We thank Dr. Larry D. Frye for editorial help with this manuscript. This work was supported by Japan Society for the Promotion of Science Grants-in-Aid for Scientific Research 24300129 (to M.L.), 25890005 (to Y.O.) and 26640025 (to Y.T.), the National Agriculture and Food Research Organization (to Y.U.), the World Premier International Research Center Initiative (WPI) from the Ministry of Education, Culture, Sports, Science, and Technology (to Y.O., Y.T., Y.U. and M.L.) and the Nestl&#233; Nutrition Council, Japan (to M.L.).

Ethics Statement: Procedures involving animal subjects have been approved by the Institutional Animal Experiment Committee at the University of Tsukuba.
1. Preparation of Electrodes and Cables for EEG/EMG Recordings
<ol>
	<li>Prepare EEG/EMG recording electrode according to the following procedure. 
	Note: The electrode is disposable and can be used only for 1 animal. Plan carefully the wiring configuration for all connectors. Place marks on the connectors for the correct orientation.
	<ol>
		<li>Solder each pin of a 4-pin header to a 2-cm stainless steel wire. In brief, hold one end of the wire to the pin, place a hot soldering iron onto the wire-pin joint, and melt some solder to ensure that just enough solder runs smoothly into the joint. Be careful not to apply too much heat to the pin; otherwise, the plastic around the pins will melt.</li>
		<li>Solder the free end of each of 2 wires attached to the pin header to the head of a 1.0-mm-diameter stainless steel screw. In brief, hold the free end of the wire to the thread below the screw head, place a hot soldering iron onto the wire-screw joint, and melt some solder to ensure that just enough solder runs smoothly into the joint. The 2 wires with screws serve as EEG recording electrodes, whereas the 2 wires without screws serve as EMG recording electrodes.</li>
		<li>Use scissors to strip off 1 mm of the insulation at the end of the EMG electrodes to increase the quality of the EMG signal.</li>
		<li>Completely cover all soldered pins with epoxy adhesive by using a thin wooden stick or tooth pick to reduce the electrical noise during EEG/EMG recordings.</li>
	</ol>
	</li>
	<li>Prepare a cable for connecting the electrode with the slip ring as described below. This cable can be reused.
	<ol>
		<li>Solder each pin of a 4-pin FFC/FPC connector with a wire of a 30-cm flat cable. In brief, hold the stripped end of the wire to the pin, place a hot soldering iron onto the wire-pin joint, and melt some solder to ensure that just enough solder runs smoothly into the joint. 
		Note: Choose the length of the flat cable which is appropriate for the height of the experimental animal cage used for EEG/EMG recordings.</li>
		<li>Solder crimp sockets to a tip of the wires on the other end of the flat cable. In brief, hold a crimp socket to the stripped free end of the wire, place a hot soldering iron onto the wire-socket joint, and melt some solder to ensure that just enough solder runs smoothly into the joint.</li>
		<li>Insert each crimp socket into a 4-position crimp housing.</li>
		<li>Completely cover the crimp sockets with epoxy adhesive by using a thin wooden stick or tooth pick.</li>
	</ol>
	</li>
</ol>
2. Implantation of Electrodes in the Mouse Head (Duration: Approx. 20 min)
<ol>
	<li>Sterilize all surgical tools in a hot bead sterilizer before the surgery. Anesthetize a male mouse (10 - 20 weeks old, 20 - 30 g) with an intraperitoneal injection of pentobarbital (50 mg/kg). After checking that the mouse is deeply anesthetized by pinching a toe, shave the hair on the head and neck with clippers.</li>
	<li>Move the mouse to the stereotaxic frame, and fix the head between the 2 ear bars. Apply petroleum jelly on the eyes to prevent dryness. Cleanse the shaved-skin with alcohol and cut it along the midline with a scalpel to expose the skull. Clip the skin to keep the surgical area open.</li>
	<li>By using a carbide cutter (drill size: 0.8-mm diameter), drill 2 holes into the skull, one over the frontal cortical area (1 mm anterior to bregma, 1.5 mm lateral to the midline) and the other over the parietal area (1 mm anterior to lambda, 1.5 mm lateral to midline) of the right hemisphere, according to stereotaxic coordinates of Paxinos and Franklin.23</li>
	<li>By using a jeweler&#39;s screw driver, place stainless steel EEG recording screws in the holes and make 2 - 2.5 turns for each screw for epidural positioning over the cortex. 
	Note: Do not insert screws too deep to prevent damage to the brain tissue. Check that the screws are tightly fixed on the skull. This is important to have stable EEG signals during a long period of multiple recordings (typically, more than 1 month). Wiggly screws produce EEG artifacts and may come off before the end of the experimental schedule.</li>
	<li>Fix the electrode assembly (cf. Section 1.1, pins turned upward) with instant glue to the skull and cover with dental cement. Make bilaterally small holes with forceps in the trapezius (neck) muscles and insert the stainless steel wires that serve as EMG electrodes into the holes. Suture the skin with a silk thread (0.1 mm diameter) to avoid exposure of the muscle.</li>
