We thank Dr. Masashi Kondo and Dr. Masanori Matsuzaki for providing virus vectors. This work was supported by Grants-in-Aid for Scientific Research (20H00481, 20A301, 20H05894, 20H05895 to S.O.), the Japan Agency for Medical Research and Development (JP19gm1310003 and JP17gm5010003 to S.O. and JP19dm0207082 to H.M.), the UTokyo Center for Integrative Science of Human Behavior (CiSHuB), the Japan Science and Technology Agency Moonshot R&#38;D (JPMJMS2024 to H.M.), Brain Science Foundation (to H. M.).

All experiments were approved by the Animal Ethics Committee and the Genetic Recombinant Experiment Safety Committee of Graduate School of Medicine and Faculty of Medicine, the University of Tokyo, and performed according to their guidelines.
1. Preparation
<ol>
	<li>Sterilize all the instruments and apparatuses necessary for the surgery using autoclave or 70% ethanol.</li>
	<li>Glass pipette preparation.
	<ol>
		<li>Firmly fix a 75 &#181;L calibrated capillary to the holder of the micropipette puller. Set the puller to the program with the recommended parameters as P (pressure) = 500, HEAT = 680, PULL = 30, VEL = 60, TIME = 180.</li>
		<li>After finishing the program (usually 4.5 to 5 s), the capillary is divided into two glass pipettes with tapering tails. Then, break the distal part of the tails with tweezers to keep the tip diameter within 30-50 &#181;m. For this, break the tails 1 cm distal to the end of the beveled part.</li>
	</ol>
	</li>
	<li>For craniotomy, prepare a cranial window by gluing a larger outer glass disc, 4 mm in diameter, 0.15 &#177; 0.02 mm in thickness, to a smaller inner glass disc, 2 mm in diameter, 0.525 &#177; 0.075 mm in thickness, using a UV light curing reagent (Figure 2A).</li>
</ol>
2. AAV injection
<ol>
	<li>Preparation of injection apparatus
	<ol>
		<li>Place a glass pipette connected to a 26 G Hamilton syringe through a tube onto a pipette holder of a stereotaxic instrument, and then tilt the pipette holder 60&#176; anteriorly from the vertical axis (Figure 1A,B).</li>
		<li>Fill the glass pipette, the syringe, and the connecting tube with liquid paraffin. Place the syringe on a microinjector.</li>
	</ol>
	</li>
	<li>Surgical procedure 
	NOTE: The following surgery was performed in a sterilized booth. A stereomicroscope was used for the surgery if necessary.
	<ol>
		<li>Anesthetize a male CX3CR1-EGFP mouse (detailed information in the Table of Materials) at 7 to 8 weeks old (bodyweight 22-26 g) with 3% isoflurane in a gas-tight chamber. After 3 min, take out the mouse from the chamber when it loses voluntary movement except breathing, and then switch to continuous anesthesia by a nasal tube with 2% isoflurane. 
		NOTE: Isoflurane is known to activate microglia. However, because imaging is performed 4 weeks after the surgery, there is no effect of isoflurane on the mice during imaging. Although isoflurane anesthesia is used for a few minutes when mice are placed in the stereotaxic instrument before imaging (step 4.2.1), we found that the effect of this short anesthesia is negligible.</li>
		<li>During surgery, keep the mouse warm with a heating pad at 37 &#176;C to prevent hypothermia. Apply eye ointment on both eyes to prevent corneal dryness and administer a meloxicam injection, subcutaneously (5 mg/kg). Attach the mouse to an auxiliary ear bar, and then fix the mouse on the stereotaxic instrument with its dorsal side up.</li>
		<li>Adjust the isoflurane concentration to an appropriate percentage (1%-2%) during the surgery while monitoring the breath and heart rate. Remove the hair using depilatory cream and disinfect the head several times in a circular motion with both an iodine-based or chlorhexidine-based scrub and alcohol.&#160;Then, inject 0.1 mL of 1% lidocaine subcutaneously and apply sterile drapes to secure the surgical site.</li>
		<li>Incise the scalp open along the midline with scissors and make the incision about 2 cm long, to ensure a good exposure of the skull above the right primary visual cortex. After the incision of the scalp, irrigate the head with saline throughout the whole procedure to prevent the exposed skull and brain from drying.</li>
		<li>Remove the periosteum on the exposed skull using forceps. Drill the skull on the stereotaxic coordinates: 3 mm lateral to the midline, 0.5 mm anterior to the lambda line, to create a small hole with approximately 0.5 mm diameter. Rinse off blood and tissue debris from the drill bit, if necessary.</li>
		<li>Fill the glass pipette with 1 &#181;L of rAAV2/1-hSyn-R-CaMP1.07 using the following procedure at a titer of 8.3 x 1012 vector genomes/mL22.
		<ol>
