We would like to thank M. Kondo and M. Matsuzaki for lending us the XLPLN25XSVMP objective lens. This work was financially supported from Japan Society for the Promotion of Science (JSPS) by Grant-in-Aid for JSPS Research Fellow (18J21331 to R.K.) and Grants-in-Aid for Scientific Research (20H00481, 20A301, 20H05894, 20H05895 to S.O.), from the Japan Agency for Medical Research and Development (JP19gm1310003 and JP17gm5010003 to S.O.), and from the Japan Science and Technology Agency by Moonshot R&#38;D (JPMJMS2024 to H.M.).

All animal experiments were approved and carried out following the policies of the Animal Ethics Committee and the Genetic Recombinant Experiment Safety Committee at the University of Tokyo. Both male and female CX3CR1-GFP mice, aged 2-4 months, were used in the experiments. For the safety of the experimenters and the maintenance of sterile conditions, all the procedures were performed with white coats, masks, and sterile gloves. All instruments were sterilized before use.&#160;
1. Preparation of instruments and animals
<ol>
	<li>Assemble a glass-bottom metal tube (Figure 1A).
	<ol>
		<li>Apply a small drop of UV-curing optical adhesive to one end of a custom-made stainless tube (outer diameter 3.0 mm, inner diameter 2.8 mm, height 1.7 mm). Spread the glue on the circular edge of the tube uniformly. 
		NOTE: A sufficient amount of adhesive should be used to cover the entire edge of the tube.</li>
		<li>Place a circular glass coverslip (3.0 mm diameter, 0.15 &#177; 0.02 mm thickness) to the glue-covered end of the metal tube, and lightly press the coverslip against the tube so that the gap between the tube and the glass is filled with the glue. Next, adjust the position of the glass to fit within the outer edge of the metal tube.</li>
		<li>Irradiate the glass with UV light for a sufficient time to cure the adhesive. 
		NOTE: Make sure that the glass and the tube are completely attached. If the adhesion is insufficient, the glass may come off after surgery, resulting in unsuccessful imaging or leakage of cerebrospinal fluid (CSF).</li>
		<li>Disinfect the assembled glass-bottom metal tube with 70% ethanol and wash with sterile saline to remove debris.</li>
	</ol>
	</li>
	<li>Sterilize all surgical instruments with a dry heat sterilizer.</li>
	<li>Prepare mice for the surgery of window implantation. 
	&#8203;NOTE: Mice should be of appropriate age. It is preferable that they are genetically engineered to express fluorescent proteins in the CA1 and the dentate gyrus (DG). These points are further explained in the Discussion section.</li>
</ol>
2. Anesthesia and head fixation
<ol>
	<li>Anesthetize a mouse by intraperitoneal injection of ketamine-xylazine (100 mg/kg ketamine and 10 mg/kg xylazine). Administer meloxicam (2 mg/kg) by subcutaneous injection for pre-operative analgesia. Confirm the disappearance of the response to painful stimuli, such as tail pinching to ensure adequate depth of anesthesia. Add a half dose of ketamine-xylazine each time the anesthesia wears off during surgery, typically 1 h after the initial administration and every 30 min thereafter. Use a heating pad under the body to provide thermal support and apply ophthalmic ointment on the eyes to prevent dryness under anesthesia. 
	NOTE: The choice of anesthesia for surgery is essential. This point is discussed in detail in the Discussion section.</li>
	<li>Apply depilatory cream&#160;to remove hair from the scalp. Disinfect the scalp several times in a circular motion with povidone-iodine scrub followed by alcohol. Place the mouse on the surgical platform prepared with a sterile waterproof pad and apply sterile drapes to secure the surgical site. Then,&#160;cut and remove the scalp&#160;circularly with a scalpel so that the parietal and the occipital bones are fully exposed on the operative side.</li>
	<li>Remove the subcutaneous connective tissue by rubbing it with a cotton swab while applying sterile saline to clear the bleeding. Gently scrape the cranial surface with a round-tipped miniature knife to remove the periosteum, which helps to strengthen the adhesion between the cement and the bone (see step 2.6).</li>
	<li>Apply dental etching material to the entire surface, wait for tens of seconds, and rinse with sufficient sterile saline.</li>
	<li>Using waterproof ink, mark the area of the bone to be removed (a circular area with a diameter of 3.0 mm, centered about 2.5 mm posterior and 2.5 mm lateral to the bregma). After marking, dry the skull surface thoroughly. 