	<li>Remove the mouse from the stereotaxic frame. Administer ampicillin (100 mg/kg) and meloxicam (1 mg/kg) intraperitoneally to the mouse to avoid bacterial infection and to reduce post-surgical pain, respectively. Keep the mouse on a heat pad and monitor it until it has regained sufficient consciousness to maintain sternal recumbency. House the mouse individually during recovery to avoid removal of electrodes by other animals, and administer meloxicam (1 mg/kg) intraperitoneally on the first day after the surgery.</li>
</ol>
3. Recording and Acquiring EEG/EMG Data
<ol>
	<li>After a 1-week recovery period, house each mouse individually in an experimental cage placed in a soundproof recording chamber. Maintain an ambient temperature of 23 &#177; 1 &#176;C and automatically control cycles of 12-hr light/12-hr dark (lights on at 08:00, illumination intensity ~ 100 lux).</li>
	<li>Connect the EEG/EMG electrode assembly on the mouse head to a recording cable. Ensure that the recording cable is connected to a slip ring (which is designed so that movements of the mouse are not restricted by twisting of the cable) and an EEG/EMG signal amplifier. Filter EEG/EMG signals (EEG, 0.5-64 Hz; EMG, 16-64 Hz), digitize at a sampling rate of 128 Hz by an analog-to-digital converter (A/D) and finally record on a computer running EEG/EMG recording software (Table &#39;Materials&#39;, 4th entry from bottom).</li>
	<li>Habituate the mouse for 2 - 3 days in the recording chamber. If the EEG/EMG recording includes intraperitoneal drug administrations, gently handle the mouse on each habituation day at the time of the drug administration.</li>
	<li>Subsequently, start EEG/EMG recording software (Table &#39;Materials&#39;, 4th entry from bottom).
	<ol>
		<li>Select the &#39;Data file information&#39; tab and click the box next to the file name. Enter a file name and click &#39;Save&#39;.</li>
		<li>Select the &#39;Recording condition&#39; tab and select all EEG/EMG channels which will be recorded.</li>
		<li>Select the sampling frequency (128 Hz) in the &#39;Recording condition&#39; tab.</li>
		<li>Check if the selected channels are displayed properly in the &#39;Channel information&#39; tab.</li>
		<li>Select the &#39;Timer setting&#39; tab and click &#39;Monitor&#39; to display EEG and EMG.</li>
		<li>Check if EEG/EMG signals are displayed correctly.</li>
		<li>Select the &#39;Timer setting&#39; tab and set the clock time for the beginning and end of the recording in the &#39;Main Timer&#39; area.</li>
		<li>Click &#39;Monitor&#39; in the &#39;Timer setting&#39; tab to start the recording.</li>
	</ol>
	</li>
	<li>Record EEG/EMG signals under baseline (i.e., sleep/wake behavior of a freely behaving mouse) and different treatment conditions (e.g., caffeine administration or sham treatment) over several days. Euthanize the mouse with an intraperitoneal injection of pentobarbital (200 mg/kg) after the last experimental day.</li>
</ol>
4. Scoring of Behavioral State Based on EEG/EMG Data
<ol>
	<li>Start the software for EEG/EMG analysis (Table &#39;Materials&#39;, 4th entry from bottom). Open the EEG/EMG raw data (.kcd file) produced under the step 3. Click the &#39;Sleep&#39; tab and select Epoch time. Select 10 sec.</li>
	<li>Click the &#39;Sleep&#39; tab and select &#39;Multi-screening&#39; to automatically score all 10-sec epochs into 3 stages (i.e., NREM and REM sleep, and wakefulness) on the basis of the amplitudes of EEG and EMG and power spectral analysis of the EEG.3</li>
	<li>Click &#39;FFT condition for EEG&#39;.</li>
	<li>Set the parameters for the power spectral analysis [256 datum points (corresponding to 2 sec of the EEG), Hanning window function, and the average of 5 spectra per epoch].3 Click &#39;OK&#39;.</li>
	<li>Click &#39;Start Screening&#39; to begin the automatic screening. Open the scored data (.raf file).</li>
	<li>Check the results of the automatic screening and, if necessary, correct them manually based on the standard criteria (see the Representative Results, Figure 1B and Table 1).2, 3 Briefly, click and hold left mouse button on an incorrectly scored epoch and drag the cursor across the string of incorrectly scored epochs. Release the left mouse button and select the correct behavioral state in the pop-up window. 
	Note: Occasionally, epochs at the transition between two vigilant states are difficult to score unambiguously to one state. In such cases, the epoch should be scored to the ostensible state and the same criteria should be applied to similar epochs throughout the experiment to ensure data reproducibility.</li>
</ol>

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The authors Yujiro Tauguchi and Sayaka Kohtoh are employees of Kissei Comtech Co., Ltd that develops SleepSign software for acquisition and automatic scoring of EEG/EMG data used in this article.

