The sensory forebrain is composed of intricately connected cell types, of which functional properties have yet to be fully elucidated. Understanding the interactions of these forebrain circuits has been aided recently by the development of optogenetic methods for light-mediated modulation of neuronal activity. Here, we describe a protocol for examining the functional organization of forebrain circuits in vitro using laser-scanning photostimulation of channelrhodopsin, expressed optogenetically via viral-mediated transfection. This approach also exploits the utility of cre-lox recombination in transgenic mice to target expression in specific neuronal cell types. Following transfection, neurons are physiologically recorded in slice preparations using whole-cell patch clamp to measure their evoked responses to laser-scanning photostimulation of channelrhodopsin expressing fibers. This approach enables an assessment of functional topography and synaptic properties. Morphological correlates can be obtained by imaging the neuroanatomical expression of channelrhodopsin expressing fibers using confocal microscopy of the live slice or post-fixed tissue. These methods enable functional investigations of forebrain circuits that expand upon more conventional approaches.
24 Related JoVE Articles!
Using Affordable LED Arrays for Photo-Stimulation of Neurons
Institutions: Institut Pasteur and Centre National de la Recherche Scientifique (CNRS).
Standard slice electrophysiology has allowed researchers to probe individual components of neural circuitry by recording electrical responses of single cells in response to electrical or pharmacological manipulations1,2
. With the invention of methods to optically control genetically targeted neurons (optogenetics), researchers now have an unprecedented level of control over specific groups of neurons in the standard slice preparation. In particular, photosensitive channelrhodopsin-2 (ChR2) allows researchers to activate neurons with light3,4
. By combining careful calibration of LED-based photostimulation of ChR2 with standard slice electrophysiology, we are able to probe with greater detail the role of adult-born interneurons in the olfactory bulb, the first central relay of the olfactory system. Using viral expression of ChR2-YFP specifically in adult-born neurons, we can selectively control young adult-born neurons in a milieu of older and mature neurons. Our optical control uses a simple and inexpensive LED system, and we show how this system can be calibrated to understand how much light is needed to evoke spiking activity in single neurons. Hence, brief flashes of blue light can remotely control the firing pattern of ChR2-transduced newborn cells.
Neuroscience, Issue 57, Adult neurogenesis, Channelrhodopsin, Neural stem cells, Plasticity, Synapses, Electrophysiology
The Spared Nerve Injury (SNI) Model of Induced Mechanical Allodynia in Mice
Institutions: Aarhus University, University of Copenhagen.
Peripheral neuropathic pain is a severe chronic pain condition which may result from trauma to sensory nerves in the peripheral nervous system. The spared nerve injury (SNI) model induces symptoms of neuropathic pain such as mechanical allodynia i.e. pain due to tactile stimuli that do not normally provoke a painful response .
The SNI mouse model involves ligation of two of the three branches of the sciatic nerve (the tibial nerve and the common peroneal nerve), while the sural nerve is left intact . The lesion results in marked hypersensitivity in the lateral area of the paw, which is innervated by the spared sural nerve. The non-operated side of the mouse can be used as a control. The advantages of the SNI model are the robustness of the response and that it doesn’t require expert microsurgical skills.
The threshold for mechanical pain response is determined by testing with von Frey filaments of increasing bending force, which are repetitively pressed against the lateral area of the paw , . A positive pain reaction is defined as sudden paw withdrawal, flinching and/or paw licking induced by the filament. A positive response in three out of five repetitive stimuli is defined as the pain threshold.
As demonstrated in the video protocol, C57BL/6 mice experience profound allodynia as early as the day following surgery and maintain this for several weeks.
Neuroscience, Issue 54, Sciatic, Injury, PNS, Mechanical allodynia, Neuropathic pain, von Frey
Homarus Americanus Stomatogastric Nervous System Dissection
With the goal of understanding how nervous systems produce activity and respond to the environment, neuroscientists turn to model systems that exhibit the activity of interest and are accessible and amenable to experimental methods. The stomatogastric nervous system (STNS) of the American lobster (Homarus americanus
; also know was the Atlantic or Maine lobster) has been established as a model system for studying rhythm generating networks and neuromodulation of networks. The STNS consists of 3 anterior ganglia (2 commissural ganglia and an oesophageal ganglion), containing modulatory neurons that project centrally to the stomatogastric ganglion (STG). The STG contains approximately 30 neurons that comprise two central pattern generating networks, the pyloric and gastric networks that underlie feeding behaviors in crustaceans1,2
. While it is possible to study this system in vivo3
, the STNS continues to produce its rhythmic activity when isolated in vitro
. Physical isolation of the STNS in a dish allows for easy access to the somata in the ganglia for intracellular electrophysiological recordings and to the nerves of the STNS for extracellular recordings. Isolating the STNS is a two-part process. The first part, dissecting the stomach from the animal, is described in an accompanying video article4
. In this video article, fine dissection techniques are used to isolate the STNS from the stomach. This procedure results in a nervous system preparation that is available for electrophysiological recordings.
