The authors are grateful for the assistance of Mrs. Sharon Spana in rodent monitoring, handling and experimentation. PhD funding support has been provided by University of Sydney and Australian Research Centre for Excellence in Vision.

This protocol follows the animal care guidelines of University of Sydney.
1. Making Electrodes
<ol>
	<li>Construct the positive electrode (the one which will contact the cornea) from a short (5 cm) length of platinum wire 1-2 mm in diameter. Fashion it into a loop a few mm in diameter. Connect this loop to a conventional lead, long enough to reach the input stage of your amplifier (see Figure 4).</li>
	<li>Construct the negative electrode (which will go in the animal&#8217;s mouth) using an Ag/AgCl pellet 1-2 mm in diameter, also connected to a convention lead (see Figure 4).</li>
	<li>As a reference electrode (which will go into the animal&#8217;s rump), use a clean hypodermic needle (23 G), also connected to a lead of appropriate length (see Figure 4).</li>
	<li>Ideally, use three-lead cables provided by instrument manufacturers, to connect the three electrodes (positive&#160;&#8594; cornea, negative &#8594; mouth, reference &#8594; rump) to the amplifier.</li>
</ol>
2. Connection and Calibration of Light Stimulus and ERG Set-up
<ol>
	<li>Create (or locate) a small recording laboratory, which can be made dark. Equip with either or both of an over-the-bench light made red or a red head-lamp.</li>
	<li>Use a lux meter to confirm that red light illuminance reaching the rat&#8217;s eye during setup does not exceed 1 lux. 
	Note: A neutral density filter can be used to reduce lamp brightness and the source of lamp light must specifically emit red light. Dark adaption will be compromised if light sources emit lower (visible) wavelengths.</li>
	<li>Seal off all stray light entering the recording laboratory (this often requires persistence with opaque tape) and prepare a neutral density filter (this can be purchased in sheets) large enough to fit over, and so dim, any computer screen you will have in the lab. 
	Note: Stray light and the light of a screen are sufficient to prejudice dark adaptation of the rat eye.</li>
	<li>Connect the amplifier to data acquisition hardware. Connect positive, negative and reference leads to the amplifier. Make sure the computer and the LED Ganzfeld power supply unit are securely connected to a ground source. 
	Note: Some labs have specialised grounding points, connected to a building ground; a water pipe is an effective alternative.</li>
	<li>Calibrate the LED light source with a research-quality radiometer. Fix the meter&#8217;s sensor in the position at which the animal&#8217;s eye will be located during an experiment.</li>
	<li>Program the Ganzfeld LEDs to run a full-field ERG protocol with step-wise increases in flash energy, flash duration, flash repetition and time between flashes, termed interstimuls interval (ISI), settings. For an example full-field protocol see Table 1. 
	Note: The full-field ERG flashes increase from repetitive dim flashes to bright flashes in a step wise fashion. The twin flash program follows on from the full-field protocol and enables isolation of rod and cone responses.</li>
</ol>
3. Day Prior to ERG Experimentation
<ol>
	<li>Dark adapt Sprague-Dawley rats for 12 hr before recording. It is convenient to do this in the recording laboratory, once stray light has been eliminated.</li>
</ol>
4. Day of ERG Experimentation
<ol>
	<li>Arrange for the animal to be gently heated while recording. We use a light metal platform built so that the animal&#8217;s head can rest at the correct point at the entry to the Ganzfeld. The platform has inbuilt tubing through which we pump water preheated to 40 &#176;C in a water bath. 
	Note: Experience shows that this keeps the animal&#8217;s core temperature at 37 &#176;C.</li>
	<li>Weigh the rat under dark conditions. Record weight and make up correct ketamine (60 mg/kg) and xylazine (5 mg/kg) dose. Restrain the rat gently and inject anaesthetic intraperitoneally.</li>
	<li>Note time of injection. Once the animal is unconscious (usually within 5 min) check depth of anaesthesia by lightly pinching one foot pad, to see if a reflex response is present. It is best to wait until this reflex is absent or weak, before proceeding.</li>
	<li>Apply a single drop of atropine and another of proxmethacaine to cornea.</li>
	<li>Cut a 10 cm length of black thread. Make a loop with a simple knot and slip the loop over the equator of the eye. Tighten it slightly; the effect is to draw the eyeball slightly forward, with minimal pressure. This keeps the cornea clear from the eyelids.</li>
	<li>Apply carbomer eye drops to the cornea surface. Ensure carbomer remains on corneal surface and does not spill onto the eyelids or the face.</li>
	<li>Place absorbent bedding on top of heated platform.</li>
	<li>Position rat on the bedding, with the head in the recommended place in the opening of the Ganzfeld.</li>
	<li>Insert internal temperature probe into the rectum. Secure temperature probe in position by taping probe cord to the tail.</li>
	<li>Insert the reference electrode (the 23 G needle) subcutaneously into the rear leg, and connect to amplifier.</li>