This protocol describes a set-up for EEG/EMG recordings that allows the assessment of sleep and wakefulness under low-noise, cost-effective, and high-throughput conditions. Due to the small size of the EEG/EMG electrode head assembly, this system can be combined with other implants for intra-brain experiments, including optogenetics (optical fiber implantation) or, in conjunction with simultaneous cannula implantation, microinfusion of drugs into the mouse brain.31 Moreover, the design of the electrode head assembly with respect to multi-pin headers offers flexibility in the number of recording channels, if the measurement of additional electrical signals (e.g., contralateral EEG, electrooculogram or local field potential) is required.
However, individual housing is required for the cable-based design described in this protocol, which therefore limits the assessment of behavioral states, i.e., sleep and wakefulness, in combination with social interaction or complex behavioral testing. In these cases, a wireless sleep monitoring system is likely more suitable, although telemetric devices are not without&#12288;their own limiting features, especially battery cost and life.
The quality of EEG/EMG signals is important for the scoring of behavioral states according to the criteria shown in Figure 1 and Table 1. Wiggly electrodes (i.e., screws) are often the reason for electric noise resulting in artifacts in the EEG and FFT analysis. Moreover, it is important for the quality of the EEG signal to completely cover the electrode screws with dental cement to avoid air bubbles between the screws and cement, resulting in increased electric noise. The quality of EEG signals can be checked in an obviously sleeping mouse by visually confirming that they have a high amplitude and low frequency.
Cost and time to implant electrodes are critical factors for many sleep research laboratories and are considered a major drawback for large-scale screening of sleep-wake behavior in mice. The EEG/EMG recording system described here can be established and operated at low-to-moderate cost, including recurring cost for electrode materials and medical supplies (approximately 2 USD per mouse) and investments for EEG/EMG recording equipment (slip ring, amplifier, and A/D converter; approximately 2,000 USD per mouse).
With respect to time, a skilled researcher can conclude the electrode implantation for 1 mouse in less than 20 min; and thus, it is possible to operate on more than 20 mice per day. Another key factor for the overall efficiency of the assessment of sleep on the basis of EEG/EMG measurement is the use of software for acquisition and automatic scoring of EEG/EMG data. For these purposes, a variety of commercial and in-house developed software is available with a high variability in pricing or scoring accuracy.

