Protocol for extracellular recordings of mechanically stimulated pentascolopidial chordotonal neurons in Drosophila larvae

Summary

Chordotonal organs are sensory structures containing ciliated neurons specialized in detecting mechanical forces. Here, we present a protocol for recording the neuronal activity of mechanically stimulated pentascolopidial chordotonal organs (lch5) of Drosophila larvae. We describe steps for dissecting larvae, measuring electrical activity of chordotonal neurons, and quantifying action currents. This protocol is useful for studying the biology of larval lch5 neurons, as well as the molecular mechanisms and components that contribute to their mechanosensitive properties.

For complete details on the use and execution of this protocol, please refer to Bormann et al.¹

Graphical abstract

Highlights

• •Steps to dissect the Drosophila larva for extracellular recordings
• •Instructions for measuring electrical activity from lch5 neurons
• •Guidance on precise piezo-driven lch5 mechanical stimulation

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Chordotonal organs are sensory structures containing ciliated neurons specialized in detecting mechanical forces. Here, we present a protocol for recording the neuronal activity of mechanically stimulated pentascolopidial chordotonal organs (lch5) of Drosophila larvae. We describe steps for dissecting larvae, measuring electrical activity of chordotonal neurons, and quantifying action currents. This protocol is useful for studying the biology of larval lch5 neurons, as well as the molecular mechanisms and components that contribute to their mechanosensitive properties.

Before you begin

Insects and crustaceans use chordotonal organs as sensors for detecting mechanical forces.^2,3 Chordotonal organs are made up of multicellular units – the scolopidia. Each scolopidium harbors at least one mono-ciliated neuron (mechanosensitive compartment), a cap cell (attachment to cuticle), a scolopale cell (encapsulates the distal dendrite), and a ligament cell (attachment to cuticle). Lateral pentascolopidial chordotonal organs (lch5 organs) contain five such units and can be found embedded within body wall muscles in each hemisegment of the Drosophila larva. In recent years lch5 organs in Drosophila have garnered attention as a powerful model system to elucidate molecular, mechanistic and cellular aspects of mechanosensation.^4,5,6,7 In Bormann et al. (2025),¹ we used electrophysiological assessment of mechanosensory lch5 neurons of third-instar Drosophila larvae to dissect the function of specific isoforms of the Ca²⁺ independent receptor of latrotoxin (ADGRL/Cirl) under physiological conditions.

The protocol presented here outlines the functional assessment of lch5 neurons. It details the dissection of the specimen, electrophysiological recording of neuronal activity generated in response to mechanical stimuli of varying intensities (applied as defined perpendicular pull lengths on the cap cells), and analysis of the neuronal lch5 response.

These extracellular recordings can be conducted on lch5 organs of wildtype or genetically modified Drosophila to explore the molecular mechanisms underlying mechanosensation and its regulation.

Preparation of micropipettes

Timing: 10–30 min

The recording electrode is a glass micropipette harboring a silver/silver chloride (Ag/AgCl) electrode used to suck in the nerve of the lch5 organ, creating a stable interface between the nerve and the electrode.

• 1.
  Fabrication of micropipettes.

  • a.
    Insert a borosilicate glass capillary into a micropipette puller (e.g. DMZ-Universal Electrode Puller, Zeitz).
  • b.
    Apply a 2-step protocol (prepull → final pull), followed by a polishing step to obtain micropipettes with a tip outer diameter of 6–10 μm (see settings in materials and equipment section) (Figure 1).

    Note: Settings to attain the desired micropipette tip size must be adjusted every time the puller is used. Consult the device manual regarding adjustments.

  • c.
    Check the micropipette using a microscope with a 32x objective and 10x ocular magnification and confirm the appropriate diameter of the tip with a micrometer scale.

    Note: The micropipette can be reused.

Figure 1.

Tip of a representative micropipette

The tip diameter of micropipettes should be 6–10 μm. Scale: 10 μm.

Preparation of dissection and recording solutions

Timing: 40–60 min

• 2.
  Dissection solution:

  • a.
    Prepare calcium-free, hemolymph-like dissection solution according to Zhang et al., 2013 (see materials and equipment section).⁸
  • b.
    Adjust the pH to 7.25 using 1 M HCl.
  • c.
    Store at 4°C and use no longer than five days.
• 3.
  Recording solution:

  • a.
    Prepare a 1 M CaCl₂ stock solution and store it at 19°C–23°C.