	<li>Place the negative electrode (the Ag/AgCl pellet) securely in the mouth. To prevent this slipping out the mouth, affix the connecting lead to a stable surface.</li>
	<li>Position the positive electrode over the centre of the cornea. Using a micromanipulator, ensure that the electrode touches the cornea gently.</li>
	<li>Check body temperature is at 37.0 - 37.5 &#176;C.</li>
	<li>Once the animal is properly positioned and electrodes are in place, drape the whole setup (Ganzfeld and animal) with an opaque material (to preserve dark adaptation). We use a soft black cloth.</li>
	<li>In the acquisition software set at a 2 KHz sampling rate with a collection time of 100-1000 msec with 5 msec of pre-collection sampling. Set the band pass filters to 1-1,000 Hz and ensure that sampling is triggered to sample the period of ~250 msec following a flash.</li>
	<li>Check the recording baseline. It should be free of extraneous noise, but show some amplifier noise and a respiratory oscillation.</li>
	<li>If the baseline shows extraneous noise, begin troubleshooting. Most problems are related to slippage in electrode position, or grounding. Use a faraday cage to ensure recordings are free of extraneous noise.</li>
	<li>Run a test flash, 0.4 log scot cd.s.m-2. An ERG waveform similar to Figure 2A should appear. In our laboratory typical responses for a 0.4 log scot cd.s.m-2 flash are (a-wave: -474 &#177; 39 &#181;V and b-wave: 1,512 &#177; 160 &#181;V, n = 11).</li>
	<li>Allow animal to dark re-adapt for 10 min. It is convenient to use these 10 min to recheck the baseline.</li>
	<li>Following confirmation of stable signal begin recording.</li>
	<li>At the end of the recording session, check that body temperature was maintained. Remove electrodes. Reapply carbomer polymer to corneas. Allow the animal to recover on a heat pad until it is fully mobile and active, before returning to animal housing.</li>
</ol>
5. Remote Ischemia
<ol>
	<li>Perform remote ischemia in either awake or anesthetised rodents.</li>
	<li>If the animal is anaesthetised, lay it on a heated platform (above) and slip the sphygmomanometer cuff over the upper part of the hind-limb, clear of the knee.</li>
	<li>If the animals are used to being handled, it is possible to perform this procedure without anaesthesia; this requires two people. One person restrains the animal gently and the second applies the sphygmomanometer cuff and operates the sphygmomanometer.</li>
	<li>For awake animals, use a piece of towel ~15 cm x 30-50 cm to gently wrap the animal, with one hindlimb free. Lay the animal on its back on (say) the left forearm, with its head tucked between the holder&#8217;s arm and torso, and place the cuff as just described.</li>
	<li>Deflate the cuff and ensure the air pressure valve is closed. Pump the cuff to 160 mmHg in anesthetised animals, and to 180 mmHg in awake animals. This exceeds systolic pressure (usually 140 mmHg and 160 mmHg respectively).</li>
	<li>Maintain these pressures as required, using the hand-held pump.</li>
	<li>After the planned time for ischemia (we use 2 periods of 5 min separated by 5 min reperfusion), deflate the cuff pressure by loosening the air pressure valve.</li>
	<li>Confirm the effect of remote ischemia with a skin temperature probe attached to the footpad. Skin temperature typically falls from 32-30 &#176;C, over 5 min and recovers on reperfusion.</li>
</ol>
6. Light Damage
<ol>
	<li>Ensure that rats are in a dark-adapted overnight, before the light damage procedure.</li>
	<li>At the appropriate time following limb ischemia (in our experiments without delay), each animal is placed singly into a plexiglass boxes, with water and food in floor-based containers. 
	Note: Light-induced damage can only be undertaken in albino animals.</li>
	<li>Switch on a pre-calibrated 1,000 lux white light at a standard time (usually 9 am) and maintain this condition for 24 hr.</li>
</ol>
7. ERG Data Extraction and Analysis
<ol>
	<li>Acquire averaged wave forms of the ERG. If required, correct for a non-zero baseline, by subtraction.</li>
	<li>Measure the amplitude of the a-wave (presented at mid- to high stimulus intensities), as the voltage difference between the baseline and the first (&#60;30 msec latency) trough (Figure 1).</li>
	<li>Measure the b-wave amplitude as the voltage difference between the peak of the a-wave and the positive of the following wave, typically occurring at a latency of 80-100 msec (Figure 1).</li>
	<li>Isolate oscillatory potentials by using a Fourier transform to filter data from 60-235 Hz, with a 90 Hz transition band7. If required the isolated oscillatory potential signal can then be subtracted from the unfiltered waveform to confirm the identity of the a-wave trough.</li>
	<li>The implicit time (latency) of the a- and b-wave peaks can also be a useful measure (Figure 1). Use the twin flashes to isolate the rod response. Subtract the cone response (flash 2) from the mixed response (flash 1) to isolate the rod response (Figure 2).</li>
	<li>Normalize individual light intensity a-wave and b-wave amplitudes (post-treatment/post-treatment-baseline) or averaged for treatment groups. Intensity-response curves plot the group amplitudes and error against flash energy.</li>
</ol>