			<li>Advance the syringe to eject 1 &#181;L of liquid paraffin from the glass pipette. Place a piece of transparent film, approximately 2 cm x 2cm, on the exposed skull of the mouse and expel a droplet of 1 &#181;L of AAV solution on the film using a pipettor.</li>
			<li>Place the glass pipette tip into the droplet of AAV solution on the film and gently pull the syringe to aspirate the AAV solution. After the ejection, turn the T-shape stopcock to release the pressure inside the tube and the glass pipette.</li>
		</ol>
		</li>
		<li>Insert the glass pipette to a depth of 500 &#181;m from the brain surface through the hole created in step 2.2.5, and then inject 0.5 &#181;L of AAV solution using the microinjector (Figure 1C). Use the injection volume flow rate of 2.0 &#181;L/h; an injection of 0.5 &#181;L takes 15 min, and then subsequently wait for 5 min.</li>
		<li>After the injection, gently withdraw the glass pipette and rinse the brain surface with saline.</li>
	</ol>
	</li>
</ol>
3. Cranial window implantation
NOTE: Cranial window implantation follows the AAV injection on the same day.
<ol>
	<li>Mark a 2.5 mm diameter circle on the skull, centered 3 mm laterally from the lambda. Create a circular groove along the mark on the skull by drilling. Clean the debris as it may adversely affect the visibility of the skull and apply saline during the drilling process to prevent heating up.</li>
	<li>To check whether the drilling depth in the circular groove is sufficient to remove the central skull fragment, gently press the central skull with forceps. If it moves vertically with little resistance, the drilling depth is sufficient.</li>
	<li>When the groove reaches enough depth, insert the tip of the forceps into the bottom of the central skull fragment and gently lift and remove it to expose the brain surface (Figure 1D).</li>
	<li>Using a 27 G needle, prick and tear the dura at the edge of the exposed brain surface. Insert the tip of the forceps through the hole made at the edge of the dura. Hold the dura and peel it off to expose the brain surface. 
	NOTE: If bleeding occurs, rinse the brain surface with saline until the bleeding stops.</li>
	<li>Using forceps, disperse hemostatic fibers one by one in saline in a 3.5 mm dish, and then place the hemostatic fibers along the margins of the hole in which the transected dura is present.</li>
	<li>Place a cranial window prepared in step 1.3 on the exposed brain surface, and then adhere it to the skull with instant glue while gently pressing the window so that the window is in close contact with the skull.</li>
	<li>Afterward, apply instant glue to the whole exposed skull, and then carefully attach a head-plate onto the skull so that the window locates at the center of the square hole of the head-plate (Figure 2C). Ensure that the surface of the head plate is parallel to the surface of the glass window.</li>
	<li>After the glue has sufficiently hardened, apply dental cement to the exposed skull to reinforce the attachment between the head and the head-plate (Figure 2). 
	NOTE: If the experiments include the presentation of visual stimuli to mice, stray light to the imaging area should be avoided. It is advisable to blacken the cement by mixing it with the activated charcoal for the reduction of light reflection.</li>
	<li>Withdraw the mouse from the anesthesia after the cement has hardened sufficiently (about 20 min). Then, put the mouse into a new cage to recover alone.</li>
	<li>After confirming its consciousness and voluntary movement, indicating full recovery, put the mouse back in the home cage. During the recovery, administer a second, or more if necessary, meloxicam dose (once every 24 hours for 1-3 days) if obvious pain-indicating behaviors occur.</li>
</ol>
4. In vivo two-photon imaging of microglia and neuronal calcium dynamics
<ol>
	<li>Habituation of mice to the microscope apparatus
	<ol>
		<li>Three weeks after the surgery, induce anesthesia in the mouse with 3% isoflurane, and set the mouse under the 25x objective lens of a two-photon microscope using a custom-made stereotaxic instrument (Figure 3C). For the setup here, the mouse is set on the top of a treadmill and allowed to run freely.</li>
		<li>Approximately 10 min later, remove the mouse from the microscope stage and return it to the home cage. Repeat the habituation at least 3 times every alternate day before two-photon image acquisition.</li>
	</ol>
	</li>
	<li>Two-photon image acquisition 
	NOTE: We recommend performing the image acquisition more than 4 weeks after surgery, as microglial activation gradually returns to the basal level.
	<ol>
