rus
Issue #5/2025
I.V.Yaminsky
SCANNING PROBE MICROSCOPY IN THE STUDY OF INSECT NEURONAL ACTIVITY: PROSPECTS AND METHODOLOGICAL APPROACHES
SCANNING PROBE MICROSCOPY IN THE STUDY OF INSECT NEURONAL ACTIVITY: PROSPECTS AND METHODOLOGICAL APPROACHES
The paper discusses the possibilities of using scanning probe microscopy (SPM) to study the neural activity of insects, including recording action potentials and visualizing the morphological features of the nervous system. Particular attention is paid to the advantages of using insects such as mosquitoes as model objects for neurophysiological research. Modern methods of recording neural impulses are discussed, including intracellular and extracellular recording, the local potential fixation method (patch clamp), as well as a comparison of action potential parameters in insects and humans. The prospects for integrating SPM with optogenetics and calcium imaging to obtain a comprehensive understanding of the functional activity of neurons in vivo are presented.
INTRODUCTION
The study of nervous system functional activity is one of the most important tasks in modern neurobiology. Development of neurophysiology methods and scanning probe microscopy opens new opportunities for analyzing electrical activity of individual neurons, neural networks, and the brain as a whole. However, studies of neural activity in higher organisms face a number of limitations related to both technical challenges and ethical aspects. This makes the use of invertebrate arthropods – insects – particularly attractive in experimental work as model systems (Fig.1).
Insects have a compact yet functionally efficient nervous system, which allows for high-resolution experiments provided by probe microscopy and minimal ethical constraints. Due to the ease of fixing insect neurons, they serve as a convenient subject for studying the generation mechanisms and nerve impulses transmission. Combination of traditional electrophysiological methods with the capabilities of scanning probe microscopy, which allows for obtaining high-precision topographic data without damaging the sample, appears particularly promising.
Scanning probe microscopy allows for obtaining three-dimensional images with nanometer resolution, which is particularly important for studying various morphological structures of insects, for example: cuticles (chitinous coverings) [1], sensilla (sensory hairs), nanostructures affecting hydrophobicity and aerodynamics of wings, and adhesion devices of legs. A series of works was carried out to study the faceted eye of insects and its structural unit – the ommatidium using atomic force microscopy methods [2].
In 2024, the results of mapping the brain of the fruit fly, obtained through electron microscopy, were published [3]. Researchers completely mapped the connections between neurons throughout the brain of the adult fruit fly. The work presented a complete picture of the structure of the adult female Drosophila melanogaster brain, containing 5 ∙ 107 chemical synapses between 139,255 neurons. With the significant capabilities achieved in electron microscopy, there remains no possibility of working with live specimens.
The only methods that allow us to study living objects with nanometer resolution in air or in liquids are the tools of scanning probe microscopy. However, to date, we have not found experimental results from probe microscopy regarding the study of the functional activity of the brain of a living being.
Before we take a closer look at the possibilities for conducting such insect studies using the mosquito as an example, we try to examine the similarities and differences between a human brain and a mosquito’s brain.
In humans, the maximum frequency of neural impulses is about 200–500 Hz (in some sensory neurons), but on average it is 10–100 Hz.
In insects, nerve cells are often larger and faster than in mammals, which compensates for their small number.
The total number of signals in an insect’s brain can be estimated as follows. Taking into account that a mosquito has about 200,000 to 400,000 neurons (for comparison: a fly has about 200,000, a bee has about 1 million), and each can generate dozens of impulses per second, then the total number of nerve signals in its brain can reach millions per second. The nervous system of insects is optimized for quick reactions, and although their brains are small, they process information very efficiently. On average: a single neuron fires 50–200 impulses/second. The entire brain processes millions of signals per second.
How do the nervous impulses of a mosquito’s brain differ from those of a human? Nervous impulses (action potentials) in mosquitoes and humans operate on similar biophysical principles, but there are key differences in speed, organization, and functionality.
Main differences:
The impulse transmission speed in mosquitoes and insects is slower.