	NOTE: The position of craniotomy depends on the size of the hippocampus and should be optimized for different ages. Useful indicators for the position of craniotomy will be presented below (see step 3.4.5, NOTE).</li>
	<li>Attach the rectangular aluminum plate with a thickness of 1.0 mm and with a 5.0 mm diameter hole at the center to the skull using dental resin cement according to the manufacturer&#39;s instructions.
	<ol>
		<li>Carefully align the hole at the center of the plate with the circular area marked with a pen (see step 2.5) so that the aluminum plate is parallel to the marked surface of the skull (Figure 1B). Ensure that the cement tightly seals all gaps between the plate and the bone. Wait for about 5 min for the cement to be sufficiently hardened.</li>
	</ol>
	</li>
	<li>Place the mouse onto the head holding device with angle adjusters via the attached plate.</li>
</ol>
3. Implantation of the observation window
NOTE: The following surgical procedures should be performed under a stereo microscope.
<ol>
	<li>Make a circular groove on the skull using a dermal punch with a diameter of 3.0 mm. Align the circular groove and the area marked with a pen in step 2.5. Twist the dermal punch with gentle pressure so that the groove depth gradually increases. Frequently check that the groove depth is constant over the entire circumference.</li>
	<li>When the dermal punch reaches the innermost layer of the cranium, before touching the underlying dura, gently lift the central bone island using a 30 G needle. 
	NOTE: Slight liquid leakage from the bottom of the groove indicates that the groove is close to the dura.</li>
	<li>Remove the dura using a dura picker. 
	NOTE: Complete removal of the dura within the cranial window is necessary and will require careful resection of the residual dura at the periphery using fine forceps.</li>
	<li>Gently aspirate the tissue to expose the alveus of the hippocampus by 23 G or 25 G blunt needles connected to the aspirator. 
	NOTE: This step is the most critical for successful imaging. Key technical points for the tissue aspiration are described in detail in the Discussion section.
	<ol>
		<li>Aspirate the pia and the pial vessels.</li>
		<li>Aspirate the cortical tissue from the surface. Keep the depth of exposed tissue surface homogeneous over the entire cranial window. Gradually deepen the aspiration until fibers of the external capsule are exposed.</li>
		<li>Carefully position the suction tip to the external capsule near the lateral ventricle (LV) located at the rostrolateral end of the cranial window. Remove the surface layer of the external capsule running in the caudomedial to rostrolateral direction, and then the inner layer running in the mediolateral direction. 
		NOTE: The external capsule near the LV can be easily removed from the underlying structure.</li>
		<li>Confirm the removal of the entire external capsule by checking the full exposure of the surface of the alveus and the dorsal hippocampal commissure (DHC). 
		NOTE: The DHC covers the caudomedial portion of the CA1. The alveus and the DHC can be distinguished from the external capsule by their fiber orientations. Namely, the fibers within the alveus and the DHC run in the rostromedial to caudolateral direction, and can be distinguished from the fibers in the external capsule. The alveus and the DHC are tightly attached to the CA1 and should not be damaged. All fragments of the external capsule should be removed, since any remaining fragments will prevent the direct contact of the glass coverslip to the alveus.</li>
		<li>Confirm that the bleeding has stopped thoroughly and fill the hole with sterile saline to the level of the skull surface. 
