Muscle sensory neurons innervating muscle spindles and Golgi tendon organs encode length and force changes essential to proprioception. Additional afferent fibers monitor other characteristics of the muscle environment, including metabolite buildup, temperature, and nociceptive stimuli. Overall, abnormal activation of sensory neurons can lead to movement disorders or chronic pain syndromes. We describe the isolation of the extensor digitorum longus (EDL) muscle and nerve for in vitro study of stretch-evoked afferent responses in the adult mouse. Sensory activity is recorded from the nerve with a suction electrode and individual afferents can be analyzed using spike sorting software. In vitro preparations allow for well controlled studies on sensory afferents without the potential confounds of anesthesia or altered muscle perfusion. Here we describe a protocol to identify and test the response of muscle spindle afferents to stretch. Importantly, this preparation also supports the study of other subtypes of muscle afferents, response properties following drug application and the incorporation of powerful genetic approaches and disease models in mice.
20 Related JoVE Articles!
The Use of Cystometry in Small Rodents: A Study of Bladder Chemosensation
Institutions: KU Leuven, Belgium, KU Leuven, Belgium, KU Leuven, Belgium.
The lower urinary tract (LUT) functions as a dynamic reservoir that is able to store urine and to efficiently expel it at a convenient time. While storing urine, however, the bladder is exposed for prolonged periods to waste products. By acting as a tight barrier, the epithelial lining of the LUT, the urothelium, avoids re-absorption of harmful substances. Moreover, noxious chemicals stimulate the bladder's nociceptive innervation and initiate voiding contractions that expel the bladder's contents. Interestingly, the bladder's sensitivity to noxious chemicals has been used successfully in clinical practice, by intravesically infusing the TRPV1 agonist capsaicin to treat neurogenic bladder overactivity1
. This underscores the advantage of viewing the bladder as a chemosensory organ and prompts for further clinical research. However, ethical issues severely limit the possibilities to perform, in human subjects, the invasive measurements that are necessary to unravel the molecular bases of LUT clinical pharmacology. A way to overcome this limitation is the use of several animal models2
. Here we describe the implementation of cystometry in mice and rats, a technique that allows measuring the intravesical pressure in conditions of controlled bladder perfusion.
After laparotomy, a catheter is implanted in the bladder dome and tunneled subcutaneously to the interscapular region. Then the bladder can be filled at a controlled rate, while the urethra is left free for micturition. During the repetitive cycles of filling and voiding, intravesical pressure can be measured via the implanted catheter. As such, the pressure changes can be quantified and analyzed. Moreover, simultaneous measurement of the voided volume allows distinguishing voiding contractions from non-voiding contractions3
Importantly, due to the differences in micturition control between rodents and humans, cystometric measurements in these animals have only limited translational value4
. Nevertheless, they are quite instrumental in the study of bladder pathophysiology and pharmacology in experimental pre-clinical settings. Recent research using this technique has revealed the key role of novel molecular players in the mechano- and chemo-sensory properties of the bladder.
Medicine, Issue 66, Physiology, Chemistry, cystometry, urodynamics, bladder function, bladder chemosensation, animal model, urinary tract
An Isolated Semi-intact Preparation of the Mouse Vestibular Sensory Epithelium for Electrophysiology and High-resolution Two-photon Microscopy
Institutions: University of Sydney, University of Newcastle.
Understanding vestibular hair cells function under normal conditions, or how trauma, disease, and aging disrupt this function is a vital step in the development of preventative approaches and/or novel therapeutic strategies. However, the majority of studies looking at abnormal vestibular function have not been at the cellular level but focused primarily on behavioral assays of vestibular dysfunction such as gait analyses and vestibulo-ocular reflex performance. While this work has yielded valuable data about what happens when things go wrong, little information is gleaned regarding the underlying causes of dysfunction. Of the studies that focus on the cellular and subcellular processes that underlie vestibular function, most have relied on acutely isolated hair cells, devoid of their synaptic connections and supporting cell environment. Therefore, a major technical challenge has been access to the exquisitely sensitive vestibular hair cells in a preparation that is least disrupted, physiologically. Here we demonstrate a semi-intact preparation of the mouse vestibular sensory epithelium that retains the local micro-environment including hair cell/primary afferent complexes.
