The authors would like to acknowledge salary awards, equipment grants, and operating funds from the Fonds de la recherche en sant&eacute; Qu&eacute;bec (FRSQ), Canadian Institutes of Health Research (CIHR), the Natural Sciences and Engineering Research Council of Canada (NSERC), and the Concordia University Research Chairs Program (CRUC), as well as the thoughtful feedback on this manuscript from Dr. Jane Stewart.

1. Animal Housing
<ol>
	<li>Cage: In order to record the running-wheel activity of an individual rodent, each cage should house a single rodent and running-wheel. Because running wheels can be considered a form of enrichment, all rodents in any study should have similar access to a running wheel.</li>
	<li>Bedding changes: Animal handling as well as changes in cages or bedding can all have non-photic effects on circadian rhythms21-23, so, cages with mesh-flooring are ideal because they minimize contact with the animal. Notwithstanding the availability of such a tray-system, bedding changes should be avoided during critical phases of an experiment. Alternatives include using longer-lasting bedding, which would allow for more infrequent cage changes, or changing cages on a pseudo-random schedule.</li>
	<li>Isolation Boxes: Cages should be kept in isolation boxes that are sound-attenuated, light-controlled, and well-ventilated. Depending on the size and configuration of the isolation boxes, the number of cages within each box will typically range from 1-8. When housing multiple cages in a single isolation box, one should be aware that various odors and sounds coming from other animals can have confounding effects on the circadian behavior of individual animals. To avoid these problems, one should attempt to house one cage per isolation box.</li>
	<li>Ventilation: Adequate air flow is imperative to making the boxes a comfortable home environment for rodents. The fan in each box should be hooded, so as to prevent light from outside the box from reaching the inside. Also, fans will typically remove air from the box and blow it into the room. Small light-tight vents allow air to enter the isolation boxes from several points, and help to avoid uncomfortable breezes. In order to verify that there is adequate ventilation, the temperature inside the isolation box (when closed for several hours, with the lights on) should be virtually identical to the temperature in the room where it is housed.</li>
	<li>Lighting: Environmental light intensity should be the same in all cages. Arrange a single light at a similar location above each cage, and always use the same brand/type of bulb. Use moderate intensity illumination (100-300 lux) at cage level. Avoid excessively high illumination levels, which are more likely to produce direct changes on behavior attributable to the light rather than the circadian system, per se (e.g. masking).</li>
	<li>Darkness/Dim red lighting: If it is necessary to handle or treat animals in the dark (e.g. in constant darkness or nighttime), night vision goggles should be used. Alternatively, because the circadian system is relatively insensitive to red-wavelengths, dim red lighting can be used. The specific red light you use should be tested to ensure it does not alter running-wheel activity (e.g. masking) or adjust the circadian clock (e.g. produce a phase shift).</li>
</ol>
2. Data Collection (See Figure 1 - Vitalview Hardware Configuration)
<ol>
	<li>Running wheels: The diameter and ergonomics of the running wheel will change the amount of use24. Thus, use smaller and lighter wheels for mice, and larger heavier wheels for rats. When washing and re-installing wheels, ensure that the wheels are able to spin unobstructed, do not "wobble", and that the recording micro-switches are activated by each turn of the wheel.</li>
	<li>Micro-switches: Each revolution of the running wheel should activate a magnetic or mechanical micro-switch. Information from the micro-switch is transmitted via a single channel and recorded by a computer which can bin the data over time (e.g. every 2, 5, 6, or 10 min).</li>
	<li>Computer hardware: Our running-wheel recordings are made with Vitalview, a hardware and software platform developed by Mini Mitter (<a href="http://www.minimitter.com/vitalview_software.cfm" target="_blank">http://www.minimitter.com/vitalview_software.cfm</a>). However, there are other recording platforms such as ClockLab, developed by Actimetrics (<a href="http://www.actimetrics.com/ClockLab/" target="_blank">http://www.actimetrics.com/ClockLab/</a>). Both platforms bring together data from many single-channel sources (e.g. a single micro-switch activated by a single running wheel) into a single computer file. Data from individual channels can then be graphed and analyzed separately at a later date.</li>
</ol>
3. Data Recordings
<ol>
	<li>Files: The above mentioned software platforms can be used to separate out single channels so that individual files are created for each running-wheel record. Such data are best visualized and graphed with specially designed programs such as Actiview (Minimitter, Bend, OR), Circadia, or Clocklab (Actimetrics, Wilmette, IL) which can all produce periodograms and actograms. However, single-channel files can also be opened and analyzed using general spreadsheet programs such as Excel (Microsoft, Redmond, WA).</li>
	<li>Calculating Circadian Time (CT): CT 12 is, by definition, the onset of running-wheel activity in nocturnal rodents. In parallel with the 24 hr day, by convention, one circadian day is broken into 24 circadian hr. Accordingly, if the circadian free-running period is 24 hr 30 min as measured by the wall clock, CT 0 will occur approximately 12 hr 15 min after CT12.</li>
</ol>

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No conflicts of interest declared.

