This work was supported by SFB 604, DFG MO 1421/2-1 and Krebshilfe 107089 (to H.M.). A.S. is recipient of a Young Investigator Award from the Children&#8217;s Tumor Foundation (New York, USA).

The present study was performed according to the Protection of Animals Act of the Federal Republic of Germany (Tierschutzgesetz der Bundesrepublik Deutschland) and was approved by the Thuringian State Office for Food Safety and Consumer Protection (Th&#252;ringer Landesamt f&#252;r Lebensmittelsicherheit und Verbraucherschutz).
1. Setting Up the Measurements
<ol>
	<li>Anesthetize the mice by isoflurane/O2 inhalation - for initiation of anesthesia 3%, for maintenance 2% isoflurane in 100% oxygen (Figure 1). Confirm sufficient anesthesia by testing simple reflexes such as movement reflexes and testing of sensitivity for low-grade pain. Use of ointment on the eyes to prevent dryness during anesthesia is recommended but not indispensable because the procedure typically takes just 30 min/animal in total. In case of survival experiments, administer timely longer acting analgesics for managing postoperative pain.</li>
	<li>Shave the fur covering the hind limbs with an electric razor and perform depilation with a commercially available hair-removal cream while animals are already under analgesia. In order to maintain pathogen-free status throughout the procedure, wear aseptic gloves and always use instruments carefully cleaned with 70% ethanol.</li>
	<li>Control body temperature stability by a feedback controlled heating plate and rectal thermo probe (Figure 2). If necessary, please use a sterile drape to cover the heating plate between the animals in order to keep a sterile experimental environment. Furthermore, it is recommended to use a heating plate, which is electronically controlled via an integrated sensor to limit the heating temperature to &#8804;40 &#176;C in order to avoid tissue damage. 
		<img alt="Figure 1" fo:content-width="4in" fo:src="/files/ftp_upload/51181/51181fig1highres.jpg" src="/files/ftp_upload/51181/51181fig1.jpg" /> 
		Figure 1. Experimental setup showing an anesthetized mouse with shaved hind limbs. </li>
	<li>Take electrocardiography (ECG) recordings to monitor the heart rate as a vital parameter. Install three electrodes for ECG recordings as follows: one electrode under the skin of each forelimb and one electrode under the skin in the neck area.</li>
	<li>Place ring electrodes using contact gel for optimal conductivity / transfer resistance. The sensing electrode (labeled in black) is placed at the position where the gastrocnemius muscle has its maximum diameter. The reference electrode (indicated in red) is placed just beneath the sensing electrode. 
		Note: Please see 'Material table' for equipment details.</li>
	<li>Mark the correct positions of measurement with predetermined but consistent distances between the proximal and distal sciatic nerve stimulation and the lead electrode. Experimental proposal: At a distance of 4 mm from the sensing electrode, the distal stimulation will take place. In a distance of 16 mm to the sensing electrode, the proximal stimulation will be carried out (Figure 2).</li>
</ol>
<img alt="Figure 2" fo:content-width="6in" fo:src="/files/ftp_upload/51181/51181fig2highres.jpg" src="/files/ftp_upload/51181/51181fig2.jpg" /> 
	Figure 2. Representative picture showing the experimental situation just prior to the beginning of the measurements. The white arrow indicates the position of the sensing (black) and reference (red) electrode at the gastrocnemius muscle of the left hind limb. The stimulation by needle electrodes will be performed at defined positions in relation to the black sensing electrode. The point of distal stimulation (black mark with "d.s." at the left hind limb) has a distance of 4 mm from the sensing electrode; the place of proximal stimulation (black mark with "p.s.") is 16 mm away. The red line on the right hind limb shows the approximate anatomical course of the sciatic nerve. Furthermore, the rough positions of relevant hind limb muscles are shown as landmarks. The asterisk indicates the rectal thermal probe.
2. Measurement
<ol>
	<li>Principle: Perform a series of nerve stimulations with repetitively generated single square-wave pulses of 0.1 msec duration by monopolar disposable 28 G needle electrodes (repetition rate 200 msec; see Figure 3A). For off-line data analysis, it is recommended to simultaneously acquire stimulation signals together with the neuromuscular function response curve (Figure 3B) due to nerve stimulation. Average a series of maximal responses ("representative neuromuscular function response") for subsequent data analysis. In order to produce reliable data, record and later average at least 3 independent, optimal response curves per stimulation site and animal. 
