This work was supported by NIH grants 5R01NS051305-14 and 5R01NS076980-08 to PJS.

1. Experimental preparation
<ol>
	<li>Collect flies as they eclose into vials, separating male and female flies. 
	NOTE: Sleep experiments are commonly conducted with female flies. It is important to collect virgin females. Mated females will lay eggs that hatch into larvae complicating the analysis of the data.</li>
	<li>House flies of a single sex in groups of &#60;20. 
	NOTE: Housing flies in a socially enriched environment (groups of &#62;50) modulates sleep drive6,13 potentially confounding measurements of rebound sleep. Further, following social enrichment, sleep will decline over a few days6. Thus, baseline sleep is not stable complicating analysis of rebound sleep. Keeping flies in groups of &#60;20 avoids this potential confound.</li>
	<li>Keep flies in vials for 3-5 days in a light and humidity controlled environment. 
	&#8203;NOTE: Age and maturity of flies strongly influence sleep. Sleep is high in one day old flies and stabilizes by 3-5 days of age4. Flies are typically maintained on a 12 h light: 12 h dark schedule at 50% humidity.</li>
</ol>
2. Preparation of tubes for sleep recording
NOTE: Sleep is monitored using locomotor activity monitors. A monitor can hold 32 flies housed individually in 5 mm diameter tubes. Typically, genotypes are analyzed in groups of 16 or 32 flies.
<ol>
	<li>Prepare an appropriate number of tubes with fly food at one end. 
	NOTE: Diet and metabolism are known to influence sleep33,34, hence it is particularly important to place flies on the same food on which they were reared.</li>
	<li>Seal the end of the tubes with wax. 
	NOTE: Sleep deprivation and rebound is a five day experiment, and food can dry out if not properly sealed. In properly sealed tubes, food can be maintained for 10 days or more. Thus, it is critical to ensure that the ends of the tubes are sealed well. Flies can also get stuck to wet food, however. Thus, it helps to make tubes 1-2 days before the start of the experiment.</li>
	<li>Individually place wake, behaving flies into 65 mm long glass tubes for sleep recording using an aspirator and plug the end of the tubes with a foam stopper. 
	&#8203;NOTE: Flies are never re-exposed to CO2 anesthesia when placing flies into tubes for sleep recording. The aspirator is made from rubber tubing with one end covered with cheesecloth and inserted into a 1 mL pipette tip.</li>
</ol>
3. Recording sleep
<ol>
	<li>Load flies in tubes into activity monitors to monitor sleep. 
	&#8203;NOTE: The SNAP rocks monitors back and forth from -60&#176; to +60&#176; every ~10 s. The monitors are held at -60&#176; for ~5.9s ; it takes ~2.9 s for the tray holding the monitors to move from -60&#176; to +60&#176; and ~1 s to move back from +60&#176; to -60&#176;. The cycle length can be altered as needed by adjusting the voltage supplied to the motor.
	<ol>
		<li>Take care to ensure that tubes are placed in activity monitors in the correct orientation. In the correct orientation, the end of the tube with food is at the top of the SNAP to ensure that flies do not get pushed into the food. In addition, the end with food is on the side of the monitor with the sleep recording jack. This allows activity monitors to be oriented correctly in the SNAP for efficient sleep deprivation while simultaneously monitoring activity.</li>
	</ol>
	</li>
	<li>Place activity monitors in the recording chamber to monitor sleep.</li>
	<li>Monitor sleep for at least two full days to estimate baseline sleep. 
	NOTE: The day flies are loaded into activity monitors is typically excluded as an adaptation day to allow flies to adapt to being housed in tubes. Baseline sleep is recorded for at least two full days (48hrs) beginning with the morning following the day flies are loaded.</li>
	<li>Save locomotor activity counts of flies in 1 min bins from the time of lights on a given day to lights on the previous day using activity recording software (e.g., from 8 AM to 8 AM).</li>
	<li>Estimate sleep from the locomotor activity data with custom macros using 5 min of inactivity as the threshold for a bout of sleep35. 