		<li>As in step 4.1, induce anesthesia in the mouse with 3% isoflurane and set the mouse under the objective lens of the two-photon microscope using a custom-made stereotaxic instrument.</li>
		<li>Install the custom-made shading device and an LCD monitor for visual stimulation as described below (Figure 3D).
		<ol>
			<li>Position the lens to focus on the brain surface, and then set this lens position as the original Z-position. Keep the x-y coordinates constant and elevate the objective lens; remove the mouse and stereotaxic instrument from the objective lens and treadmill.</li>
			<li>Attach a shading device on the top of the head plate using silicone, which is blackened by mixing with charcoal powder, and ensure the space between the head plate and the shading device is well-sealed.</li>
			<li>Fill the shading device with distilled water, and then fix the mouse and the stereotaxic frame again under the objective and on the treadmill. Carefully reset the focal plane at the brain surface, checking the depth of the objective lens. 
			NOTE: To avoid the risk of damaging the objective lens, set the xyz coordinates first, and then apply the shading device of the objective lens. It is also reasonable to install the cover first, and then adjust the coordinates, but cautious handling is necessary for this option.</li>
			<li>Cover the objective lens with black aluminum foil to avoid light contamination from the LCD monitor used for visual stimulation through the objective lens (Figure 3D). Set a 10-inch LCD monitor at 12.5 cm in front of the mouse&#39;s eyes to present visual stimuli (Figure 3D).</li>
		</ol>
		</li>
		<li>Configure the fluorescence emission collection filters to EGFP fluorescence (525/50 nm emission filter) and R-CaMP fluorescence (593/46 nm emission filter) and the excitation wavelength to 1,000 nm. Acquire images with the spatial resolution of 0.25 &#181;m/pixel using a 25x 1.1 NA objective and GaAsP PMTs.</li>
		<li>Find the imaging region in which R-CaMP(+) neurons and EGFP(+) microglia can be simultaneously imaged in layer 2/3. For this setup, data obtained in Figure 4 and Video 1 were at a depth of 100 &#181;m from the pial surface, using a live image preview. Keep the power of the excitation laser as low as possible to avoid photo-bleaching and damage caused by increased local temperature in the imaging region.</li>
		<li>Acquire images at the frame rate of 30 Hz. Simultaneously with the image acquisition, present a drifting-grating visual stimuli at 100% contrast, 1.5 Hz, 0.04 cycles per degree in 12 directions at 6 orientations from 0&#176; to 150&#176; in 30&#176; steps to the mouse17.</li>
		<li>After the image acquisition, remove the mouse from the microscope stage, detach the shading device and the stereotaxic instrument from the mouse, and return the mouse to its home cage.</li>
	</ol>
	</li>
	<li>Data registration
	<ol>
		<li>Convert and save the acquired image data (nd2 format in this case) as a series of tiff image files for each color channel, using Fiji or the software supplied with the microscope.</li>
		<li>Apply Turboreg, a plugin of Fiji, to perform registration with mode = translation. Use averaged image as the reference image with the tiff image files of EGFP channel.</li>
		<li>Perform the following processing for each frame using MATLAB. 
		NOTE: Although MATLAB is used in the following processing, the description mainly focuses on the intention of each step so that data can be analyzed by other programming software such as Python.
		<ol>
			<li>Use the imread function to read two EGFP tiff images before and after registration of the same frame, respectively. Use the imread function to read the R-CaMP tiff image of the same frame.</li>
			<li>Use the normcorr2 function to calculate the correlation function between the vectorized variables of the two GFP images read in step 4.3.3.1.</li>
			<li>Use the max function to detect the linear index with the highest cross-correlation of the correlation function, and then use the ind2sub function to calculate the movement position of the registered image from its original image.</li>
			<li>Use the imwarp function to translate the R-CaMP image to the position calculated in step 4.3.3.3, and then use the imwrite function to save the registered image of R-CaMP.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Generation of calcium traces
	<ol>
		<li>Perform the following processing using MATLAB. Use the imread function to read all the registered R-CaMP tiff images generated in step 4.3.3.4. If the registered images have been already loaded as a variable in workspace, omit this step.</li>