		<img alt="Figure 3" fo:content-width="6in" fo:src="/files/ftp_upload/51181/51181fig3highres.jpg" src="/files/ftp_upload/51181/51181fig3.jpg" /> 
		Figure 3. Procedure for data acquisition and analysis (schematic presentation). Repetitively generated pulses are applied to the sciatic nerve via needle electrodes (upper row in Figure 3A). Simultaneously, several corresponding neuromuscular response curves are recorded (lower row in Figure 3A). When averaged and magnified, neuromuscular response curves due to stimulation pulses (upper row in Figure 3B) show the following characteristic properties (lower row in Figure 3B): Latency of the signal response as well as duration and amplitude of the signal are indicated and can be obtained for subsequent calculations and statistics. Irregular signal conduction and / or suboptimal recordings usually result in various signal deformations with more than just one positive and one negative deflection or signal deformation (bimodal shape) with reduced amplitude (Figure 3C).</li>
	<li>Perform proximal stimulation with a needle electrode at determined position ("p.s." in Figure 2).
		<ol>
			<li>In order to achieve best recording conditions with maximum amplitudes, visualize the actual response curves simultaneous to the stimulation process. By doing so, the experimenter is able to immediately assess the shape of the response curves as well as the size of the amplitude.</li>
			<li>If necessary, slightly and carefully manipulate the position and / or the angle of the stimulation needle with respect to the sciatic nerve. This gentle optimization of stimulation conditions allows reaching constant amplitudes with the largest possible value and a response curve of typical biphasic shape (Figure 4).</li>
		</ol>
	</li>
	<li>Perform distal stimulation with a needle electrode at determined position ("d.s." in Figure 2).</li>
	<li>After completion of measurement, transfer the tested mouse to a separate cage until it has regained sufficient consciousness to maintain sternal recumbency. Do not leave an animal unattended and in the company of other animals until it has fully recovered from anesthesia. Administer timely longer acting analgesics for managing postoperative pain. Systemic administration of nonsteroidal anti-inflammatory drugs (NSAIDs) and opioids are recommended for 1-3 days.</li>
	<li>Alternatively, sacrifice the still anesthetized mouse in a quick and humane fashion without any further pain for the animal, e.g. by neck dislocation.</li>
</ol>
<img alt="Figure 4" fo:content-width="6in" fo:src="/files/ftp_upload/51181/51181fig4highres.jpg" src="/files/ftp_upload/51181/51181fig4.jpg" /> 
	Figure 4. Illustration to determine the CMAP recordings with maximum amplitudes. A complete registration series is presented. (a) Insertion point with minimal CMAP response. (b) Slight stimulation needle movement results in CMAP recordings with maximum amplitudes. (c) Additional changes in needle placement produce CMAP recordings with different amplitudes including near-maximum amplitudes. (d) Stimulation needle replacement with serial CMAP recordings of near-maximum amplitudes. Note: Typical decrement in CMAP amplitudes can occur during repetitive stimulation at optimal stimulation site12,13. Asterisks indicate CMAP recordings with maximum amplitudes depicted for averaging.
3. Analysis
<ol>
	<li>Extract nerve conduction parameters based on the representative neuromuscular function response data set using an appropriate software package (e.g. AtisaPro). 
		Note: Please handle time-related data determination with special care because inflection point determination of compound motor action potential (CMAP) onset and termination can be difficult. A procedure with verified reproducibility is given in Figure 3B, where tangents on signal deflections after onset and before termination are used.</li>
	<li>Calculate 'signal latency' and 'CMAP' values.
		<ol>
			<li>Latency' represents the time delay between stimulation and CMAP onset, whereas the time span between CAMP onset of initial negative deflection to initial return to baseline is called 'CMAP duration'. Use the difference between distal and proximal latencies to calculate conduction velocity together with the distance between distal and proximal stimulation sites. 
				conduction velocity = latency / distance between stimulation sites</li>
			<li>CMAP' (compound motor action potential) amplitude represents the magnitude between maximum positive and negative turnaround point of the CMAP signal (given in mV). 
				CMAP = value 'positive turnaround point' - value 'negative turnaround point'</li>
		</ol>
	</li>
</ol>