	&#8203;NOTE: A number of sleep metrics are computed from the locomotor activity counts. These include sleep in min/h over 24 h, total sleep time in 24 h, average and maximum daytime and nighttime sleep bout lengths36.</li>
</ol>
4. Sleep deprivation and recovery
<ol>
	<li>As flies can be sleep deprived for variable lengths of time (e.g., 12 h, 24 h and 36 h) and recovery sleep can also be evaluated at various intervals (e.g., 6 h, 12 h, 24 h and 48 h), determine the duration of recovery by experimental need. Sleep recovery can be visualized using a sleep gain/loss plot or by examining percent sleep recovered over a predetermined interval (e.g., 6 h).</li>
	<li>If sleep is stable over the two baseline days, on the third day, place activity monitors into the SNAP for overnight sleep deprivation. 
	NOTE: Flies will exhibit a robust sleep rebound over a range of sleep times8,32,37,38, but sleep has to be stable to reliably evaluate rebound sleep. Sleep is stable when the difference in sleep between baseline days is &#177; 100 min.</li>
	<li>Make sure activity monitors are secured in place with monitor holder pins, monitor cords plugged in, and monitors oriented correctly with the end with food at the back, and plastic barriers in front (Figure 1). 
	NOTE: The SNAP is designed so the cam rotates once every 10 s (Figure 1). The plastic insert resets the tubes by pushing the tubes back when the apparatus is in the &#34;up&#34; position. Resetting the tubes is important to ensure that all tubes have the full range of motion at the beginning of each cycle.</li>
	<li>Unplug activity monitors, and take monitors out of the SNAP immediately upon lights on following overnight sleep deprivation. 
	NOTE: It is critical that sleep deprivation is terminated, and flies are placed in recovery immediately upon lights on following 12 h of overnight sleep deprivation. Even a 20-30 min delay in placing flies into recovery can interfere with the extent of rebound sleep.</li>
	<li>Place flies in a recording chamber where they will be undisturbed for two days (48 h) to monitor recovery sleep. 
	NOTE: If the recording chamber is being used for other experiments, extra care must be taken to avoid stimulating recovering flies.</li>
	<li>Calculate the amount of sleep lost. For each individual fly, calculate the hourly difference between sleep obtained during sleep deprivation and the corresponding hour during baseline; sum the hourly differences to calculate total sleep lost.</li>
	<li>Calculate the amount of sleep recovered. For each individual fly, calculate the hourly difference between sleep obtained during recovery and the corresponding hour during baseline; sum the hourly differences to calculate total sleep gained. 
	NOTE: Whether a fly is actually sleep deprived is empirical. Thus, the experimenter should examine percent sleep lost. If the fly has not lost a sufficient amount of sleep it can be excluded from the analysis. Although this might be required for other sleep deprivation approaches, it is rarely if ever required for the SNAP. More commonly, sleep may not be stable in a given fly prior to the initiation of sleep deprivation. If sleep is not stable, homeostasis cannot be calculated. We accept a maximum difference of &#177; 100 min of sleep calculated prior to the initiation of sleep deprivation as candidates for inclusion. On occasion, an individual fly&#39;s sleep is distributed unevenly across the 24 h day (e.g., some individuals may obtain 60-70% of their sleep quota during the day and thus only lose a small proportion of their 24 h sleep quota when deprived for 12 h at night). These flies can be evaluated separately.</li>
	<li>Calculate the average percentage of sleep recovered (relative to baseline) over 12 h, 24 h and 48 h of the recovery period for each genotype.</li>
	<li>From sleep data, compute the average and maximum daytime sleep bout length on baseline, and recovery days for each genotype. 
	NOTE: Rebound sleep in flies is characterized by increased sleep amount, and increased sleep depth in the recovery days. Sleep consolidation is used as measure of sleep depth. Arousal thresholds could also be used as a measure of sleep depth.</li>
</ol>