		<li>Use the mean function to generate a time-projection image. Use the imshow function to show the averaged image as a figure; use the roipoly function to generate a binary ROI mask of the target structure (dendritic spine in this data).</li>
		<li>Using the mean function with the images obtained by multiplying the binary mask with images of each frame as input variables, calculate the averaged fluorescence intensity within the ROI for each frame.</li>
		<li>As described previously17, to the variable obtained in step 4.4.3, apply the butter worth frequency filter at cutoff = 1.6 s, for cutting high-frequency noise, and then apply the sliding median filter at the time window = 200 s, for cutting low-frequency noise.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We describe the protocol of AAV injection and craniotomy for simultaneous imaging of microglial dynamics and neuronal activity in awake mice as well as data processing. This technique can uncover the coordination between microglial dynamics and neuronal activity on time scales ranging from sub-second to tens of seconds.
The surgery protocol involves several technically demanding steps. AAV injection is one of the critical steps. Unsuccessful AAV injection may cause a significant reduction in the expression of R-CaMP. There are two main reasons: clogged glass pipettes and tissue damage. The clogging of glass pipettes reduces or completely blocks the ejection of AAV solution. This situation can be avoided by coloring the AAV solution with a dye such as fast green and visually confirming the success of the injection. Tissue damage caused by AAV injection prevents exogenous gene expression near the center of the injection site. The damage induced by AAV injection is likely to be caused by a sudden increase in the efflux from the tip of the glass pipette after the accumulation of the pressure inside the pipette. Therefore, the outflow of AAV solution from the glass pipette should be constant during the injection. Selecting glass pipettes with a slightly wider diameter at their tips would solve this problem. In cranial window implantation, the speed of surgery is critical. If the surgery takes too long, the brain tissue can be severely damaged. Accordingly, repeated practice is necessary to ensure the speed and smoothness of surgery. In addition, during the 4 weeks from surgery to imaging, the quality of imaging may be reduced by tissue regeneration between the cranial window and the brain tissue by the conventional method17. The method here overcomes this issue by applying thick glasses for the inner glass disc of the cranial windows to inhibit tissue regeneration and keep the window clear. In the system described here, this effect was evident by using an inner glass disc with a thickness of 0.525 &plusmn; 0.075 &#181;m.
The method can be applied successfully to mice older than 4 weeks, but the application to younger mice may be problematic. In juvenile mice, the skull shows rapid and prominent growth, which induces a mismatch between the cranial bone and the glass window.
In some cutting-edge studies, in vivo two-photon imaging was used to study microglia-neuron interactions12,21,23. Especially in the pioneering work by Merlini et al., they performed simultaneous in vivo imaging of microglial dynamics and neural activity within the cell bodies21. In this method, by using thicker inner glasses for cranial windows, we could suppress motion artifacts in the depth orientation and achieve the stable measurement of neuronal activity in microstructures, such as dendritic spines. This method would help investigate synapse-microglial process interactions in awake mice.
Recently, in vitro study demonstrated that thin filopodia-like microglial processes have faster motility than thick processes, suggesting the importance of their faster motility in surveillance19. The system here can track the motility of thin microglial processes with a temporal resolution from a sub-second to a few tens of seconds. This property helps elucidate the functional significance of their motility for surveillance in vivo.
In the future, the combination of this imaging technique with interventions, such as optogenetics24 or chemogenetics25, applied to local neural circuits or interregional neural connections would shed light on novel microglial functions in synaptic development and plasticity which sculpt characteristics of neural circuits. Also, further integration of imaging, intervention, and behavioral tasks would contribute to revealing the coordination of microglia and neurons underlying specific behaviors.