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	<li>Kimura, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3rd+ed.">3rd ed.</a> Electrodiagnosis in Diseases of Nerve and Muscle. , (2001).</li><li>Rutkove, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+temperature+on+neuromuscular+electrophysiology.">Effects of temperature on neuromuscular electrophysiology.</a> Muscle Nerve. 24, 867-882 (2001).</li><li>Xia, R. H., Yosef, N., Ubogu, E. 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The authors have nothing to disclose.

The described protocol provides an easy and reliable method to determine sciatic nerve conduction properties on anesthetized mice without the need to expose the nerve of interest. Nevertheless, this experimental procedure causes tissue injury by needle puncture. It is therefore a reasonable option to sacrifice the animals after finishing the recordings. However, compared to other more invasive procedures, which require the exposure of the nerve prior to recordings, tissue damage is comparably small 3,14. There...

Electrophysiological measurements are an indispensable tool for investigating the functional integrity of peripheral nerves in both clinical and laboratory environments. In humans, a large number of neuromuscular disorders and neuropathies diagnostically rely on electrophysiological measurements. By measuring nerve properties as conduction velocity or potential amplitudes of the signal, it is possible to characterize the rough origin of peripheral nerve diseases.
The nerve conduction velocity is highly dependent on rapid signal propagation enabled by myelination. Therefore, demyelinating processes generally show decreased conduction velocities4. The compound motor action potential (CMAP) - correlating with the number of functional axons - is an indicator for axonal damage when significantly reduced5.
Thus, by means of electrophysiological methods the etiology of peripheral nerve damage can be discriminated, such as for hereditary neuropathies6,7, diabetic neuropathy8,9, chronic inflammatory demyelinating polyneuropathies (CIDP)10, or metabolic neuropathies11.
Normally, in the human application noninvasive recordings on the sural or ulnar nerve are preferred. In mice, it is straightforward to analyze nerve properties of sciatic nerves, the largest nerve of the peripheral nervous system (PNS) containing both large - and small-caliber axons of the motoric and sensory system.
The procedure as demonstrated here is a quick, easy and reliable method to measure all standard values relevant for electrophysiology on peripheral nerves in the intact mouse. By taking recordings from a preserved organism, physiological conditions of the nerve environment are guaranteed.

<table border="0" cellpadding="0" cellspacing="0" width="917">
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:321px;">Concentric Needle Electrodes (Stimulation)</td>
			<td style="width:311px;">Natus Medical Incorporated 
			San Carlos, CA 94070, USA</td>
			<td style="width:119px;">9013S0901</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Digital Ring Electrodes (Recording)</td>
			<td style="width:311px;">Natus Medical Incorporated 
			San Carlos, CA 94070, USA</td>
			<td>9013S0302</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">ToM - Tower of Measurement (A/D converter)</td>
			<td>GJB Datentechnik GmbH, Langewiesen, Germany</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">AtisaPro, Data acquisition &#38; analysis software</td>
			<td>GJB Datentechnik GmbH, Langewiesen, Germany</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HSE-Stimulator T</td>
			<td colspan="2">Hugo Sachs Elektronik, Hugstetten, Germany</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo electrophysiological measurements on mouse sciatic nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological techniques, in the understanding of the underlying pathophysiology1. Here we describe a method to perform electrophysiological studies on mouse sciatic nerves in vivo.
The animals are anesthetized with isoflurane in order to ensure analgesia for the tested mice and undisturbed working environment during the measurements that take about 30 min/animal. A constant body temperature of 37 &#176;C is maintained by a heating plate and continuously measured by a rectal thermo probe2. Additionally, an electrocardiogram (ECG) is routinely recorded during the measurements in order to continuously monitor the physiological state of the investigated animals.
Electrophysiological recordings are performed on the sciatic nerve, the largest nerve of the peripheral nervous system (PNS), supplying the mouse hind limb with both motoric and sensory fiber tracts. In our protocol, sciatic nerves remain in situ and therefore do not have to be extracted or exposed, allowing measurements without any adverse nerve irritations along with actual recordings. Using appropriate needle electrodes3 we perform both proximal and distal nerve stimulations, registering the transmitted potentials with sensing electrodes at gastrocnemius muscles. After data processing, reliable and highly consistent values for the nerve conduction velocity (NCV) and the compound motor action potential (CMAP), the key parameters for quantification of gross peripheral nerve functioning, can be achieved.