Neuroscience, Issue 27, lobster, stomach, neural network, dissection, central pattern generator
Long-term Behavioral Tracking of Freely Swimming Weakly Electric Fish
Institutions: University of Ottawa, University of Ottawa, University of Ottawa.
Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and social interactions can be best studied by observing unrestrained, freely behaving animals. Weakly electric fish (WEF) display readily observable exploratory and social behaviors by emitting electric organ discharge (EOD). Here, we describe three effective techniques to synchronously measure the EOD, body position, and posture of a free-swimming WEF for an extended period of time. First, we describe the construction of an experimental tank inside of an isolation chamber designed to block external sources of sensory stimuli such as light, sound, and vibration. The aquarium was partitioned to accommodate four test specimens, and automated gates remotely control the animals' access to the central arena. Second, we describe a precise and reliable real-time EOD timing measurement method from freely swimming WEF. Signal distortions caused by the animal's body movements are corrected by spatial averaging and temporal processing stages. Third, we describe an underwater near-infrared imaging setup to observe unperturbed nocturnal animal behaviors. Infrared light pulses were used to synchronize the timing between the video and the physiological signal over a long recording duration. Our automated tracking software measures the animal's body position and posture reliably in an aquatic scene. In combination, these techniques enable long term observation of spontaneous behavior of freely swimming weakly electric fish in a reliable and precise manner. We believe our method can be similarly applied to the study of other aquatic animals by relating their physiological signals with exploratory or social behaviors.
Neuroscience, Issue 85, animal tracking, weakly electric fish, electric organ discharge, underwater infrared imaging, automated image tracking, sensory isolation chamber, exploratory behavior
Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae
Institutions: University of Michigan, University of Michigan, University of Michigan, University of Michigan, University of Michigan.
Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila
larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila
larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd
instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.
Bioengineering, Issue 84, Drosophila melanogaster, Live Imaging, Microfluidics, axonal injury, axonal degeneration, calcium imaging, photoconversion, laser microsurgery
Flat-floored Air-lifted Platform: A New Method for Combining Behavior with Microscopy or Electrophysiology on Awake Freely Moving Rodents
Institutions: University of Helsinki, Neurotar LTD, University of Eastern Finland, University of Helsinki.
It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal’s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g.
, learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.
Empty Value, Issue 88, awake, in vivo two-photon microscopy, blood vessels, dendrites, dendritic spines, Ca2+ imaging, intrinsic optical imaging, patch-clamp
Intravital Imaging of Axonal Interactions with Microglia and Macrophages in a Mouse Dorsal Column Crush Injury
Institutions: Case Western Reserve University, Case Western Reserve University, Case Western Reserve University.
Traumatic spinal cord injury causes an inflammatory reaction involving blood-derived macrophages and central nervous system (CNS)-resident microglia. Intra-vital two-photon microscopy enables the study of macrophages and microglia in the spinal cord lesion in the living animal. This can be performed in adult animals with a traumatic injury to the dorsal column. Here, we describe methods for distinguishing macrophages from microglia in the CNS using an irradiation bone marrow chimera to obtain animals in which only macrophages or microglia are labeled with a genetically encoded green fluorescent protein. We also describe a injury model that crushes the dorsal column of the spinal cord, thereby producing a simple, easily accessible, rectangular lesion that is easily visualized in an animal through a laminectomy. Furthermore, we will outline procedures to sequentially image the animals at the anatomical site of injury for the study of cellular interactions during the first few days to weeks after injury.
Cellular Biology, Issue 93, Intravital, spinal cord crush injury, chimera, microglia, macrophages, dorsal column crush, axonal dieback
Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex
Institutions: Washington State University.
Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1
. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4
. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6
. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7
. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8
; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.
Neuroscience, Issue 70, Physiology, Anatomy, Medicine, Pharmacology, Surgery, Sleep, rapid eye movement, glucose, glycolysis, pyramidal neurons, channelrhodopsin, optogenetics, optogenetic stimulation, electroencephalogram, EEG, EMG, brain, animal model
Constructing a Low-budget Laser Axotomy System to Study Axon Regeneration in C. elegans
Institutions: University of Utah.
Laser axotomy followed by time-lapse microscopy is a sensitive assay for axon regeneration phenotypes in C. elegans1
. The main difficulty of this assay is the perceived cost ($25-100K) and technical expertise required for implementing a laser ablation system2,3
. However, solid-state pulse lasers of modest costs (<$10K) can provide robust performance for laser ablation in transparent preparations where target axons are "close" to the tissue surface. Construction and alignment of a system can be accomplished in a day. The optical path provided by light from the focused condenser to the ablation laser provides a convenient alignment guide. An intermediate module with all optics removed can be dedicated to the ablation laser and assures that no optical elements need be moved during a laser ablation session. A dichroic in the intermediate module allows simultaneous imaging and laser ablation. Centering the laser beam to the outgoing beam from the focused microscope condenser lens guides the initial alignment of the system. A variety of lenses are used to condition and expand the laser beam to fill the back aperture of the chosen objective lens. Final alignment and testing is performed with a front surface mirrored glass slide target. Laser power is adjusted to give a minimum size ablation spot (<1um). The ablation spot is centered with fine adjustments of the last kinematically mounted mirror to cross hairs fixed in the imaging window. Laser power for axotomy will be approximately 10X higher than needed for the minimum ablation spot on the target slide (this may vary with the target you use). Worms can be immobilized for laser axotomy and time-lapse imaging by mounting on agarose pads (or in microfluidic chambers4
). Agarose pads are easily made with 10% agarose in balanced saline melted in a microwave. A drop of molten agarose is placed on a glass slide and flattened with another glass slide into a pad approximately 200 um thick (a single layer of time tape on adjacent slides is used as a spacer). A "Sharpie" cap is used to cut out a uniformed diameter circular pad of 13mm. Anesthetic (1ul Muscimol 20mM) and Microspheres (Chris Fang-Yen personal communication) (1ul 2.65% Polystyrene 0.1 um in water) are added to the center of the pad followed by 3-5 worms oriented so they are lying on their left sides. A glass coverslip is applied and then Vaseline is used to seal the coverslip and prevent evaporation of the sample.
Neuroscience, Issue 57, laser axotomy, regeneration, growth cone, time lapse, C. elegans, neuroscience, Nd:Yag laser
Selective Viral Transduction of Adult-born Olfactory Neurons for Chronic in vivo Optogenetic Stimulation
Institutions: Institut Pasteur and Centre National de la Recherche Scientifique (CNRS).
Local interneurons are continuously regenerated in the olfactory bulb of adult rodents1-3
. In this process, called adult neurogenesis, neural stem cells in the walls of the lateral ventricle give rise to neuroblasts that migrate for several millimeters along the rostral migratory stream (RMS) to reach and incorporate into the olfactory bulb. To study the different steps and the impact of adult-born neuron integration into preexisting olfactory circuits, it is necessary to selectively label and manipulate the activity of this specific population of neurons. The recent development of optogenetic technologies offers the opportunity to use light to precisely activate this specific cohort of neurons without affecting surrounding neurons4,5
. Here, we present a series of procedures to virally express Channelrhodopsin2(ChR2)-YFP in a temporally restricted cohort of neuroblasts in the RMS before they reach the olfactory bulb and become adult-born neurons. In addition, we show how to implant and calibrate a miniature LED for chronic in vivo
stimulation of ChR2-expressing neurons.