Neurobiology, Issue 76, Neuroscience, Cellular Biology, Molecular Biology, Biomedical Engineering, Anatomy, Physiology, Surgery, Vestibular, Hair cells, Epithelium, two-photon microscopy, isolated, semi-intact, electrophysiology, electroporation, microscopy, tissue, isolation, animal model
Physiological, Morphological and Neurochemical Characterization of Neurons Modulated by Movement
Institutions: University of Maryland.
The role of individual neurons and their function in neuronal circuits is fundamental to understanding the neuronal mechanisms of sensory and motor functions. Most investigations of sensorimotor mechanisms rely on either examination of neurons while an animal is static1,2
or record extracellular neuronal activity during a movement.3,4
While these studies have provided the fundamental background for sensorimotor function, they either do not evaluate functional information which occurs during a movement or are limited in their ability to fully characterize the anatomy, physiology and neurochemical phenotype of the neuron. A technique is shown here which allows extensive characterization of individual neurons during an in vivo
movement. This technique can be used not only to study primary afferent neurons but also to characterize motoneurons and sensorimotor interneurons. Initially the response of a single neuron is recorded using electrophysiological methods during various movements of the mandible followed by determination of the receptive field for the neuron. A neuronal tracer is then intracellularly injected into the neuron and the brain is processed so that the neuron can be visualized with light, electron or confocal microscopy (Fig. 1). The detailed morphology of the characterized neuron is then reconstructed so that neuronal morphology can be correlated with the physiological response of the neuron (Figs. 2,3). In this communication important key details and tips for successful implementation of this technique are provided. Valuable additional information can be determined for the neuron under study by combining this method with other techniques. Retrograde neuronal labeling can be used to determine neurons with which the labeled neuron synapses; thus allowing detailed determination of neuronal circuitry. Immunocytochemistry can be combined with this method to examine neurotransmitters within the labeled neuron and to determine the chemical phenotypes of neurons with which the labeled neuron synapses. The labeled neuron can also be processed for electron microscopy to determine the ultrastructural features and microcircuitry of the labeled neuron. Overall this technique is a powerful method to thoroughly characterize neurons during in vivo
movement thus allowing substantial insight into the role of the neuron in sensorimotor function.
Neuroscience, Issue 50, neurophysiology, sensory neuron, motor control, proprioception, neurotransmitter, sensorimotor integration, rat
Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones
Institutions: The University of Iowa, Roy J. and Lucille A. Carver College of Medicine, The University of Iowa.
Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.
Neuroscience, Issue 59, pain, immunostaining, sensory nerve fiber, skin, bone, plantar punch, CGRP, NF200, TRPV1, Tubulin
Identification of Specific Sensory Neuron Populations for Study of Expressed Ion Channels
Institutions: AT Still University of Health Sciences.
Sensory neurons transmit signals from various parts of the body to the central nervous system. The soma for these neurons are located in the dorsal root ganglia that line the spinal column. Understanding the receptors and channels expressed by these sensory afferent neurons could lead to novel therapies for disease. The initial step is to identify the specific subset of sensory neurons of interest. Here we describe a method to identify afferent neurons innervating the muscles by retrograde labeling using a fluorescent dye DiI (1,1'-dioctadecyl-3,3,3',3'-tetramethylindocarbocyanine perchlorate). Understanding the contribution of ion channels to excitation of muscle afferents could help to better control excessive excitability induced by certain disease states such as peripheral vascular disease or heart failure. We used two approaches to identify the voltage dependent ion channels expressed by these neurons, patch clamp electrophysiology and immunocytochemistry. While electrophysiology plus pharmacological blockers can identify functional ion channel types, we used immunocytochemistry to identify channels for which specific blockers were unavailable and to better understand the ion channel distribution pattern in the cell population. These techniques can be applied to other areas of the nervous system to study specific neuronal groups.