In invertebrates, there are no myelinated fibers (like in vertebrates), so the transmission speed is approximately 1–5 m/s (in thick axons). The frequency of impulses in individual neurons can reach up to 200–500 Hz (in sensory neurons at peak activity);
In humans, the speed of nerve signal transmission occurs faster. Myelinated fibers transmit signals at speeds of up to 120 m/s. The frequency of impulses can reach 500–1000 Hz (in some neurons), but on average it is 10–200 Hz.
Organization of the nervous system
Mosquito:
The decentralized system consists of the brain (supraesophageal ganglion) + the ventral nerve cord with ganglia. The mosquito’s brain is made up of several divisions:
Protocerebrum – is responsible for processing visual information (coming from the compound eyes) and complex behavioral responses;
Deitocerabrum – processes signals from the antennas (organs of smell, touch, and thermoception);
The tritocerebrum is connected to the subesophageal ganglion and controls the mouthparts.
In addition, mosquitoes have an abdominal nerve chain with ganglia that control the movements of their limbs and wings.
In insects, there are fewer neurons (~200,000–400,000), but a high efficiency is manifested in simple reactions (searching for prey, avoiding threats).
In a human:
A centralized brain (86 billion neurons) with a hierarchical structure (cortex, subcortical nuclei, etc.). Complex cognitive functions (memory, learning, abstract thinking).
Types of neurons and synaptic transmission
Mosquitoes: Neurotransmitters: acetylcholine, glutamate, GABA (as in humans, but receptors may differ). Less synaptic plasticity – learning is limited (e.g. mosquitoes can remember dangerous odours but not complex associations).
Human: More complex neurotransmitter systems (dopamine, serotonin, noradrenaline – regulate emotions, motivation). High synaptic plasticity – formation of long-term memory, complex neural networks.
Energy efficiency
Mosquitoes: Nervous system consumes minimal energy as it is optimised for survival. Rapid response to key stimuli (CO2, heat, blood odour).
Human: Brain consumes ~20% of body energy as it supports complex computations.
Sensitivity to neurotoxins
Mosquitoes: Their neurons are more resistant to some poisons (e.g. nicotine) but sensitive to insecticides (pyrethroids block sodium channels).
Humans: Neurons are more vulnerable to neurotoxins (mercury, lead, botulinum toxin).
DISCUSSION
Mosquito and human nerve impulses are similar at the cellular level, but differ in speed, organisation and complexity of information processing. Mosquitoes are fast, energy efficient, but primitive reactions. Humans are slower in individual neurons, but have complex thinking and adaptive behaviour.
In brief: a mosquito is a "microcomputer" for finding blood, while a human is a "supercomputer" with advanced intelligence.
How do nerve impulses in the mosquito brain differ from human nerve impulses? Nerve impulses (action potentials) in mosquitoes and humans operate on similar biophysical principles, but there are key differences in speed, organisation and functionality. The key differences are:
The action potential (AP) in mosquitoes and humans is based on the same fundamental principles (changes in the ionic permeability of the neuron membrane), but there are key differences in the parameters. A detailed comparison is shown in Table 1.
Key Differences:
Ionuc crystals
Mosquitoes: Sodium (Na⁺) and potassium (K⁺) channels are less selective than in mammals. Some channels are resistant to tetrodotoxin (TTX), which blocks AP in vertebrates.
Human: Channels are more specialised (e.g. fast Naᵥ1.1-Naᵥ1.9). Sensitive to TTX and other neurotoxins.
Action potential transit velocity
Mosquitoes: No myelination, so, speed 1–5 m/s (up to 10 m/s in thick axons, e.g. in giant cockroach neurons).
Human: Myelinated fibres, so, speed up to 120 m/s (e.g. motor neurons).
Energy assumption
Mosquitoes: AP requires less energy (ATP) because ionic gradients are recovered more slowly.
Human: Up to 60% of neuronal energy is spent to maintain Na⁺/K⁺-ATPase.
Temperature dependence
Mosquitoes: AP is possible at lower temperatures (some insects are active at +5 °C).