Monitoring daily activity rhythms using running wheels is the most commonly used and reliable method for assessing the output of the master circadian clock in nocturnal rodents. Wheel-running activity, however, is only one of many aspects of behavior and physiology that can be monitored continuously. Although the vast majority of running-wheel activity occurs during the night, over 30% of the total wakefulness occurs during the daytime25,26. Other endpoints can be used to assess circadian rhythms, including ge...

<table border="1">
	<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Vitalview Card &amp; Software</td>
 <td>Mini Mitter </td>
 <td>#855-0030-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>DP24 Dataport</td>
 <td>Mini Mitter </td>
 <td>#840-0024-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>QA4-Module </td>
 <td>Mini Mitter </td>
 <td>#130-0050-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Magnetic Switch</td>
 <td>Mini Mitter </td>
 <td>#130-0015-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>C-50 Cable assembly</td>
 <td>Mini Mitter </td>
 <td>#060-0045-10</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Rat running wheel assembly</td>
 <td>Mini Mitter </td>
 <td>#640-0700-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Cage and tray support</td>
 <td>Mini Mitter </td>
 <td>#640-0400-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Useable cut away cage</td>
 <td>Mini Mitter </td>
 <td>#664-2154-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Grid floor for cage</td>
 <td>Mini Mitter </td>
 <td>#676-2154-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Waste tray</td>
 <td>Mini Mitter </td>
 <td>#684-2154-00</td>
 <td>(Bend, OR, USA)</td>
 </tr>
 <tr>
 <td>Lamp housing</td>
 <td>Microlites Scientific </td>
 <td>#R-101</td>
 <td>(Toronto, ON, Canada)</td>
 </tr>
 <tr>
 <td>4W Fluorescent lamps</td>
 <td>Microlites Scientific </td>
 <td>#F4T5/CW</td>
 <td>(Toronto, ON, Canada)</td>
 </tr>
 <tr>
 <td>Isolation chambers</td>
 <td>Custom built</td>
 <td></td>
 <td>28"H x 20"W x 28"D &frac12;" Black Melamine.</td>
 </tr>
	 </tbody>
 </table>

recording and analysis of circadian rhythms in running-wheel activity in rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

<ol>
	<li>Computer programs: Specialized computer programs are typically used in the generation of actograms and the calculation of circadian period. These programs include, but are not limited to, Actiview (Minimitter, Bend, OR) and Circadia.</li>
	<li>Actograms: Actograms provide a graphic illustration of the daily patterns of running-wheel activity. There are single-plotted (x-axis = 24 hr) and double-plotted (x-axis = 48 hr) actograms. Both methods plot sequential days from top to bottom, but double-plott...

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Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

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21.1K Views.  McGill University. Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

Video: Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) pacemakers or clocks. Malfunctions in the human circadian system are associated with numerous diseases or disorders. Much progress towards our understanding of the mechanisms underlying circadian rhythms has emerged from genetic screens whereby an easily measured behavioral rhythm is used as a read-out of clock function. Studies using Drosophila have made seminal contributions to our understanding of the cellular and biochemical bases underlying circadian rhythms. The standard circadian behavioral read-out measured in Drosophila is locomotor activity. In general, the monitoring system involves specially designed devices that can measure the locomotor movement of Drosophila. These devices are housed in environmentally controlled incubators located in a darkroom and are based on using the interruption of a beam of infrared light to record the locomotor activity of individual flies contained inside small tubes. When measured over many days, Drosophila exhibit daily cycles of activity and inactivity, a behavioral rhythm that is governed by the animal's endogenous circadian system. The overall procedure has been simplified with the advent of commercially available locomotor activity monitoring devices and the development of software programs for data analysis. We use the system from Trikinetics Inc., which is the procedure described here and is currently the most popular system used worldwide. More recently, the same monitoring devices have been used to study sleep behavior in Drosophila. Because the daily wake-sleep cycles of many flies can be measured simultaneously and only 1 to 2 weeks worth of continuous locomotor activity data is usually sufficient, this system is ideal for large-scale screens to identify Drosophila manifesting altered circadian or sleep properties.

Assaying Locomotor Activity to Study Circadian Rhythms and Sleep Parameters in Drosophila

We describe procedures for recording daily locomotor activity rhythms of Drosophila and subsequent data analysis. Locomotor activity rhythms are a reliable behavioral output of animal circadian clocks and are used as the standard readout of clock function when studying circadian mutants or examining how the environment regulates the circadian system.