Technical advances have often precipitated quantum leaps in the understanding of neurobiological processes. For example, Hans Berger&#39;s discovery in 1929 that electrical potentials recorded from the human scalp took the form of sinusoidal waves, the frequency of which was directly related to the level of wakefulness of the subject, led to rapid advances in the understanding of sleep-wake regulation, in both animals and humans alike.1 To this day the electroencephlogram (EEG), in conjunction with the electromyogram (EMG), i.e., electrical activity produced by skeletal muscles, represents the data &#34;backbone&#34; of nearly every experimental and clinical assessment that seeks to correlate behavior and physiology with the activity of cortical neurons in behaving animals, including humans. In most basic sleep research laboratories these EEG recordings are performed by using a cable-based system (Figure 1) wherein acquired data is subjected off-line to pattern and spectrum analysis [e.g., applying a fast Fourier transform (FFT) algorithm] to determine the vigilance state of the subject being recorded.2, 3 Sleep consists of rapid-eye movement (REM) and non-REM (NREM) sleep. REM sleep is characterized by a rapid low-voltage EEG, random eye movement, and muscle atonia, a state in which the muscles are effectively paralyzed. REM sleep is also known as paradoxical sleep, because the brain activity resembles that of wakefulness, whereas the body is largely disconnected from the brain and appears to be in deep sleep. By contrast, motor neurons are stimulated during NREM sleep but there is no eye movement. Human NREM sleep can be divided into 4 stages, whereby stage 4 is called deep sleep or slow-wave sleep and is identified by large, slow brain waves with delta activity between 0.5 - 4 Hz in the EEG. On the other hand, a subdivision between phases of NREM sleep in smaller animals, like rats and mice, has not been established, mostly because they do not have long consolidated periods of sleep as seen in humans.
Over the years, and on the basis of EEG interpretation, several models of sleep-wake regulation, both circuit- and humoral-based, have been proposed. The neural and cellular basis of the need for sleep or, alternatively, &#34;sleep drive,&#34; remains unresolved, but has been conceptualized as a homeostatic pressure that builds during the waking period and is dissipated by sleep. One theory is that endogenous somnogenic factors accumulate during wakefulness and that their gradual accumulation is the underpinning of sleep homeostatic pressure. While the first formal hypothesis that sleep is regulated by humoral factors has been credited to Rosenbaum&#39;s work published in 18924, it was Ishimori5, 6 and Pieron7 who independently, and over 100 years ago, demonstrated the existence of sleep-promoting chemicals. Both researchers proposed, and indeed proved, that hypnogenic substances or &#39;hypnotoxins&#39; were present in the cerebral spinal fluid (CSF) of sleep-deprived dogs.8 Over the past century several additional putative hypnogenic substances implicated in the sleep homeostatic process have been identified (for review, see ref. 9), including prostaglandin (PG) D2,10 cytokines,11 adenosine,12 anandamide,13 and the urotensin II peptide.14
Experimental work by Economo15, 16, Moruzzi and Magoun17, and others in the early and mid-20th century produced findings that inspired circuit-based theories of sleep and wakefulness and, to a certain degree, overshadowed the then prevailing humoral theory of sleep. To date, several &#34;circuit models&#34; have been proposed, each informed by data of varying quality and quantity (for review, see ref. 18). One model, for example, proposes that slow-wave sleep is generated through adenosine-mediated inhibition of acetylcholine release from cholinergic neurons in the basal forebrain, an area mainly consisiting of the nucleus of the horizontal limb of the diagonal band of Broca and the substantia inominata.19 Another popular model of sleep/wake regulation describes a flip-flop switch mechanism on the basis of mutually inhibitory interactions between sleep-inducing neurons in the ventrolateral preoptic area and wake-inducing neurons in the hypothalamus and brain stem.18, 20, 21 Moreover, for the switching in and out of REM sleep, a similar reciprocally inhibitory interaction has been proposed for areas in the brain stem, that is the ventral periaqueductal gray, lateral pontine tegmentum, and sublaterodorsal nucleus.22 Collectively, these models have proven valuable heuristics and afforded important interpretative frameworks for studies in sleep research; however, a yet fuller understanding of the molecular mechanisms and circuits regulating the sleep-wake cycle will require a more complete knowledge of its components. The system for polygraphic recording detailed below should aid in this goal.

<table border="0" cellpadding="0" cellspacing="0" width="729">
	<tbody>
		<tr height="19">
			<td height="19" style="height:19px;width:216px;">4-pin header</td>
			<td style="width:143px;">Hirose</td>
			<td style="width:167px;">A3B-4PA-2DSA(71)</td>
			<td style="width:205px;"></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:216px;">Ampicillin</td>
			<td style="width:143px;">Meiji Seika</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Analog-to-digital converter</td>
			<td style="width:143px;">Contec</td>
			<td style="width:167px;">AD16-16U(PCIEV)</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Caffeine</td>
			<td style="width:143px;">Sigma</td>
			<td style="width:167px;">C0750</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Carbide cutter</td>
			<td style="width:143px;">Minitor</td>
			<td style="width:167px;">B1055</td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:216px;">Crimp housing</td>
			<td style="width:143px;">Hirose</td>
			<td style="width:167px;">DF11-4DS-2C</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Crimp socket</td>
			<td style="width:143px;">Hirose</td>
			<td style="width:167px;">DF11-30SC</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Dental cement (Toughron Rebase)</td>
			<td style="width:143px;">Miki Chemical Product</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Epoxy adhesive</td>
			<td style="width:143px;">Konishi</td>
			<td style="width:167px;">#16351</td>
			<td></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:216px;">FFC/FPC connector</td>
			<td style="width:143px;">Honda Tsushin Kogyo</td>
			<td style="width:167px;">FFC-10BMEP1(B)</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Flat cable</td>
			<td style="width:143px;">Hitachi Cable</td>
			<td style="width:167px;">20528-ST LF</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:216px;">Instant glue (Aron Alpha A)</td>
			<td style="width:143px;">Toagosei</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Meloxicam</td>
			<td style="width:143px;">Boehringer Ingelheim</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Pentobarbital</td>
			<td style="width:143px;">Kyoritsu Seiyaku</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Signal amplifier</td>
			<td style="width:143px;">Biotex</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sleep recording chamber</td>
			<td style="width:143px;">APL</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">SleepSign software</td>
			<td style="width:143px;">Kissei Comtec</td>
			<td style="width:167px;">N/A</td>
			<td>for EEG/EMG recording/analysis</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Slip ring</td>
			<td style="width:143px;">Biotex</td>
			<td style="width:167px;">N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stainless steel screw</td>
			<td style="width:143px;">Yamazaki</td>
			<td style="width:167px;">N/A</td>
			<td>&#966;1.0&#215;2.0</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Stainless steel wire</td>
			<td style="width:143px;">Cooner Wire</td>
			<td style="width:167px;">AS633</td>
			<td></td>
		</tr>
	</tbody>
</table>