Humans: When cooling below +20 °C, pulse conduction is impaired.
What accounts for these differences? In insects, as a result of evolutionary adaptation, the AP is optimised to minimise energy for a small body size. In humans, it is optimised for speed and precision in a complex nervous system.
Neuronal size: Mosquito axons are shorter, hence no myelination is needed. Note that some insects (e.g. grasshoppers) have giant axons (up to 50 µm in diameter) that conduct signals at speeds of up to 25 m/s – a record among invertebrates!
The following conclusions can be drawn:
The action potential in mosquito and human is similar in mechanism but different in mosquito:
Smaller amplitude (by 10–20 mV);
Longer PD (1.5–2 times);
Resistance to toxins (e.g., TTX);
Slower conduction due to lack of myelin.
These features reflect a general principle: the insect nervous system sacrifices "precision" for energy efficiency and compactness.
REGISTRATION METHODS
How to experimentally record the nerve impulse potential of the mosquito brain? Experimental recording of action potentials (APs) in the mosquito nervous system is a difficult task due to the small size of neurons (often less than 5 µm in diameter). However, modern neurophysiological methods make it possible.
The main approaches are:
Intracellular recording
How it works. Uses a glass microelectrode (tip diameter <1 µm) filled with electrolyte (KCl) that is injected directly into the neuron.
Measures changes in membrane potential with high accuracy (down to 0.1 mV).
Application to mosquito: Suitable for large neurons (e.g. in the abdominal nerve chain or optic ganglia).
Difficulties:
The mosquito’s neurons are easily damaged.
Requires fixation of the insect (often with wax or special glue).
Extracellular recording
How it works. An electrode is placed next to a neuron or axon, recording extracellular currents.
Methods: Succinylcholine electrodes – for recording antennal nerve activity.
Microcontact arrays – for simultaneous recording from multiple neurons.
Examples: Recording the response of olfactory neurons to CO2 and blood odours.
Study of neuronal activity in ganglia of the ventral nerve chain.
Advantages: Less invasive method. Allows for long-term recordings.
Patch-clamp
How it works. A microelectrode is tightly adhered to the membrane of a neuron, recording ionic currents through individual channels.
Options: Cell-attached is recording without damaging the cell. Whole-cell means full access to intracellular environment.
Application: Study of properties of sodium and potassium channels in mosquito neurons.
Example: Study of resistance to insecticides (e.g. pyrethroids).
Limitations: Requires isolation of individual neurons (e.g. from cultured larval brain cells).
Optogenetics and calcium imaging
How it works. Calcium dyes (e.g. GCaMP) – fluoresce when bound to Ca²⁺, which enters the neuron during AP.
Optogenetics – introduction of light-sensitive ion channels (Channelrhodopsin) to activate neurons by light. Examples: Real-time visualisation of olfactory neuron activity. Control of wing motor neurons in genetically modified mosquitoes. Pros: Does not require mechanical contact with the neuron. Allows for the study of neural networks.
Scanning probe microscopy
Scanning probe microscopy (SPM) is a unique tool for studying structure and functional activity of biological objects at the nanoscale. Unlike electron microscopy, which allows obtaining high-resolution images but requires harsh conditions (vacuum, metallisation), SPM works under ambient conditions – in air or in liquid, which makes it particularly useful when working with living samples.
One of the most well-known representatives of SPM is atomic force microscopy (AFM). It allows not only to obtain three-dimensional images of cell and tissue surfaces with nanometre resolution, but also to record mechanical properties of the surface, such as stiffness, adhesion and viscoelasticity. These parameters can be related to changes in state of the neuron membrane during the action potential.
SPM opportunities:
obtaining topographic maps of neurons and axons;
studying morphological changes during neuron excitation;
registration of local mechanical oscillations associated with pulse generation;
combination with optical methods (e.g. fluorescence microscopy);
study of dynamic processes in real time.
