This study was supported by the Merck Investigator Studies Program (#54843), a KAKENHI Grant-in-Aid for Scientific Research on Innovative Areas, &#34;WillDynamics&#34; (16H06401) (T.S.), and a KAKENHI Grant-in-Aid for Exploratory Research on Innovative Areas (T.S.) (18H02595).

All experiments here were approved by the Animal Experiment and Use Committee of the University of Tsukuba, complying with NIH guidelines.
1. Animal Surgery, Virus Injection, Electrode for EEG/EMG, and Optical Fiber Implantation
CAUTION: Appropriate protection and handling techniques should be selected based on the biosafety level of the virus to be used. AAV should be used in an isolated P1A graded room for injection, and the tube carrying AAV must be sterilized with an autoclave after all the volume is used up. The surgical site and all implanted material should be clean and sterile during use. 
NOTE: See Figure 1.
<ol>
	<li>Disinfect the surgical equipment with the autoclave.</li>
	<li>Anesthetize mice with isoflurane using an anesthetic vaporizer. Observe until the mouse has reached the desired depth of anesthesia, determined by loss of response to pinching the tail with forceps. Apply ophthalmic ointment to the eyes to prevent them drying.</li>
	<li>Disinfect the surgical field with iodine solution or 70% EtOH (3x) and dry sufficiently. Allow the virus to thaw on ice as the surgery is being performed. Cover the surgical area with absorbent lab bench paper.</li>
	<li>Fix the mouse&#39;s head in the stereotactic apparatus with ear bars and a nose pinch. After confirming the head is held stably, make a midsagittal incision in the scalp to ensure the positions of the bregma and lambda are located at the same level on a horizontal line.</li>
	<li>To avoid a positioning gap, appropriately adjust the levels of the nose pinch and ear bars up and down. The bregma and lambda refer to the intersection between the sutura saggitalis and sutura coronalis or sutura lambdoidal, respectively (Figure 2).</li>
	<li>Use Serafin clamps to hold the skin to maintain access to the cranium. After exposure of the skull, disinfect the surface of the skull with iodine or 5% H2O2, to enable the cranial sutures including the bregma and lambda to be visualized more clearly.</li>
	<li>Prepare AAV vector injection:
	<ol>
		<li>Wash the inside of a 10 mL syringe (see Table of Materials) sequentially with 70% EtOH, 100% EtOH, and sterilized water, 5 times each. Secure the syringe in the clamp of a microinjection pump arm and make sure all solution in the syringe is discharged.</li>
		<li>Carefully aspirate 2 &#181;L of mineral oil without air bubbles, then aspirate the designated volume of the virus solution. After aspiration, manipulate the plunger button and confirm that virus solution emerges at the tip of the needle. 
		NOTE: The injection volume of the virus solution was determined in pilot experiments using the same mouse strain and same virus product. The relation between the volume of virus solution and extent of the infection area should be estimated in advance.</li>
	</ol>
	</li>
	<li>Inject AAV vector:
	<ol>
		<li>Adjust the tip of the microinjection needle on the bregma and note the coordinates as the original point. Move the tip to the designated injection site (for the BNST: anteroposterior + 0.2 mm, mediolateral &#177; 1.0 mm, dorsoventral - 4.2 mm) and place the tip of the needle onto the position. Put a mark on the skull and drill holes of approximately 2 mm in diameter using a dental drill with a 0.7 mm carbide cutter. Be careful not to damage the dura or brain tissue.</li>
		<li>After removing blood from around the holes with a cotton swab, slowly move the needle into the position of the BNST. Slowly inject the designated amount of virus solution (0.07 &#181;l/min) with a mechanical microinjector. After completing the injection, leave the needle for 5 min to allow the solution to sufficiently infiltrate the BNST tissue. Carefully take out the needle.</li>
		<li>For bilateral injection, repeat steps 1.8.1-1.8.2 on the other side. Throughout the procedure, keep the skull moist with applications of sterile saline. 
		NOTE: We used custom EEG/EMG implants (W: 5 mm, D: 7 mm, H: 1 mm) with four EEG electrodes (4 mm), two EMG electrodes (2 mm; cut the 4 mm electrode to 2 mm with nippers) and 6 electrodes (4.5 mm) (Figure 2A).</li>
	</ol>