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The authors have nothing to disclose.

Sleep in Drosophila was independently characterized in 2000, by two groups4,5. In these pioneering studies, flies were deprived of sleep by gentle handling (i.e., hand deprivation) and shown to exhibit a robust homeostatic response to overnight sleep deprivation. Importantly, with any sleep deprivation experiment it is crucial to control for potential confounding effects of the method used to keep the animal awake. Hand deprivation studies set the benchm...

Sleep is near universal in animals, yet its function remains unclear. Sleep homeostasis, the compensatory increase in sleep following sleep deprivation, is a defining property of sleep, that has been used to characterize sleep states in a number of animals1,2,3,4,5.
Sleep in the fly has many similarities with human sleep, including a robust homeostatic response to sleep loss4,5. Numerous studies of sleep in the fly have used sleep deprivation both to infer sleep function by examining the adverse consequences that accrue from extended waking, and to understand sleep regulation by determining the neurobiological mechanisms controlling the homeostatic regulation of sleep. Thus sleep deprived flies were shown to exhibit impairments in learning and memory6,7,8,9,10,11,12, structural plasticity13,14,15, visual attention16, recovery from neuronal injury17,18, mating and aggressive behaviors19,20, cell proliferation21, and responses to oxidative stress22,23 to name a few. Further, investigations into the neurobiological mechanisms controlling rebound sleep have yielded critical insights into the neuronal machinery that constitutes the sleep homeostat8,9,23,24,25,26,27,28,29. Finally, in addition to revealing fundamental insights into sleep function in healthy animals, sleep deprivation studies have also informed insights into sleep function in diseased states30,31.
While sleep deprivation is undeniably a powerful tool, with any sleep deprivation experiment, it is important to distinguish phenotypes that result from extended waking, from those induced by the stimulus used to keep the animal awake. Sleep deprivation by hand deprivation or gentle handling, is generally regarded as setting the standard for minimally disruptive sleep deprivation. Here we describe a protocol for sleep depriving flies using the Sleep Nullifying Apparatus (SNAP). The SNAP is a device that delivers a mechanical stimulus to flies every 10s, keeping flies awake by inducing negative geotaxis (Figure 1). The SNAP efficiently deprives flies of &#62;98% of night-time sleep, even in flies with high sleep drive8,32. The SNAP has been calibrated on bang sensitive flies, agitation of flies in the SNAP does not harm flies; sleep deprivation with the SNAP induces a rebound comparable with that obtained by hand deprivation7. The SNAP is thus a robust method to sleep deprive flies while controlling for the effects of the arousing stimulus.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Locomotor activity tubes</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fisher Tissue Prep Wax</td>
			<td>Thermo Fisher</td>
			<td>13404-122</td>
			<td>Wax used for sealing tubes</td>
		</tr>
		<tr>
			<td>Glass tubes</td>
			<td>Wale Apparatus</td>
			<td>244050</td>
			<td>We cut 5mm diameter Pyrex glass&#160;tubes into 65mm long tubes&#160;to record sleep. Pre-cut tubes&#160;can also be purchased.</td>
		</tr>
		<tr>
			<td>Nutri Fly Bloomington Formulation fly food</td>
			<td>Genesee Scientific</td>
			<td>66-113</td>
			<td>Labs might use their own fly food recipe. It is important that sleep be recorded on the same food that flies were reared in.</td>
		</tr>
		<tr>
			<td>Rotary glass cutting tool</td>
			<td>Dremel Multi Pro</td>
			<td>395</td>
			<td>Used to cut 65mm long glass tubes&#160;</td>
		</tr>
		<tr>
			<td>Monitoring Sleep</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DAM System and DAMFileScan software</td>
			<td>Trikinetics</td>
			<td></td>
			<td>Software used to acquire data from DAM monitors and save the acquired data in an appropriate format</td>
		</tr>
		<tr>
			<td>Data acquisition computer</td>
			<td>Lenovo</td>
			<td>Idea Centre AIO3</td>
			<td>A equivalent computer from any manufacturer can substitute</td>
		</tr>
		<tr>
			<td>Drosophila Activity Monitors</td>
			<td>Trikinetics</td>
			<td>DAM2</td>
			<td>These monitors are used to record flies&#39; locomotor activity</td>
		</tr>
		<tr>
			<td>Environment Monitor</td>
			<td>Trikinetics</td>
			<td>DEnM</td>
			<td>Not essential, but an easy way to monitor environmental conditions in the chamber where sleep is recorded</td>
		</tr>
		<tr>
			<td>Light Controller</td>
			<td>Trikinetics</td>
			<td>LC4</td>
			<td>A convenient way to control the timing of when the SNAP is turned on and off</td>
		</tr>
		<tr>
			<td>Power Supply Interface Unit for DAM</td>
			<td>Trikinetics</td>
			<td>PSIU-9</td>
			<td>Required for data acquisition computers to record Trikinetics locomotor acitvity data</td>
		</tr>
		<tr>
			<td>RJ11 connector</td>
			<td>7001-64PC</td>
			<td>Multicomp</td>
			<td>DAM monitors accept RJ11 jacks</td>
		</tr>
		<tr>
			<td>Splitters</td>
			<td>Trikinetics</td>
			<td>SPLT5</td>
			<td>Used to connect upto 5 DAM monitors</td>
		</tr>
		<tr>
			<td>Telephone cable wire</td>
			<td>Radioshack</td>
			<td>278-367</td>
			<td>Phone cables to acquire data from DAM monitors</td>
		</tr>
		<tr>
			<td>Sleep Deprivation</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Power supply</td>
			<td>Gw INSTEK</td>
			<td>GPS-30300</td>
			<td>Power supply for the SNAP</td>
		</tr>
		<tr>
			<td>Sleep Nullifying Apparatus</td>
			<td>Washington University School of Medicine machine shop</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

