMP (Figure 3C,F; repeated measures ANOVA and Tukey's post-test) were followed by a significant reduction of the whole-cell input resistance (IR) on AVPV/PeN^Kisspeptin (0.9 ± 0.1 GΩ to 0.7 ± 0.1 GΩ during hGH application, p = 0.02; Figure 3D), and on ARH^kisspeptin(1.7 ± 0.1 GΩ to 1.0 ± 0.1 GΩ during hGH application, p < 0.0001; repeated measures ANOVA and Tukey's post-test; Figure 3G) neurons. Additionally, the frequency of spontaneous APs (fAPs) of hGH-hyperpolarized neurons decreased in both populations of cells (0.1 Hz ± 0.06 Hz to 0.0 Hz ± 0.0 Hz in AVPV/PeN^Kisspeptin and 1.0 Hz ± 0.5 Hz to 0.2 Hz ± 0.1 Hz in ARH^kisspeptin neurons). However, the extent of this decrease failed to reach a level of statistical significance (p > 0.05, Mann-Whitney test). After the hGH washout, the RMP and IR were restored to baseline (Figure 3A,C,D,F,G). The remaining kisspeptin neurons were unresponsive to hGH administration.

We have previously demonstrated that pGH induces no effects on hypothalamic kisspeptin neuron activity (please refer to Silveira et al.²⁵; Figure 3H). Of note, it is known that hGH can activate prolactin (PRL) receptors in addition to GH receptors³⁷^,³⁸. Besides, PRL indirectly depolarizes only ≈20% of AVPV/PeN^Kisspeptin neurons in mice²⁴. In contrast, PRL does not modulate the fast synaptic transmission of the ARH^kisspeptincells²⁴. Therefore, the hGH-induced hyperpolarization effect reported here seems to be nonspecific. The observed differences may depend on variables such as drug concentration, species difference (human vs. mouse), or even the presence of salts in the composition of the used drugs, as previously reported²⁸.

Figure 1: Schematic diagram summarizing the whole-cell patch-clamp technique's contribution to the knowledge of the kisspeptin neurons' activity. Kisspeptin neurons (shown in green) are located in the anteroventral periventricular and rostral periventricular nuclei (AVPV/PeN) and arcuate nucleus of the hypothalamus (ARH). The AVPV/PeN^Kisspeptin and ARH^kisspeptin cells send direct connections to gonadotrophin-releasing hormone (GnRH) neurons' soma located in the preoptic area (POA) and their terminals at the median eminence (ME), culminating in the modulation of the hypothalamus-pituitary-axis (HPG). Different neuromodulators, such as hormones, have been shown to differentially modulate the activity of the AVPV/PeN^Kisspeptin and ARH^kisspeptin neurons. Possible effects on the resting membrane potential are schematically demonstrated by representative tracings obtained using the whole-cell path-clamp technique and current-clamp recordings. The red color indicates that a specific neurotransmitter induces the depolarization of the resting membrane potential (RMP)¹⁷^,²²^,²⁴^,²⁶^,²⁸^,²⁹^,³⁰^,³¹^,³²; the blue color indicates no effect on RMP²⁴^,²⁵^,²⁶^,²⁷^,³⁰. The dashed line indicates the RMP. Please click here to view a larger version of this figure.

Figure 2: Basic steps to obtain the sealing of the cell of interest by the whole-cell patch-clamp technique. (A,B) Representative photomicrographs of brain slices (250 µm) containing kisspeptin cells at the anteroventral periventricular nucleus (AVPV) and arcuate nucleus of the hypothalamus (ARH). Kisspeptin neurons were identified by green fluorescent protein (GFP) expression. (C) Photomicrograph demonstrating a micropipette (containing electrolyte solution [internal solution]) close enough to the cell to create a dimple in the plasma membrane to perform the seal. (D,E) Mild negative pressure (mouth suction performed on a tube attached to the headstage and micropipette) is required to seal the cell membrane to the micropipette (D). A second application of negative pressure (mild and brief) is necessary to induce the plasma membrane rupture (E). (F) The registration of the cell activity is performed by a mechanical setup used for patch-clamp experiments. After breaking the plasma membrane, currents flowing through the ionic channels in the patched cell can be recorded by an electrode connected to a highly sensitive amplifier. A feedback resistor generates the current needed for voltage-clamp (G) or current-clamp (H) recordings. Abbreviations: 3V = third ventricle; ME = median eminence. Scale bars: A = 130 µm, B = 145 µm, C = 20 µm, D = 15 µm. Please click here to view a larger version of this figure.