    Note: Always check the 1 M CaCl₂ stock solution before use for precipitates and renew if necessary.

  • b.
    On the day of the experiment, prepare the recording solution by adjusting the dissection solution to 2 mM CaCl₂ (e.g. 28 μl 1 M CaCl₂ to 14 ml dissection solution).
  • c.
    Keep the solution at 19°C–23°C.

Prepare the instruments for dissection

Timing: 30–45 min + one 24-h step

• 4.Scissors: Refine/sharpen the tip size of the dissection scissors using a fine sharpening stone and surgical instrument oil.

Note: This step ensures that the tip of the scissors cuts effortlessly, minimizing mechanical strain inflicted on the nerve and the lch5 organ.

• 5.
  Dissection support: Dissection support stabilizes the petri dish lid in which the dissection of the larvae is done (Figure 2).

  • a.
    Attach six elastic buffers to the bottom of a round acrylic glass plate (3 mm thickness, 16 cm diameter).
  • b.
    Cut a petri dish lid into halves and attach them to the top of the acrylic glass plate fitting another ‘intact’ petri dish and stabilizing it.

    Note: Using the lid of a petri dish instead of the petri dish itself facilitates handling of the scissors during the dissection due to the lower edges of the lid, but using the petri dish would work as well.

  • c.
    Place a piece of black paper into the cut petri dish halves to increase the contrast between the background and the larva.
• 6.
  Sylgard pad:

  • a.
    Mix the base and the curing agent in a 10:1 ratio in a petri dish (e.g. 3 g base and 0.3 g curing agent in a 5.5 cm petri dish) and let it dry at 19°C–23°C for 24 h.

    Note: This results in a 1–2 mm thick Sylgard layer. Sylgard can also be thicker; however, it must remain sufficiently thin to ensure that the working distance of the condenser is adequate to bring the sample into the focal plane, thereby enabling proper Köhler illumination.

  • b.
    Use a scalpel to cut a pad of about 1 x 0.5 cm size out of this hardened Sylgard (Figure 2B).
  • c.
    Place it in a clean and dry petri dish.

    Note: The petri dish and Sylgard can be reused for several months.

• 7.Prepare minutien pins for fixation of larvae: Cut 6–8 pins (∼12.5 μm in tip-diameter) with a fine side cutter to a maximal length of 2 mm to fix the larvae on the Sylgard pad.

CRITICAL: Shortening of the pin prevents damaging the dip-in objective used in later steps.

Figure 2.

Dissection support base and chamber for dissection

A Petri dish containing a Sylgard pad is stabilized by a base made from acrylic glass and two halves of another Petri dish.

(A) Side view of the base.

(B) Top view of the Sylgard pad in the Petri dish. Inset shows a clipped minutien pin. Scale bar: 500 μm.

Preparation of the electrophysiology setup

Timing: 3–4 h

• 8.
  Preparation of a stimulation hook:

  • a.
    Under microscopic observation, carefully manipulate the distal extremity of ultra-fine pins (tip diameter approximately 12.5 μm) using precision forceps, such as size 4 or 5 from Fine Science Tools, to induce controlled deformation.

    Note: Alternatively, the tip of a pin can be bent by cautiously pushing it against a hard edge (e.g. glass). The resulting bent tips are somewhat random. Therefore several attempts are usually needed to get an optimal shape.

  • b.
    Next, use super glue to attach the pin to a micropipette.
  • c.
    Couple the micropipette, including the hook, to a piezo element using a short acrylic tube and a plastic screw (Figure 3).
  • d.
    Ground the piezo element.

    Note: The hooks can be used indefinitely unless they are damaged.

• 9.
  Preparation of a plastic clamp: This clamp is used to fix the Sylgard pad in place.

  • a.
    Attach a magnet to the tip of a bent cable tie (Figure 4).
• 10.
  Preparation of Ag/AgCl recording electrode:

  • a.
    Rinse the Ag wire with 70% ethanol.
  • b.
    Immerse the Ag wire (high purity, 0.25 mm diameter) into a 1 M KCl solution. This serves as the electrolyte for the chloridation process.
  • c.
    Connect the Ag wire to the anode of a DC power supply and place another Ag wire in the KCl solution to provide the cathode.
  • d.
    Apply DC current until a brown/gray layer of AgCl forms on the silver surface, i.e., no shiny silver surface is visible.

    Note: We use the Chlorinator CHG1 as a power source and apply a 3 mA current for around 45 s.