Sample preparation for scanning probe microscopy: Immobilisation of the insect: Chilling or CO2-narcosis. Fixation in a special holder (e.g. wax substrate);
Access to the nervous system:
For recording from the brain, opening the head capsule. For peripheral neurons – exposure of nerves (e.g. antennal nerve).
Example of an experiment (AP registration in the antennal nerve):
Fix the mosquito, open the antennal segment. Bring the extracellular electrode to the nerve. Apply a stimulus (blood odour or heat). Record bursts of activity.
Such methods help to study how mosquitoes find prey by smell and why insecticides stop working, as well as how the insects’ "brain" works.
With scanning probe microscopy, you can start with extracellular recording – this is the easiest way to "hear" the mosquito’s nerve impulses! When measuring an action potential, one electrode (atomic force microscope probe) touches the neuron, but where is the second electrode placed? In action potential (AP) recording, the second electrode plays a critical role – it serves as a reference (ground) electrode and closes the electrical circuit. It is placed outside the cell but close to the neuron (e.g., in insect haemolymph or physiological solution). Chlorinated silver electrode (Ag/AgCl) is often used to minimise interference. Important: Both electrodes must be in the same ionic environment to avoid diffusion potentials.
Control experiments. To exclude artefacts check that the signal disappears when the active electrode is removed from the neuron, then make sure that the reference electrode does not create noise (e.g. using low impedance electrodes).
Why is this important? Without a reference electrode it is not possible to measure the potential difference – the circuit will remain open. Incorrect placement of the reference adds noise (e.g. due to differences in the ionic composition of the media). The analogue is as in a voltmeter – one probe is connected to the measuring point, the other to the ground potential. The right choice of reference electrode is the key to clear AP registration in microscopic mosquito neurons!
To maintain cells viability and tissues of insects outside the body, a modified Ringer’s solution adapted to their haemolymph is used. Approximate composition (in mM):
NaCl – 120–150 (main osmotic component);
KCl – 3–5 (membrane potential maintenance);
CaCl2 – 1–2 (for membrane stabilisation and synaptic transmission);
MgCl2 – 1–4 (enzyme cofactor);
Glucose – 5–10 (energy source);
Buffer (HEPES or Tris) – 5–10 (sustaining pH on 7.0–7.4 level).
The use of Ringer’s solution is essential for scanning capillary microscopy when surface imaging is performed in an electrolyte that conducts an electric current.
Observation of the Drosophila eye using scanning capillary microscopy was carried out in the group of Yu.Korchev [4]. It was shown that capillary microscopy allows obtaining topographic data without damaging the specimen, but with lower resolution than electron microscopy. Electron microscopy gives clearer images of nanostructures (e.g. ommatidia of the eye), but requires vacuum and metallisation. Capillary microscopy is better suited for dynamic studies of living cells. Significantly, scanning capillary microscopy allows obtaining three-dimensional images of neurons and maps of local stiffness distribution of their membrane. Additionally, the aforementioned patch-clamp, extracellular and intracellular recordings of nerve signals can be performed with the capillary microscope.
CONCLUSIONS
Based on the presented data, it is possible to note significant methodological advantages of neurophysiological studies using insects. In particular, it is their high availability and compact size, which is important for scanning probe microscopy. Insects can be induced into an immobile state – anabiosis – by reducing the temperature or increasing the concentration of carbon dioxide. This also greatly simplifies the experiment. The percentage ratio of the number of neurons and other cells in insects is much higher than in humans, which gives certain advantages in studies with atomic force microscopy. Unlike other methods – patch-clamp, optogenetics, electron microscopy – probe microscopy makes it possible to visualise the insect brain in 3D with nanometre resolution in a natural environment. Thus, combining electrophysiological methods with the capabilities of scanning probe microscopy allows:
obtain information on structure and function of neurons;
study dynamics of action potentials;
study the influence of external factors on the passage of nerve impulses (temperature, chemicals, insecticides);
develop new approaches to insect population management and biosensors.
This research is not only of fundamental importance but also has practical applications, from gaining new insights to developing neural interfaces based on biological systems.