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Jonathan Stone is the director of CSCM Pty Ltd

The dark-adapted flash ERG method described above is a reliable method for assessing retinal function in rats. Both the a-wave and b-wave were reduced by light damage. Remote ischemic preconditioning mitigated light damage-induced reductions in the a-wave and b-wave. This preservation of retinal function suggests that remote ischemic preconditioning has induced neuroprotection, resembling other forms of protective preconditioning such as hypoxia, ischemia and exercise8-10. The ERG signal recorded is determined...

The ERG is an electrical potential generated by the retina in response to light, and recorded from the corneal surface of the eye. When conditions of recording are managed carefully, the ERG can be used in a variety of ways to assess retinal function. Here we described how to record the &#8216;flash ERG&#8217;, the potential generated when the retina is exposed to a brief, bright flash presented in a Ganzfeld background. The Ganzfeld disperses the light homogenously and the flash of light reaches the whole retina approximately uniformly. If the retina is dark adapted before recording, and the dark-adaptation is maintained as the animal is prepared for recording, the ERG obtained is generated by both rod and cone photoreceptors.
The dark-adapted flash ERG has a characteristic waveform, which has been analysed in two ways. First, early and late components of the ERG waveform have been distinguished, and related to the sequence of neuronal activation in the retina. The earliest component is a short-latency negative-going potential, the a-wave (Figure 1). This is followed by a positive-going potential, called the b-wave. The rising phase of the b-wave shows oscillations, which are considered a separate component (oscillatory potentials or OPs). The a-wave is considered to be generated by photoreceptors, the b-wave by cells of the inner nuclear layer, and the OPs by amacrine cells1.
Based on the stimulus strength, responses to very dim flashes termed the scotopic threshold response are possible. The scotopic threshold response is understood to be generated from the retinal ganglion cells2-4. Second, the flash ERG can be separated by light adaptation, or by a two flash protocol described below, into rod- and cone-driven components. Under photopic conditions, the a-wave is not detectable in rats, because the cone population is low, but OPs and a b-wave are clear5. In primates, whose retinas have higher cone populations, both rod- and cone- pathways generate a detectable a-wave6.
Two useful measures often extracted from the flash ERG are the amplitudes of the a- and b-waves, measured as in Figure 1, with typical flash responses shown in Figure 2. When the photoreceptor population is reduced, for example by exposure to damagingly bright light, all components of the ERG are reduced. Neuroprotective interventions, such as remote ischemic preconditioning (RIP), can be validated by the preservation of the amplitudes of the a- and b-waves (Figure 3). In summary, the analysis of the ERG enables comparisons between healthy, light damaged and neuroprotected retina.