Neuroscience, Issue 82, DiI, patch clamp, sensory neurons, muscle afferent neurons, immunocytochemistry, electrophysiology
Preparation of Acute Hippocampal Slices from Rats and Transgenic Mice for the Study of Synaptic Alterations during Aging and Amyloid Pathology
Institutions: University of Kentucky College of Public Health, University of Kentucky College of Medicine, University of Kentucky College of Medicine.
The rodent hippocampal slice preparation is perhaps the most broadly used tool for investigating mammalian synaptic function and plasticity. The hippocampus can be extracted quickly and easily from rats and mice and slices remain viable for hours in oxygenated artificial cerebrospinal fluid. Moreover, basic electrophysisologic techniques are easily applied to the investigation of synaptic function in hippocampal slices and have provided some of the best biomarkers for cognitive impairments. The hippocampal slice is especially popular for the study of synaptic plasticity mechanisms involved in learning and memory. Changes in the induction of long-term potentiation and depression (LTP and LTD) of synaptic efficacy in hippocampal slices (or lack thereof) are frequently used to describe the neurologic phenotype of cognitively-impaired animals and/or to evaluate the mechanism of action of nootropic compounds. This article outlines the procedures we use for preparing hippocampal slices from rats and transgenic mice for the study of synaptic alterations associated with brain aging and Alzheimer's disease (AD)1-3
. Use of aged rats and AD model mice can present a unique set of challenges to researchers accustomed to using younger rats and/or mice in their research. Aged rats have thicker skulls and tougher connective tissue than younger rats and mice, which can delay brain extraction and/or dissection and consequently negate or exaggerate real age-differences in synaptic function and plasticity. Aging and amyloid pathology may also exacerbate hippocampal damage sustained during the dissection procedure, again complicating any inferences drawn from physiologic assessment. Here, we discuss the steps taken during the dissection procedure to minimize these problems. Examples of synaptic responses acquired in "healthy" and "unhealthy" slices from rats and mice are provided, as well as representative synaptic plasticity experiments. The possible impact of other methodological factors on synaptic function in these animal models (e.g. recording solution components, stimulation parameters) are also discussed. While the focus of this article is on the use of aged rats and transgenic mice, novices to slice physiology should find enough detail here to get started on their own studies, using a variety of rodent models.
Neuroscience, Issue 49, aging, amyloid, hippocampal slice, synaptic plasticity, Ca2+, CA1, electrophysiology
Simultaneous Long-term Recordings at Two Neuronal Processing Stages in Behaving Honeybees
Institutions: University of Würzburg.
In both mammals and insects neuronal information is processed in different higher and lower order brain centers. These centers are coupled via convergent and divergent anatomical connections including feed forward and feedback wiring. Furthermore, information of the same origin is partially sent via parallel pathways to different and sometimes into the same brain areas. To understand the evolutionary benefits as well as the computational advantages of these wiring strategies and especially their temporal dependencies on each other, it is necessary to have simultaneous access to single neurons of different tracts or neuropiles in the same preparation at high temporal resolution. Here we concentrate on honeybees by demonstrating a unique extracellular long term access to record multi unit activity at two subsequent neuropiles1
, the antennal lobe (AL), the first olfactory processing stage and the mushroom body (MB), a higher order integration center involved in learning and memory formation, or two parallel neuronal tracts2
connecting the AL with the MB. The latter was chosen as an example and will be described in full. In the supporting video the construction and permanent insertion of flexible multi channel wire electrodes is demonstrated. Pairwise differential amplification of the micro wire electrode channels drastically reduces the noise and verifies that the source of the signal is closely related to the position of the electrode tip. The mechanical flexibility of the used wire electrodes allows stable invasive long term recordings over many hours up to days, which is a clear advantage compared to conventional extra and intracellular in vivo
Neuroscience, Issue 89, honeybee brain, olfaction, extracellular long term recordings, double recordings, differential wire electrodes, single unit, multi-unit recordings
Preparation of Parasagittal Slices for the Investigation of Dorsal-ventral Organization of the Rodent Medial Entorhinal Cortex
Institutions: University of Edinburgh , University of Edinburgh .