ACKNOWLEDGEMENTS
The study was conducted mainly on an initiative basis. This work was partially performed under the state order of the Lomonosov Moscow State University. FemtoScan Online software is provided by Advanced Technologies Center, www.femtoscan.ru
PEER REVIEW INFO
Editorial board thanks the anonymous reviewer(s) for their contribution to the peer review of this work. It is also grateful for their consent to publish papers on the journal’s website and SEL eLibrary eLIBRARY.RU.
Declaration of Competing Interest. The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
The study of nervous system functional activity is one of the most important tasks in modern neurobiology. Development of neurophysiology methods and scanning probe microscopy opens new opportunities for analyzing electrical activity of individual neurons, neural networks, and the brain as a whole. However, studies of neural activity in higher organisms face a number of limitations related to both technical challenges and ethical aspects. This makes the use of invertebrate arthropods – insects – particularly attractive in experimental work as model systems (Fig.1).
Insects have a compact yet functionally efficient nervous system, which allows for high-resolution experiments provided by probe microscopy and minimal ethical constraints. Due to the ease of fixing insect neurons, they serve as a convenient subject for studying the generation mechanisms and nerve impulses transmission. Combination of traditional electrophysiological methods with the capabilities of scanning probe microscopy, which allows for obtaining high-precision topographic data without damaging the sample, appears particularly promising.
Scanning probe microscopy allows for obtaining three-dimensional images with nanometer resolution, which is particularly important for studying various morphological structures of insects, for example: cuticles (chitinous coverings) [1], sensilla (sensory hairs), nanostructures affecting hydrophobicity and aerodynamics of wings, and adhesion devices of legs. A series of works was carried out to study the faceted eye of insects and its structural unit – the ommatidium using atomic force microscopy methods [2].
In 2024, the results of mapping the brain of the fruit fly, obtained through electron microscopy, were published [3]. Researchers completely mapped the connections between neurons throughout the brain of the adult fruit fly. The work presented a complete picture of the structure of the adult female Drosophila melanogaster brain, containing 5 ∙ 107 chemical synapses between 139,255 neurons. With the significant capabilities achieved in electron microscopy, there remains no possibility of working with live specimens.
The only methods that allow us to study living objects with nanometer resolution in air or in liquids are the tools of scanning probe microscopy. However, to date, we have not found experimental results from probe microscopy regarding the study of the functional activity of the brain of a living being.
Before we take a closer look at the possibilities for conducting such insect studies using the mosquito as an example, we try to examine the similarities and differences between a human brain and a mosquito’s brain.
In humans, the maximum frequency of neural impulses is about 200–500 Hz (in some sensory neurons), but on average it is 10–100 Hz.
In insects, nerve cells are often larger and faster than in mammals, which compensates for their small number.
The total number of signals in an insect’s brain can be estimated as follows. Taking into account that a mosquito has about 200,000 to 400,000 neurons (for comparison: a fly has about 200,000, a bee has about 1 million), and each can generate dozens of impulses per second, then the total number of nerve signals in its brain can reach millions per second. The nervous system of insects is optimized for quick reactions, and although their brains are small, they process information very efficiently. On average: a single neuron fires 50–200 impulses/second. The entire brain processes millions of signals per second.
How do the nervous impulses of a mosquito’s brain differ from those of a human? Nervous impulses (action potentials) in mosquitoes and humans operate on similar biophysical principles, but there are key differences in speed, organization, and functionality.
Main differences:
The impulse transmission speed in mosquitoes and insects is slower.
In invertebrates, there are no myelinated fibers (like in vertebrates), so the transmission speed is approximately 1–5 m/s (in thick axons). The frequency of impulses in individual neurons can reach up to 200–500 Hz (in sensory neurons at peak activity);
In humans, the speed of nerve signal transmission occurs faster. Myelinated fibers transmit signals at speeds of up to 120 m/s. The frequency of impulses can reach 500–1000 Hz (in some neurons), but on average it is 10–200 Hz.