the sleep nullifying apparatus: a highly efficient method of sleep depriving drosophila

Sleep homeostasis, the increase in sleep observed following sleep loss, is one of the defining criteria used to identify sleep throughout the animal kingdom. As a consequence, sleep deprivation and sleep restriction are powerful tools that are commonly used to provide insight into sleep function. Nonetheless, sleep deprivation experiments are inherently problematic in that the deprivation stimulus itself may be the cause of observed changes in physiology and behavior. Accordingly, successful sleep deprivation techniques should keep animals awake and, ideally, result in a robust sleep rebound without also inducing a large number of unintended consequences. Here, we describe a sleep deprivation technique for Drosophila melanogaster. The Sleep Nullifying Apparatus (SNAP) administers a stimulus every 10s to induce negative geotaxis. Although the stimulus is predictable, the SNAP effectively prevents &#62;95% of nighttime sleep even in flies with high sleep drive. Importantly, the subsequent homeostatic response is very similar to that achieved using hand-deprivation. The timing and spacing of the stimuli can be modified to minimize sleep loss and thus examine non-specific effects of the stimulus on physiology and behavior. The SNAP can also be used for sleep restriction and to assess arousal thresholds. The SNAP is a powerful sleep disruption technique that can be used to better understand sleep function.

Canton S (Cs) was used as a wild-type strain. Flies were maintained on a 12 h light: 12 h dark schedule, and sleep deprived for 12 hours overnight. Inspection of the sleep profiles of Cs flies on the baseline day (bs), sleep deprivation day (sd), and two recovery days (rec1 and rec2) (Figure 2A) suggests that flies were effectively sleep deprived in the SNAP, and recovered sleep during the day consistent with observed reports in the literature<sup class="xref...

Watch this Scientific Journal Video about The Sleep Nullifying Apparatus: A Highly Efficient Method of Sleep Depriving Drosophila at JoVE.com

The Sleep Nullifying Apparatus: A Highly Efficient Method of Sleep Depriving Drosophila

Sleep deprivation is a powerful tool to investigate sleep function and regulation. We describe a protocol to sleep deprive Drosophila using the Sleep Nullifying Apparatus, and to determine the extent of rebound sleep induced by deprivation.

The Sleep Nullifying Apparatus: A Highly Efficient Method of Sleep Depriving Drosophila

Sleep homeostasis, the increase in sleep observed following sleep loss, is one of the defining criteria used to identify sleep throughout the animal ...

the-sleep-nullifying-apparatus-highly-efficient-method-sleep

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Neuroscience

3.3K Views.  Washington University School of Medicine. Sleep deprivation is a powerful tool to investigate sleep function and regulation. We describe a protocol to sleep deprive Drosophila using the Sleep Nullifying Apparatus, and to determine the extent of rebound sleep induced by deprivation.