  • e.
    Inspect the electrode and ensure that it is coated evenly with an AgCl layer. If the coating appears uneven, repeat chloridation.
  • f.
    Rinse the Ag wire with 70% ethanol.

    Note: Rechloride your electrodes once a week or when you spot shiny, silver surfaces on the Ag/AgCl electrode.

• 11.
  Recording electrode:

  • a.
    Place the manipulator holding the headstage with the recording electrode at a 30° angle to the horizontal plane (Figure 5C).
  • b.
    Turn the recording chamber and position the electrode in one line with the anterior-ventral side of the lch5 organ, i.e., the electrode points to where the nerve leaves the organ.
  • c.
    Connect the recording electrode holder to a 50 ml syringe using flexible silicone tubing for the application of negative pressure to suck in the nerve bundle into the recording electrode.
  • d.
    Connect the bath electrode (chloride as described in step 10) to the headstage.
  • e.
    Couple the headstage with the recording amplifier.
  • f.
    Link the scaled output of the amplifier to the analog input of the analog-digital converter (Figure 5).
• 12.
  Stimulation hook:

  • a.
    Mount the piezo element with the attached stimulation hook on a micromanipulator that faces the recording chamber at a 30° angle to the horizontal plane (Figure 5A). The orientation of the stimulation faces the cap cells at a 90° angle to deliver mechanical forces perpendicularly to the cap cell (Figure 10).
  • b.
    Couple the piezo element to the piezo amplifier.
  • c.
    To control the piezo, its amplifier control input is connected to the analog output of the analog-digital converter via a BNC cable (Figure 6).
  • d.
    The same output channel is connected to the analog input of the analog-digital converter to monitor the applied voltage jumps, which control the movement of the piezo element.
• 13.Ground the recording amplifier. Adjust the recording amplifier to voltage-clamp mode, whole cell (β = 1) configuration, 10x output gain and a 1 kHz low-pass Bessel filter.
• 14.
  First, test different voltage jumps to verify whether the piezo element drives movement of the stimulation hook at the desired pull lengths.

  • a.
    Using a microscope and a camera, adjust the voltage jumps to get 3 and 1 μm pull lengths.
  • b.
    Extrapolate the applied voltage to obtain 0.3 μm, 0.1 μm and 0.03 μm pulls (see materials and equipment section for respective voltage jumps applied).
• 15.
  Set up the pCLAMP software for electrophysiological recordings.

  • a.
    Install pCLAMP 10 (Clampex and Clampfit software) on your PC.
  • b.
    Connect Clampex 10 to the analog-digital converter (Configure → Digitizer).
  • c.
    Create and save protocols in Clampex 10 for the simultaneous control of the piezo element and the recording electrode (Figure 7B; Acquire → New Protocol).

    Note: Each protocol contains three 500 ms stimulation periods of the respective pull distance. The periods are separated by a 1 s pause. At the end of each protocol a 7 s pause is included to separate different pull lengths (structure of a protocol: 1 s pause, 500 ms pull, 1 s pause, 500 ms pull, 1 s pause, 500 ms pull, 7 s pause) (Figure 7A).

    • i.
      Generate protocols for five different pull lengths, 0.03 μm, 0.1 μm, 0.3 μm, 1 μm and 3 μm.
    • ii.
      For each protocol, set the sampling rate to 10 kHz and the sweep duration to 10.5 s.
    • iii.
      Generate six steps for each protocol (use the tab ‘Waveform’). Set the duration and voltage of the first, third and fifth step to 1 s and 0 V, respectively. For the second, fourth and sixth step, set the duration to 500 ms and the voltage according to the pull length (see substep 14, Figure 7B).
  • d.
    Program the automatic sequential start of protocols using the sequencing keys option, beginning with the 0.03 μm pull followed by the next larger pull length (Figure 8).

    • i.
      Go to: Sequencing keys → Add → assign a key (e.g. Alt+Shift+1).
    • ii.
      Set the Protocol Action option to ‘record’, the Protocol File option to the path of the appropriate protocol for the specific pull length and the Repetition Count to 1.
    • iii.
      In the Sequencing tab, set the Next key option to the key with the protocol of the next longer pull length (e.g. Alt+Shift+2) and the After option to ‘minimum’.
    • iv.
      Assign a key to all five protocols. For the last protocol (3 μm pull) set the next key option to ‘None’.

Figure 3.

Stimulation hook coupled to the piezo element

(A) The piezo element is attached to a micropipette with a custom-made acrylic tube (indicated by the arrow) and a screw (indicated by an arrowhead). The hook is glued to the micropipette.