Neuroscience, Issue 58, Olfactory bulb, Olfactory neurons, in vivo, viral transduction, mouse, LED
Optogenetic Perturbation of Neural Activity with Laser Illumination in Semi-intact Drosophila Larvae in Motion
Institutions: The University of Tokyo, The University of Tokyo.
larval locomotion is a splendid model system in developmental and physiological neuroscience, by virtue of the genetic accessibility of the underlying neuronal components in the circuits1-6
. Application of optogenetics7,8
in the larval neural circuit allows us to manipulate neuronal activity in spatially and temporally patterned ways9-13
. Typically, specimens are broadly illuminated with a mercury lamp or LED, so specificity of the target neurons is controlled by binary gene expression systems such as the Gal4-UAS system14,15
. In this work, to improve the spatial resolution to "sub-genetic resolution", we locally illuminated a subset of neurons in the ventral nerve cord using lasers implemented in a conventional confocal microscope. While monitoring the motion of the body wall of the semi-intact larvae, we interactively activated or inhibited neural activity with channelrhodopsin16,17
, respectively. By spatially and temporally restricted illumination of the neural tissue, we can manipulate the activity of specific neurons in the circuit at a specific phase of behavior. This method is useful for studying the relationship between the activities of a local neural assembly in the ventral nerve cord and the spatiotemporal pattern of motor output.
Neuroscience, Issue 77, Molecular Biology, Neurobiology, Developmental Biology, Bioengineering, Cellular Biology, Motor Neurons, Neurosciences, Drosophila, Optogenetics, Channelrhodopsin-2, Halorhodopsin, laser, confocal microscopy, animal model
An In Vitro Preparation for Eliciting and Recording Feeding Motor Programs with Physiological Movements in Aplysia californica
Institutions: Case Western Reserve University , Case Western Reserve University , Case Western Reserve University .
Multifunctionality, the ability of one peripheral structure to generate multiple, distinct behaviors1
, allows animals to rapidly adapt their behaviors to changing environments. The marine mollusk Aplysia californica
provides a tractable system for the study of multifunctionality. During feeding, Aplysia
generates several distinct types of behaviors using the same feeding apparatus, the buccal mass. The ganglia that control these behaviors contain a number of large, identified neurons that are accessible to electrophysiological study. The activity of these neurons has been described in motor programs that can be divided into two types, ingestive and egestive programs, based on the timing of neural activity that closes the food grasper relative to the neural activity that protracts or retracts the grasper2
. However, in isolated ganglia, the muscle movements that would produce these behaviors are absent, making it harder to be certain whether the motor programs observed are correlates of real behaviors. In vivo
, nerve and muscle recordings have been obtained corresponding to feeding programs2,3,4
, but it is very difficult to directly record from individual neurons5
. Additionally, in vivo
, ingestive programs can be further divided into bites and swallows1,2
, a distinction that is difficult to make in most previously described in vitro
The suspended buccal mass preparation (Figure 1
) bridges the gap between isolated ganglia and intact animals. In this preparation, ingestive behaviors - including both biting and swallowing - and egestive behaviors (rejection) can be elicited, at the same time as individual neurons can be recorded from and stimulated using extracellular electrodes6
. The feeding movements associated with these different behaviors can be recorded, quantified, and related directly to the motor programs. The motor programs in the suspended buccal mass preparation appear to be more similar to those observed in vivo
than are motor programs elicited in isolated ganglia. Thus, the motor programs in this preparation can be more directly related to in vivo
behavior; at the same time, individual neurons are more accessible to recording and stimulation than in intact animals. Additionally, as an intermediate step between isolated ganglia and intact animals, findings from the suspended buccal mass can aid in interpretation of data obtained in both more reduced and more intact settings. The suspended buccal mass preparation is a useful tool for characterizing the neural control of multifunctionality in Aplysia
Neuroscience, Issue 70, Physiology, Biomedical Engineering, Anatomy, Marine Biology, Aplysia, Aplysia californica, California sea slug, invertebrate, feeding, neurobiology, buccal mass, semi-intact preparation, extracellular electrodes, extracellular recording, neurons, animal model
A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo
Institutions: University of Colorado Boulder, University of Colorado Boulder.
Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g.
, channelrhodopsin-2, ChR2) or silence (e.g.
, halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2
transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e.
, pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo
preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.
Neuroscience, Issue 79, Genetic Techniques, Genetics, Behavioral, Biological Science Disciplines, Neurosciences, genetics (animal and plant), Investigative Techniques, Behavior and Behavior Mechanisms, Behavioral Disciplines and Activities, Natural Science Disciplines, Optogenetics, prefrontal cortex, subiculum, virus injection, in vivo recording, Neurophysiology, prelimbic, optrode, molecular neurogenetics, Gene targeting
Optogenetic Activation of Zebrafish Somatosensory Neurons using ChEF-tdTomato
Institutions: University of California, Los Angeles .