There is growing evidence that the internal state of the body constantly influences the brain functions in animals1,2,3,4,5. Accordingly, to gain a deeper understanding of brain functions, it is crucial to elucidate how glial cells in the brain monitor biological conditions in the periphery and relay the information to neurons.
Microglia, immune cells in the brain, are involved in synaptic development and plasticity, which sculpt characteristics of neural circuits in the brain6,7,8,9. For instance, the pioneering work by Wake et al. demonstrated that microglial processes make contact with synapses in a neuronal activity-dependent manner in the mouse neocortex and that artificial ischemia induces synapse loss following prolonged contact with microglia10. Tremblay et al. found that alteration of visual experience changes the modality of microglial interaction with synapses. During the critical period when dendritic spine turnover in the primary visual cortex (V1) is increased by binocular deprivation, dark adaptation reduces the motility of microglial processes and increases both their contact frequency with synaptic clefts and the number of cellular inclusions in microglia11. These results suggest that microglial processes sense neural activity and their surrounding environment to remodel neural circuits. Furthermore, a recent study reported that microglial surveillance differs between awake and anesthetized conditions, suggesting the importance of experiments using awake mice for investigating neuron-microglia communication under physiological conditions12.
In vivo two-photon calcium imaging is a powerful tool for the recording of calcium dynamics, reflecting ongoing neuronal firing, in hundreds of neurons simultaneously in a living animal13,14,15. Calcium imaging in neurons generally requires a temporal resolution of more than a few Hz to track rapid neuronal responses16,17. In contrast, previous studies tracking microglial dynamics have sampled microglial structures at a relatively low temporal resolution of less than 0.1 Hz18,19,20. One recent study applied simultaneous two-photon imaging to understand neuron-microglia communication21. However, it is still unclear how dynamic microglial processes respond to the surrounding neural activity at a temporal resolution of more than a few Hz in awake mice. In order to address this issue, we describe a simultaneous in vivo two-photon imaging method of neural activity and microglial dynamics with a temporal resolution higher than 1 Hz in awake mice. This method allows us to achieve stable imaging in awake mice at a higher frame rate (maximum, 30 Hz with the pixel frame size of 512 x 512 pixels) and provides a more favorable way to investigate the surveillance behavior of microglia or their interaction with neuronal activity in awake mice.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10-inch LCD monitor</td>
			<td>EIZO</td>
			<td>DuraVision FDX1003</td>
			<td>For presenting grating visual stimuli</td>
		</tr>
		<tr>
			<td>26G Hamilton syringe</td>
			<td>Hamilton</td>
			<td>701N</td>
			<td></td>
		</tr>
		<tr>
			<td>27G needle</td>
			<td>Terumo</td>
			<td>NN-2719S</td>
			<td></td>
		</tr>
		<tr>
			<td>AAV-hSyn-R-CAMP1.07</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Kindly gifted from Prof. Matsuzaki&#39;s laboratory&#160;</td>
		</tr>
		<tr>
			<td>Activated charcoal powder</td>
			<td>Nacalai tesque</td>
			<td>07909-65</td>
			<td></td>
		</tr>
		<tr>
			<td>Black aluminum foil</td>
			<td>THORLABS</td>
			<td>BKF12</td>
			<td></td>
		</tr>
		<tr>
			<td>CX3CR1-EGFP mouse</td>
			<td>Jackson laboratory</td>