Computation in the brain relies on neurons responding appropriately to their synaptic inputs. Neurons differ in their complement and distribution of membrane ion channels that determine how they respond to synaptic inputs. However, the relationship between these cellular properties and neuronal function in behaving animals is not well understood. One approach to this problem is to investigate topographically organized neural circuits in which the position of individual neurons maps onto information they encode or computations they carry out1
. Experiments using this approach suggest principles for tuning of synaptic responses underlying information encoding in sensory and cognitive circuits2,3
The topographical organization of spatial representations along the dorsal-ventral axis of the medial entorhinal cortex (MEC) provides an opportunity to establish relationships between cellular mechanisms and computations important for spatial cognition. Neurons in layer II of the rodent MEC encode location using grid-like firing fields4-6
. For neurons found at dorsal positions in the MEC the distance between the individual firing fields that form a grid is on the order of 30 cm, whereas for neurons at progressively more ventral positions this distance increases to greater than 1 m. Several studies have revealed cellular properties of neurons in layer II of the MEC that, like the spacing between grid firing fields, also differ according to their dorsal-ventral position, suggesting that these cellular properties are important for spatial computation2,7-10
Here we describe procedures for preparation and electrophysiological recording from brain slices that maintain the dorsal-ventral extent of the MEC enabling investigation of the topographical organization of biophysical and anatomical properties of MEC neurons. The dorsal-ventral position of identified neurons relative to anatomical landmarks is difficult to establish accurately with protocols that use horizontal slices of MEC7,8,11,12
, as it is difficult to establish reference points for the exact dorsal-ventral location of the slice. The procedures we describe enable accurate and consistent measurement of location of recorded cells along the dorsal-ventral axis of the MEC as well as visualization of molecular gradients2,10
. The procedures have been developed for use with adult mice (> 28 days) and have been successfully employed with mice up to 1.5 years old. With adjustments they could be used with younger mice or other rodent species. A standardized system of preparation and measurement will aid systematic investigation of the cellular and microcircuit properties of this area.
Neuroscience, Issue 61, Parasagittal slice, Medial Entorhinal Cortex, Stellate cell, Grid cell, Synaptic integration, Topographic map
Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes
Institutions: Vanderbilt University Medical Center.
is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila
have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila
ovaries and testes3-12
. Herein, methods for the dissection and preparation of Drosophila
testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila
mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.
Basic Protocol, Issue 83, Drosophila melanogaster, dissection, testes, spermatogenesis, meiosis, germ cells, phase-contrast microscopy, immunofluorescence
Mapping Inhibitory Neuronal Circuits by Laser Scanning Photostimulation
Institutions: University of California, Irvine, University of California, Irvine.
Inhibitory neurons are crucial to cortical function. They comprise about 20% of the entire cortical neuronal population and can be further subdivided into diverse subtypes based on their immunochemical, morphological, and physiological properties1-4
. Although previous research has revealed much about intrinsic properties of individual types of inhibitory neurons, knowledge about their local circuit connections is still relatively limited3,5,6
. Given that each individual neuron's function is shaped by its excitatory and inhibitory synaptic input within cortical circuits, we have been using laser scanning photostimulation (LSPS) to map local circuit connections to specific inhibitory cell types. Compared to conventional electrical stimulation or glutamate puff stimulation, LSPS has unique advantages allowing for extensive mapping and quantitative analysis of local functional inputs to individually recorded neurons3,7-9
. Laser photostimulation via glutamate uncaging selectively activates neurons perisomatically, without activating axons of passage or distal dendrites, which ensures a sub-laminar mapping resolution. The sensitivity and efficiency of LSPS for mapping inputs from many stimulation sites over a large region are well suited for cortical circuit analysis.