Larval zebrafish are emerging as a model for describing the development and function of simple neural circuits. Due to their external fertilization, rapid development, and translucency, zebrafish are particularly well suited for optogenetic approaches to investigate neural circuit function. In this approach, light-sensitive ion channels are expressed in specific neurons, enabling the experimenter to activate or inhibit them at will and thus assess their contribution to specific behaviors. Applying these methods in larval zebrafish is conceptually simple but requires the optimization of technical details. Here we demonstrate a procedure for expressing a channelrhodopsin variant in larval zebrafish somatosensory neurons, photo-activating single cells, and recording the resulting behaviors. By introducing a few modifications to previously established methods, this approach could be used to elicit behavioral responses from single neurons activated up to at least 4 days post-fertilization (dpf). Specifically, we created a transgene using a somatosensory neuron enhancer, CREST3
, to drive the expression of the tagged channelrhodopsin variant, ChEF-tdTomato. Injecting this transgene into 1-cell stage embryos results in mosaic expression in somatosensory neurons, which can be imaged with confocal microscopy. Illuminating identified cells in these animals with light from a 473 nm DPSS laser, guided through a fiber optic cable, elicits behaviors that can be recorded with a high-speed video camera and analyzed quantitatively. This technique could be adapted to study behaviors elicited by activating any zebrafish neuron. Combining this approach with genetic or pharmacological perturbations will be a powerful way to investigate circuit formation and function.
Neuroscience, Issue 71, Developmental Biology, Molecular Biology, Cellular Biology, Biochemistry, Bioengineering, Anatomy, Physiology, Zebrafish, Behavior, Animal, Touch, optogenetics, channelrhodopsin, ChEF, sensory neuron, Rohon-Beard, Danio rerio, somatosensory, neurons, microinjection, confocal microscopy, high speed video, animal model
Optogenetic Stimulation of Escape Behavior in Drosophila melanogaster
Institutions: Stanford University .
A growing number of genetically encoded tools are becoming available that allow non-invasive manipulation of the neural activity of specific neurons in Drosophila melanogaster1
. Chief among these are optogenetic tools, which enable the activation or silencing of specific neurons in the intact and freely moving animal using bright light. Channelrhodopsin (ChR2) is a light-activated cation channel that, when activated by blue light, causes depolarization of neurons that express it. ChR2 has been effective for identifying neurons critical for specific behaviors, such as CO2
avoidance, proboscis extension and giant-fiber mediated startle response2-4
. However, as the intense light sources used to stimulate ChR2 also stimulate photoreceptors, these optogenetic techniques have not previously been used in the visual system. Here, we combine an optogenetic approach with a mutation that impairs phototransduction to demonstrate that activation of a cluster of loom-sensitive neurons in the fly's optic lobe, Foma-1 neurons, can drive an escape behavior used to avoid collision. We used a null allele of a critical component of the phototransduction cascade, phospholipase C-β, encoded by the norpA
gene, to render the flies blind and also use the Gal4-UAS transcriptional activator system to drive expression of ChR2 in the Foma-1 neurons. Individual flies are placed on a small platform surrounded by blue LEDs. When the LEDs are illuminated, the flies quickly take-off into flight, in a manner similar to visually driven loom-escape behavior. We believe that this technique can be easily adapted to examine other behaviors in freely moving flies.
Neurobiology, Issue 71, Neuroscience, Genetics, Anatomy, Physiology, Molecular Biology, Cellular Biology, Behavior, optogenetics, channelrhodopsin, ChR2, escape behavior, neurons, fruit fly, Drosophila melanogaster, animal model
Extracellularly Identifying Motor Neurons for a Muscle Motor Pool in Aplysia californica
Institutions: Case Western Reserve University , Case Western Reserve University , Case Western Reserve University .
In animals with large identified neurons (e.g.
mollusks), analysis of motor pools is done using intracellular techniques1,2,3,4
. Recently, we developed a technique to extracellularly stimulate and record individual neurons in Aplysia californica5
. We now describe a protocol for using this technique to uniquely identify and characterize motor neurons within a motor pool.