	</li>
	<li>Solder two stainless steel wires (see Table of Materials) from which 1 mm of the insulation is stripped off both ends to the EMG electrodes. Adjust the center of the electrodes to the bregma and mark the position of each EEG electrode (anteroposterior &#177; 1.5 mm, mediolateral &#177; 1.0 mm), and determine the position of the implant (Figure 2B).</li>
	<li>Implant optical fibers:
	<ol>
		<li>Attach an optic fiber ferrule to the manipulator and rotate the manipulator arm so that it has an angle of &#177;30&#176; against a horizontal line (this process is only needed to avoid interference between electrodes and optic fibers, such as in the case of BNST stimulation). Put the fiber tip on the bregma and record the coordinates.</li>
		<li>Move the tip to the targeted insertion line and mark the position on the skull. Also put additional marks near the insertion site for the anchor screws. Drill the skull on each site with a dental drill to insert the optic fiber and fix the screw. Fix the screw on the skull. Be careful not to break the dura or damage any tissue by screw.</li>
		<li>Insert the optic fiber gently until reaching above the BNST with a manipulator. The ferrule should rest on the remaining cranium (Figure 1B).</li>
		<li>Apply photocurable dental cement (see Table of Materials) to cover the fiber and the screw. The reaction time to solidify the glue should be specified by the manufacturer&#39;s manual (Our material needs exposure to light for at least 10 s with specific wave length photo-generator. It is unnecessary to dry the glue after this).</li>
		<li>In this step, make sure that no materials (screw or glue) occupy the mounting space for the electrodes. In addition, avoid making any interruption in the cement for the ferrule connecting to the optic fiber and cable. Repeat steps 1.10.1-1.10.4 on the opposite side for bilateral stimulation.</li>
	</ol>
	</li>
	<li>Drill holes for EEG/EMG electrodes. Insert the tips of the electrodes into the holes. Hold the implant and apply cyanoacrylate adhesive to the space between the skull and the electrodes. Insert again with attention not to interfere with any materials.</li>
	<li>Cover the circumference of the electrodes and optic fibers with cyanoacrylate adhesive followed by application of cyanoacrylate accelerant on the adhesive. This step avoids causing any interruption at the ferrule-to-optic cable and electrode-to-lead wire connecting zone (Figure 1C). 
	NOTE: Cyanoacrylate adhesive and its accelerant are harmful to the mouse eye. Pay attention not to cause spillage of these chemical substances. Also, be careful not to strongly touch the electrodes and the fibers in order to avoid unexpected deviation immediately after adhesive solidification.</li>
	<li>Expose the mouse neck muscles and insert the wires for the EMG electrode under the muscle. Adjust the length of EMG electrode so that it locates just under the nuchal muscles. Light connection between tip of electrode and muscle fascia is enough to catch the EMG signal.</li>
	<li>Apply cyanoacrylate adhesive to fill the implants and solidify the adhesive with acceleration liquid. Then, put the mouse on a heat pad for recovery until the postural reflex appears. Adjust heat pad temperature to animal resting body temperature (36.0 &#176;C in ZT 0-12 in case of C57BL6 mice; do not exceed 38.0 &#176;C). 
	NOTE: An antibiotic is not required for sterile surgery.</li>
	<li>Follow your local institutional guidelines for post-operative analgesia. Keep the mice in a home cage for a recovery period of at least 7 days.</li>
</ol>
2. EEG/EMG Monitoring with Photo-excitation of Targeted Neurons in Specific Sleep States
CAUTION: This protocol includes use of class 3B laser equipment or LED devices. Experimenters should be aware of safety information. Protective eye goggles are required.
<ol>
	<li>Before connecting the laser cable to the optic fiber, adjust laser intensity with a scaler. Tether the tip of the laser cable to an unused optic fiber with a ferrule and confirm that there is no space at the junction between the fiber and the cable.</li>
	<li>Turn on the main switch of the laser and wait 20 min for it to warm up.</li>