Here we introduce the technique of LSPS combined with whole-cell patch clamping for local inhibitory circuit mapping. Targeted recordings of specific inhibitory cell types are facilitated by use of transgenic mice expressing green fluorescent proteins (GFP) in limited inhibitory neuron populations in the cortex3,10
, which enables consistent sampling of the targeted cell types and unambiguous identification of the cell types recorded. As for LSPS mapping, we outline the system instrumentation, describe the experimental procedure and data acquisition, and present examples of circuit mapping in mouse primary somatosensory cortex. As illustrated in our experiments, caged glutamate is activated in a spatially restricted region of the brain slice by UV laser photolysis; simultaneous voltage-clamp recordings allow detection of photostimulation-evoked synaptic responses. Maps of either excitatory or inhibitory synaptic input to the targeted neuron are generated by scanning the laser beam to stimulate hundreds of potential presynaptic sites. Thus, LSPS enables the construction of detailed maps of synaptic inputs impinging onto specific types of inhibitory neurons through repeated experiments. Taken together, the photostimulation-based technique offers neuroscientists a powerful tool for determining the functional organization of local cortical circuits.
Neuroscience, Issue 56, glutamate uncaging, whole cell recording, GFP, transgenic, interneurons
Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals
Institutions: University of Illinois at Chicago.
Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+
into the presynaptic terminal, through voltage-gated calcium channels (VGCCs) located in the release face membrane, is the trigger for vesicle fusion and neurotransmitter release. Crucial to the rapidity of synaptic transmission is the spatial and temporal synchrony between the arrival of the action potential, VGCCs and the neurotransmitter release machinery. The ability to directly record Ca2+
currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca2+
and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca2+
entry has been characterized using imaging techniques and macroscopic current measurements – techniques that do not have sufficient temporal resolution to visualize Ca2+
entry. The characterization of VGCCs directly at single presynaptic terminals has not been possible in central synapses and has thus far been successfully achieved only in the calyx-type synapse of the chick ciliary ganglion and in rat calyces. We have successfully addressed this problem in the giant reticulospinal synapse of the lamprey spinal cord by developing an acutely dissociated preparation of the spinal cord that yields isolated reticulospinal axons with functional presynaptic terminals devoid of postsynaptic structures. We can fluorescently label and identify individual presynaptic terminals and target them for recording. Using this preparation, we have characterized VGCCs directly at the release face of individual presynaptic terminals using immunohistochemistry and electrophysiology approaches. Ca2+
currents have been recorded directly at the release face membrane of individual presynaptic terminals, the first such recording to be carried out at central synapses.
Neuroscience, Issue 92, reticulospinal synapse, reticulospinal axons, presynaptic terminal, presynaptic calcium, voltage-gated calcium channels, vesicle fusion, synaptic transmission, neurotransmitter release, spinal cord, lamprey, synaptic vesicles, acute dissociation
Postsynaptic Recordings at Afferent Dendrites Contacting Cochlear Inner Hair Cells: Monitoring Multivesicular Release at a Ribbon Synapse
Institutions: The Johns Hopkins School of Medicine, Consejo Nacional de Investigaciones Científicas y Técnicas.
The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the postsynaptic activity of a single ribbon synapse 1-4
. Ribbon synapses of sensory cells release neurotransmitter continuously, the rate of which is modulated in response to graded changes in IHC membrane potential 5
. Ribbon synapses have been shown to operate by multivesicular release, where multiple vesicles can be released simultaneously to evoke excitatory postsynaptic currents (EPSCs) of varying amplitudes 1, 4, 6-11
. Neither the role of the presynaptic ribbon, nor the mechanism underlying multivesicular release is currently well understood.
The IHC is innervated by 10-20 auditory nerve fibers, and every fiber contacts the IHC with a unmyelinated single ending to form a single ribbon synapse. The small size of the afferent boutons contacting IHCs (approximately 1 μm in diameter) enables recordings with exceptional temporal resolution to be made. Furthermore, the technique can be adapted to record from both pre- and postsynaptic cells simultaneously, allowing the transfer function at the synapse to be studied directly 2
. This method therefore provides a means by which fundamental aspects of neurotransmission can be studied, from multivesicular release to the elusive function of the ribbon in sensory cells.
Neuroscience, Issue 48, electrophysiology, whole-cell recording, patch clamp, synaptic transmission, ribbon synapse, multivesicular, dendrite, auditory nerve, hearing, hair cell.