Figure 3: Testing drug specificity. (A) Representative current-clamp recording demonstrating that human recombinant growth hormone (hGH) induced a hyperpolarization of the resting membrane potential (RMP) of the kisspeptin neurons located at the arcuate nucleus of the hypothalamus (ARH^kisspeptin). (B–G) Bar graphs demonstrating the average change in the resting membrane potential (RMP) (B,C,E,F) and average input resistance (IR) (D,G) of hGH-responsive kisspeptin neurons located at the anteroventral periventricular and rostral periventricular nuclei (AVPV/PeN^Kisspeptin) (B–D) or ARH (E–G). Representative current-clamp recording demonstrating that porcine growth hormone (pGH) induced no effect on the RMP of ARH^kisspeptin neurons, as previously reported²⁵ (H). The significance tests used are the Mann-Whitney test for (B) and (E), and repeated measures ANOVA and Tukey's post-test for (C,D,F,G). The dashed line indicates the RMP. *p = 0.02; **p = 0.004,***p = 0.0003; ****p < 0.0001. Please click here to view a larger version of this figure.

Internal Solution (100 mL)
Salt	FW (g/mol)	Concentration	Weight (g)
K-gluconate	234.2	120 mM	2.81
NaCl	58.4	1.0 mM	0.006
KCl	74.5	10 mM	0.074
HEPES	238.3	10 mM	0.24
EGTA	380.3	5.0 mM	0.19
CaCl₂	147.0	1.0 mM	0.015
MgCl2	203.0	1.0 mM	0.02
KOH	56.11	3.0 mM	0.017
ATP	507.18	4.0 mM	0.20
pH =7.3 / osmolarity = 275 – 280 mOsm

Table 1: Reagents for the preparation of the internal solution. The table contains the molecular weight (FW), desired concentrations, and the calculated weight of the salts for the preparation of 100 mL of solution.

aCSF/Slicing Solution (250 mL)
Salt	FW	Concentration	Weight (g)
Sucrose	342.3	238 mM	18.5
KCL	74.5	2.5 mM	0.046
NaHCO₃	84.0	26 mM	0.546
NaH₂PO₄	120.0	1.0 mM	0.03
MgCl₂	203.0	5 mM	0.254
D-glucose	180.2	10 mM	0.450
CaCl₂	147.0	1.0 mM	0.037
pH = 7.3 / osmolarity = 290 – 295 mOsm

Table 2: Reagents to prepare the slicing solution. The table contains the molecular weight (FW), desired concentrations, and calculated weight of the salts for the preparation of 250 mL of solution. The brain is submerged in this solution to be sliced.

aCSF for recording (1 L)
Salt	FW	Concentration	Weight (g)
NaCl	58.4	135 mM	7.88
KCL	74.5	3.5 mM	0.261
NaHCO₃	84.0	26 mM	2,184
NaH₂PO₄	120.0	1.25 mM	0.150
MgSO₄	246.5	1.2 mM	0.296
D-glucose	180.2	10 mM	1,802
CaCl₂	147.0	1.0 mM	0.148
pH = 7.3 / osmolarity = 290-300 mOsm

Table 3: Reagents to prepare aCSF for recordings. The table contains the molecular weight (FW), desired concentrations, and calculated weight of the salts for the preparation of 1 L of solution.

Supplementary Figure 1: Example of an in-house made recovery chamber. (A,B) 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. (C) This adapted base can be placed inside a 500 mL beaker containing artificial cerebrospinal spinal fluid (aCSF) constantly saturated with carbogen (95% O2 and 5% CO2). (D) The beaker holding the recovery chamber is kept in the water bath during experimentation. (E,F) An acrylic transfer pipette is used to transfer the brain slices to the recovery chamber. Please click here to download this File.