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			<td>PC computer</td>
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			<td>Powerlab, 4 channel acquistion hardware</td>
			<td>AD Instruments</td>
			<td>PL 35044</td>
			<td>Acquistion of ERG</td>
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			<td>Animal Bio Amp</td>
			<td>AD Instruments</td>
			<td>FE 136</td>
			<td>Amplifier for ERG</td>
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			<td>Lab chart</td>
			<td>AD Instruments</td>
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			<td>Signal collection software</td>
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			<td>Ganzfield</td>
			<td>Photometric solutions</td>
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			<td>Connect postive, negative ground electrodes to amplifier</td>
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			<td>Lead wires -aligator</td>
			<td>AD Instruments</td>
			<td></td>
			<td>ground ganzfield and acquistion hardware to computer</td>
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			<td>Platinum wire 95%</td>
			<td>A&#38;E metals</td>
			<td></td>
			<td>postive electrode</td>
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			<td>Mouth electrode Ag/AgCl Pellet</td>
			<td>SDR</td>
			<td>E205</td>
			<td>negative electode</td>
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			<td>26 gauge needle</td>
			<td>BD</td>
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			<td>ground electode</td>
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			<td>Water pump</td>
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			<td>Water bath</td>
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			<td>Tubing</td>
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			<td>Homeothermic blanket system with flexible probe</td>
			<td>Harvard Appartus</td>
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			<td>Bausch &#38; Lomb</td>
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			<td>topical mydriasis</td>
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			<td>Proxmethycaine 0.5% w/v</td>
			<td>Bausch &#38; Lomb</td>
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			<td>topical anaesthetic</td>
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			<td>Visco tears eye drops</td>
			<td>Novartis</td>
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			<td>carbomer polymer</td>
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			<td>Blu tac</td>
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			<td>Troy Laboratories Pty Ltd</td>
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			<td>Scale</td>
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using the electroretinogram to assess function in the rodent retina and the protective effects of remote limb ischemic preconditioning

The ERG is the sum of all retinal activity. The ERG is usually recorded from the cornea, which acts as an antenna that collects and sums signals from the retina. The ERG is a sensitive measure of changes in retinal function that are pan-retinal, but is less effective for detecting damage confined to a small area of retina. In the present work we describe how to record the &#8216;flash&#8217; ERG, which is the potential generated when the retina is exposed to a brief light flash. We describe methods of anaesthesia, mydriasis and corneal management during recording; how to keep the retina dark adapted; electrode materials and placement; the range and calibration of stimulus energy; recording parameters and the extraction of data. We also describe a method of inducing ischemia in one limb, and how to use the ERG to assess the effects of this remote-from-the-retina ischemia on retinal function after light damage. A two-flash protocol is described which allows isolation of the cone-driven component of the dark-adapted ERG, and thereby the separation of the rod and cone components. Because it can be recorded with techniques that are minimally invasive, the ERG has been widely used in studies of the physiology, pharmacology and toxicology of the retina. We describe one example of this usefulness, in which the ERG is used to assess the function of the light-damaged retina, with and without a neuroprotective intervention; preconditioning by remote ischemia.

The protocol can be used to measure visual function of rodent retina in vivo. The a-wave, a measure of photoreceptor function, and the b-wave, a measure of inner retina function, are annotated in Figure 1.
The rod-dominated ERG signal increases with the increasing light stimulus, as shown in Figure 2A. The a-wave becomes apparent at ~0.4 log scot cd.s.m-2 and the amplitude of the a-wave increases until saturation at 2.5 log scot cd.s.m...

Watch this Scientific Journal Video about Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning at JoVE.com

Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning

The electroretinogram (ERG) is an electrical potential generated by the retina in response to light. This paper describes how to use the ERG to assess retinal function, in dark-adapted rats, and how it can be can be used to assess a neuroprotective intervention, in the present case remote ischemic preconditioning.

The ERG is the sum of all retinal activity. The ERG is usually recorded from the cornea, which acts as an antenna that collects and sums signals from the ...

using-electroretinogram-to-assess-function-rodent-retina-protective

Research

JoVE Journal

Neuroscience

16.0K vistas.  University of Sydney. The electroretinogram (ERG) is an electrical potential generated by the retina in response to light. This paper describes how to use the ERG to assess retinal function, in dark-adapted rats, and how it can be can be used to assess a neuroprotective intervention, in the present case remote ischemic preconditioning.