We conducted a series of in vivo electrophysiological measurements on sciatic nerves of 12 mice in total for this study: 6 animals of each gender. The measurements were performed with the presented protocol and delivered the following results:
Both male and female mice display a mean sciatic nerve conduction velocity of approximately 20 m/sec (Figure 5). This is consistent with other measurements in the literature. Furthermore, it shows that there are no relevant diff...

Watch this Scientific Journal Video about In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves at JoVE.com

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Measurements of nerve conduction properties in vivo exemplify a powerful tool to characterize various animal models of neuromuscular diseases. Here, we present an easy and reliable protocol by which electrophysiological analysis on sciatic nerves of anesthetized mice can be performed.

In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological studies allow a rational classification of various neuromuscular diseases and are of help, together with neuropathological ...

in-vivo-electrophysiological-measurements-on-mouse-sciatic-nerves

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Neuroscience

33.1K 次观看.  Fritz Lipmann Institute.  体内神经传导特性的测量体现了强大的工具来表征神经肌肉疾病的多种动物模型。在这里，我们提出了一个简单而可靠的协议，其中对麻醉小鼠坐骨神经电生理分析可以执行。 

视频: In Vivo Electrophysiological Measurements on Mouse Sciatic Nerves

Electrophysiological Measurements from a Moth Olfactory System

Insect olfactory systems provide unique opportunities for recording odorant-induced responses in the forms of electroantennograms (EAG) and single sensillum recordings (SSR), which are summed responses from all odorant receptor neurons (ORNs) located on the antenna and from those housed in individual sensilla, respectively. These approaches have been exploited for getting a better understanding of insect chemical communication. The identified stimuli can then be used as either attractants or repellents in management strategies for insect pests.

Insect olfactory systems provide unique opportunities for recording odorant-induced responses in the forms of electroantennograms (EAG) and single sensillum recordings (SSR), which are summed responses from all odorant receptor neurons (ORNs) located on the antenna and from those housed in individual sensilla, respectively.

Insect olfactory systems provide unique opportunities for recording odorant-induced responses in the forms of electroantennograms (EAG) and single ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Electrophysiological Measurements and Analysis of Nociception in Human Infants

Pain is an unpleasant sensory and emotional experience. Since infants cannot verbally report their experiences, current methods of pain assessment are based on behavioural and physiological body reactions, such as crying, body movements or changes in facial expression. While these measures demonstrate that infants mount a response following noxious stimulation, they are limited: they are based on activation of subcortical somatic and autonomic motor pathways that may not be reliably linked to central sensory processing in the brain. Knowledge of how the central nervous system responds to noxious events could provide an insight to how nociceptive information and pain is processed in newborns.
The heel lancing procedure used to extract blood from hospitalised infants offers a unique opportunity to study pain in infancy. In this video we describe how electroencephalography (EEG) and electromyography (EMG) time-locked to this procedure can be used to investigate nociceptive activity in the brain and spinal cord.
This integrative approach to the measurement of infant pain has the potential to pave the way for an effective and sensitive clinical measurement tool.

The assessment and treatment of pain in infants is difficult because infants cannot verbally report their experience. In this video we describe quantitative electrophysiological methods and analysis techniques that can be used to measure the response to noxious events from the infant nervous system.