			<td>IMSR_JAX: 005582</td>
			<td>CX3CR-1EGFP/+ knock-in/knock-out mice expressing EGFP in microglia in the brain under the control of the endogenous Cx3cr1 locus.</td>
		</tr>
		<tr>
			<td>DENT SILICONE-V</td>
			<td>Shofu Inc</td>
			<td>N/A</td>
			<td>For the attachment of a shading device to a head-plate</td>
		</tr>
		<tr>
			<td>Dental cement (quick resin, liquid)</td>
			<td>Shofu Inc</td>
			<td>N/A</td>
			<td>AB</td>
		</tr>
		<tr>
			<td>Dental cement(quick resin, powder)</td>
			<td>Shofu Inc</td>
			<td>N/A</td>
			<td>B Color 3</td>
		</tr>
		<tr>
			<td>Drill</td>
			<td>Toyo Associates</td>
			<td>HP-200</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass capillary pipette</td>
			<td>Drummond Scientific Company</td>
			<td>2-000-075</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass disc (large)</td>
			<td>Matsunami</td>
			<td>N/A</td>
			<td>4mm in diameter, 0.15&#177;0.02mm in thickness</td>
		</tr>
		<tr>
			<td>Glass disc (small)</td>
			<td>Matsunami</td>
			<td>N/A</td>
			<td>2mm in diameter, 0.525&#177;0.075mm in thickness</td>
		</tr>
		<tr>
			<td>Head-plate</td>
			<td>Customized</td>
			<td>N/A</td>
			<td>Material: SUS304, thickness: 0.5mm, see Figure2B for the shape</td>
		</tr>
		<tr>
			<td>Hemostatic fiber</td>
			<td>Davol Inc</td>
			<td>1010090</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ Fiji software</td>
			<td></td>
			<td>Free software</td>
			<td>For data registration</td>
		</tr>
		<tr>
			<td>Instant glue (Aron alpha)</td>
			<td>Daiichi Sankyo</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Pfizer</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Lidocaine</td>
			<td>Astrazeneca</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>MATLAB 2017b</td>
			<td>MathWorks</td>
			<td>N/A</td>
			<td>For data registration and processing</td>
		</tr>
		<tr>
			<td>Meloxicam</td>
			<td>Tokyo Chemical Industry</td>
			<td>M1959</td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjector</td>
			<td>KD scienfitic</td>
			<td>KDS-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instrument Company</td>
			<td>P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Multi-photon excitation microscope</td>
			<td>NIKON</td>
			<td>N/A</td>
			<td>The commercial name is &#34;A1MP+&#34;.</td>
		</tr>
		<tr>
			<td>Objective lens</td>
			<td>NIKON</td>
			<td>N/A</td>
			<td>The commercial name is &#34;CFI75 apochromat 25xC W&#34;.</td>
		</tr>
		<tr>
			<td>Paraffin Liquid</td>
			<td>Nacalai tesque</td>
			<td>26132-35</td>
			<td></td>
		</tr>
		<tr>
			<td>Psychtoolbox</td>
			<td></td>
			<td>Free software</td>
			<td>For presenting grating visual stimuli</td>
		</tr>
		<tr>
			<td>Shading device</td>
			<td>Customized</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>M165 FC</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic instrument</td>
			<td>Narishige Scientific Instrument</td>
			<td>SR-611</td>
			<td>For the surgery</td>
		</tr>
		<tr>
			<td>Stereotaxic instrument</td>
			<td>Customized</td>
			<td>N/A</td>
			<td>For fixing mice under the two-photon microscope</td>
		</tr>
		<tr>
			<td>Stopcock</td>
			<td>ISIS</td>
			<td>VXB1079</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical silk</td>
			<td>Ethicon</td>
			<td>K881H</td>
			<td></td>
		</tr>
		<tr>
			<td>Treadmill</td>
			<td>Customized</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>UV light curing agent</td>
			<td>Norland Products Inc</td>
			<td>NOA 65</td>
			<td></td>
		</tr>
		<tr>
			<td>Vaporizer</td>
			<td>Penlon</td>
			<td>Sigma Delta</td>
			<td>Anesthetic machine</td>
		</tr>
	</tbody>
</table>