(B) Microscopic image of the hook. Compared to the thickness of the chordotonal organ, the hook is still quite large. Nevertheless, stimulation is very reliable. Scale: 20 μm.

Figure 4.

Sylgard pad-stabilizing clamp

(A) A magnet (indicated by the arrow) is glued to a cable tie.

(B) The clamp prevents floating of the Sylgard pad.

Figure 5.

Arrangement of micromanipulators

(A and C) The piezo-element attached to the hook (A) and the recording electrode (C) are installed in a 30° angle to the horizontal plane.

(B) The hook and recording electrode are arranged in a 90° angle to each other.

Figure 10.

Setup for electrophysiological measurements

(A and B) Setup of the instruments for the electrophysiological recordings. Micromanipulators holding the recording electrode and hook are set up at a 90° angle to each other. A clamp fixes the Sylgard pad. The bath electrode is immersed in the recording solution.

(C) Schematic depiction of lch5 organ. Figure reprinted and adapted with permission from Bormann et al., 2025.¹

(D) Placement of micropipette and hook at the lch5 organ. Scale: 20 μm.

Figure 6.

Connectome of electrophysiological setup devices

The Clampex software (version 10.2) is installed on the Windows 10-operated computer, which is connected to the analog-digital converter via USB. The analog-digital converter receives input from the recording amplifier, which is linked to the recording electrode through the headstage. The headstage is also connected to the bath electrode. The output signal from the analog-digital converter is split and directed to the piezo amplifier and back to the analog-digital converter in order to monitor the applied voltage jumps. The piezo amplifier is connected to the piezo element.

Figure 7.

Stimulation protocol

(A) Schematic representation of the stimulation protocol (top) and a simplified depiction of the expected neuronal response elicited by mechanical stimulation (bottom).

(B) A protocol is saved in Clampex for each deflection length. Used parameters are indicated. The ‘First level (V)’ needs to be adjusted to the respective deflection length (see materials and equipment section). Note: The 1 s interval prior to the first stimulation is set to 911 ms in each protocol due to a delay in the start of the protocol.

Figure 8.

Arrangement of sequencing keys for the Clampex protocols

Parameters for the automatic sequential start of Clampex protocols.

Drosophila crosses

Timing: 15 min

• 16.
  Handling of Drosophila:

  • a.
    Set up crosses with 20 virgin female and 10 male flies in a vial containing standard cornmeal food.
  • b.
    Transfer to a new vial every other day.
  • c.
    Keep the vials at 25°C and a 12:12 h light-dark cycle.
  • d.
    Renew the crosses every week.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Chemicals, peptides, and recombinant proteins
NaCl	Merck	Cat#106404
KCl	Merck	Cat#104936
MgCl2 x 6H2O	Merck	Cat#105833
TES	Sigma-Aldrich	Cat#T6541
Sucrose	Sigma-Aldrich	Cat#S9378
D-(+)-glucose	Sigma-Aldrich	Cat#G7528
D-(+)-trehalose	Sigma-Aldrich	Cat#T9531
NaHCO3	Sigma-Aldrich	Cat#S6297
NaH2PO4	Sigma-Aldrich	Cat#71507
CaCl2	Merck	Cat#102382
HCl	Carl Roth	Cat#6331.3
Ethanol (70%)	N/A	N/A
Experimental models: Organisms/strains
Drosophila melanogaster, 3rd instar larvae (male or female), applicable to various genotypes	N/A	N/A
Software and algorithms
pClamp	Molecular Devices	RRID:SCR_011323
Prism v.10	GraphPad Software, Inc	RRID:SCR_002798
Hokawo v.3	Hamamatsu Photonics	N/A
Other
Minutien pins	Fine Science Tools	Cat#26002-10
Petri dish, 55 mm diameter	Greiner Bio-One	N/A
Vannas-Tübingen spring scissors	Fine Science Tools	Cat#15005-08
Dumont forceps #4	Fine Science Tools	Cat#11241-30 (Dumoxel)
Dumont forceps #5	Fine Science Tools	Cat#11251-20 (Inox)
Dumont forceps #55	Fine Science Tools	Cat#11255-20 (Inox-Biology)
Bath chamber (μ-Dish 35 mm, low)	ibidi	Cat#80136
SYLGARD 184	Dow Corning	Cat#1673921
Dissection microscope	Leica Microsystems	S8 APO
Electrophysiology microscope	Leica Microsystems	DM6 FS
Table Vision IsoStation	Newport	04SI37545
Axopatch 200B recording amplifier (with headstage CV 203BU)	Molecular Devices	N/A
Axon Digidata 1550B analog-digital converter	Molecular Devices	N/A
Micromanipulator system	Sutter Instrument Company	Cat#MPC-385
Micromanipulator	NARISHIGE Group	Cat#NMN-25
Piezo element	Physik Instrumente	Cat#P-840.30
Piezo amplifier	Physik Instrumente	Cat#E-663.00
DMZ-universal electrode puller	Zeitz-Instruments	N/A
Borosilicate glass capillaries	Science Products	Cat#GB150-8P
Ag/AgCl bath electrode (0.5 mm)	self-fabricated from Merck silver wire	Cat#265586
Ag/AgCl recording electrode (0.25 mm)		Cat#327034
10x dry objective	Leica Microsystems	Cat#506505
63x dip-in objective	Leica Microsystems	Cat#506148
Elastic buffers (Bumpon SJ5302)	3M	N/A
Plate (Acrylglas XT 3 mm transparent, round; 16 cm diameter)	Nordic Panel GmbH	Cat#C10000-AC03
Surgical instrument oil	Fine Science Tools	Cat#29055-00
Grindstone/fine sharpening stone	Fine Science Tools	Cat#29008-01
Chlorinator CHG1	AD-Elektronik	Cat#CHG1
Syringe (50 ml) with silicone tubing (2 mm outer diameter)	N/A	N/A
Plastic screw to attach hook to piezo element	N/A	N/A
Short acrylic tube to attach hook to piezo element	N/A	N/A
Super glue	Pattex, Henkel	Cat#2804584
pH meter	Knick	Cat#766
Beakers to prepare solutions	N/A	N/A
Volumetric flasks to prepare solutions	N/A	N/A
Black paper	N/A	N/A
Scalpel	N/A	N/A
Fine side cutter	N/A	N/A
Cables for grounding electrophysiology setup	N/A	N/A
BNC cables to connect electronic devices	N/A	N/A
Magnet (diameter 7 mm)	N/A	N/A
Cable tie	N/A	N/A
Windows computer	N/A	N/A
ORCA-flash4.0 sCMOS	Hamamatsu Photonics	Cat#C13440
Scale	Kern	AEJ 200-5CM

Materials and equipment

Recipe for dissection solution according to Zhang et al., 2013⁸

Reagent	Final concentration	Amount
NaCl	103 mM	3.010 g
KCl	3 mM	0.112 g
MgCl2	4 mM	0.407 g
TES	5 mM	0.573 g
Sucrose	7 mM	1.198 g
Glucose	10 mM	0.901 g
Trehalose	10 mM	1.892 g
NaHCO3	26 mM	1.092 g
NaH2PO4	1 mM	0.069 g
Milli-Q water	–	fill up to 500 ml

Adjust pH to 7.25 with 1 M HCl.

Note: Store at 4°C for up to five days.

Parameters of the DMZ Universal puller for obtaining the micropipettes

	H	F(TH)	s(TH)	t(H)	s(H)	t(F1)	F1	s(F2)	F2	AD
P(A)	800	016	023∗	010	030	000	000	000	000	021
P(B)	027∗	031∗	040	006	000	049	015	003	010	520

∗Need to be adjusted depending on the condition of the heater filament, room temperature and other external parameters.

Voltage jumps applied to the piezo amplifier and the respective pull lengths of the hook

Values in Clampex protocol	−0.013 V	−0.043 V	−0.13 V	−0.42 V	−1 V
Applied voltage jump	−0.13 V	−0.43 V	−1.3 V	−4.2 V	−10 V
Pull length	0.03 μm	0.1 μm	0.3 μm	1 μm	3 μm

Values that need to be set in the Clampex protocols are 10x lower than the resulting voltage jumps due to the 10x gain of the piezo amplifier.

Step-by-step method details

Each of the following steps are critical for the successful outcome of the recording, which is why each step should be carried out as described.

Dissection of Drosophila larvae to expose the chordotonal organ

Timing: 5–10 min per larva

This section describes the dissection of Drosophila larvae to enable access of the recording electrode and stimulation hook to the lch5 organ (Methods videos S1 and S2).

• 1.
  Select non-wandering third-instar larvae (6–7 days after egg laying).

  Note: Due to possible sex-specific differences, we only use male larvae.