	<li>Emit the laser to the intensity checker and adjust laser intensity to 10 mW/mm2. Change the laser mode to transistor logic and confirm that light pulses are emitted from the fiber controlled by the pattern regulator which is set at 10 ms for duration, 40 ms for rest, 20 times for cycle, and 20 times repeat (that is, 20 Hz of 10 ms light pulses for 20 s).</li>
	<li>After the recovery period, move the mice to the experimental chamber for recording EEG/EMG. House mice at a constant 23 &#176;C with a 12 h light/dark cycle with food and water available ad libitum.</li>
	<li>Connect the implanted electrode and cable adaptor which is tethered to a slip ring to avoid entanglement. &#65279;It is recommended to cover the junction with light-impermeable material such as aluminum foil to prevent laser leakage. If a bilateral experiment is required, use a slip ring with a bifurcate attachment for the cables.</li>
	<li>In this protocol, we assess latency to wakefulness from NREM sleep or REM sleep, so the recording time should be limited in optimized zeitgeber time (ZT0 is defined as the time when the light is on). This protocol was conducted between ZT4 - ZT10. Let the mice stay freely in the experimental chamber for at least 1 h as acclimatization.</li>
	<li>During the experimental period, monitor EEG and EMG signals in the same monitor and evaluate the mouse&#39;s state as wakefulness, NREM sleep or REM sleep. Use the gain control for each wave to make it easier to distinguish each state.</li>
	<li>For measurement of NREM sleep to wakefulness latency, observe stable NREM sleep for 40 s or stable REM sleep for 30 s, then turn on the switch of the pattern generator for photostimulation (this protocol generates 20 Hz of 10 ms light pulses for 20 s). Confirm laser emission to the implanted optic fibers.</li>
	<li>Record EEG/EMG signals until the sleeping state changes to wakefulness. If two or more experimental trials are needed, limit optogenetic manipulation to once a day because photostimulation is an artificial intervention which might affect sleep/wakefulness architecture.</li>
	<li>After the experiment, deeply anesthetize and perfuse with sterile saline and paraformaldehyde (PFA) for sampling the whole brain for immunohistochemical analysis26.</li>
</ol>
3. Analysis of Latency Time from NREM Sleep to Wakefulness
<ol>
	<li>After recording EEG/EMG signals, transfer the signal data to a computer for scoring sleep/wakefulness states. This protocol describes the method for EEG analysis with recording software (see Table of Materials).</li>
	<li>Start the sleep sign application and click the File tab and select Open to choose the recorded data (.kcd file). Click the Sleep tab to select Epoch time for adjusting the time window for each epoch (we use 1 epoch/4 sec).</li>
	<li>Manually score sleep/wakefulness states based on EEG/EMG signals, according to the following criteria: wakefulness, high EMG and low EEG voltage with high frequency; NREM, low EMG tone and high EEG voltage with high &#948; (0.5-4 Hz) frequency; and REM, EMG indicates muscle atonia and low EEG voltage with high &#952; (6-9 Hz) frequency. A state that does not consecutively continue for 16 s (i.e. 4 epochs) is not defined as a state change because it is not a stable state.</li>
	<li>Click and hold the left mouse button on the first epoch and drag the cursor until the specific trend of more than 16 s (4 epochs) is ended. Then, release the left mouse button and choose the appropriate state (wakefulness or NREM sleep or REM sleep) in the pop-up window. Repeat this procedure to score all EEG/EMG signals recorded in the file.</li>
	<li>Find the exact time of stimulation in the epoch where the EEG shows NREM or REM sleep, and the epoch showing state transition following the stimulation point. Count the number of epochs between the periods just after stimulation and just before state transition.</li>
	<li>Then multiply counted number of epochs by 4 s (A). In the epoch of stimulation, take a screen shot and measure the width between the stimulation point and the end of the epoch. Then, divide the measured length by the entire epoch length and multiply by 4 s (B). Similarly, calculate the duration of NREM sleep in the epoch of state change, which means the time between the start of the epoch and the state change point (C).</li>
	<li>Sum A, B and C to obtain the latency from NREM sleep to wakefulness. The same procedure is used for analysis of state transition from REM sleep to wakefulness.</li>
</ol>