simultaneous imaging of microglial dynamics and neuronal activity in awake mice

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in the brain sense various biological conditions in the periphery and transmit the signals to neurons. Microglia, immune cells in the brain, are involved in synaptic development and plasticity. Therefore, the contribution of microglia to neural circuit construction in response to the internal state of the body should be tested critically by intravital imaging of the relationship between microglial dynamics and neuronal activity.
Here, we describe a technique for the simultaneous imaging of microglial dynamics and neuronal activity in awake mice. Adeno-associated virus encoding R-CaMP, a gene-encoded calcium indicator of red fluorescence protein, was injected into layer 2/3 of the primary visual cortex in CX3CR1-EGFP transgenic mice expressing EGFP in microglia. After viral injection, a cranial window was installed onto the brain surface of the injected region. In vivo two-photon imaging in awake mice 4 weeks after the surgery demonstrated that neural activity and microglial dynamics could be recorded simultaneously at the sub-second temporal resolution. This technique can uncover the coordination between microglial dynamics and neuronal activity, with the former responding to peripheral immunological states and the latter encoding the internal brain states.

We performed the AAV injection and cranial window implantation in V1 of an 8-week-old CX3XR1-EGFP transgenic mouse as described in this protocol. Four weeks after the surgery, we performed simultaneous in vivo two-photon imaging of R-CaMP-based neural activity and microglial dynamics in layer 2/3 of V1 (Figure 4 and Video 1). During the imaging, the mouse was placed on a treadmill and allowed to run freely. Images were acquired at the frame rate of 30 Hz with the spatial resolution of 0.25 &#181;m/pixel using a 25x, 1.1 NA objective and GaAsP PMTs. Both EGFP and R-CaMP were excited at a wavelength of 1,000 nm, and EGFP fluorescence (525/50 nm emission filter) and R-CaMP fluorescence (593/46 nm emission filter) were collected simultaneously. After the registration of acquired data to minimize motion artifacts, every four frames were averaged into one frame to reduce background noise. Grating visual stimuli were presented to the mouse, and the visual responses of neurons and individual dendritic spines were analyzed (Figure 4D). Microglial processes showed fast dynamics and changed their morphology within 10 s (Figure 4E&#160;and Video 1). As detailed in the Discussion section, when the AAV injection failed, the expression of R-CaMP could not be detected. If the surgery was not successful, the signals of EGFP and R-CaMP could not be observed.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64111/64111fig01.jpg" /> 
Figure 1: AAV injection. (A) A glass pipette was connected to a 26 G Hamilton syringe using a silicon tube. (B) The set-up for AAV injection. (C) AAV solution was injected via the glass pipette tilted 60&#176; anteriorly from the vertical axis. (D)&#160;Schematic diagram of the procedure of open skull (described in step 3.3). <a href="https://www.jove.com/files/ftp_upload/64111/64111fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64111/64111fig02v2.jpg" /> 
Figure 2: Cranial window and head-plate. (A) Schematic diagram of cranial window implantation. (B) Custom-made head-plates were used. For imaging in the right hemisphere, the shorter arm should be used on the right side. (C) Dorsal view of a mouse with a cranial window and an implanted head plate. (D) High magnification view of the cranial window shown in Figure 2C. (E) Dorsal view of a mouse fixed with the custom-made stereotaxic instrument. <a href="https://www.jove.com/files/ftp_upload/64111/64111fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64111/64111fig03.jpg" /> 
Figure 3: The set-up for in vivo two-photon imaging. (A) Dorsal view of shading device. (B) Side view of shading device. (C)&#160;View of a mouse with the shading device on the head plate under the objective lens. The mouse is placed on the treadmill. (D) View under the two-photon excitation microscope. A monitor presenting visual stimuli is placed on the right side of the figure. The objective lens is covered with the shading device and black aluminum foil to avoid light from the monitor to the objective lens. <a href="https://www.jove.com/files/ftp_upload/64111/64111fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64111/64111fig04.jpg" /> 
Figure 4: Microglial dynamics and visual responses in a dendritic spine. (A-C) Time-averaged images of EGFP(+) microglia and R-CaMP(+) neurons in layer 2/3 of V1 in a 12-week-old CX3XR1-EGFP transgenic mouse. Signal intensity in each channel was normalized independently. (D) Visual responses in a dendritic spine. Stripe diagrams with black arrows indicate the direction of the grating stimuli presented in the timing indicated by gray columns. In calcium traces, gray lines indicate individual responses, and a black line indicates their average. In the right inset, a yellow arrowhead indicates the dendritic spine in which the region of interest (ROI) was generated to acquire the calcium traces. (E) Microglial process (cyan arrowhead) showed rapid retraction in 10 s. <a href="https://www.jove.com/files/ftp_upload/64111/64111fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Two-photon imaging of neuronal activity and microglial dynamics. Imaging data of EGFP(+) microglia and R-CaMP(+) neurons in layer 2/3 of V1 in a 12-week-old CX3XR1-EGFP transgenic mouse. On the left side of the movie, magenta indicates R-CaMP in neurons, and green indicates EGFP in microglia. The center of the movie corresponds to the EGFP signal, and the right side corresponds to the R-CaMP signal. Signal intensity in each channel was normalized independently. The frame rate of the movie is 40 times faster than the actual rate. Scale bar = 10 &#181;m <a href="https://www.jove.com/files/ftp_upload/64111/220330CatMovieForJoVE_IntNormalized-2.mp4" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice at JoVE.com

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Here, we describe a protocol combining adeno-associated virus injection with cranial window implantation for simultaneous imaging of microglial dynamics and neuronal activity in awake mice.