polygraphic recording procedure for measuring sleep in mice

Recording of the epidural electroencephalogram (EEG) and electromyogram (EMG) in small animals, like mice and rats, has been pivotal to study the homeodynamics and circuitry of sleep-wake regulation. In many laboratories, a cable-based sleep recording system is used to monitor the EEG and EMG in freely behaving mice in combination with computer software for automatic scoring of the vigilance states on the basis of power spectrum analysis of EEG data. A description of this system is detailed herein. Steel screws are implanted over the frontal cortical area and the parietal area of 1 hemisphere for monitoring EEG signals. In addition, EMG activity is monitored by the bilateral placement of wires in both neck muscles. Non-rapid eye movement (Non-REM; NREM) sleep is characterized by large, slow brain waves with delta activity below 4 Hz in the EEG, whereas a shift from low-frequency delta activity to a rapid low-voltage EEG in the theta range between 6 and 10 Hz can be observed at the transition from NREM to REM sleep. By contrast, wakefulness is identified by low- to moderate-voltage brain waves in the EEG trace and significant EMG activity.

Figure 1B illustrates examples of the mouse EEG in the different vigilance states. As shown in Table 1, epochs are classified as NREM sleep if the EEG shows large, slow brain waves with a delta rhythm below 4 Hz and the EMG has only a weak or no signal. Epochs are classified as REM sleep if the EEG shows rapid low-voltage brain waves in the theta range between 6 and 10 Hz and the EMG shows low amplitude. Other epochs should be classified as wakefulness (i.e., low-to-moderate voltage EEG and occurrence of EMG activity).
For instance, the EEG/EMG recording set-up described in this protocol can be used to determine the sleep amount and sleep/wake profile of C57BL/6 mice under baseline conditions or after treatment with caffeine (Figures 2 and 3).
Under baseline conditions, mice, which are nocturnal animals, exhibited a clear circadian sleep-wake rhythm, as seen in these figures, with larger amounts of sleep during the light period than during the dark one (Figure 2A). During the 12-hr light period, the mice showed 6.7 hr and 0.9 hr of NREM and REM sleep, respectively; whereas during the 12-hr dark period, wakefulness was predominant (Figure 2B). On the other hand, the quality of sleep can be evaluated on the basis of the vigilance state and EEG power spectrum analysis (Figure 2C-F). Typically, polygraphic recordings of EEG and EMG can be used to determine episode duration distribution, mean duration, and stage transition number for each vigilance state (Figure 2C-E). Moreover, the EEG power spectrum for NREM and REM sleep in mice during the light and dark period (Figure 2F) shows strong EEG power density in the frequency range of 0.5 - 4 Hz and 6 - 10 Hz, respectively. It should be noted that the EEG during REM sleep includes small amounts of delta waves (0.5 - 4 Hz), since the intermingled states of NREM and REM sleep are sometimes a contaminant.
To assess drug effects on the sleep-wake behavior of mice,24-30 EEG and EMG are typically recorded for 2 consecutive days. To determine, for example, the arousal effect of caffeine on C57BL/6 mice,24 the mice were treated with vehicle (10 ml/kg saline; intraperitoneally) on day 1 at 10:00 A.M. in the early phase of the light period. The animals were then treated with caffeine (15 mg/kg) 24 hr later, and the vigilance states were classified offline into waking, REM sleep, and NREM sleep. Figure 3A shows typical examples of EEG, EMG, and hypnograms after the administration of caffeine (lower polysomnographic panels) or vehicle (upper polysomnographic panels) in a C57BL/6 mouse. Caffeine increased the amount of wakefulness in C57BL/6 mice 2.8-fold for 3 hr after the injection (Figure 3B).
<img alt="Figure 1" src="/files/ftp_upload/53678/53678fig1.jpg" /> 
Figure 1. Sleep Bioassay System for Mice.
(A)&#160;To monitor EEG signals, stainless steel screws are implanted epidurally over the frontal cortical and parietal areas of 1 hemisphere. In addition, EMG activity is monitored by stainless steel wires placed bilaterally within the trapezius muscles. (B) Typical examples of EEG, EMG, and EEG power density for 10 sec during NREM or REM sleep or wakefulness in a mouse. In NREM sleep, EEG shows high-amplitude waves; and the delta band (0.5 - 4 Hz) is dominant (left). In REM sleep, EEG shows low amplitude waves, with the theta band (6 - 10 Hz) being dominant (middle). In wakefulness, the EEG shows low-amplitude waves, with no frequency being dominant (right). EMG signals are lower in both NREM and REM sleep than in wakefulness. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53678/53678fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53678/53678fig2.jpg" /> 
Figure 2. Sleep-wake Profiles under Baseline Conditions in C57BL/6 Mice Assessed by EEG/EMG Recordings. 
(A) Time-course changes in the hourly amount of each behavioral stage. White and black bars above the x-axes indicate the light and dark periods, respectively. (B) Total amount of each stage for 12 hr shows a larger amount of NREM and REM sleep during the light period compared with that in the dark period. (C) Distribution of episode duration of each stage. (D) Mean duration of each stage is longer for wakefulness in the dark period. (E) Stage-transition number of each stage shows more frequent transitions during the light period. (F) EEG power spectrum during NREM and REM sleep shows essentially no power-density differences between light and dark periods. Data are presented as the mean &#177; SEM (n = 5). *p &#60;0.05, **p &#60;0.01, as assessed by paired two-tailed Student&#39;s t test. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53678/53678fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" height="890" src="/files/ftp_upload/53678/53678fig3.jpg" width="700" /> 
Figure 3. Arousal Effect of Caffeine Assessed by EEG/EMG Recordings. 
(A) Typical examples of EEG, EMG, and hypnograms after administration of vehicle (upper panel) or caffeine at a dose of 15 mg/kg (lower panel). (B) Time courses of wakefulness in mice treated with caffeine. (C) Wakefulness over a 3-hr period after injection of caffeine. Data are presented as the mean &#177; SEM (n = 5). **p &#60;0.01 compared to vehicle injection, as assessed by paired two-tailed Student&#39;s t test. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53678/53678fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0" width="519">
	<colgroup>
		<col />
		<col span="2" />
		<col />
	</colgroup>
	<tbody>
		<tr height="20">
			<td height="20" style="height:20px;width:156px;"></td>
			<td style="width:139px;">NREM sleep</td>
			<td style="width:139px;">REM sleep</td>
			<td style="width:87px;">Wakefulness</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EEG amplitude</td>
			<td>High</td>
			<td>Low</td>
			<td>Low</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dominant EEG frequency</td>
			<td>Delta band (0.5 - 4 Hz)</td>
			<td>Theta band (6 - 10 Hz)</td>
			<td>None</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">EMG amplitude</td>
			<td>Low</td>
			<td>Low</td>
			<td>High</td>
		</tr>
	</tbody>
</table>
Table 1:General Criteria to Score Behavioral States by EEG/EMG Signals.