Most life forms exhibit daily rhythms in cellular, physiological and behavioral phenomena that are driven by endogenous circadian (&equiv;24 hr) ...

Measuring Circadian and Acute Light Responses in Mice using Wheel Running Activity

Circadian rhythms are physiological functions that cycle over a period of approximately 24 hours (circadian- circa: approximate and diem: day)1, 2. They are responsible for timing our sleep/wake cycles and hormone secretion. Since this timing is not precisely 24-hours, it is synchronized to the solar day by light input. This is accomplished via photic input from the retina to the suprachiasmatic nucleus (SCN) which serves as the master pacemaker synchronizing peripheral clocks in other regions of the brain and peripheral tissues to the environmental light dark cycle3-7. The alignment of rhythms to this environmental light dark cycle organizes particular physiological events to the correct temporal niche, which is crucial for survival8. For example, mice sleep during the day and are active at night. This ability to consolidate activity to either the light or dark portion of the day is referred to as circadian photoentrainment and requires light input to the circadian clock9. Activity of mice at night is robust particularly in the presence of a running wheel. Measuring this behavior is a minimally invasive method that can be used to evaluate the functionality of the circadian system as well as light input to this system. Methods that will covered here are used to examine the circadian clock, light input to this system, as well as the direct influence of light on wheel running behavior.

This article will review methods that can be used to determine circadian function and light responsiveness in mice.

Circadian rhythms are physiological functions that cycle over a period of approximately 24 hours (circadian- circa: approximate and diem: day)1, 2.  They ...

Large-scale Recording of Neurons by Movable Silicon Probes in Behaving Rodents

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of neurons and circuits requires methods with the spatial selectivity and temporal resolution appropriate for mechanistic analysis of neural ensembles in the behaving animal, i.e. recording of representatively large samples of isolated single neurons. Ensemble monitoring of neuronal activity has progressed remarkably in the past decade in both small and large-brained animals, including human subjects1-11. Multiple-site recording with silicon-based devices are particularly effective because of their scalability, small volume and geometric design.
Here, we describe methods for recording multiple single neurons and local field potential in behaving rodents, using commercially available micro-machined silicon probes with custom-made accessory components. There are two basic options for interfacing silicon probes to preamplifiers: printed circuit boards and flexible cables. Probe supplying companies (<a href="http://www.neuronexustech.com/" >http://www.neuronexustech.com/</a>; <a href="http://www.sbmicrosystems.com/">http://www.sbmicrosystems.com/</a>; <a href="http://www.acreo.se/">http://www.acreo.se/</a>) usually provide the bonding service and deliver probes bonded to printed circuit boards or flexible cables. Here, we describe the implantation of a 4-shank, 32-site probe attached to flexible polyimide cable, and mounted on a movable microdrive. Each step of the probe preparation, microdrive construction and surgery is illustrated so that the end user can easily replicate the process.

We describe methods for large-scale recording of multiple single units and local field potential in behaving rodents with silicon probes. Drive fabrication, probe attachment to the drive and probe implantation processes are illustrated in sufficient details for easy replication.

A major challenge in neuroscience is linking behavior to the collective activity of neural assemblies. Understanding of input-output relationships of ...

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Multi-unit Recording Methods to Characterize Neural Activity in the Locust (Schistocerca Americana) Olfactory Circuits

Detection and interpretation of olfactory cues are critical for the survival of many organisms. Remarkably, species across phyla have strikingly similar olfactory systems suggesting that the biological approach to chemical sensing has been optimized over evolutionary time1. In the insect olfactory system, odorants are transduced by olfactory receptor neurons (ORN) in the antenna, which convert chemical stimuli into trains of action potentials. Sensory input from the ORNs is then relayed to the antennal lobe (AL; a structure analogous to the vertebrate olfactory bulb). In the AL, neural representations for odors take the form of spatiotemporal firing patterns distributed across ensembles of principal neurons (PNs; also referred to as projection neurons)2,3. The AL output is subsequently processed by Kenyon cells (KCs) in the downstream mushroom body (MB), a structure associated with olfactory memory and learning4,5. Here, we present electrophysiological recording techniques to monitor odor-evoked neural responses in these olfactory circuits.
First, we present a single sensillum recording method to study odor-evoked responses at the level of populations of ORNs6,7. We discuss the use of saline filled sharpened glass pipettes as electrodes to extracellularly monitor ORN responses. Next, we present a method to extracellularly monitor PN responses using a commercial 16-channel electrode3. A similar approach using a custom-made 8-channel twisted wire tetrode is demonstrated for Kenyon cell recordings8. We provide details of our experimental setup and present representative recording traces for each of these techniques.

