In Vivo Whole-cell Patch-clamp Recording of Hypothalamic Peptidergic Neurons in Larval Zebrafish

Summary

This article presents a protocol for in vivo recording of electrical activity of hypothalamic peptidergic neurons using whole-cell patch-clamp electrophysiology in intact larval zebrafish.

Abstract

The hypothalamus is an ancient brain region that regulates diverse aspects of physiology and behavior, including sleep and wakefulness, appetite, energy homeostasis, anxiety, depression, and social interaction. Specific neuronal populations in the hypothalamus exert their effects via the release of neurotransmitters and neuropeptides. Whole-cell patch-clamp recording is an indispensable approach for studying the roles of these factors in synaptic transmission and brain function. However, it is challenging to access hypothalamic neurons for electrophysiological recordings in intact mammals due to their location deep within the brain. As a result, our understanding of the intrinsic properties and physiological functions of hypothalamic neurons is limited. The larval zebrafish is a useful alternative model to study hypothalamic neurons due to its transparent and small, but well-conserved, vertebrate brain. Here, we present a protocol for in vivo whole-cell patch clamp recordings of hypothalamic neurons in intact larval zebrafish. Using this technique, we can record from peptidergic neurons in the hypothalamus, examine the responses of these neurons to sensory stimuli, and explore their effects on downstream neurons. This experimental technique thus provides a useful approach to study the physiological functions of hypothalamic neuropeptidergic neurons in intact animals.

Introduction

As a popular vertebrate animal model, zebrafish (Danio rerio) are widely used in neuroscience research and have several advantages¹. Their complement of genes and brain architecture is highly conserved with mammals, and they have a rich behavioral repertoire, making them useful to study genetic and neuronal mechanisms that underlie behaviors such as sleep, anxiety, depression, and social interaction^2,3,4,5,6. Their small size and low maintenance costs make them ideal for high-throughput screening of genes or drugs aimed at treating neuropsychiatric diseases^7,8. Finally, the optical transparency of larval zebrafish, combined with advanced light-sheet microscope techniques, and the availability of multiple fluorescent reporters, makes them particularly well-suited for monitoring neuronal and astroglial activity at cellular resolution across the entire brain^9,10,11.

The zebrafish hypothalamus, located deep in the ventral diencephalon, is anatomically and molecularly conserved with the mammalian hypothalamus, but is 2-3 orders of magnitude smaller, providing a simpler system to study hypothalamus function¹². Hypothalamic neurons form extensive synaptic connections with multiple brain regions, including the thalamus, brainstem, pituitary gland, and telencephalon, through which they regulate physiological homeostasis, neuroendocrine signaling, and autonomic behaviors^13,14. Such modulation of behavior and physiology largely relies on neuropeptides, which are short-chain amino acids that act by binding to G-protein-coupled receptors (GPCRs)¹⁵. In contrast to fast-acting amino acid neurotransmitter release at the synapse, neuropeptides may diffuse over longer distances via volume transmission, and mediate slow modulatory effects on synaptic transmission and neuronal excitability¹⁶.

Recent studies in zebrafish have revealed diverse functions of hypothalamic neuropeptides through multidisciplinary approaches such as whole-brain calcium imaging, single-cell RNA sequencing, high-throughput behavioral analysis, and neuronal circuit mapping^4,12,17,18. These emerging techniques have transformed our ability to characterize cellular diversity at a large scale, bridge molecular mechanisms with circuit-level neuropeptide functions, and map neural circuits that underlie specific behaviors^19,20. However, these approaches remain limited in their ability to resolve the rapid physiological dynamics and precise synaptic mechanisms that underlie neuropeptide function.

The patch-clamp technique, developed by Neher and Sakmann, remains the gold standard for investigating neuronal physiology due to its unparalleled temporal resolution and biophysical precision, enabling direct measurement of membrane potential fluctuations, synaptic currents, and ion channel dynamics^21,22. This technique is particularly invaluable for studying neuropeptidergic cells, whose characteristic bursting patterns and neuromodulatory influences require direct electrophysiological measurement to fully understand their roles in behavior and homeostasis^15,16,23.