Organization of the nervous system
Mosquito:
The decentralized system consists of the brain (supraesophageal ganglion) + the ventral nerve cord with ganglia. The mosquito’s brain is made up of several divisions:
Protocerebrum – is responsible for processing visual information (coming from the compound eyes) and complex behavioral responses;
Deitocerabrum – processes signals from the antennas (organs of smell, touch, and thermoception);
The tritocerebrum is connected to the subesophageal ganglion and controls the mouthparts.
In addition, mosquitoes have an abdominal nerve chain with ganglia that control the movements of their limbs and wings.
In insects, there are fewer neurons (~200,000–400,000), but a high efficiency is manifested in simple reactions (searching for prey, avoiding threats).
In a human:
A centralized brain (86 billion neurons) with a hierarchical structure (cortex, subcortical nuclei, etc.). Complex cognitive functions (memory, learning, abstract thinking).
Types of neurons and synaptic transmission
Mosquitoes: Neurotransmitters: acetylcholine, glutamate, GABA (as in humans, but receptors may differ). Less synaptic plasticity – learning is limited (e.g. mosquitoes can remember dangerous odours but not complex associations).
Human: More complex neurotransmitter systems (dopamine, serotonin, noradrenaline – regulate emotions, motivation). High synaptic plasticity – formation of long-term memory, complex neural networks.
Energy efficiency
Mosquitoes: Nervous system consumes minimal energy as it is optimised for survival. Rapid response to key stimuli (CO2, heat, blood odour).
Human: Brain consumes ~20% of body energy as it supports complex computations.
Sensitivity to neurotoxins
Mosquitoes: Their neurons are more resistant to some poisons (e.g. nicotine) but sensitive to insecticides (pyrethroids block sodium channels).
Humans: Neurons are more vulnerable to neurotoxins (mercury, lead, botulinum toxin).
DISCUSSION
Mosquito and human nerve impulses are similar at the cellular level, but differ in speed, organisation and complexity of information processing. Mosquitoes are fast, energy efficient, but primitive reactions. Humans are slower in individual neurons, but have complex thinking and adaptive behaviour.
In brief: a mosquito is a "microcomputer" for finding blood, while a human is a "supercomputer" with advanced intelligence.
How do nerve impulses in the mosquito brain differ from human nerve impulses? Nerve impulses (action potentials) in mosquitoes and humans operate on similar biophysical principles, but there are key differences in speed, organisation and functionality. The key differences are:
The action potential (AP) in mosquitoes and humans is based on the same fundamental principles (changes in the ionic permeability of the neuron membrane), but there are key differences in the parameters. A detailed comparison is shown in Table 1.
Key Differences:
Ionuc crystals
Mosquitoes: Sodium (Na⁺) and potassium (K⁺) channels are less selective than in mammals. Some channels are resistant to tetrodotoxin (TTX), which blocks AP in vertebrates.
Human: Channels are more specialised (e.g. fast Naᵥ1.1-Naᵥ1.9). Sensitive to TTX and other neurotoxins.
Action potential transit velocity
Mosquitoes: No myelination, so, speed 1–5 m/s (up to 10 m/s in thick axons, e.g. in giant cockroach neurons).
Human: Myelinated fibres, so, speed up to 120 m/s (e.g. motor neurons).
Energy assumption
Mosquitoes: AP requires less energy (ATP) because ionic gradients are recovered more slowly.
Human: Up to 60% of neuronal energy is spent to maintain Na⁺/K⁺-ATPase.
Temperature dependence
Mosquitoes: AP is possible at lower temperatures (some insects are active at +5 °C).
Humans: When cooling below +20 °C, pulse conduction is impaired.
What accounts for these differences? In insects, as a result of evolutionary adaptation, the AP is optimised to minimise energy for a small body size. In humans, it is optimised for speed and precision in a complex nervous system.
Neuronal size: Mosquito axons are shorter, hence no myelination is needed. Note that some insects (e.g. grasshoppers) have giant axons (up to 50 µm in diameter) that conduct signals at speeds of up to 25 m/s – a record among invertebrates!