Video: Using the Electroretinogram to Assess Function in the Rodent Retina and the Protective Effects of Remote Limb Ischemic Preconditioning

Strategies for Study of Neuroprotection from Cold-preconditioning

Neurological injury is a frequent cause of morbidity and mortality from general anesthesia and related surgical procedures that could be alleviated by development of effective, easy to administer and safe preconditioning treatments. We seek to define the neural immune signaling responsible for cold-preconditioning as means to identify novel targets for therapeutics development to protect brain before injury onset. Low-level pro-inflammatory mediator signaling changes over time are essential for cold-preconditioning neuroprotection. This signaling is consistent with the basic tenets of physiological conditioning hormesis, which require that irritative stimuli reach a threshold magnitude with sufficient time for adaptation to the stimuli for protection to become evident.
Accordingly, delineation of the immune signaling involved in cold-preconditioning neuroprotection requires that biological systems and experimental manipulations plus technical capacities are highly reproducible and sensitive. Our approach is to use hippocampal slice cultures as an in vitro model that closely reflects their in vivo counterparts with multi-synaptic neural networks influenced by mature and quiescent macroglia / microglia. This glial state is particularly important for microglia since they are the principal source of cytokines, which are operative in the femtomolar range. Also, slice cultures can be maintained in vitro for several weeks, which is sufficient time to evoke activating stimuli and assess adaptive responses. Finally, environmental conditions can be accurately controlled using slice cultures so that cytokine signaling of cold-preconditioning can be measured, mimicked, and modulated to dissect the critical node aspects. Cytokine signaling system analyses require the use of sensitive and reproducible multiplexed techniques. We use quantitative PCR for TNF-&alpha; to screen for microglial activation followed by quantitative real-time qPCR array screening to assess tissue-wide cytokine changes. The latter is a most sensitive and reproducible means to measure multiple cytokine system signaling changes simultaneously. Significant changes are confirmed with targeted qPCR and then protein detection. We probe for tissue-based cytokine protein changes using multiplexed microsphere flow cytometric assays using Luminex technology. Cell-specific cytokine production is determined with double-label immunohistochemistry. Taken together, this brain tissue preparation and style of use, coupled to the suggested investigative strategies, may be an optimal approach for identifying potential targets for the development of novel therapeutics that could mimic the advantages of cold-preconditioning.

We seek to define the neural immune signaling responsible for cold-preconditioning as means to identify novel targets for therapeutics development to protect brain before injury onset. We present strategies for such work that require biological systems, experimental manipulations plus technical capacities that are highly reproducible and sensitive.

Neurological injury is a frequent cause of morbidity and mortality from general anesthesia and related surgical procedures that could be alleviated by ...

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

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.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Assessment of Vascular Regeneration in the CNS Using the Mouse Retina

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). It is increasingly being recognized that several neurodegenerative diseases such as Alzheimer&#8217;s, multiple sclerosis, and amyotrophic lateral sclerosis present elements of vascular compromise. In addition, the most prominent causes of blindness in pediatric and working age populations (retinopathy of prematurity and diabetic retinopathy, respectively) are characterized by vascular degeneration and failure of physiological vascular regrowth. The aim of this technical paper is to provide a detailed protocol to study CNS vascular regeneration in the retina. The method can be employed to elucidate molecular mechanisms that lead to failure of vascular growth after ischemic injury. In addition, potential therapeutic modalities to accelerate and restore healthy vascular plexuses can be explored. Findings obtained using the described approach may provide therapeutic avenues for ischemic retinopathies such as that of diabetes or prematurity and possibly benefit other vascular disorders of the CNS.

The rodent retina has long been recognized as an accessible window to the brain. In this technical paper we provide a protocol that employs the mouse model of oxygen-induced retinopathy to study the mechanisms that lead to failure of vascular regeneration within the central nervous system after ischemic injury. The described system can also be harnessed to explore strategies to promote regrowth of functional blood vessels within the retina and CNS.

The rodent retina is perhaps the most accessible mammalian system in which to investigate neurovascular interplay within the central nervous system (CNS). ...

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

Imaging Serotonergic Fibers in the Mouse Spinal Cord Using the CLARITY/CUBIC Technique

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal cord of the majority of these fiber systems have not been thoroughly investigated in any species. Serotonergic fibers, which project to the spinal cord, have been studied in rats and opossums on histological sections and their functional significance has been deduced based on their fiber termination pattern in the spinal cord. With the development of CLARITY and CUBIC techniques, it is possible to investigate this fiber system and its distribution in the spinal cord, which is likely to reveal previously unknown features of serotonergic supraspinal pathways. Here, we provide a detailed protocol for imaging the serotonergic fibers in the mouse spinal cord using the combined CLARITY and CUBIC techniques. The method involves perfusion of a mouse with a hydrogel solution and clarification of the tissue with a combination of clearing reagents. Spinal cord tissue was cleared in just under two weeks, and the subsequent immunofluorescent staining against serotonin was completed in less than ten days. With a multi-photon fluorescent microscope, the tissue was scanned and a 3D image was reconstructed using Osirix software.