Discussion

The development of the whole-cell patch-clamp technique had a significant impact on the scientific community, being considered of paramount importance for developing scientific research and enabling several discoveries. Its impact on science was enough to culminate in the Nobel Prize in Medicine in 1991, as this discovery opened the door to a better understanding of how ion channels function under physiological and pathological conditions, as well as the identification of potential targets for therapeutic agents¹¹^,³⁹^,⁴⁰^,⁴¹. In the field of medicine, one of the outstanding findings made using this technique was that several substances used clinically interact directly with ion channels (e.g., local anesthetics, antiarrhythmics, antidiabetics, and muscle relaxants)⁴². Therefore, its applicability is evident in clinical institutes and basic research departments. This article describes the protocol for the basic preparation to perform whole-cell patch-clamp experiments on brain slices containing hypothalamic kisspeptin neurons. We outline the basic steps and highlight notable parameters that must be controlled to obtain tissue and prepare solutions for the patch-clamp technique. However, this article cannot fully describe the complexity of this technique and the mechanisms involved in each type of recording, especially the analysis. For further theoretical learning, some books and reviews on the patch-clamp technique are recommended¹⁰^,⁴³^,⁴⁴^,⁴⁵^,⁴⁶.

Each solution used in the patch-clamp method has a specific purpose; therefore, its specific compositions and criteria must be rigorously adopted during preparation. For example, the internal solution's composition should be determined according to the experiment's goal, since it varies depending on the type of current to be measured. The chloride-based solution mentioned here, in which Cl^– (15 mM) mimics the physiological concentration of the cell cytoplasm⁴⁷, is utilized to study active and passive neuronal properties or responses to synaptic input. It is important to emphasize that the chloride content in the internal solution keeps the chloride equilibrium potential at optimal levels for cell recording (in the mentioned solution E_CL, it is about -59 mV). The intracellular chloride concentration can be estimated by evaluating the reversal potential for GABA-induced current (E_GABA), assuming that all current through the GABA_A receptor is carried by Cl^–²¹^,⁴⁸. For the study of hypothalamic kisspeptin neurons, one must consider that the intracellular Cl^– concentration for ARH^kisspeptin is higher than for AVPV/PeN^Kisspeptin cells²¹. This characteristic must be considered when planning an experiment. The osmolarity of the internal solution is recommended to be at least 5% lower than that of aCSF for recordings to avoid loss of seal due to possible swelling and/or cell weakening³³^,⁴⁷. If there is a need to add an intracellular compound that can be further used as a cell marker to the internal solution, such as biocytin or an intracellular dye (e.g., Alexa Fluor 594), K⁺ levels may be reduced so that osmolarity can be constantly maintained²⁰^,⁴⁷^,⁴⁹^,⁵⁰.

Performing a rapid brain dissection and maintaining a low temperature (0-2 °C) during slicing with an appropriate slicing solution is crucial. The slicing solutions may differ according to the type of cell and/or brain region being evaluated³³^,⁴⁷. The aCSF solution used to obtain the brain slices (i.e., slicing solution) usually has a different composition from the aCSF for recordings (for comparison, see Table 2 and Table 3). The cation (Ca²⁺ and Mg²⁺) concentrations can be adjusted to modify the excitability of neurons, which can also affect the firing threshold and neurotransmitter release⁴⁷. Low Ca²⁺ and high Mg²⁺ media are widely utilized for minimizing possible excitotoxic processes (for more information, see⁵¹). In addition, low Ca²⁺ and high Mg²⁺ media can stimulate hypothalamic neuron activity⁵². Regarding the aCSF for recordings, some characteristics are similar. The buffer system is based on NaHCO₃, high NaCl concentrations (generally >100 mM) and low KCl concentrations (generally <5 mM), relative concentrations of Ca²⁺ and Mg²⁺ (2:1 is often used) are usually around 2 mM, and glucose levels can range from 1 to 10 mM⁴⁷. Besides variations of the solutions' osmolarity, pH, or temperature changes (digitally controlled to be between 30-32 °C), it is important to highlight that other issues may occur during the experimental protocol that substantially interfere with the experimentation. Electrophysiological measurements must be stable during the recordings, and any variations must have a critical reading. For example, SR changes account for cellular access in whole-cell mode and have considerable consequences on measured currents and potentials. Furthermore, it is important to emphasize that a relevant limitation related to the patch-clamp methodology that must be considered is mechanical overstimulation of the cell by excessive suction/pressure protocols that can induce morphological and functional changes⁵³. This bias is often difficult to control in protocols where suction is performed by mouth rather than a device that can precisely control the suction intensity applied. Additionally, tissue-related issues, which culminate with a high percentage of cell mortality, may occur due to inadequate dissection, hypoxia, or the mouse's health condition that a researcher cannot identify by observation.