Here, we have developed an optimized protocol that enables reliable whole-cell patch-clamp recordings of larval zebrafish hypothalamic peptidergic neurons, despite the technical challenges posed by the deep anatomical location of these neurons. This methodology provides direct access to the electrical activity of these cells, permitting detailed investigation of their responses to visual and auditory stimulus (Figure 1). By integrating patch-clamp electrophysiology with optogenetics, we can precisely characterize functional connectivity between hypothalamic circuits and downstream neurons (Figure 2 and Figure 3), facilitating the study of neuropeptides in physiological homeostasis and behavioral control. This approach bridges a critical gap in neuroscience research, allowing mechanistic exploration of hypothalamic neuron function at the biophysical, cellular, and circuit levels simultaneously.

Protocol

All experiments were approved by the Institutional Care and Use Committee (IACUC) at Shanghai University (animal protocol YS 2025-169) and IACUC at California Institute of Technology (animal protocol 1836). Zebrafish from 5 to 7 days post-fertilization (dpf) were fed with rotifers and used for experiments. At this stage of development, sex is not determined. Adult zebrafish on a nacre [mitfa(w2/w2)] background²⁴ were used for breeding.

1. Solutions and recipes

1. Prepare E3 medium containing 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl₂, and 0.33 mM MgSO₄. Adjust the solution by NaOH to pH 7.2. Store at room temperature for use within 1 month.
2. Prepare the extracellular solution containing 134 mM NaCl, 2.9 mM KCl, 2.1 mM CaCl₂, 1.2 mM MgCl₂, 10 mM HEPES, and 10 mM glucose. The osmolality of the extracellular solution is ~290 mOsmol L^-1. Adjust the solution by NaOH to pH 7.8. Store at 4 °C for use within 1 month.
3. Prepare the intracellular solution containing 100 mM K-gluconate, 10 mM KCl, 2 mM CaCl₂, 2 mM Mg-ATP, 0.3 mM Na-GFP, 10 mM HEPES, and 10 mM EGTA. The osmolality of the intracellular solution is ~280 mOsmol L^-1. Adjust the solution by KOH to pH 7.4. Aliquot 100 µL of the internal solution into microfuge tubes and store at -20 °C for use within 1 year.
   NOTE: The addition of Mg-ATP and Na-GTP is recommended for prolonged and stable recordings. Intracellular solution needs to be filtered through a 0.22 µm membrane, which removes particulates and microbial contaminants that could compromise seal formation or lead to pipette clogging.
4. α-bungarotoxin: Dissolve 1 mg of α-bungarotoxin in 1 mL of E3 medium to a final concentration of 1 mg/mL. Aliquot 20 µL of the solution into microfuge tubes and store at -20 °C for use within 1 year.
5. Low-melting point agarose: Dissolve 0.75 g of agarose in 50 mL of E3 medium in a glass bottle to a final concentration of 1.5%. Heat the mixture in a microwave oven until the agarose is melted and the solution appears clear and homogeneous. Maintain the melted agarose at 42-45 °C in a water bath for use within 1 month.