Bladder Smooth Muscle Strip Contractility as a Method to Evaluate Lower Urinary Tract Pharmacology
Institutions: University of Pittsburgh School of Medicine, University of Pittsburgh School of Medicine.
We describe an in vitro
method to measure bladder smooth muscle contractility, and its use for investigating physiological and pharmacological properties of the smooth muscle as well as changes induced by pathology. This method provides critical information for understanding bladder function while overcoming major methodological difficulties encountered in in vivo
experiments, such as surgical and pharmacological manipulations that affect stability and survival of the preparations, the use of human tissue, and/or the use of expensive chemicals. It also provides a way to investigate the properties of each bladder component (i.e.
smooth muscle, mucosa, nerves) in healthy and pathological conditions.
The urinary bladder is removed from an anesthetized animal, placed in Krebs solution and cut into strips. Strips are placed into a chamber filled with warm Krebs solution. One end is attached to an isometric tension transducer to measure contraction force, the other end is attached to a fixed rod. Tissue is stimulated by directly adding compounds to the bath or by electric field stimulation electrodes that activate nerves, similar to triggering bladder contractions in vivo
. We demonstrate the use of this method to evaluate spontaneous smooth muscle contractility during development and after an experimental spinal cord injury, the nature of neurotransmission (transmitters and receptors involved), factors involved in modulation of smooth muscle activity, the role of individual bladder components, and species and organ differences in response to pharmacological agents. Additionally, it could be used for investigating intracellular pathways involved in contraction and/or relaxation of the smooth muscle, drug structure-activity relationships and evaluation of transmitter release.
The in vitro
smooth muscle contractility method has been used extensively for over 50 years, and has provided data that significantly contributed to our understanding of bladder function as well as to pharmaceutical development of compounds currently used clinically for bladder management.
Medicine, Issue 90, Krebs, species differences, in vitro, smooth muscle contractility, neural stimulation
A Method of Nodose Ganglia Injection in Sprague-Dawley Rat
Institutions: University of Illinois at Chicago, University of Illinois at Chicago, University of Illinois at Chicago.
Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located in the organs of the abdomen and thorax. The vagus nerve communicates information from stimuli such as heart rate, blood pressure, bronchopulmonary irritation, and gastrointestinal distension to the nucleus of solitary tract of the medulla. The cell bodies of the vagus nerve are located in the nodose and petrosal ganglia, of which the majority are located in the former. The nodose ganglia contain a wealth of receptors for amino acids, monoamines, neuropeptides, and other neurochemicals that can modify afferent vagus nerve activity. Modifying vagal afferents through systemic peripheral drug treatments targeted at the receptors on nodose ganglia has the potential of treating diseases such as sleep apnea, gastroesophageal reflux disease, or chronic cough. The protocol here describes a method of injection neurochemicals directly into the nodose ganglion. Injecting neurochemicals directly into the nodose ganglia allows study of effects solely on cell bodies that modulate afferent nerve activity, and prevents the complication of involving the central nervous system as seen in systemic neurochemical treatment. Using readily available and inexpensive equipment, intranodose ganglia injections are easily done in anesthetized Sprague-Dawley rats.
Neuroscience, Issue 93, neuroscience, nodose ganglia, vagus nerve, EMG, serotonin, apnea, genioglossus, cannabinoids
Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates
Institutions: University of California Riverside, University of California Riverside, University of California Riverside.
Close to two decades of research has established that astrocytes in situ
and in vivo
express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+
indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+
events using confocal microscopy. In essence, a “calcium roadmap” is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.
Neuroscience, Issue 85, astrocyte, plasticity, mGluRs, neuronal Firing, electrophysiology, Gq GPCRs, Bolus-loading, calcium, microdomains, acute slices, Hippocampus, mouse
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
Paired Whole Cell Recordings in Organotypic Hippocampal Slices
Institutions: University of Auckland, Stanford University.