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This project was partly supported financially by Merck &#38; Co.

We here presented a method to evaluate the effect of optogenetic stimulation of neurons with particular chemical identities on state transitions of sleep/wakefulness and gave an example of manipulation of GABABNST neurons. Our data showed that optogenetic excitation of GABABNST neurons results in immediate transition from NREM sleep to wakefulness.
Various experimental designs are available because of the development of numerous types of optogenetic tools. It is possible ...

Sleep is essential for optimal cognitive function.Recent findings also suggest that disturbances in sleep are associated with a wide range of diseases1,2,3. Although the functions of sleep are as yet largely unresolved, substantial progress has been made recently in understanding the neural circuits and mechanisms that control sleep/wakefulness states4. In mammals, there are three states of vigilance: wakefulness, non-rapid eye movement (NREM) sleep, and rapid eye movement (REM) sleep. Wakefulness is characterized by fast EEG oscillations (5-12 Hz) of low amplitude with purposeful and sustained motor activity. NREM sleep is defined by slow oscillations (1-4 Hz) of high amplitude (delta waves), with lack of consciousness and purposeful motor activity. REM sleep is characterized by relatively fast oscillations (6-12 Hz) of low amplitude and almost complete bilateral muscle atonia5.
Borbely proposed a theory of sleep-wakefulness regulation known as the two process model6,7. A homeostatic process, also referred to as process S, represents sleep pressure that accumulates during wakefulness and dissipates during sleep. Another process, referred to as process C, is a circadian process, which explains why vigilance levels fluctuate in the 24 h cycle. In addition to these two processes, allostatic factors are also important for regulation of sleep/wakefulness8,9. Allostatic factors include nutritional states and emotion. Fear and anxiety are usually accompanied by an increase in arousal along with autonomic and neuroendocrine responses10,11,12. The limbic system is believed to play a role in regulation of fear and anxiety, and mechanisms underlying autonomic and neuroendocrine responses have been studied extensively, but the pathway by which the limbic system influences sleep/wakefulness states has not yet been revealed. A large number of recent studies using opto- and pharmacogenetics have suggested that neurons and neuronal circuits that regulate sleep/wakefulness states are distributed throughout the brain, including the cortices, basal forebrain, thalamus, hypothalamus, and brain stem. In particular, recent advances in optogenetics have allowed us to stimulate or inhibit specific neural circuits in vivo with high spatial and temporal resolutions. This technique will allow progress in our understanding of the neural substrates of sleep and wakefulness, and how sleep/wakefulness states are regulated by circadian processes, sleep pressure, and allostatic factors, including emotion. This paper aims to introduce how to use optogenetic manipulation combined with sleep/wake recording, which could have the potential to update our understanding of the connectomes and mechanisms in the brain that play a role in the regulation of NREM sleep, REM sleep, and wakefulness. Understanding of this mechanism by which the limbic system regulates sleep/wakefulness states is of paramount importance to health, because insomnia is usually associated with anxiety or fear of being unable to sleep (somniphobia).
The BNST is thought to play an essential role in anxiety and fear. GAD 67-expressing GABAergic neurons are a major population of the BNST12,13. We examined the effect of optogenetic manipulation of these neurons (GABABNST) on sleep/wakefulness states. One of the greatest advances in neuroscience in recent years has been methods that enable manipulation of neurons with particular chemical identities in vivo, with high spatial and temporal resolutions. Optogenetics is highly useful for demonstrating causal links between neural activity and specific behavioral responses14. We describe optogenetics as a method to examine the functional connectivity of defined neural circuits in the regulation of sleep/wakefulness states. By utilizing this technique, great progress has been achieved in understanding the neuronal circuits that regulate sleep/wakefulness states15,16,17,18,19. In many cases, opsins are specifically introduced in&#160;neurons with particular chemical identities in selective brain regions by a combination of Cre-driver mice and Cre-inducible AAV-mediated gene transfer.&#160;Further, focal expression of photo-sensitive opsins such as channelrhodopsin 2 (ChR2)20 or archaerhodopsin (ArchT)21 combined with a Cre-loxP or Flp-FRT system allows us to manipulate a selective neuronal population and specific neural pathway22.
We describe here experiments on GABAergic neurons in the BNST as an example. To express opsins in a designated neuronal population, appropriate Cre driver mice and Cre-dependent virus vectors are most frequently used. Transgenic or knock-in lines in which opsins are expressed in particular neuronal populations are also useful. In the following experiments, we used GAD67-Cre knock-in mice23 in which only GABAergic neurons express Cre recombinase with a C57BL/6J genetic background, and an AAV vector which contains ChR2 (hChR2 H134R) fused with EYFP or EYFP as a control with a &#34;FLEx (Flip-excision) switch&#34;24. The procedure specifically describes optogenetic excitation of GABAergic neurons in the BNST during monitoring of sleep/wakefulness states25.