This extracellular technique has advantages. First, extracellular electrodes can stimulate and record neurons through the sheath5
, so it does not need to be removed. Thus, neurons will be healthier in extracellular experiments than in intracellular ones. Second, if ganglia are rotated by appropriate pinning of the sheath, extracellular electrodes can access neurons on both sides of the ganglion, which makes it easier and more efficient to identify multiple neurons in the same preparation. Third, extracellular electrodes do not need to penetrate cells, and thus can be easily moved back and forth among neurons, causing less damage to them. This is especially useful when one tries to record multiple neurons during repeating motor patterns that may only persist for minutes. Fourth, extracellular electrodes are more flexible than intracellular ones during muscle movements. Intracellular electrodes may pull out and damage neurons during muscle contractions. In contrast, since extracellular electrodes are gently pressed onto the sheath above neurons, they usually stay above the same neuron during muscle contractions, and thus can be used in more intact preparations.
To uniquely identify motor neurons for a motor pool (in particular, the I1/I3 muscle in Aplysia
) using extracellular electrodes, one can use features that do not require intracellular measurements as criteria: soma size and location, axonal projection, and muscle innervation4,6,7
. For the particular motor pool used to illustrate the technique, we recorded from buccal nerves 2 and 3 to measure axonal projections, and measured the contraction forces of the I1/I3 muscle to determine the pattern of muscle innervation for the individual motor neurons.
We demonstrate the complete process of first identifying motor neurons using muscle innervation, then characterizing their timing during motor patterns, creating a simplified diagnostic method for rapid identification. The simplified and more rapid diagnostic method is superior for more intact preparations, e.g.
in the suspended buccal mass preparation8
or in vivo9
. This process can also be applied in other motor pools10,11,12
or in other animal systems2,3,13,14
Neuroscience, Issue 73, Physiology, Biomedical Engineering, Anatomy, Behavior, Neurobiology, Animal, Neurosciences, Neurophysiology, Electrophysiology, Aplysia, Aplysia californica, California sea slug, invertebrate, feeding, buccal mass, ganglia, motor neurons, neurons, extracellular stimulation and recordings, extracellular electrodes, animal model
Membrane Potentials, Synaptic Responses, Neuronal Circuitry, Neuromodulation and Muscle Histology Using the Crayfish: Student Laboratory Exercises
Institutions: University of Kentucky, University of Toronto.
The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that influence a cell's membrane potential and which we address in these experiments are: (1) Ion concentration of K+
on the outside of the membrane, and (2) the permeability of the membrane to specific ions. The crayfish abdominal extensor muscles are in groupings with some being tonic (slow) and others phasic (fast) in their biochemical and physiological phenotypes, as well as in their structure; the motor neurons that innervate these muscles are correspondingly different in functional characteristics. We use these muscles as well as the superficial, tonic abdominal flexor muscle to demonstrate properties in synaptic transmission. In addition, we introduce a sensory-CNS-motor neuron-muscle circuit to demonstrate the effect of cuticular sensory stimulation as well as the influence of neuromodulators on certain aspects of the circuit. With the techniques obtained in this exercise, one can begin to answer many questions remaining in other experimental preparations as well as in physiological applications related to medicine and health. We have demonstrated the usefulness of model invertebrate preparations to address fundamental questions pertinent to all animals.
Neuroscience, Issue 47, Invertebrate, Crayfish, neurophysiology, muscle, anatomy, electrophysiology
Optogenetic Stimulation of the Auditory Nerve
Institutions: University Medical Center Goettingen, University of Goettingen, University Medical Center Goettingen, University of Goettingen, University of Guanajuato.
Direct electrical stimulation of spiral ganglion neurons (SGNs) by cochlear implants (CIs) enables open speech comprehension in the majority of implanted deaf subjects1-6
. Nonetheless, sound coding with current CIs has poor frequency and intensity resolution due to broad current spread from each electrode contact activating a large number of SGNs along the tonotopic axis of the cochlea7-9
. Optical stimulation is proposed as an alternative to electrical stimulation that promises spatially more confined activation of SGNs and, hence, higher frequency resolution of coding. In recent years, direct infrared illumination of the cochlea has been used to evoke responses in the auditory nerve10
. Nevertheless it requires higher energies than electrical stimulation10,11
and uncertainty remains as to the underlying mechanism12
. Here we describe a method based on optogenetics to stimulate SGNs with low intensity blue light, using transgenic mice with neuronal expression of channelrhodopsin 2 (ChR2)13
or virus-mediated expression of the ChR2-variant CatCh14
. We used micro-light emitting diodes (µLEDs) and fiber-coupled lasers to stimulate ChR2-expressing SGNs through a small artificial opening (cochleostomy) or the round window. We assayed the responses by scalp recordings of light-evoked potentials (optogenetic auditory brainstem response: oABR) or by microelectrode recordings from the auditory pathway and compared them with acoustic and electrical stimulation.