Video: The Sleep Nullifying Apparatus: A Highly Efficient Method of Sleep Depriving Drosophila

Novel Apparatus and Method for Drug Reinforcement

Animal models of reinforcement have proven to be useful for understanding the neurobiological mechanisms underlying drug addiction. Operant drug self-administration and conditioned place preference (CPP) procedures are expansively used in animal research to model various components of drug reinforcement, consumption, and addiction in humans. For this study, we used a novel approach to studying drug reinforcement in rats by combining traditional CPP and self-administration methodologies. We assembled an apparatus using two Med Associate operant chambers, sensory stimuli, and a Plexiglas-constructed neutral zone. These modifications allowed our experiments to encompass motivational aspects of drug intake through self-administration and drug-free assessment of drug/cue conditioning strength with the CPP test. In our experiments, rats self-administered cocaine (0.75 mg/kg/inj, i.v.) during either four (e.g., the "short-term") or eight (e.g., the "long-term") alternating-day sessions in an operant environment containing distinctive sensory cues (e.g., olfactory and visual). On the alternate days, in the other (differently-cued) operant environment, saline was available for self-infusion (0.1 ml, i.v.). Twenty-four hours after the last self-administration/cue-pairing session, a CPP test was conducted. Consistent with typical CPP findings, there was a significant preference for the chamber associated with cocaine self-administration. In addition, in animals undergoing the long-term experiment, a significant positive correlation between CPP magnitude and the number of cocaine-reinforced lever responses. In conclusion, this apparatus and approach is time and cost effective, can be used to examine a wide array of topics pertaining to drug abuse, and provides more flexibility in experimental design than CPP or self-administration methods alone.

Operant drug self-administration and conditioned place preference (CPP) procedures are expansively used in research to model various components of drug reinforcement, consumption, and addiction in humans. In this report, we combined traditional CPP and self-administration methods as a novel approach to studying drug reinforcement and addiction in rats.

Animal models of reinforcement have proven to be useful for understanding the neurobiological mechanisms underlying drug addiction.  Operant drug ...

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) ...

Cryopreservation of Cortical Tissue Blocks for the Generation of Highly Enriched Neuronal Cultures

In this study, we outline a standardized protocol for the successful cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. For this protocol the freezing medium used is 10% dimethyl sulfoxide (DMSO) diluted in Hank's Buffered Salt Solution (HBSS). Blocks of cortical tissue are transferred to cryovials containing the freezing medium and slowly frozen at -1&deg;C/min in a rate-controlled freezing container. Post-thaw processing and dissociation of frozen tissue blocks consistently produced neuronal-enriched cultures which exhibited rapid neuritic growth during the first 5 days in culture and significant expansion of the neuronal network within 10 days. Immunocytochemical staining with the astrocytic marker glial fibrillary acidic protein (GFAP) and the neuronal marker beta-tubulin class III, revealed high numbers of neurons and astrocytes in the cultures. Generation of neural precursor cell cultures after tissue block dissociation resulted in rapidly expanding neurospheres, which produced large numbers of neurons and astrocytes under differentiating conditions. This simple cryopreservation protocol allows for the rapid, efficient, and inexpensive preservation of cortical brain tissue blocks, which grants increased flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

Here, we describe a method for efficient cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. This simple protocol provides flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

In this study, we outline a standardized protocol for the successful cryopreservation and thawing of cortical brain tissue blocks to generate highly ...

Non-Laser Capture Microscopy Approach for the Microdissection of Discrete Mouse Brain Regions for Total RNA Isolation and Downstream Next-Generation Sequencing and Gene Expression Profiling

As technological platforms, approaches such as next-generation sequencing, microarray, and qRT-PCR have great promise for expanding our understanding of the breadth of molecular regulation. Newer approaches such as high-resolution RNA sequencing (RNA-Seq)1 provides new and expansive information about tissue- or state-specific expression such as relative transcript levels, alternative splicing, and micro RNAs2-4. Prospects for employing the RNA-Seq method in comparative whole transcriptome profiling5 within discrete tissues or between phenotypically distinct groups of individuals affords new avenues for elucidating molecular mechanisms involved in both normal and abnormal physiological states. Recently, whole transcriptome profiling has been performed on human brain tissue, identifying gene expression differences associated with disease progression6. However, the use of next-generation sequencing has yet to be more widely integrated into mammalian studies.
Gene expression studies in mouse models have reported distinct profiles within various brain nuclei using laser capture microscopy (LCM) for sample excision7,8. While LCM affords sample collection with single-cell and discrete brain region precision, the relatively low total RNA yields from the LCM approach can be prohibitive to RNA-Seq and other profiling approaches in mouse brain tissues and may require sub-optimal sample amplification steps. Here, a protocol is presented for microdissection and total RNA extraction from discrete mouse brain regions. Set-diameter tissue corers are used to isolate 13 tissues from 750-&mu;m serial coronal sections of an individual mouse brain. Tissue micropunch samples are immediately frozen and archived. Total RNA is obtained from the samples using magnetic bead-enabled total RNA isolation technology. Resulting RNA samples have adequate yield and quality for use in downstream expression profiling. This microdissection strategy provides a viable option to existing sample collection strategies for obtaining total RNA from discrete brain regions, opening possibilities for new gene expression discoveries.