<table><tbody><tr><td>1x1 Fiber-optic Rotary Joints</td><td>Doric</td><td>FRJ 1x1 FC-FC</td><td>for optogenetics</td></tr><tr><td>6-pin header</td><td>KEL corporation</td><td>DSP02-006-431G</td><td></td></tr><tr><td>6-pin socket</td><td>Hirose</td><td>21602X3GSE</td><td></td></tr><tr><td>A/D converter</td><td>Nippon koden</td><td>N/A</td><td>Analog to digital converter</td></tr><tr><td>AAV&lt;sub&gt;10&lt;/sub&gt;-EF1a-DIO-ChR2-EYFP</td><td></td><td></td><td>3.70&amp;times;10&lt;sup&gt;13&lt;/sup&gt;(genomic copies/ml)</td></tr><tr><td>AAV&lt;sub&gt;10&lt;/sub&gt;-EF1a-DIO-EYFP</td><td></td><td></td><td>5.82&amp;times;10&lt;sup&gt;13&lt;/sup&gt;(genomic copies/ml)</td></tr><tr><td>Ampicillin</td><td>Fuji film</td><td>014-23302</td><td></td></tr><tr><td>Amplifier</td><td>Nippon koden</td><td>N/A</td><td>for EEG/EMG recording</td></tr><tr><td>Anesthetic vaporizer</td><td>Muromachi</td><td>MK-AT-210D</td><td></td></tr><tr><td>Automatic injecter</td><td>KD scientific</td><td>780311</td><td></td></tr><tr><td>Carbide cutter</td><td>Minitor</td><td>B1055</td><td>&amp;phi;0.7 mm. Reffered as dental drill, used with high speed rotary micromotor&amp;nbsp;</td></tr><tr><td>Cyanoacrylate adhesion&amp;nbsp; (Aron alpha A) and acceleration</td><td>Konishi</td><td>#30533</td><td></td></tr><tr><td>Dental curing light</td><td>3M</td><td>Elipar S10</td><td></td></tr><tr><td>Epoxy adhesive</td><td>Konishi</td><td>#04888</td><td>insulation around the solder of 6-pin and shielded cable</td></tr><tr><td>Fiber optic patch cord (branching)</td><td>Doric</td><td>BFP(#)_50/125/900-0.22</td><td></td></tr><tr><td>&lt;em&gt;Gad67&lt;/em&gt;-Cre mice</td><td>provided by Dr. Kenji Sakimura</td><td></td><td>Cre recombinase gene is knocked-in in the Gad67 allele</td></tr><tr><td>Hamilton syringe</td><td>Hamilton</td><td>65461-01</td><td></td></tr><tr><td>High speed rotary micromotor kit</td><td>FOREDOM</td><td>K.1070</td><td>Used with carbide cutter</td></tr><tr><td>Interconnecting sleeve</td><td>Thorlab</td><td>ADAF1</td><td>&amp;phi;2.5 mm Ceramic&amp;nbsp;</td></tr><tr><td>Isoflurane</td><td>Pfizer</td><td>871119</td><td></td></tr><tr><td>Laser&amp;nbsp;&amp;nbsp;</td><td>Rapp OptoElectronic</td><td>N/A</td><td>473 nm wave length</td></tr><tr><td>Laser intesity checker</td><td>COHERENT</td><td>1098293</td><td></td></tr><tr><td>Laser stimulator</td><td>Bio research center</td><td>STO2</td><td>reffered as pulse generator in text</td></tr><tr><td>Optic fiber with ferrule&amp;nbsp;</td><td>Thorlab</td><td>FP200URT-CANNULA-SP-JP</td><td></td></tr><tr><td>pAAV2-rh10</td><td>provided by PennVector Core</td><td></td><td></td></tr><tr><td>pAAV-EF1a-DIO-EYFP-WPRE-HGHpA</td><td>Addgene</td><td>plasimid # 20296</td><td></td></tr><tr><td>pAAV-EF1a-DIO-hChR2(H134R)-EYFP-WPRE-HGHpA</td><td>provided by Dr. Karl Deisseroth</td><td></td><td></td></tr><tr><td>Patch cord</td><td>Doric</td><td>D202-9089-0.4</td><td>0.4m length, laser conductor between laser and rotary joint</td></tr><tr><td>&lt;em&gt;pH&lt;/em&gt;elper</td><td>Stratagene</td><td></td><td></td></tr><tr><td>Photocurable dental cement</td><td>3M</td><td>56846</td><td></td></tr><tr><td>Serafin clamp</td><td>Stoelting</td><td>52120-43P</td><td></td></tr><tr><td>Shielded cable</td><td>mogami</td><td>W2780</td><td>Soldering to 6-pin socket for EEG/EMG recording</td></tr><tr><td>Sleep recording chamber</td><td>N/A</td><td>N/A</td><td>Custum-made (21 cm&amp;times; 29 cm &amp;times; 19 cm) with water tank holder</td></tr><tr><td>Sleep sign software</td><td>KISSEI COMTEC</td><td>N/A</td><td>for EEG/EMG analysis</td></tr><tr><td>Slip ring</td><td>neuroscience,inc</td><td>N/A</td><td>for EEG/EMG analysis</td></tr><tr><td>Stainless screw</td><td>Yamazaki</td><td>N/A</td><td>&amp;phi;1.0 x 2.0</td></tr><tr><td>Stainless wire</td><td>Cooner wire</td><td>AS633</td><td>&amp;nbsp;0.0130 inch diameter</td></tr><tr><td>Stereotaxic frame with digital console</td><td>Koph</td><td>N/A</td><td>Model 940</td></tr><tr><td>Syringe needle</td><td>Hamilton</td><td>7803-05</td><td></td></tr><tr><td>Vital recorder software</td><td>KISSEI COMTEC</td><td>N/A</td><td>for EEG/EMG recording</td></tr></tbody></table>

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.