The main advantage of the whole-cell patch-clamp technique is the ability to record neurons in specific brain regions of interest precisely. This technique has tremendously benefited from the creation of genetically modified animals. Our group typically works with a mouse model that expresses the hrGFP under the Kiss1 gene promoter or another that expresses Cre-recombinase under the Kiss1 gene promoter and GFP under the Cre-conditional expression. However, there are many validated animal models nowadays²³^,²⁶^,³⁶. In addition, the absence of the blood-brain barrier and the fact that the extracellular and intracellular environment can be easily controlled and manipulated represents advantages of this method; however, they do not necessarily represent a physiological condition. Among the limitations of this technique compared to in vivo preparations, or other recording types aiming to preserve the cytoplasmatic ionic concentration, such as on-cell or perforated-patch recordings, it is important to mention that the invasiveness of the whole-cell configuration causes dialysis of the cytoplasm content⁵⁴. Dialysis may cause the interruption of molecular aspects necessary for some phenomena to develop or be expressed. By recording from slices, it needs to be remembered that most of the neurons' projections are sectioned. Thus, it is out of the technique's scope to evaluate how much this disruption impacts the observed effects or a physiological condition. As mentioned, coronal brain slices of 200-300 μm are usually performed to study the activity of hypothalamic kisspeptin neurons¹⁷^,¹⁹^,²⁰^,²¹^,³⁴. The limitation of using a thicker brain slice section to study kisspeptin cells and different slice angles maintaining specific AVPV/PeN or ARH connectivity⁵⁵ needs further investigation. In addition, by testing the effect of a hormone/drug, synthetic or not, on the RMP of a neuron, several studies are based on the drug EC₅₀ (if it is known) or on published data that demonstrate that a specific concentration is effective in activating firing rate or changing [Ca²⁺]_i levels. However, one should be aware of the composition/specificity of the drug to be used in the experiments, as it is possible that purified synthetic drugs, compared to other similar drugs, may have antagonistic effects²⁸. As demonstrated previously²⁵, while purified pGH induces no effect on hypothalamic kisspeptin neuron activity, the hGH produced controversial data (see Figure 3). Similar results were demonstrated when insulin effects on the ARH were assessed²⁸. Therefore, drug-specificity should be considered when planning an experiment and interpreting the obtained results.

The patch-clamp technique is an excellent tool for obtaining data about neuron electrical activity and has significantly contributed to the knowledge of several neuronal populations, such as the kisspeptin neurons. Most of the details provided here are generally used for recordings of hypothalamic neurons, as we have reported previously²⁵^,⁵⁰^,⁵⁶^,⁵⁷^,⁵⁸^,⁵⁹^,⁶⁰. Importantly, for recording other neuronal populations besides kisspeptin neurons, one must know or determine the electrophysiological measurements that can help identify cell types, such as cell capacitance, SR, input resistance, cell firing pattern, and other parameters. These properties vary between different brain cells, brain nuclei, and physiological or induced conditions, such as circulating sex steroid levels¹⁶^,¹⁹^,²⁰^,²¹^,⁶¹, which can substantially interfere with the critical analysis of the results. In addition, it is necessary to understand the possible variables involved in tissue preparation and their associated advantages and limitations to work with this technique. All the steps described here must be conducted judiciously, as any change in the variables involved in the protocol may drastically interfere with the results.