Non-Laser Capture Microscopy Approach for the Microdissection of Discrete Mouse Brain Regions for Total RNA Isolation and Downstream Next-Generation Sequencing and Gene Expression Profiling  |

RNA expression profiling of discrete mouse brain regions requires a precise and repeatable tissue collection strategy. A protocol that uses both coronal brain sectioning and tissue corer-assisted microdissection is described here. The yield and quality of total RNA obtained from the resulting samples confirms the utility of the outlined method.

As technological platforms, approaches such as next-generation sequencing, microarray, and qRT-PCR have great promise for expanding our understanding of ...

Simultaneous Electroencephalography, Real-time Measurement of Lactate Concentration and Optogenetic Manipulation of Neuronal Activity in the Rodent Cerebral Cortex

Although the brain represents less than 5% of the body by mass, it utilizes approximately one quarter of the glucose used by the body at rest1. The function of non rapid eye movement sleep (NREMS), the largest portion of sleep by time, is uncertain. However, one salient feature of NREMS is a significant reduction in the rate of cerebral glucose utilization relative to wakefulness2-4. This and other findings have led to the widely held belief that sleep serves a function related to cerebral metabolism. Yet, the mechanisms underlying the reduction in cerebral glucose metabolism during NREMS remain to be elucidated.
	One phenomenon associated with NREMS that might impact cerebral metabolic rate is the occurrence of slow waves, oscillations at frequencies less than 4 Hz, in the electroencephalogram5,6. These slow waves detected at the level of the skull or cerebral cortical surface reflect the oscillations of underlying neurons between a depolarized/up state and a hyperpolarized/down state7. During the down state, cells do not undergo action potentials for intervals of up to several hundred milliseconds. Restoration of ionic concentration gradients subsequent to action potentials represents a significant metabolic load on the cell8; absence of action potentials during down states associated with NREMS may contribute to reduced metabolism relative to wake.
	Two technical challenges had to be addressed in order for this hypothetical relationship to be tested. First, it was necessary to measure cerebral glycolytic metabolism with a temporal resolution reflective of the dynamics of the cerebral EEG (that is, over seconds rather than minutes). To do so, we measured the concentration of lactate, the product of aerobic glycolysis, and therefore a readout of the rate of glucose metabolism in the brains of mice. Lactate was measured using a lactate oxidase based real time sensor embedded in the frontal cortex. The sensing mechanism consists of a platinum-iridium electrode surrounded by a layer of lactate oxidase molecules. Metabolism of lactate by lactate oxidase produces hydrogen peroxide, which produces a current in the platinum-iridium electrode. So a ramping up of cerebral glycolysis provides an increase in the concentration of substrate for lactate oxidase, which then is reflected in increased current at the sensing electrode. It was additionally necessary to measure these variables while manipulating the excitability of the cerebral cortex, in order to isolate this variable from other facets of NREMS.
	We devised an experimental system for simultaneous measurement of neuronal activity via the elecetroencephalogram, measurement of glycolytic flux via a lactate biosensor, and manipulation of cerebral cortical neuronal activity via optogenetic activation of pyramidal neurons. We have utilized this system to document the relationship between sleep-related electroencephalographic waveforms and the moment-to-moment dynamics of lactate concentration in the cerebral cortex. The protocol may be useful for any individual interested in studying, in freely behaving rodents, the relationship between neuronal activity measured at the electroencephalographic level and cellular energetics within the brain.

A procedure is described for manipulating the activity of cerebral cortical pyramidal neurons optogenetically while the electroencephalogram, electromyogram, and cerebral lactate concentration are monitored. Experimental recordings are performed on cable-tethered mice while they undergo spontaneous sleep/wake cycles. Optogenetic equipment is assembled in our laboratory; recording equipment is commercially available.