The present study showed the effect of optogenetic excitation of GABABNST neurons on sleep state transition. ChR2-EYFP was focally expressed in GABA neurons in the BNST. An in situ hybridization histochemical study showed that ChR2-EYFP was colocalized in neurons expressing GAD 67 mRNA signals, indicating that these are GABAergic neurons. Immunohistochemical slice samples confirmed the position of the optic fiber, whose tip was just above the BNST25....

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

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 ...

optogenetic-manipulation-neural-circuits-during-monitoring

Research

JoVE Journal

Neuroscience

9.7K Views.  Kanazawa University. Deze procedure kan ons helpen bij het identificeren van de functie van specifieke neuro-circuit die betrokken zijn bij de slaap wakkerheid verordening met minder invasieve manipulatie en hoge temporele resolutie. Deze methode stelt ons in staat de monitoring oppervlak EEG in real time in combinatie met manipulatie van een specifieke neuro-circuit om de causale relatie tussen circuits en slaap wakkerheid staten te onderzoeken. Shota Kodani gaat de procedure demonstreren.Na bevestiging v...

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

Noninvasive, High-throughput Determination of Sleep Duration in Rodents

Traditionally, sleep is monitored by an electroencephalogram (EEG). EEG studies in rodents require surgical implantation of the electrodes followed by a long recovery period. To perform an EEG recording, the animal is connected to a receiver, creating an unnatural tether to the head-mount. EEG monitoring is time consuming, carries risk to the animal, and is not a completely natural setting for the measurement of sleep. Alternative methods to detect sleep, particularly in a high-throughput fashion, would greatly advance the field of sleep research. Here, we describe a validated method for detecting sleep via activity-based home-cage monitoring. Previous studies have shown that sleep assessed via this method has a high degree of agreement with sleep defined by traditional EEG-based measures. Whereas this method is validated for total sleep time, it is important to note that sleep bout duration should be assessed by an EEG which has better temporal resolution. The EEG can also differentiate rapid eye movement (REM) and non-REM sleep, giving more detail about the exact nature of sleep. Nevertheless, activity-based sleep determination can be used to analyze multiple days of undisturbed sleep and to assess sleep as a response to an acute event (like stress). Here, we show the power of this system to detect the response of mice to daily intraperitoneal injections.

We describe a high-throughput method of measuring sleep by means of activity-based home-cage monitoring. This method offers advantages over traditional EEG-based methods. It is well validated for the determination of total sleep duration and can be a powerful tool to monitor sleep in rodent models of human disease.

Traditionally, sleep is monitored by an electroencephalogram (EEG). EEG studies in rodents require surgical implantation of the electrodes followed by a ...

Behavior

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 for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

Optogenetic methods have emerged as a powerful tool for elucidating neural circuit activity underlying a diverse set of behaviors across a broad range of species. Optogenetic tools of microbial origin consist of light-sensitive membrane proteins that are able to activate (e.g., channelrhodopsin-2, ChR2) or silence (e.g., halorhodopsin, NpHR) neural activity ingenetically-defined cell types over behaviorally-relevant timescales. We first demonstrate a simple approach for adeno-associated virus-mediated delivery of ChR2 and NpHR transgenes to the dorsal subiculum and prelimbic region of the prefrontal cortex in rat. Because ChR2 and NpHR are genetically targetable, we describe the use of this technology to control the electrical activity of specific populations of neurons (i.e., pyramidal neurons) embedded in heterogeneous tissue with high temporal precision. We describe herein the hardware, custom software user interface, and procedures that allow for simultaneous light delivery and electrical recording from transduced pyramidal neurons in an anesthetized in vivo preparation. These light-responsive tools provide the opportunity for identifying the causal contributions of different cell types to information processing and behavior.

A Method for High Fidelity Optogenetic Control of Individual Pyramidal Neurons In vivo

The recent development of neuroscience tools that combine genetics and optics, termed "optogenetics", enables control over neural circuit activity with an unprecedented level of spatial and temporal resolution. Here we provide a protocol for integrating in vivo recording with optogenetic manipulation of genetically-defined subsets of prefrontal cortical and subicular pyramidal neurons.

Optogenetic methods  have emerged as a powerful tool for elucidating neural circuit activity  underlying a diverse set of behaviors across a broad range ...