  • a.
    Place a larva in the petri dish containing the Sylgard pad.
  • b.
    Add ice-cold, Ca²⁺-free dissection solution (∼15 ml).

    Note: Using a large volume of dissection solution is advisable to prevent warming up and the resulting movement of the larva.

• 2.
  Dissection:

  Note: The dissection is carried out using three different forceps and fine scissors.

  • a.
    Fix the larva on the Sylgard pad with the dorsal side upwards using one pin near the posterior and one near the anterior end (Figure 9A).
  • b.
    Perform small transverse incisions at both ends and then carefully cut through the dorsal midline between the main tracheae (Figure 9B).

    Note: Be careful not to injure the innards.

  • c.
    Open up the body wall, gently stretch it to the sides and fix the body wall with four more pins (Figure 9C).
  • d.
    Sever tracheal and nervous connections to the body wall:

    • i.
      Cut the intestine close to the anus using scissors.
    • ii.
      Gently remove the innards and the brain (Figure 9D) without touching the body wall muscles (see Methods video S1).
  • e.
    Expose the lch5 organ of the third abdominal hemisegment (see Methods video S2):

    Note: We always dissect the hemisegment on the right-hand side because of the orientation of the micromanipulators of our electrophysiology setup. The left hemisegments could otherwise be recorded from as well.

    Note: Instead of the third abdominal hemisegment also other abdominal hemisegments can be used, however, we recommend to always use the same hemisegment to reduce variability.

    • i.
      Cut the layer of the superficial longitudinal muscles (Figures 9G and 9H).
    • ii.
      Remove the remaining anterior parts of the muscles to enable easier access of the scissor to the lch5 nerve bundle and muscles 21–23.
    • iii.
      Cut the nerve bundle close to the anterior, ventral corner of muscle 21 (Figure 9J).
    • iv.
      Cut muscles 21–23 (Figure 9K).

      Note: Avoid damaging the lch5 organ by keeping the lower blade of the scissors away from the organ.

      Note: When cutting muscles 21–23, start cutting closer to the dorsal side of muscle 21 as it might help to prevent touching the lch5 organ.

    • v.
      Remove any remaining parts of muscles 21–23 that may obstruct access of the stimulation hook to the cap cells (Figure 9M).

Figure 9.

Dissection of larvae for extracellular recordings from lch5 neurons

(A) A third-instar larva is pinned (blue asterisks) to the Sylgard pad. Small incisions are made at the anterior and posterior end.

(B) A longitudinal cut between the main tracheae connects the posterior and anterior incisions.

(C) The larval body wall is opened to the sides.

(D) The internal organs are removed. Scale: 500 μm.

(E and F) Confocal image of lch5 organ expressing GFP. The GFP signal helps to visualize the organs and can be used for practicing dissection. (E) Scale: 500 μm. (F) Arrowhead points to the lch5 organ. Scale: 100 μm.

(G, I, and L) Schematic depiction of body wall muscles of an abdominal hemisegment. Muscles which are cut in the respective step are marked in blue.

(H, J, K, M, and N) Images of the steps of the dissection of the third abdominal hemisegment. (G and H) Longitudinal muscles are cut, and the remaining parts at the anterior end are removed. (I, J, and K) The lch5 nerve bundle (black line) is severed close to the ventral side of muscle 21 (J). Muscles 21, 22 and 23 are cut (K). (L and M) The remaining parts of muscles 21, 22 and 23, which might impede the placement of the stimulation hook, are removed. (N) Finished dissection. Scale: 100 μm. (Related to Methods videos S1 and S2).

Electrophysiological recordings from lch5 neurons

Timing: 10 min per larva

This section outlines the steps for electrophysiological recordings of action currents of the lch5 nerve bundle during mechanical stimulation with a hook that displaces the cap cells of the lch5 organs at minuscule distances.

• 3.Fill the recording chamber with 2 mM CaCl₂-containing recording solution (3.5 ml).

Note: The room temperature should be between 19°C and 23°C.

• 4.
  Transfer the Sylgard pad with the dissected larva to the recording chamber.

  • a.
    Turn the chamber to align the recording electrode in parallel with the lch5 organ and the hook perpendicular to the lch5 organ.
  • b.
    Fix the Sylgard pad with the clamp (Figures 10A and 10B).
• 5.Move the recording electrode into the recording solution using the micromanipulator.
• 6.Suck recording solution into the micropipette of the recording electrode to immerse the Ag/AgCl electrode in the solution (∼0.5–1 cm of the Ag/AgCl electrode should be immersed in solution).
• 7.Place the recording electrode and the stimulation hook near the dissected lch5 organ under visual control using the 10x dry objective.
• 8.Place the bath electrode into the recording chamber.
• 9.Change to the 63x dip-in objective. Visually confirm the structural integrity of the lch5 organ (troubleshooting 1).