2. Zebrafish preparation and dissection

1. Place single male and female adult fish containing a desired transgene, e.g., Tg(hcrt:RFP) fish²⁵, in a mating tank and separate them by a clear divider overnight. In the morning of the second day, remove the divider to allow the fish breed. Collect fertilized embryos at the bottom of the tank and return adult fish to their home tanks by early afternoon.
2. Raise the embryos in a 9 cm diameter Petri dish with E3 medium including 0.0005% methylene blue. Place the embryos in an incubator at 28.5 °C with 14:10 h light:dark cycles. Change the E3 solution every day.
   NOTE: Methylene blue in E3 medium is used to prevent fungal growth until 3 days post-fertilization (dpf), but is replaced with E3 medium alone at 3 dpf, since methylene blue may affect cellular metabolism and mitochondrial function at later stages of development²⁶.
3. Screen larval fish under an epi-fluorescence stereomicroscope with maximum magnification (~115x) starting at around 3 dpf. Place individual fish in water droplets (larger than their body size) to facilitate the sorting. For the fish older than 3 dpf, anesthetize the fish using 3 mL of E3 medium containing 0.01% tricaine to suppress spontaneous locomotor activity. Collect larvae with bright fluorophore expression in the hypothalamus.
   NOTE: Wear nitrile gloves, safety glasses, and a lab coat when handling tricaine. Excess tricaine must be disposed of as chemical waste according to animal protocols.
4. After the screening, wash out the tricaine using E3 medium immediately, and put these larvae back in the incubator. Add ~5 mL of feeding solution containing rotifers into the Petri dish starting at 5 dpf until the day of the experiments. Change the E3 solution and feed the fish with rotifers every day.
5. At 5-7 dpf, collect larval fish that appear healthy and show active locomotion. Transfer them to a small petri dish (3.5 cm in diameter) using a glass pipette.
6. Remove the water around the fish, and add 20 µL of 1 mg/mL α-bungarotoxin onto the fish for ~30 s. Then wash out the drug by adding ~3 mL of E3 medium into the Petri dish.
   NOTE: Wear nitrile gloves, safety glasses, and a lab coat when handling α-bungarotoxin. Excess α-bungarotoxin must be disposed of as chemical waste according to animal protocols.
7. Cover the Petri dish with aluminum foil to keep the fish in the dark and gradually become paralyzed.
   NOTE: α-bungarotoxin is an acetylcholine receptor blocker that specifically acts on the neuromuscular junction, thereby paralyzing the fish without affecting the nervous system. The fish can still sense environmental stimuli, and brain function is normal, after paralysis^9,27.
8. Check for paralysis after ~15 min using a stereomicroscope and select a paralyzed fish that shows a normal heartbeat and robust blood circulation in the brain and body.
9. Transfer the fish into a custom-made recording chamber, which is a 35 mm Petri dish with a 10 mm diameter hole in the center and a 20 x 20 mm glass coverslip glued at the bottom to cover the hole. Remove the water and add ~200 µL of 1.5% low-melting agarose.
10. Position the fish dorsal side up in the agarose using forceps (Dumont #5). When the agarose has fully solidified, add the extracellular solution to cover the agarose. It takes 5-10 min for the agarose to fully solidify.
11. Cut and remove the agarose above the fish's head using a micro knife. Then make a small incision in the skin above the brain ventricle between the optic tectum and cerebellum using a glass micropipette with ~1 µm tip opening for pipette advancing into the brain next step.
12. Remove tissue debris or blood clots around the dissection hole using a broken glass pipette.