Optogenetic Manipulation of Neuronal Activity to Modulate Behavior in Freely Moving Mice

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of single neurons and region-specific transmitter release, up to whole neuronal circuitries in the central nervous system, and allows the direct measurement of behavioral outcomes. Neurons express optogenetic tools via an injection of viral vectors carrying the DNA of choice, such as Channelrhodopsin2 (ChR2). Light is brought into specific brain regions via chronic optical implants that terminate directly above the target region. After two weeks of recovery and proper tool-expression, mice can be repeatedly used for behavioral tests with optogenetic stimulation of the neurons of interest.
Optogenetic modulation has a high temporal and spatial resolution that can be accomplished with high cell specificity, compared to the commonly used methods such as chemical or electrical stimulation. The light does not harm neuronal tissue and can therefore be used for long-term experiments as well as for multiple behavioral experiments in one mouse. The possibilities of optogenetic tools are nearly unlimited and enable the activation or silencing of whole neurons, or even the manipulation of a specific receptor type by light.
The results of such behavioral experiments with integrated optogenetic stimulation directly visualizes changes in behavior caused by the manipulation. The behavior of the same animal without light stimulation as a baseline is a good control for induced changes. This allows a detailed overview of neuronal types or neurotransmitter systems involved in specific behaviors, such as anxiety. The plasticity of neuronal networks can also be investigated in great detail through long-term stimulation or behavioral observations after optical stimulation. Optogenetics will help to enlighten neuronal signaling in several kinds of neurological diseases.

With optogenetic manipulation of specific neuronal populations or brain regions, behavior can be modified with high temporal and spatial resolution in freely moving animals. By using different optogenetic tools in combination with chronically implanted optical fibers, a variety of neuronal modulations and behavioral testing can be performed.

Optogenetic modulation of neuronal circuits in freely moving mice affects acute and long-term behavior. This method is able to perform manipulations of ...

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 ...

Quantifying Infra-slow Dynamics of Spectral Power and Heart Rate in Sleeping Mice

Three vigilance states dominate mammalian life: wakefulness, non-rapid eye movement (non-REM) sleep, and REM sleep. As more neural correlates of behavior are identified in freely moving animals, this three-fold subdivision becomes too simplistic. During wakefulness, ensembles of global and local cortical activities, together with peripheral parameters such as pupillary diameter and sympathovagal balance, define various degrees of arousal. It remains unclear the extent to which sleep also forms a continuum of brain states-within which the degree of resilience to sensory stimuli and arousability, and perhaps other sleep functions, vary gradually-and how peripheral physiological states co-vary. Research advancing the methods to monitor multiple parameters during sleep, as well as attributing to constellations of these functional attributes, is central to refining our understanding of sleep as a multifunctional process during which many beneficial effects must be executed. Identifying novel parameters characterizing sleep states will open opportunities for novel diagnostic avenues in sleep disorders.
We present a procedure to describe dynamic variations of mouse non-REM sleep states via the combined monitoring and analysis of electroencephalogram (EEG)/electrocorticogram (ECoG), electromyogram (EMG), and electrocardiogram (ECG) signals using standard polysomnographic recording techniques. Using this approach, we found that mouse non-REM sleep is organized into cycles of coordinated neural and cardiac oscillations that generate successive 25-s intervals of high and low fragility to external stimuli. Therefore, central and autonomic nervous systems are coordinated to form behaviorally distinct sleep states during consolidated non-REM sleep. We present surgical manipulations for polysomnographic (i.e., EEG/EMG combined with ECG) monitoring to track these cycles in the freely sleeping mouse, the analysis to quantify their dynamics, and the acoustic stimulation protocols to assess their role in the likelihood of waking up. Our approach has already been extended to human sleep and promises to unravel common organizing principles of non-REM sleep states in mammals.

Here, we present experimental and analytical procedures to describe the temporal dynamics of the neural and cardiac variables of non-REM sleep in mice, which modulate sleep responsiveness to acoustic stimuli.

Three vigilance states dominate mammalian life: wakefulness, non-rapid eye movement (non-REM) sleep, and REM sleep. As more neural correlates of behavior ...

Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools to selectively manipulate brain rhythms. Here we describe an approach combining projection-specific optogenetics with extracellular electrophysiology for high-fidelity control of hippocampal theta oscillations (5-10 Hz) in behaving mice. The specificity of the optogenetic entrainment is achieved by targeting channelrhodopsin-2 (ChR2) to the GABAergic population of medial septal cells, crucially involved in the generation of hippocampal theta oscillations, and a local synchronized activation of a subset of inhibitory septal afferents in the hippocampus. The efficacy of the optogenetic rhythm control is verified by a simultaneous monitoring of the local field potential (LFP) across lamina of the CA1 area and/or of neuronal discharge. Using this readily implementable preparation we show efficacy of various optogenetic stimulation protocols for induction of theta oscillations and for the manipulation of their frequency and regularity. Finally, a combination of the theta rhythm control with projection-specific inhibition addresses the readout of particular aspects of the hippocampal synchronization by efferent regions.