Neuroscience, Issue 92, hearing, cochlear implant, optogenetics, channelrhodopsin, optical stimulation, deafness
Proprioception and Tension Receptors in Crab Limbs: Student Laboratory Exercises
Institutions: University of Kentucky, University of Kentucky, University of Oregon.
The primary purpose of these procedures is to demonstrate for teaching and research purposes how to record the activity of living primary sensory neurons responsible for proprioception as they are detecting joint position and movement, and muscle tension. Electrical activity from crustacean proprioceptors and tension receptors is recorded by basic neurophysiological instrumentation, and a transducer is used to simultaneously measure force that is generated by stimulating a motor nerve. In addition, we demonstrate how to stain the neurons for a quick assessment of their anatomical arrangement or for permanent fixation. Staining reveals anatomical organization that is representative of chordotonal organs in most crustaceans. Comparing the tension nerve responses to the proprioceptive responses is an effective teaching tool in determining how these sensory neurons are defined functionally and how the anatomy is correlated to the function. Three staining techniques are presented allowing researchers and instructors to choose a method that is ideal for their laboratory.
Neuroscience, Issue 80, Crustacean, joint, Muscle, sensory, teaching, educational, neuroscience
Utilizing Transcranial Magnetic Stimulation to Study the Human Neuromuscular System
Institutions: Ohio University.
Transcranial magnetic stimulation (TMS) has been in use for more than 20 years 1
, and has grown exponentially in popularity over the past decade. While the use of TMS has expanded to the study of many systems and processes during this time, the original application and perhaps one of the most common uses of TMS involves studying the physiology, plasticity and function of the human neuromuscular system. Single pulse TMS applied to the motor cortex excites pyramidal neurons transsynaptically 2
(Figure 1) and results in a measurable electromyographic response that can be used to study and evaluate the integrity and excitability of the corticospinal tract in humans 3
. Additionally, recent advances in magnetic stimulation now allows for partitioning of cortical versus spinal excitability 4,5
. For example, paired-pulse TMS can be used to assess intracortical facilitatory and inhibitory properties by combining a conditioning stimulus and a test stimulus at different interstimulus intervals 3,4,6-8
. In this video article we will demonstrate the methodological and technical aspects of these techniques. Specifically, we will demonstrate single-pulse and paired-pulse TMS techniques as applied to the flexor carpi radialis (FCR) muscle as well as the erector spinae (ES) musculature. Our laboratory studies the FCR muscle as it is of interest to our research on the effects of wrist-hand cast immobilization on reduced muscle performance6,9
, and we study the ES muscles due to these muscles clinical relevance as it relates to low back pain8
. With this stated, we should note that TMS has been used to study many muscles of the hand, arm and legs, and should iterate that our demonstrations in the FCR and ES muscle groups are only selected examples of TMS being used to study the human neuromuscular system.
Medicine, Issue 59, neuroscience, muscle, electromyography, physiology, TMS, strength, motor control. sarcopenia, dynapenia, lumbar
The Swimmeret System of Crayfish: A Practical Guide for the Dissection of the Nerve Cord and Extracellular Recordings of the Motor Pattern
Institutions: University of Cologne.
Here we demonstrate the dissection of the crayfish abdominal nerve cord. The preparation comprises the last two thoracic ganglia (T4, T5) and the chain of abdominal ganglia (A1 to A6). This chain of ganglia includes the part of the central nervous system (CNS) that drives coordinated locomotion of the pleopods (swimmerets): the swimmeret system. It is known for over five decades that in crayfish each swimmeret is driven by its own independent pattern generating kernel that generates rhythmic alternating activity 1-3
. The motor neurons innervating the musculature of each swimmeret comprise two anatomically and functionally distinct populations 4
. One is responsible for the retraction (power stroke, PS) of the swimmeret. The other drives the protraction (return stroke, RS) of the swimmeret. Motor neurons of the swimmeret system are able to produce spontaneously a fictive motor pattern, which is identical to the pattern recorded in vivo 1
The aim of this report is to introduce an interesting and convenient model system for studying rhythm generating networks and coordination of independent microcircuits for students’ practical laboratory courses. The protocol provided includes step-by-step instructions for the dissection of the crayfish’s abdominal nerve cord, pinning of the isolated chain of ganglia, desheathing the ganglia and recording the swimmerets fictive motor pattern extracellularly from the isolated nervous system.