Watch this Scientific Journal Video about Polygraphic Recording Procedure for Measuring Sleep in Mice at JoVE.com

Polygraphic Recording Procedure for Measuring Sleep in Mice

The recording of electroencephalogram (EEG) and electromyogram (EMG) in freely behaving mice is a critical step to correlate behavior and physiology with sleep and wakefulness. The experimental protocol described herein provides a cable-based system for acquiring EEG and EMG recordings in mice.

Recording of the epidural electroencephalogram (EEG) and electromyogram (EMG) in small animals, like mice and rats, has been pivotal to study the ...

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23.3K 조회수.  University of Tsukuba. The recording of electroencephalogram (EEG) and electromyogram (EMG) in freely behaving mice is a critical step to correlate behavior and physiology with sleep and wakefulness. The experimental protocol described herein provides a cable-based system for acquiring EEG and EMG recordings in mice.

비디오: Polygraphic Recording Procedure for Measuring Sleep in Mice

Measuring Motor Coordination in Mice

Mice are increasingly being used in behavioral neuroscience, largely replacing rats as the behaviorist's animal of choice. Before aspects of behavior such as emotionality or cognition can be assessed, however, it is vital to determine whether the motor capabilities of e.g. a mutant or lesioned mouse allow such an assessment. Performance on a maze task requiring strength and coordination, such as the Morris water maze, might well be impaired in a mouse by motor, rather than cognitive, impairments, so it is essential to selectively dissect the latter from the former. For example, sensorimotor impairments caused by NMDA antagonists have been shown to impair water maze performance2. 
Motor coordination has traditionally been assessed in mice and rats by the rotarod test, in which the animal is placed on a horizontal rod that rotates about its long axis; the animal must walk forwards to remain upright and not fall off. Both set speed and accelerating versions of the rotarod are available.
The other three tests described in this article (horizontal bar, static rods and parallel bars) all measure coordination on static apparatus. The horizontal bar also requires strength for adequate performance, particularly of the forelimbs as the mouse initially grips the bar just with the front paws. 
Adult rats do not perform well on tests such as the static rods and parallel bars (personal observations); they appear less well coordinated than mice. I have only tested male rats, however, and male mice seem generally less well coordinated than females. Mice appear to have a higher strength:weight ratio than rats; the Latin name, Mus musculus, seems entirely appropriate. The rotarod, the variations of the foot fault test12 or the Catwalk (Noldus)15 apparatus are generally used to assess motor coordination in rats.