We describe the use of optogenetics and electrophysiological recordings for selective manipulations of hippocampal theta oscillations (5-10 Hz) in behaving mice. The efficacy of the rhythm entrainment is monitored using local field potential. A combination of opto- and pharmacogenetic inhibition addresses the efferent readout of hippocampal synchronization.

Extensive data on relationships of neural network oscillations to behavior and organization of neuronal discharge across brain regions call for new tools ...

Optogenetics Identification of a Neuronal Type with a Glass Optrode in Awake Mice

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the identification of the neuronal type in in vivo electrophysiological experiments in broad brain regions. In optogenetics experiments, it is critical to deliver the light to the recording site. However, it is often hard to deliver the stimulation light to the deep brain regions from the brain&#39;s surface. Especially, it is difficult for the stimulation light to reach the deep brain regions when the optical transparency of the brain surface is low, as is often the case with recordings from awake animals. Here, we describe a method to record spike responses to the light from an awake mouse using a custom-made glass optrode. In this method, the light is delivered through the recording glass electrode so that it is possible to reliably stimulate the recorded neuron with light in the deep brain regions. This custom-made optrode system consists of accessible and inexpensive materials and is easy to assemble.

This work introduces a method to perform an optogenetic single-unit recording reliably from an awake mouse using a custom-made glass optrode.

It is a major concern in neuroscience how different types of neurons work in neural circuits. Recent advances in optogenetics have enabled the ...

Electrocorticographic Recording of Cerebral Cortex Areas Manipulated Using an Adeno-Associated Virus Targeting Cofilin in Mice

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. Adeno-associated viruses (AAVs) are increasingly used to improve understanding of brain circuits and their functions. The AAV-mediated manipulation of specific cell populations and/or of precise molecular components has been tremendously useful to identify new sleep regulatory circuits/molecules and key proteins contributing to the adverse effects of sleep loss. For instance, inhibiting activity of the filamentous actin-severing protein&#160;cofilin&#160;using AAV prevents sleep deprivation-induced memory impairment. Here, a protocol is&#160;described that combines the manipulation of cofilin function in a cerebral cortex area with the recording of ECoG activity to examine whether cortical cofilin modulates the wakefulness and sleep ECoG signals. AAV injection is performed during the same surgical procedure as the implantation of ECoG and electromyographic (EMG) electrodes in adult male and female mice. Mice are anesthetized, and their heads are shaved. After skin cleaning and incision, stereotaxic coordinates of the motor cortex are determined, and the skull is pierced at this location. A cannula prefilled with an AAV expressing cofilinS3D, an inactive form of cofilin, is slowly positioned in the cortical tissue. After AAV infusion, gold-covered screws (ECoG electrodes) are screwed through the skull and cemented to the skull with gold wires inserted in the neck muscles (EMG electrodes). The animals are allowed three weeks to recover and to ensure sufficient expression of cofilinS3D. The infected area and cell type are verified using immunohistochemistry, and the ECoG is analyzed using visual identification of vigilance states and spectral analysis. In summary, this combined methodological approach allows the investigation of the precise contribution of molecular components regulating neuronal morphology and connectivity to the regulation of synchronized cerebral cortex activity during wakefulness and sleep.

This article describes a protocol for the manipulation of molecular targets in the cerebral cortex using adeno-associated viruses and for monitoring the effects of this manipulation during wakefulness and sleep using electrocorticographic recordings.

The use of electrocorticographic (ECoG) recordings in rodents is relevant to sleep research and to the study of a wide range of neurological conditions. ...

Selective Manipulation of Theta Oscillations in a Mouse Model

Source: Bender, F. et al., <a href="https://app.jove.com/v/57349">Optogenetic Entrainment of Hippocampal Theta Oscillations in Behaving Mice.</a> J. Vis. Exp (2018)
This video demonstrates a protocol to modulate theta oscillations in a genetically engineered mouse model with light-sensitive ion channels in its GABAergic medial septum neurons. By delivering light pulses to the hippocampus, the protocol aims to synchronize neuronal activity and generate theta oscillations, which are recorded as local field potentials.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
PV-Cre ...

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

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