Supraspinal projections are important for pain perception and other behaviors, and serotonergic fibers are one of these fiber systems. The present study focused on the application of the combined CLARITY/CUBIC protocol to the mouse spinal cord in order to investigate the termination of these serotonergic fibers.

Long descending fibers to the spinal cord are essential for locomotion, pain perception, and other behaviors. The fiber termination pattern in the spinal ...

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 ...

Effects of Blast-induced Neurotrauma on Pressurized Rodent Middle Cerebral Arteries

Though there have been studies on the histopathological and behavioral effects of blast exposure, fewer have been dedicated to blast&#39;s cerebral vascular effects. Impact (i.e., non-blast) traumatic brain injury (TBI) is known to decrease pressure autoregulation in the cerebral vasculature in both humans and experimental animals. The hypothesis that blast-induced traumatic brain injury (bTBI), like impact TBI, results in impaired cerebral vascular reactivity was tested by measuring myogenic dilatory responses to reduced intravascular pressure in rodent middle cerebral arterial (MCA) segments from rats subjected to mild bTBI using an Advanced Blast Simulator (ABS) shock tube. Adult, male Sprague-Dawley rats were anesthetized, intubated, ventilated and prepared for Sham bTBI (identical manipulation and anesthesia except for blast injury) or mild bTBI. Rats were randomly assigned to receive Sham bTBI or mild bTBI followed by sacrifice 30 or 60 min post-injury. Immediately after bTBI, righting reflex (RR) suppression times were assessed, euthanasia at the time points post-injury was completed, the brain was harvested and the individual MCA segments were collected, mounted and pressurized. As the intraluminal pressure perfused through the arterial segments was reduced in 20 mmHg increments from 100 to 20 mmHg, MCA diameters were measured and recorded. With decreasing intraluminal pressure, MCA diameters steadily increased significantly above baseline in the Sham bTBI groups while MCA dilator responses were significantly reduced (p &#60; 0.05) in both bTBI groups as evidenced by the impaired, smaller MCA diameters recorded for the bTBI groups. In addition, RR suppression in the bTBI groups was significantly (p &#60; 0.05) higher than in the Sham bTBI groups. MCA&#39;s collected from the Sham bTBI groups exhibited typical vasodilatory properties to decreases in intraluminal pressure while MCA&#39;s collected following bTBI exhibited significantly impaired myogenic vasodilatory responses to reduced pressure that persisted for at least 60 min after bTBI.

Here, we present a protocol to describe methods for ex vivo vascular reactivity determination following a primary blast traumatic brain injury (bTBI) using isolated, pressurized, rodent middle cerebral arterial (MCA) segments. bTBI induction is accomplished using a shock tube, also known as an Advanced Blast Simulator (ABS) device.

Though there have been studies on the histopathological and behavioral effects of blast exposure, fewer have been dedicated to blast&#39;s cerebral ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Combining Behavior and EEG to Study the Effects of Mindfulness Meditation on Episodic Memory

Although there has been recent interest in how mindfulness meditation can affect episodic memory as well as brain structure and function, no study has examined the behavioral and neural effects of mindfulness meditation on episodic memory. Here we present a protocol that combines mindfulness meditation training, an episodic memory task, and EEG to examine how mindfulness meditation changes behavioral performance and the neural correlates of episodic memory. Subjects in a mindfulness meditation experimental group were compared to a waitlist control group. Subjects in the mindfulness meditation experimental group spent four weeks training and practicing mindfulness meditation. Mindfulness was measured before and after training using the Five Facet Mindfulness Questionnaire (FFMQ). Episodic memory was measured before and after training using a source recognition task. During the retrieval phase of the source recognition task, EEG was recorded. The results showed that mindfulness, source recognition behavioral performance, and EEG theta power in right frontal and left parietal channels increased following mindfulness meditation training. In addition, increases in mindfulness correlated with increases in theta power in right frontal channels. Therefore, results obtained from combining mindfulness meditation training, an episodic memory task, and EEG reveal the behavioral and neural effects of mindfulness meditation on episodic memory.

Here we present a protocol for combining mindfulness meditation training, an episodic memory task, and EEG to understand the behavioral and neural effects of mindfulness meditation on episodic memory.

Although there has been recent interest in how mindfulness meditation can affect episodic memory as well as brain structure and function, no study has ...