Optional: Instead of a 63x dip-in objective also a dip-in objective with a smaller magnification can be used. However, we recommend to use a magnification not lower than 40x.

• 10.Gently suck the end of the cut nerve bundle into the recording electrode and move it closer to the lch5 soma while applying suction (Figures 10C and 10D).

CRITICAL: Ensure the lch5 nerve bundle is not overstretched during suction to avoid neuron damage.

• 11.Check the electrophysiological measurement for spontaneous activity of the lch5 neurons and for the quality of the recording.

CRITICAL: If the lch5 neurons are intact, they show irregular spontaneous electrical activity, i.e., action currents (ACurr). The absence of spontaneously occurring ACurr indicates that the animal/organ lost its physiological function. Start again at step 1. Also, check the quality of the measurement, e.g. whether too much electrical noise is present (troubleshooting 2, 3).

• 12.Apply 50 V to the piezo by turning the respective knob of the piezo amplifier to extend the piezo element. Thus, applying negative voltage jumps will lead to the contraction of the piezo and a defined pull length of the hook.
• 13.Place the stimulation hook at the ventral side of the cap cells about 30–40 μm distal to the dendrites (Figures 10C and 10D).

CRITICAL: Make sure to touch, but do not pre-stretch the cap cells with the hook.

• 14.Adjust the pipette offset to ideally ∼0 pA.
• 15.Start the pre-saved Clampex protocols, beginning with the shortest pull and finishing with the longest pull.
• 16.After the measurement, visually check that neither the larva nor the hook drifted during the measurement.
• 17.For each experimental group, recordings are performed from lch5 organs (segment A3) of 10 different animals.

Pause Point: Recordings are saved and can be analyzed later.

Data analysis

Timing: 15–20 min per larva

Analysis of electrical activity recorded from lch5 neurons is performed using Clampfit by manually counting the ACurr number, followed by statistical analysis using software such as Prism (GraphPad) or SigmaPlot (Systat).

• 18.
  Open each file of the measurements separately in Clampfit 10.

  • a.
    For analyzing the spontaneous activity of the lch5, manually count the ACurr within 500 ms prior to the first 0.03 μm pull stimulation.
  • b.
    For mechanically evoked activity, count the ACurr occurring in the first 50 ms of each pull (50 ms ON) and 50 ms after each pull (50 ms OFF) (Figure 11).

Note: Using the cursors in Clampfit helps setting the boundaries of the 500 ms or 50 ms time frame. In Bormann et al. 2025¹ only ‘50 ms ON’ events were analyzed.

• 19.For each condition, i.e., 50 ms ON and 50 ms OFF, calculate the mean of the ACurr number from all three stimulations of the same pull length to obtain N=1.
• 20.To convert the ACurr numbers to ACurr frequency, divide by the time period 0.5 s or 0.05 s for spontaneous and mechanically-evoked activity, respectively.
• 21.Statistical analysis can be carried out to compare different experimental groups (e.g. different genotypes) or to compare data points within a group using Prism (GraphPad) or other programs.

Figure 11.

Exemplary trace of electrical activity recorded from larval lch5 neurons

The top line represents the stimulation protocol containing three pulls displayed as rectangles. The representative electrophysiological recording shows the response of lch5 neurons to the stimulation with 30 nm pulls in a w¹¹¹⁸ larva. For analysis of spontaneous activity, the action currents in the 500 ms (grey box) before the first stimulation with the 30 nm pull are counted. Mechanically-evoked activity is assessed by counting the action currents in the first 50 ms of each pull (ON, blue box) or 50 ms after each pull (OFF, magenta box). The mean ACurr number of all three pulls is calculated to obtain N=1.

Expected outcomes

Pulling at the cap cells mechanically stimulates the lch5 neurons. Lch5 organs are differential sensors, and thus, a neuronal response, i.e., a change in the ACurr frequency, is expected to follow a change in stimulus intensity and to return to baseline levels even when the stimulus pesists (Figure 11). Moreover, it is expected that the ACurr frequency scales with stimulation intensity (30 nm → 3 μm pull length = increase in ACurr).