The following conclusions can be drawn:
The action potential in mosquito and human is similar in mechanism but different in mosquito:
Smaller amplitude (by 10–20 mV);
Longer PD (1.5–2 times);
Resistance to toxins (e.g., TTX);
Slower conduction due to lack of myelin.
These features reflect a general principle: the insect nervous system sacrifices "precision" for energy efficiency and compactness.
REGISTRATION METHODS
How to experimentally record the nerve impulse potential of the mosquito brain? Experimental recording of action potentials (APs) in the mosquito nervous system is a difficult task due to the small size of neurons (often less than 5 µm in diameter). However, modern neurophysiological methods make it possible.
The main approaches are:
Intracellular recording
How it works. Uses a glass microelectrode (tip diameter <1 µm) filled with electrolyte (KCl) that is injected directly into the neuron.
Measures changes in membrane potential with high accuracy (down to 0.1 mV).
Application to mosquito: Suitable for large neurons (e.g. in the abdominal nerve chain or optic ganglia).
Difficulties:
The mosquito’s neurons are easily damaged.
Requires fixation of the insect (often with wax or special glue).
Extracellular recording
How it works. An electrode is placed next to a neuron or axon, recording extracellular currents.
Methods: Succinylcholine electrodes – for recording antennal nerve activity.
Microcontact arrays – for simultaneous recording from multiple neurons.
Examples: Recording the response of olfactory neurons to CO2 and blood odours.
Study of neuronal activity in ganglia of the ventral nerve chain.
Advantages: Less invasive method. Allows for long-term recordings.
Patch-clamp
How it works. A microelectrode is tightly adhered to the membrane of a neuron, recording ionic currents through individual channels.
Options: Cell-attached is recording without damaging the cell. Whole-cell means full access to intracellular environment.
Application: Study of properties of sodium and potassium channels in mosquito neurons.
Example: Study of resistance to insecticides (e.g. pyrethroids).
Limitations: Requires isolation of individual neurons (e.g. from cultured larval brain cells).
Optogenetics and calcium imaging
How it works. Calcium dyes (e.g. GCaMP) – fluoresce when bound to Ca²⁺, which enters the neuron during AP.
Optogenetics – introduction of light-sensitive ion channels (Channelrhodopsin) to activate neurons by light. Examples: Real-time visualisation of olfactory neuron activity. Control of wing motor neurons in genetically modified mosquitoes. Pros: Does not require mechanical contact with the neuron. Allows for the study of neural networks.
Scanning probe microscopy
Scanning probe microscopy (SPM) is a unique tool for studying structure and functional activity of biological objects at the nanoscale. Unlike electron microscopy, which allows obtaining high-resolution images but requires harsh conditions (vacuum, metallisation), SPM works under ambient conditions – in air or in liquid, which makes it particularly useful when working with living samples.
One of the most well-known representatives of SPM is atomic force microscopy (AFM). It allows not only to obtain three-dimensional images of cell and tissue surfaces with nanometre resolution, but also to record mechanical properties of the surface, such as stiffness, adhesion and viscoelasticity. These parameters can be related to changes in state of the neuron membrane during the action potential.
SPM opportunities:
obtaining topographic maps of neurons and axons;
studying morphological changes during neuron excitation;
registration of local mechanical oscillations associated with pulse generation;
combination with optical methods (e.g. fluorescence microscopy);
study of dynamic processes in real time.
Sample preparation for scanning probe microscopy: Immobilisation of the insect: Chilling or CO2-narcosis. Fixation in a special holder (e.g. wax substrate);
Access to the nervous system:
For recording from the brain, opening the head capsule. For peripheral neurons – exposure of nerves (e.g. antennal nerve).
Example of an experiment (AP registration in the antennal nerve):
Fix the mosquito, open the antennal segment. Bring the extracellular electrode to the nerve. Apply a stimulus (blood odour or heat). Record bursts of activity.
Such methods help to study how mosquitoes find prey by smell and why insecticides stop working, as well as how the insects’ "brain" works.