Multi-unit Recording Methods to Characterize Neural Activity in the Locust Schistocerca Americana Olfactory Circuits

We demonstrate variations of the extracellular multi-unit recording technique to characterize odor-evoked responses in the first three stages of the invertebrate olfactory pathway. These techniques can easily be adapted to examine ensemble activity in other neural systems as well.

Detection and interpretation of olfactory cues are critical for the survival  of many organisms. Remarkably, species across phyla have strikingly similar ...

Deep Brain Stimulation with Simultaneous fMRI in Rodents

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for using blood oxygen level dependent (BOLD) functional magnetic resonance imaging (fMRI) to image rodents with simultaneous DBS. DBS fMRI presents a number of technical challenges, including accuracy of electrode implantation, MR artifacts created by the electrode, choice of anesthesia and paralytic to minimize any neuronal effects while simultaneously eliminating animal motion, and maintenance of physiological parameters, deviation from which can confound the BOLD signal.&#160;Our laboratory has developed a set of procedures that are capable of overcoming most of these possible issues. For electrical stimulation, a homemade tungsten bipolar microelectrode is used, inserted stereotactically at the stimulation site in the anesthetized subject. In preparation for imaging, rodents are fixed on a plastic headpiece and transferred to the magnet bore. For sedation and paralysis during scanning, a cocktail of dexmedetomidine and pancuronium is continuously infused, along with a minimal dose of isoflurane; this preparation minimizes the BOLD ceiling effect of volatile anesthetics. In this example experiment, stimulation of the subthalamic nucleus (STN) produces BOLD responses which are observed primarily in ipsilateral cortical regions, centered in motor cortex. Simultaneous DBS and fMRI allows the unambiguous modulation of neural circuits dependent on stimulation location and stimulation parameters, and permits observation of neuronal modulations free of regional bias. This technique may be used to explore the downstream effects of modulating neural circuitry at nearly any brain region, with implications for both experimental and clinical DBS.

This protocol describes a standard method for simultaneous functional magnetic resonance imaging and deep brain stimulation in the rodent. The combined use of these experimental tools allows for the exploration of global downstream activity in response to electrical stimulation at virtually any brain target.

In order to visualize the global and downstream neuronal responses to deep brain stimulation (DBS) at various targets, we have developed a protocol for ...

In Vitro Recording of Mesenteric Afferent Nerve Activity in Mouse Jejunal and Colonic Segments

Afferent nerves not only convey information concerning normal physiology, but also signal disturbed homeostasis and pathophysiological processes of the different organ systems from the periphery towards the central nervous system. As such, the increased activity or &#39;sensitization&#39; of mesenteric afferent nerves has been allocated an important role in the pathophysiology of visceral hypersensitivity and abdominal pain syndromes.
Mesenteric afferent nerve activity can be measured in vitro in an isolated intestinal segment that is mounted in a purpose-built organ bath and from which the splanchnic nerve is isolated, allowing researchers to directly assess nerve activity adjacent to the gastrointestinal segment. Activity can be recorded at baseline in standardized conditions, during distension of the segment or following the addition of pharmacological compounds delivered intraluminally or serosally. This technique allows the researcher to easily study the effect of drugs targeting the peripheral nervous system in control specimens; besides, it provides crucial information on how neuronal activity is altered during disease. It should be noted however that measuring afferent neuronal firing activity only constitutes one relay station in the complex neuronal signaling cascade, and researchers should bear in mind not to overlook neuronal activity at other levels (e.g., dorsal root ganglia, spinal cord or central nervous system) in order to fully elucidate the complex neuronal physiology in health and disease.
Commonly used applications include the study of neuronal activity in response to the administration of lipopolysaccharide, and the study of afferent nerve activity in animal models of irritable bowel syndrome. In a more translational approach, the isolated mouse intestinal segment can be exposed to colonic supernatants from IBS patients. Furthermore, a modification of this technique has been recently shown to be applicable in human colonic specimens.

In Vitro Recording of Mesenteric Afferent Nerve Activity in Mouse Jejunal and Colonic Segments

Mesenteric afferent nerves convey information from the gastrointestinal tract towards the brain regarding normal homeostasis as well as pathophysiology. Gastrointestinal afferent nerve activity can be assessed by mounting isolated intestinal segments with attached afferent nerves into an organ bath, isolating the nerve, and assessing basal as well as stimulated activity.

Afferent nerves not only convey information concerning normal physiology, but also signal disturbed homeostasis and pathophysiological processes of the ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.