3 Whole-cell patch-clamp recording

1. Prepare recording micropipettes with ~1 µm tip opening and ~20 MΩ resistance using borosilicate capillaries on the day of the experiment. Store pulled micropipettes in a container and use them on the same day to prevent contamination.
2. Turn on the computer, CCD camera, monitor, amplifier, digitizer, micromanipulator, and associated software. Amplifier and digitizer must be turned on prior to opening software.
3. Carefully move the fish to an electrophysiological rig where the patch clamp experiment will be performed.
4. Perfuse the fish with oxygenated extracellular solution at ~2 mL per min using a peristaltic pump to maintain fish health and cell viability.
5. Move the recording chamber using an X-Y translator (custom-made) to center the fish in the visual field of a 4x objective under an upright microscope.
6. Switch to a 60x objective and move the translator to visualize the target hypothalamus region. Adjust Nomarski differential interference contrast (DIC) with 900 nm infrared light to get the best visualization of the cells.
7. Turn on the fluorescence excitation light source (e.g., mercury bulb) and use it to visualize fluorescent neurons in the hypothalamus. Locate healthy neurons that show bright fluorescence and clear cell boundaries, which often indicate good cellular health and facilitate high-quality electrophysiological recordings.
8. Slowly back-fill a recording micropipette with intracellular solution using a fine microloader tip.
9. Affix the electrode to the headstage of the amplifier and tighten the knob to prevent air leakage. Apply a slight positive pressure (~150-200 mBar) to the inside of the micropipette through the tubing connected to the holder, which prevents contaminants at the air-solution interface from entering the electrode tip.
10. Lower the micropipette into the bath solution using a micromanipulator in fast mode and position the micropipette tip above the dissected hole under the 4x objective.
11. Switch the micromanipulator to slow mode, and change the objective to the 60x one. Advance the tip of the glass pipette slowly through the dissection hole, and along the ventricle between the two tectal hemispheres.
12. Bring the pipette tip orthogonally towards a target fluorescent cell viewed using both DIC and fluorescence. Once the pipette tip touches the cell membrane, a small indentation is formed.
    NOTE: To visualize deep tissues like the hypothalamus, 900 nm IR-DIC is used together with polarizer and analyzer filters that are finely adjusted for IR-DIC.
13. Release of the positive pressure quickly when the tip touches a cell enables the cell to form a seal with the pipette. Simultaneously, apply a brief negative pressure via the tubing connected to the holder.
    NOTE: The value of resistance on the software should quickly increase to > 1 GΩ, indicating a high-resistance seal is formed. When the fluorescent hypothalamic cell is recorded successfully, there should be fluorescent cell membrane within the tip of the recording micropipette.
14. Clamp the cell at -60 mV and wait for 1-2 min until the baseline and access resistance are stable. Apply a brief electrical pulse or gentle suction by mouth to rupture the cell membrane beneath the micropipette tip.
15. Compensate for the capacitive current and series resistance in an electrophysiologic software configuration. Discard the data if the series resistance is larger than 100 MΩ and varies > 20% in the recordings.
16. Apply a 1 s -10 mV voltage pulse in voltage mode to monitor intrinsic membrane resistance before and after the experiments. Discard the cell if the membrane resistance or membrane potential varies a lot.
17. First record in voltage-clamp (VC) mode when the cell is held at a negative value. After establishing a stable recording, switch to current-clamp (IC) mode if needed.
    NOTE: The giga-seal may not be stable shortly after the rupture, so clamping the cell at -60 mV under VC mode for a few minutes at the beginning of an experiment can provide better cellular conditions.
18. Process the analysis of electrophysiological data and spike detection offline using MATLAB after the experiment is complete.
19. After the experiments, euthanize the larval fish using 1.5% Sodium hypochlorite (NaClO) and dispose of them according to institutional animal care protocols.

Results

In this manuscript, we present an improved in vivo whole-cell patch-clamp recording technique for investigating hypothalamic neurons in intact zebrafish, particularly focusing on hypocretin (Hcrt) neurons.

As presented in Figure 1, our methodology enables direct electrophysiological recordings from these neurons deep within the brain in an intact animal (Figure 1A), overcoming limitations of traditional in vitro slic...

Discussion

The protocol described here enables patch-clamp recordings of peptidergic neurons in the larval zebrafish hypothalamus, one of the deepest and most technically challenging brain regions to access. Due to the inherent difficulty of this preparation, successful patch-clamp recordings require meticulous attention to a few critical parameters, namely pipette quality, approach technique, solution purity, and tissue health. These factors collectively determine the likelihood of achieving and maintaining stable giga-seal record...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Amplifier	Axon	700B
Borosilicate glass capillaries	Sutter	BF100-58-10
CCD camera	Dage-MTI	IR-1000
Computers for electrophysiological recordings	Dell	Precision 3660
Digidata	Axon	1440A
Faraday Cage	Custom-made
Forceps	F.S.T.	Dumont #5
Incubator	Lonroy	GZP-150B
Membrane filter	Millipore Sigma	SLGV004SL
Micro knife	F.S.T.	10318-14
Objective	Olympus	Mplan 5X/0.1; UMPlanFI/IR 60X/0.9w
Peristaltic pump	Longer	BT100-1L
Pipette holder	Narishige	H-7	for dissection
Puller	Sutter Instrument	p-97
Stereomicroscope for fluorescent screening and dissection	Olympus	SZX16
Stimulator	A.M.P.I	Master8
Three-dimensional micromanipulator	Sutter Instrument	MPC-325
Upright infrared DIC microscope	Olympus	BX51WI
Vibration isolator table	TMC	61-541-06
Video monitor	SUNSPO	SP-717
Water bath	Yiheng	HWS-12
X-Y translator	Custom-made