Pain is an unpleasant sensory and emotional experience. Since infants cannot verbally report their experiences, current methods of pain assessment are ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous system. SC-selective genetic manipulation in living animals is a powerful technique for studying the molecular and cellular mechanisms of SC myelination and demyelination in vivo. While knockout/knockin and transgenic mice are powerful tools for studying SC biology, these methods are costly and time consuming. Viral vector-mediated transgene introduction into the sciatic nerve is a simpler and less laborious method. However, viral methods have limitations, such as toxicity, transgene size constraints, and infectivity restricted to certain developmental stages. Here, we describe a new method that allows selective transfection of myelinating SCs in the rodent sciatic nerve using electroporation. By applying electric pulses to the sciatic nerve at the site of plasmid DNA injection, genes of interest can be easily silenced or overexpressed in SCs in both neonatal and more mature animals. Furthermore, this in vivo electroporation method allows for highly efficient simultaneous expression of multiple transgenes. Our novel technique should enable researchers to efficiently manipulate SC gene expression, and facilitate studies on SC development and function.

In Vivo Gene Transfer to Schwann Cells in the Rodent Sciatic Nerve by Electroporation

Here, we present an in vivo technique for gene transfer to Schwann cells (SCs) in the rodent sciatic nerve. This simple technique is useful for investigating signaling mechanisms involved in the development and maintenance of myelinating SCs.

The formation of the myelin sheath by Schwann cells (SCs) is essential for rapid conduction of nerve impulses along axons in the peripheral nervous ...

Analysis of Immune Cells in Single Sciatic Nerves and Dorsal Root Ganglion from a Single Mouse Using Flow Cytometry

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of the PNS are affected by injury and disease, affecting the nerve function and the capacity for regeneration. Neuronal immune cells are commonly analyzed by immunofluorescence (IF). While IF is essential for determining the location of the immune cells in the nerve, IF is only semi-quantitative and the method is limited to the number of markers that can be analyzed simultaneously and the degree of surface expression. In this study, flow cytometry was used for quantitative analysis of leukocyte infiltration into sciatic nerves or dorsal root ganglions (DRGs) of individual mice. Single cell analysis was performed using DAPI and several proteins were analyzed simultaneously for either surface or intracellular expression. Both sciatic nerves from one mouse that were treated according to this protocol generated &#8805; 30,000 single nucleated events. The proportion of leukocytes in the sciatic nerves, determined by expression of CD45, was approximately 5% of total cell content in the sciatic nerve and approximately 5-10% in the DRG. Although this protocol focuses primarily on the immune cell population within the PNS, the flexibility of flow cytometry to measure a number of markers simultaneously means that the other cells populations present within the nerve, such as Schwann cells, pericytes, fibroblasts, and endothelial cells, can also be analyzed using this method. This method therefore provides a new means for studying systemic effects on the PNS, such as neurotoxicology and genetic models of neuropathy or in chronic diseases, such as diabetes.

Quantitative analysis of cell content within the murine sciatic nerve is difficult due to the scarcity of the tissue. This protocol describes a method for tissue digestion and preparation that provides sufficient cells for flow cytometry analysis of immune cell populations from nerves of individual mice.

Nerve-resident immune cells in the peripheral nervous system (PNS) are essential to maintaining neuronal integrity in a healthy nerve. The immune cells of ...

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings provide a sensitive approach to measure nerve conduction in humans and rodent models. To broaden the technical possibilities for electromyography in mice, the measurement of compound muscle action potentials (CMAPs) from the brachial plexus nerve in the forelimb using needle electrodes is described here. CMAP recordings after stimulating the sciatic nerve in hindlimbs have been previously described. The newly introduced method here allows for the evaluation of the nerve conductivity at an additional site, and thus provides a more profound overview of the neuromuscular functionality. The technique provides information on both the relative number of functional axons and the myelination level. Thereby, this method can be applied to assess both axonal diseases as well as demyelinating conditions. This minimally invasive method does not require extraction of the nerve and therefore it is suitable for repeated measurements for longitudinal follow-up in the same animal. Similar recordings are performed in clinical setups to emphasize the translational relevance of the method.

In Vivo Electrophysiological Measurement of Compound Muscle Action Potential from the Forelimbs in Mouse Models of Motor Neuron Degeneration

The measurement of nerve conduction is a useful tool to assess mouse models of neurodegeneration but it is frequently only applied to stimulate the sciatic nerve in hindlimbs. Here, we describe a technique to measure compound muscle action potential (CMAP) in vivo in the mouse forelimb muscles innervated by the brachial plexus.

Assessing the functionality of the nerve axon provides detailed information on the progression of neuromuscular disorders. Electrophysiological recordings ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...