Additionally, we can monitor the activity of swimmeret neurons recorded intracellularly from dendrites. Here we also describe briefly these techniques and provide some examples. Furthermore, the morphology of swimmeret neurons can be assessed using various staining techniques. Here we provide examples of intracellular (by iontophoresis) dye filled neurons and backfills of pools of swimmeret motor neurons. In our lab we use this preparation to study basic functions of fictive locomotion, the effect of sensory feedback on the activity of the CNS, and coordination between microcircuits on a cellular level.
Neurobiology, Issue 93, crustacean, dissection, extracellular recording, fictive locomotion, motor neurons, locomotion
An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time
Institutions: University of Louisville, University of Calgary.
Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo
living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo
spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo
spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g.,
calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo
); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.
Neuroscience, Issue 93, spinal cord injury, axon, myelin, two-photon excitation microscopy, Nile Red, axonal degeneration, axonal dieback, axonal retraction
Axoplasm Isolation from Rat Sciatic Nerve
Institutions: Weizmann Institute of Science.
Isolation of pure axonal cytoplasm (axoplasm) from peripheral nerve is crucial for biochemical studies of many biological processes. In this article, we demonstrate and describe a protocol for axoplasm isolation from adult rat sciatic nerve based on the following steps: (1) dissection of nerve fascicles and separation of connective tissue; (2) incubation of short segments of nerve fascicles in hypotonic medium to release myelin and lyse non-axonal structures; and (3) extraction of the remaining axon-enriched material. Proteomic and biochemical characterization of this preparation has confirmed a high degree of enrichment for axonal components.
Neuroscience, Issue 43, Axoplasm, nerve, isolation, method, rat
Electrophysiological Methods for Recording Synaptic Potentials from the NMJ of Drosophila Larvae
Institutions: Columbia University College of Physicians and Surgeons.
In this video, we describe the electrophysiological methods for recording synaptic transmission at the neuromuscular junction (NMJ) of Drosophila larva. The larval neuromuscular system is a model synapse for the study of synaptic physiology and neurotransmission, and is a valuable research tool that has defined genetics and is accessible to experimental manipulation. Larvae can be dissected to expose the body wall musculature, central nervous system, and peripheral nerves. The muscles of Drosophila and their innervation pattern are well characterized and muscles are easy to access for intracellular recording. Individual muscles can be identified by their location and orientation within the 8 abdominal segments, each with 30 muscles arranged in a pattern that is repeated in segments A2 - A7. Dissected drosophila larvae are thin and individual muscles and bundles of motor neuron axons can be visualized by transillumination1
. Transgenic constructs can be used to label target cells for visual identification or for manipulating gene products in specific tissues. In larvae, excitatory junction potentials (EJP’s) are generated in response to vesicular release of glutamate from the motoneurons at the synapse. In dissected larvae, the EJP can be recorded in the muscle with an intracellular electrode. Action potentials can be artificially evoked in motor neurons that have been cut posterior to the ventral ganglion, drawn into a glass pipette by gentle suction and stimulated with an electrode. These motor neurons have distinct firing thresholds when stimulated, and when they fire simultaneously, they generate a response in the muscle. Signals transmitted across the NMJ synapse can be recorded in the muscles that the motor neurons innervate. The EJP’s and minature excitatory junction potentials (mEJP’s) are seen as changes in membrane potential. Electrophysiological responses are recorded at room temperature in modified minimal hemolymph-like solution2
(HL3) that contains 5 mM Mg2+
and 1.5 mM Ca2+
. Changes in the amplitude of evoked EJP’s can indicate differences in synaptic function and structure. Digitized recordings are analyzed for EJP amplitude, mEJP frequency and amplitude, and quantal content.
Neuroscience, Issue 24, Neuromuscular junction, synaptic transmission, Drosophila larvae, electrophysiology