Limitations

The ACurr frequencies measured in Bormann et al. 2025,¹ follow roughly the Weber-Fechner law, indicating that the pull lengths are within a reasonable window of error. However, the exact displacement of the shorter pulls (0.03, 0.1 and 0.3 μm) could not be confirmed visually.

We measure extracellularly in voltage clamp mode and thus ACurr amplitude depends on how tightly the micropipette seals the nerve bundle. Therefore, ACurr amplitudes should only be investigated after ensuring reproducible resistance of the ‘loose patch’ (i.e., the fit between the diameter of the recording electrode tip and the nerve). Determining resistance can be done by applying a controlled current, measuring the resulting voltage, and calculating the resistance using Ohm’s law (V = IR, where V is voltage, R is resistance, and I is current). However, this approach was not implemented in the current protocol. The presented electrophysiological measurement can be affected by environmental factors, e.g. noise, vibration or movement of the electrophysiological setup might provoke activity of lch5 neurons. Hence, it is essential to prevent such unspecific/unwanted stimulation.

Troubleshooting

Problem 1

The lch5 organ is not intact, shows no spontaneous activity or shows abnormal activity, i.e., large amplitude ACurr-like currents with a very regular unchanging frequency and amplitude (step 2, substep 11).

Potential solution

• •Sharpen the scissors. When cutting the muscles, keep the blades of the scissors away from the lch5 organ to prevent contact. Aim to cut muscles 21–23 closer to the dorsal end of the muscles.
• •Make sure to not touch the body wall when removing the innards from the opened larvae.
• •For learning the dissection, it is helpful to express a fluorophore in the lch5 neurons to ease visualization of the nerve bundle. This can be done using any binary expression system in combination with driver lines expressing a reporter in lch5 neurons, e.g. 20xUAS-6XmCherry-HA (BDSC#52268), or simply utilizing the GFP signal derived from the second-chromosomal CyOGFPw^- balancer.

Problem 2

It is difficult to distinguish the ACurr from the background noise (step 2, substep 11).

Potential solution

• •Decrease the micropipette tip diameter of the recording electrode to augment the signal (see ‘before you begin’ substep 1).
• •When reusing a recording electrode, background electrical noise might increase, presumably due to the build-up of a salt crust within the electrode. Consider using a new micropipette/recording electrode if noise persists.
• •Ensure that the lch5 nerve bundle is sucked into the micropipette of the recording electrode properly (substep 10) and that the pipette opening is located very close to the soma. A tight fit increases the amplitude of ACurr, thus increasing the signal-to-noise ratio.

Problem 3

High variations of electrical activity are observed in the same genotype (step 2, substep 17 & step 3, substep 18).

Potential solution

• •Keep room temperature between 19°C–23°C.
• •When pinning the body wall to the side during dissection, make sure to only slightly stretch the body wall. The stretch of the body wall should be comparable in every dissection.
• •Touch the cap cells gently with the stimulation hook (substep 13), but be careful not to stretch the lch5 organ with the hook when positioning it at the cap cells.
• •Ensure that no external factors like noise, vibration, or movement stimulate the lch5, thus, distorting the electrophysiological recording.

Resource availability

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Dr. Nicole Scholz (scholzlab@gmail.com).

Technical contact

Technical questions on executing this protocol should be directed to and will be answered by the technical contact, Dr. Dmitrij Ljaschenko (dmitrij.ljaschenko@medizin.uni-leipzig.de).

Materials availability

This study did not generate new unique reagents.

Data and code availability

This study did not generate/analyze datasets/code.

Acknowledgments

We thank Amit Alon for his help in producing the film clip showing the larval fillet preparation. This work was supported by grants from the Deutsche Forschungsgemeinschaft (DFG) to N.S. through FOR2149, project P01 (project number 265903901), and CRC 1423 project B06 (project number 421152132), as well as N.S. and D.L. through Junior research grants from the Medical Faculty, Leipzig University. Due to affiliation with Leipzig University, the open access publication costs were partly covered under the agreement between the DEAL consortium and Elsevier.

Author contributions

M.B.K.: methodology, validation, formal analysis, investigation, visualization, and manuscript preparation. M.M.: visualization and manuscript editing. D.L.: visualization, conceptualization, methodology, supervision, funding acquisition, and manuscript preparation. N.S.: visualization, conceptualization, supervision, project administration, funding acquisition, and manuscript preparation.

Declaration of interests

The authors declare no competing interests.