Disclosures

The authors have nothing to disclose.

Acknowledgements

This study was supported by the São Paulo Research Foundation [FAPESP grant numbers: 2021/11551-4 (JNS), 2015/20198-5 (TTZ), 2019/21707/1 (RF); and by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) – Finance Code 001" (HRV).

Materials

Compounds for aCSF, internal and slicing solutions
ATP	Sigma Aldrich/various	A9187
CaCl₂	Sigma Aldrich/various	C7902
D-(+)-Glucose	Sigma Aldrich/various	G7021
EGTA	Sigma Aldrich/various	O3777
HEPES	Sigma Aldrich/various	H3375
KCL	Sigma Aldrich/various	P5405
K-gluconate	Sigma Aldrich/various	G4500
KOH	Sigma Aldrich/various	P5958
MgCl₂	Sigma Aldrich/various	M9272
MgSO₄	Sigma Aldrich/various	230391
NaCl	Sigma Aldrich/various	S5886
NaH₂PO₄	Sigma Aldrich/various	S5011
NaHCO₃	Sigma Aldrich/various	S5761
nitric acid	Sigma Aldrich/various	225711	CAUTION
Sucrose	Sigma Aldrich/various	S1888
Equipments
Air table	TMC	63-534
Amplifier	Molecular Devices	Multiclamp 700B
Computer	various	–
DIGIDATA 1440 LOW-NOISE DATA ACQUISITION SYSTEM	Molecular Devices	DD1440
Digital peristaltic pump	Ismatec	ISM833C
Faraday cage	TMC	81-333-03
Imaging Camera	Leica	DFC 365 FX
Micromanipulator	Sutter Instruments	Roe-200
Micropipette Puller	Narishige	PC-10
Microscope	Leica	DM6000 FS
Osteotome	Bonther equipamentos & Tecnologia/various	128
Recovery chamber	Warner Instruments/Harvard apparatus	–	can be made in-house
Recording chamber	Warner Instruments	640277
Spatula	Fisher Scientific /various	FISH-14-375-10; FISH-21-401-20
Vibratome	Leica	VT1000 S
Water Bath	Fisher Scientific /various	Isotemp
Software and systems
AxoScope 10 software	Molecular Devices	–	Commander Software
LAS X wide field system	Leica	–	Image acquisition and analysis
MultiClamp 700B	Molecular Devices	MULTICLAMP 700B	Commander Software
PCLAMP 10 SOFTWARE FOR WINDOWS	Molecular Devices	Pclamp 10 Standard
Tools
Ag/AgCl electrode, pellet, 1.0 mm	Warner Instruments	64-1309
Curved hemostatic forcep	various	–
cyanoacrylate glue	LOCTITE/various	–
Decapitation scissors	various	–
Filter paper	various	–
Glass capillaries (micropipette)	World Precision Instruments, Inc	TW150F-4
Iris scissors	Bonther equipamentos & Tecnologia/various	65-66
Pasteur glass pipette	Sigma Aldrich/various	CLS7095B9-1000EA
Petri dish	various	–
Polyethylene tubing	Warner Instruments	64-0756
Razor blade for brain dissection	TED PELLA	TEDP-121-1
Razor blade for the vibratome	TED PELLA	TEDP-121-9
Scissors	Bonther equipamentos & Tecnologia/various	71-72, 48,49;
silicone teat	various	–
Slice Anchor	Warner Instruments	64-0246
Syringe filters	Merck Millipore Ltda	SLGVR13SL	Millex-GV 0.22 μm
Tweezers	Bonther equipamentos & Tecnologia/various	131, 1518

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Silva, J. d. N., Zampieri, T. T., Vieira, H. R., Frazao, R. Hypothalamic Kisspeptin Neurons as a Target for Whole-Cell Patch-Clamp Recordings. J. Vis. Exp. (193), e64989, doi:10.3791/64989 (2023).

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