Although the brain represents less than 5% of the body  by mass, it utilizes approximately one quarter of the glucose used by the body  at rest1. The ...

A Method of Nodose Ganglia Injection in Sprague-Dawley Rat

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located in the organs of the abdomen and thorax. The vagus nerve communicates information from stimuli such as heart rate, blood pressure, bronchopulmonary irritation, and gastrointestinal distension to the nucleus of solitary tract of the medulla. The cell bodies of the vagus nerve are located in the nodose and petrosal ganglia, of which the majority are located in the former. The nodose ganglia contain a wealth of receptors for amino acids, monoamines, neuropeptides, and other neurochemicals that can modify afferent vagus nerve activity. Modifying vagal afferents through systemic peripheral drug treatments targeted at the receptors on nodose ganglia has the potential of treating diseases such as sleep apnea, gastroesophageal reflux disease, or chronic cough. The protocol here describes a method of injection neurochemicals directly into the nodose ganglion. Injecting neurochemicals directly into the nodose ganglia allows study of effects solely on cell bodies that modulate afferent nerve activity, and prevents the complication of involving the central nervous system as seen in systemic neurochemical treatment. Using readily available and inexpensive equipment, intranodose ganglia injections are easily done in anesthetized Sprague-Dawley rats.

Afferent vagal signaling transmits important information to central nervous system from receptors located in organs of the abdomen and thorax. The nodose ganglia of vagus nerves contain many types of receptors that modulate vagal activity. This protocol describes a method of local injections of neurochemicals into the nodose ganglia.

Afferent signaling via the vagus nerve transmits important general visceral information to the central nervous system from many diverse receptors located ...

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies using Drosophila have contributed important breakthroughs in understanding neurological processes. Thus, with the goal of understanding the cellular and molecular basis of TBI pathologies in humans, we developed the High Impact Trauma (HIT) device to inflict closed head TBI in flies. Flies subjected to the HIT device display phenotypes consistent with human TBI such as temporary incapacitation and progressive neurodegeneration. The HIT device uses a spring-based mechanism to propel flies against the wall of a vial, causing mechanical damage to the fly brain. The device is inexpensive and easy to construct, its operation is simple and rapid, and it produces reproducible results. Consequently, the HIT device can be combined with existing experimental tools and techniques for flies to address fundamental questions about TBI that can lead to the development of diagnostics and treatments for TBI. In particular, the HIT device can be used to perform large-scale genetic screens to understand the genetic basis of TBI pathologies.

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Here we describe a method to inflict closed head traumatic brain injury (TBI) in Drosophila. This method provides a gateway to investigate the cellular and molecular mechanisms that underlie TBI pathologies using the vast array of experimental tools and techniques available for flies.

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies ...

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.

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 ...

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.

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

How to Obtain Reliable Visual Event-related Potentials in Newborns

The present study discusses the characteristics of visual event-related potentials (VEPs) and outlines methodological steps for obtaining reliable measurements in newborns. Obtaining high-quality, reliable VEPs is crucial for the early detection of abnormal development of the central nervous system in at-risk newborns, and for implementing successful early interventions. Recommendations are based on a previous study which showed that when post-conceptional age, polysomnography-identified sleep stages, and light-emitting diodes (LEDs) googles as the luminous source are controlled, no more than 4 repetitions of VEP averages are required to obtain replicable recordings, variability decreases, and reliable VEPs can be obtained. By controlling for these sources of variability and using statistical analyses, we were able to clearly and reliably identify the amplitude and latency of three main components (NII, PII and NIII) present in 100% of newborns (n = 20) during active sleep. Recording VEPs during awake states, quiet sleep and transitional sleep is not recommended because VEP morphology may differ significantly from one average to the next, leading to the risk of misleading clinical prognoses. Moreover, it is easier to obtain VEPs during active sleep because this state can be clearly and reliably identified at this stage of development, sleep cycles are short enough to allow measurements to be taken in a reasonable time, and the method does not require new o expensive equipment.

Several important points for obtaining high-quality reliable visual evoked potentials (VEPs) in newborns while minimizing variability and the risk of misleading prognoses are presented.