Since brain functions are under the continuous influence of the signals derived from peripheral tissues, it is critical to elucidate how glial cells in ...

simultaneous-imaging-microglial-dynamics-neuronal-activity-awake

Research

JoVE Journal

Neuroscience

2.1K Views.  The University of Tokyo. Here, we describe a protocol combining adeno-associated virus injection with cranial window implantation for simultaneous imaging of microglial dynamics and neuronal activity in awake mice.

Video: Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Functional Mapping with Simultaneous MEG and EEG

We use magnetoencephalography (MEG) and electroencephalography (EEG) to locate and determine the temporal evolution in brain areas involved in the processing of simple sensory stimuli. We will use somatosensory stimuli to locate the hand somatosensory areas, auditory stimuli to locate the auditory cortices, visual stimuli in four quadrants of the visual field to locate the early visual areas. These type of experiments are used for functional mapping in epileptic and brain tumor patients to locate eloquent cortices. In basic neuroscience similar experimental protocols are used to study the orchestration of cortical activity. The acquisition protocol includes quality assurance procedures, subject preparation for the combined MEG/EEG study, and acquisition of evoked-response data with somatosensory, auditory, and visual stimuli. We also demonstrate analysis of the data using the equivalent current dipole model and cortically-constrained minimum-norm estimates. Anatomical MRI data are employed in the analysis for visualization and for deriving boundaries of tissue boundaries for forward modeling and cortical location and orientation constraints for the minimum-norm estimates.

We use magnetoencephalography (MEG) and electroencephalography (EEG) to map brain areas involved in the processing of simple sensory stimuli.

We use magnetoencephalography (MEG) and electroencephalography (EEG) to locate and determine the temporal evolution in brain areas involved in the ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

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 ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Deep Brain Stimulation with Simultaneous fMRI in Rodents

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

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

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

Super-resolution Imaging of Neuronal Dense-core Vesicles

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and medical applications. Strengths of fluorescence include its sensitivity, specificity, and compatibility with live imaging. Unfortunately, conventional forms of fluorescence microscopy suffer from one major weakness, diffraction-limited resolution in the imaging plane, which hampers studies of structures with dimensions smaller than ~250 nm. Recently, this limitation has been overcome with the introduction of super-resolution fluorescence microscopy techniques, such as photoactivated localization microscopy (PALM). Unlike its conventional counterparts, PALM can produce images with a lateral resolution of tens of nanometers. It is thus now possible to use fluorescence, with its myriad strengths, to elucidate a spectrum of previously inaccessible attributes of cellular structure and organization.
Unfortunately, PALM is not trivial to implement, and successful strategies often must be tailored to the type of system under study. In this article, we show how to implement single-color PALM studies of vesicular structures in fixed, cultured neurons. PALM is ideally suited to the study of vesicles, which have dimensions that typically range from ~50-250 nm. Key steps in our approach include labeling neurons with photoconvertible (green to red) chimeras of vesicle cargo, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a high-resolution PALM image. We also demonstrate the efficacy of our approach by presenting exceptionally well-resolved images of dense-core vesicles (DCVs) in cultured hippocampal neurons, which refute the hypothesis that extrasynaptic trafficking of DCVs is mediated largely by DCV clusters.

We describe how to implement photoactivated localization microscopy (PALM)-based studies of vesicles in fixed, cultured neurons. Key components of our protocol include labeling vesicles with photoconvertible chimeras, collecting sparsely sampled raw images with a super-resolution microscopy system, and processing the raw images to produce a super-resolution image.

Detection of fluorescence provides the foundation for many widely utilized and rapidly advancing microscopy techniques employed in modern biological and ...

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.

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

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

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 ...