With scanning probe microscopy, you can start with extracellular recording – this is the easiest way to "hear" the mosquito’s nerve impulses! When measuring an action potential, one electrode (atomic force microscope probe) touches the neuron, but where is the second electrode placed? In action potential (AP) recording, the second electrode plays a critical role – it serves as a reference (ground) electrode and closes the electrical circuit. It is placed outside the cell but close to the neuron (e.g., in insect haemolymph or physiological solution). Chlorinated silver electrode (Ag/AgCl) is often used to minimise interference. Important: Both electrodes must be in the same ionic environment to avoid diffusion potentials.
Control experiments. To exclude artefacts check that the signal disappears when the active electrode is removed from the neuron, then make sure that the reference electrode does not create noise (e.g. using low impedance electrodes).
Why is this important? Without a reference electrode it is not possible to measure the potential difference – the circuit will remain open. Incorrect placement of the reference adds noise (e.g. due to differences in the ionic composition of the media). The analogue is as in a voltmeter – one probe is connected to the measuring point, the other to the ground potential. The right choice of reference electrode is the key to clear AP registration in microscopic mosquito neurons!
To maintain cells viability and tissues of insects outside the body, a modified Ringer’s solution adapted to their haemolymph is used. Approximate composition (in mM):
NaCl – 120–150 (main osmotic component);
KCl – 3–5 (membrane potential maintenance);
CaCl2 – 1–2 (for membrane stabilisation and synaptic transmission);
MgCl2 – 1–4 (enzyme cofactor);
Glucose – 5–10 (energy source);
Buffer (HEPES or Tris) – 5–10 (sustaining pH on 7.0–7.4 level).
The use of Ringer’s solution is essential for scanning capillary microscopy when surface imaging is performed in an electrolyte that conducts an electric current.
Observation of the Drosophila eye using scanning capillary microscopy was carried out in the group of Yu.Korchev [4]. It was shown that capillary microscopy allows obtaining topographic data without damaging the specimen, but with lower resolution than electron microscopy. Electron microscopy gives clearer images of nanostructures (e.g. ommatidia of the eye), but requires vacuum and metallisation. Capillary microscopy is better suited for dynamic studies of living cells. Significantly, scanning capillary microscopy allows obtaining three-dimensional images of neurons and maps of local stiffness distribution of their membrane. Additionally, the aforementioned patch-clamp, extracellular and intracellular recordings of nerve signals can be performed with the capillary microscope.
CONCLUSIONS
Based on the presented data, it is possible to note significant methodological advantages of neurophysiological studies using insects. In particular, it is their high availability and compact size, which is important for scanning probe microscopy. Insects can be induced into an immobile state – anabiosis – by reducing the temperature or increasing the concentration of carbon dioxide. This also greatly simplifies the experiment. The percentage ratio of the number of neurons and other cells in insects is much higher than in humans, which gives certain advantages in studies with atomic force microscopy. Unlike other methods – patch-clamp, optogenetics, electron microscopy – probe microscopy makes it possible to visualise the insect brain in 3D with nanometre resolution in a natural environment. Thus, combining electrophysiological methods with the capabilities of scanning probe microscopy allows:
obtain information on structure and function of neurons;
study dynamics of action potentials;
study the influence of external factors on the passage of nerve impulses (temperature, chemicals, insecticides);
develop new approaches to insect population management and biosensors.
This research is not only of fundamental importance but also has practical applications, from gaining new insights to developing neural interfaces based on biological systems.
ACKNOWLEDGEMENTS
The study was conducted mainly on an initiative basis. This work was partially performed under the state order of the Lomonosov Moscow State University. FemtoScan Online software is provided by Advanced Technologies Center, www.femtoscan.ru
PEER REVIEW INFO
Editorial board thanks the anonymous reviewer(s) for their contribution to the peer review of this work. It is also grateful for their consent to publish papers on the journal’s website and SEL eLibrary eLIBRARY.RU.
Declaration of Competing Interest. The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.
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