Protocols are presented for two established motor coordination tasks, the accelerating rotarod and horizontal bar, also two tests developed in Oxford recently, the static rods and parallel bars. These tests can detect motor impairments potentially of interest in their own right, as well as being possible variables in tests of other areas of behavior.

마우스에 모터 조정을 측정

Mice are increasingly being used in behavioral neuroscience, largely replacing rats as the behaviorist's animal of choice. Before aspects of behavior such ...

연구

신경과학

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.

역행 형광 라벨이 확인 된 신경 세포의 표적 세포 외 단일 단위 기록을 허용<em&gt; 생체 내</em

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

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.

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.

쥐에 활동 륜 실행에 녹음 및 Circadian 리듬 분석

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, ...

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of primary afferent fibers mediated by GABAergic axo-axonal synapses (primary afferent depolarization). The strength of primary afferent depolarization can be measured by recording of volume-conducted potentials at the dorsal root (dorsal root potentials, DRP). Pathological changes of presynaptic inhibition are crucial in the abnormal central processing of certain pain conditions and in some disorders of motor hyperexcitability. Here, we describe a method of recording DRP in vivo in mice. The preparation of spinal cord dorsal roots in the anesthetized animal and the recording procedure using suction electrodes are explained. This method allows measuring GABAergic DRP and thereby estimating spinal presynaptic inhibition in the living mouse. In combination with transgenic mouse models, DRP recording may serve as a powerful tool to investigate disease-associated spinal pathophysiology. In vivo recording has several advantages compared to ex vivo isolated spinal cord preparations, e.g. the possibility of simultaneous recording or manipulation of supraspinal networks and induction of DRP by stimulation of peripheral nerves.

Measuring Spinal Presynaptic Inhibition in Mice By Dorsal Root Potential Recording In Vivo

GABAergic presynaptic inhibition is a powerful inhibitory mechanism in the spinal cord important for motor and sensory signal integration in spinal cord networks. Underlying primary afferent depolarization can be measured by recording of dorsal root potentials (DRP). Here we demonstrate a method of in vivo recording of DRP in mice.

배근 잠재적 기록하여 생쥐의 척수 시냅스 억제를 측정<em&gt; 생체</em

Presynaptic inhibition is one of the most powerful inhibitory mechanisms in the spinal cord. The underlying physiological mechanism is a depolarization of ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

마우스에 콜드 민감도 및 적응을 결정하기위한 간단하고 저렴한 방법

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Extracellular Recording of Neuronal Activity Combined with Microiontophoretic Application of Neuroactive Substances in Awake Mice

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and awake animals. Therefore, methods allowing the manipulation of synaptic systems in awake animals are required in order to determine the contribution of synaptic inputs to neuronal processing unaffected by anesthetics. Here, we present methodology for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. By combining these procedures, we performed microiontophoretic injections of gabazine to selectively block GABAA receptors in neurons of the inferior colliculus of head-restrained mice. Gabazine successfully modified neural response properties such as the frequency response area and stimulus-specific adaptation. Thus, we demonstrate that our methods are suitable for recording single-unit activity and for dissecting the role of specific neurotransmitter receptors in auditory processing.
The main limitation of the described procedure is the relatively short recording time (~3 hr), which is determined by the level of habituation of the animal to the recording sessions. On the other hand, multiple recording sessions can be performed in the same animal. The advantage of this technique over other experimental procedures used to manipulate the level of neurotransmission or neuromodulation (such as systemic injections or the use of optogenetic models), is that the drug effect is confined to the local synaptic inputs to the target neuron. In addition, the custom-manufacture of electrodes allows adjustment of specific parameters according to the neural structure and type of neuron of interest (such as the tip resistance for improving the signal-to-noise ratio of the recordings).

We present methods for the construction of electrodes to simultaneously record extracellular neural activity and release multiple neuroactive substances at the vicinity of the recording sites in awake mice. This technique allows the detailed analysis of putative local synaptic inputs to the neuron of interest.

깨어 마우스의 신경 활성 물질의 Microiontophoretic 응용 프로그램과 결합하여 신경 활동의 세포 외 기록

Differences in the activity of neurotransmitters and neuromodulators, and consequently different neural responses, can be found between anesthetized and ...