Pair recordings involve simultaneous whole cell patch clamp recordings from two synaptically connected neurons, enabling not only direct electrophysiological characterization of the synaptic connections between individual neurons, but also pharmacological manipulation of either the presynaptic or the postsynaptic neuron. When carried out in organotypic hippocampal slice cultures, the probability that two neurons are synaptically connected is significantly increased. This preparation readily enables identification of cell types, and the neurons maintain their morphology and properties of synaptic function similar to that in native brain tissue. A major advantage of paired whole cell recordings is the highly precise information it can provide on the properties of synaptic transmission and plasticity that are not possible with other more crude techniques utilizing extracellular axonal stimulation. Paired whole cell recordings are often perceived as too challenging to perform. While there are challenging aspects to this technique, paired recordings can be performed by anyone trained in whole cell patch clamping provided specific hardware and methodological criteria are followed. The probability of attaining synaptically connected paired recordings significantly increases with healthy organotypic slices and stable micromanipulation allowing independent attainment of pre- and postsynaptic whole cell recordings. While CA3-CA3 pyramidal cell pairs are most widely used in the organotypic slice hippocampal preparation, this technique has also been successful in CA3-CA1 pairs and can be adapted to any neurons that are synaptically connected in the same slice preparation. In this manuscript we provide the detailed methodology and requirements for establishing this technique in any laboratory equipped for electrophysiology.
Neuroscience, Issue 91, hippocampus, paired recording, whole cell recording, organotypic slice, synapse, synaptic transmission, synaptic plasticity
Deriving the Time Course of Glutamate Clearance with a Deconvolution Analysis of Astrocytic Transporter Currents
Institutions: National Institutes of Health.
The highest density of glutamate transporters in the brain is found in astrocytes. Glutamate transporters couple the movement of glutamate across the membrane with the co-transport of 3 Na+
and 1 H+
and the counter-transport of 1 K+
. The stoichiometric current generated by the transport process can be monitored with whole-cell patch-clamp recordings from astrocytes. The time course of the recorded current is shaped by the time course of the glutamate concentration profile to which astrocytes are exposed, the kinetics of glutamate transporters, and the passive electrotonic properties of astrocytic membranes. Here we describe the experimental and analytical methods that can be used to record glutamate transporter currents in astrocytes and isolate the time course of glutamate clearance from all other factors that shape the waveform of astrocytic transporter currents. The methods described here can be used to estimate the lifetime of flash-uncaged and synaptically-released glutamate at astrocytic membranes in any region of the central nervous system during health and disease.
Neurobiology, Issue 78, Neuroscience, Biochemistry, Molecular Biology, Cellular Biology, Anatomy, Physiology, Biophysics, Astrocytes, Synapses, Glutamic Acid, Membrane Transport Proteins, Astrocytes, glutamate transporters, uptake, clearance, hippocampus, stratum radiatum, CA1, gene, brain, slice, animal model
Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology
Institutions: University of Washington.
The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes >10 μm in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.
Physiology, Issue 76, Biophysics, Molecular Biology, Biochemistry, Genetics, Cellular Biology, Proteins, Membranes, Artificial, Lipid Bilayers, Liposomes, Phospholipids, biochemistry, Lipids, Giant Unilamellar Vesicles, liposome, electrophysiology, electroformation, reconstitution, patch clamp
Pyrosequencing: A Simple Method for Accurate Genotyping
Institutions: Washington University in St. Louis.
Pharmacogenetic research benefits first-hand from the abundance of information provided by the completion of the Human Genome Project. With such a tremendous amount of data available comes an explosion of genotyping methods. Pyrosequencing(R) is one of the most thorough yet simple methods to date used to analyze polymorphisms. It also has the ability to identify tri-allelic, indels, short-repeat polymorphisms, along with determining allele percentages for methylation or pooled sample assessment. In addition, there is a standardized control sequence that provides internal quality control. This method has led to rapid and efficient single-nucleotide polymorphism evaluation including many clinically relevant polymorphisms. The technique and methodology of Pyrosequencing is explained.
Cellular Biology, Issue 11, Springer Protocols, Pyrosequencing, genotype, polymorphism, SNP, pharmacogenetics, pharmacogenomics, PCR