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.

준비 및 Xenopus laevis Tectal 신경의 전체 셀 패치 클램프 기록에 대 한 프로토콜

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

Recording Spatially Restricted Oscillations in the Hippocampus of Behaving Mice

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed electrical field of a large volume of brain tissue, extracting information about local activity is challenging. Studying neuronal microcircuits, however, requires a reliable distinction between truly local events and volume-conducted signals originating in distant brain areas. Current source density (CSD) analysis offers a solution for this problem by providing information about current sinks and sources in the vicinity of the electrodes. In brain areas with laminar cytoarchitecture such as the hippocampus, one-dimensional CSD can be obtained by estimating the second spatial derivative of the LFP. Here, we describe a method to record multilaminar LFPs using linear silicon probes implanted into the dorsal hippocampus. CSD traces are computed along individual shanks of the probe. This protocol thus describes a procedure to resolve spatially restricted neuronal network oscillations in the hippocampus of freely moving mice.

This protocol describes the recording of local field potentials with multi-shank linear silicon probes. Conversion of the signals using current source density analysis allows the reconstruction of local electrical activity in the mouse hippocampus. With this technique, spatially restricted brain oscillations can be studied in freely moving mice.

행동 하는 쥐의 해 마에서 진동 제한 공간 기록

The local field potential (LFP) emerges from ion movements across neural membranes. Since the voltage recorded by LFP electrodes reflects the summed ...

Optogenetic Manipulation of Neural Circuits During Monitoring Sleep/wakefulness States in Mice

In recent years, optogenetics has been widely used in many fields of neuroscientific research. In many cases, an opsin, such as channel rhodopsin 2 (ChR2), is expressed by a virus vector in a particular type of neuronal cells in various Cre-driver mice. Activation of these opsins is triggered by application of light pulses which are delivered by laser or LED through optic cables, and the effect of activation is observed with very high time resolution. Experimenters are able to acutely stimulate neurons while monitoring behavior or another physiological outcome in mice. Optogenetics can enable useful strategies to evaluate function of neuronal circuits in the regulation of sleep/wakefulness states in mice. Here we describe a technique for examining the effect of optogenetic manipulation of neurons with a specific chemical identity during electroencephalogram (EEG) and electromyogram (EMG) monitoring to evaluate the sleep stage of mice. As an example, we describe manipulation of GABAergic neurons in the bed nucleus of the stria terminalis (BNST). Acute optogenetic excitation of these neurons triggers a rapid transition to wakefulness when applied during NREM sleep. Optogenetic manipulation along with EEG/EMG recording can be applied to decipher the neuronal circuits that regulate sleep/wakefulness states.

Here, we describe methods of optogenetic manipulation of particular types of neurons during monitoring of sleep/wakefulness states in mice, presenting our recent work on the bed nucleus of the stria terminalis as an example.

마우스에서 수면/각성 상태를 모니터링하는 동안 신경 회로의 광유전학 적 조작

In recent years, optogenetics has been widely used in many fields of neuroscientific research. In many cases, an opsin, such as channel rhodopsin 2 ...

Evaluation of Hemisphere Lateralization with Bilateral Local Field Potential Recording in Secondary Motor Cortex of Mice

This article demonstrates complete, detailed procedures for both in vivo bilateral recording and analysis of local field potential (LFP) in the cortical areas of mice, which are useful for evaluating possible laterality deficits, as well as for assessing brain connectivity and coupling of neural network activities in rodents. The pathological mechanisms underlying Alzheimer&#39;s disease (AD), a common neurodegenerative disease, remain largely unknown. Altered brain laterality has been demonstrated in aging people, but whether or not abnormal lateralization is one of the early signs of AD has not been determined. To investigate this, we recorded bilateral LFPs in 3-5-month-old AD model mice, APP/PS1, together with littermate wild type (WT) controls. The LFPs of the left and right secondary motor cortex (M2), specifically in the gamma band, were more synchronized in APP/PS1 mice than in WT controls, suggesting a declined hemispheric asymmetry of bilateral M2 in this AD mouse model. Notably, the recording and data analysis processes are flexible and easy to carry out, and can also be applied to other brain pathways when conducting experiments that focus on neuronal circuits.

We present in vivo electrophysiological recording of the local field potential (LFP) in bilateral secondary motor cortex (M2) of mice, which can be applied to evaluate hemisphere lateralization. The study revealed altered levels of synchronization between the left and right M2 in APP/PS1 mice compared to WT controls.

마우스의 보조 모터 피질에서 양측 지역 필드 전위 기록을 가진 반구 측면화의 평가

This article demonstrates complete, detailed procedures for both in vivo bilateral recording and analysis of local field potential (LFP) in the cortical ...