		NOTE: The field of view here helps to judge whether the hole is in the appropriate position for the specific age of the mouse. Ideally, the alveus and the underlying CA1 should occupy most of the central area. A part of the DHC is visible in the caudomedial area. The opening of LV is detectable at the rostrolateral edge (Figure 3A).</li>
	</ol>
	</li>
	<li>Insert the glass-bottom metal tube (see step 1.1) vertically into the hole while aspirating the excess water that overflows from the sides of the tube until the glass bottom lightly presses against the alveus and partially flattens the underlying CA1. 
	NOTE: This pressure needs to be adjusted appropriately. If it is too strong, the CA1 will be damaged. If it is too weak, the spherical aberration due to the curvature of the CA1 will impair imaging resolution in deep structures.</li>
	<li>Using dental resin cement, fix the wall of the inserted tube to the surrounding skull (Figure 1C). Ensure that the entire circumference is tightly sealed and that there is no leakage of air or CSF. Wait for about 5 min for the cement to harden. 
	NOTE: If the cement is placed when its viscosity is low, it may leak into the brain parenchyma through the gap between the tube and the skull and damage the brain. Therefore, the cement should be applied after polymerization initiation, which increases the viscosity and reduces leakage through the gap.</li>
	<li>Wash the entire plate, the skull, and the inside of the tube with sterile saline to remove debris.</li>
</ol>
4. Two-photon microscopy immediately after surgery for quality check of the surgery
<ol>
	<li>Set the mouse in the head-holding device under the objective lens of the two-photon microscope and on the motorized XY scanning stage. Use a heating pad for thermal support. Monitor and adjust the depth of anesthesia as described in Step 2.1, as this quality check may take up to 30 mins.&#160; 
	NOTE: The water-immersion objective lens with a working distance (WD) longer than 3 mm and a high numerical aperture (NA) is recommended. A correction collar of the objective lens can improve the resolution by compensating for the refractive index mismatch between the glass and the biomaterials.</li>
	<li>Fill the space between the glass and the objective lens with water, taking special care to avoid air bubbles. If necessary, expand the area of the metal plate using a plastic film, which holds more water, to cover the entire front lens of the objective.</li>
	<li>Turn on a femtosecond pulsed laser of 920 nm wavelength for the excitation of the fluorescent probes expressed in the brain and start image acquisition software.</li>
	<li>Adjust the focus to the CA1 with the use of the motorized stage and the motorized focus system. Position the target structure with the guidance of the fluorescence emitted from the brain parenchyma and the reflected light from the edge of the metal tube under continuous illumination by the pulsed laser.</li>
	<li>Measure the depths of the brain parenchyma touching the glass bottom by adjusting the focus with the motorized focus system. Compare the depths at different horizontal locations to judge the alignment of the glass bottom and the imaging plane.
	<ol>
		<li>Adjust the angle of the mouse head by tilting the head holding device until the glass bottom is set to be parallel to the imaging plane.</li>
	</ol>
	</li>
	<li>Adjust the correction collar of the objective to achieve the highest resolution at the depth of the target structure in the CA1.</li>
	<li>Confirm that the fluorescent cells in the molecular layer (ML) of the DG upper blade at a depth of 500 &#181;m from the glass bottom can be imaged in the entire field of view. 
	NOTE: This step is important in judging the quality of surgery. If the CA1 is damaged, focal brain edema prevents fluorescence detection at a depth of more than 200 &#181;m from the surface. Only when there is no damage to the CA1, and the curvature of the CA1 layers is properly flattened by the overlying coverslip, the DG signals can be detected immediately after surgery (Figure 2A-D). The Representative Results section provides a simple way to determine the boundary between the CA1 and the DG.</li>
</ol>
5. Postoperative care
<ol>
	<li>Aspirate the water inside the tube and cover it with a plastic seal to prevent debris from entering.</li>
	<li>Keep the mouse warm and wait for recovery from the anesthesia.</li>
	<li>Administer meloxicam (2 mg/kg) subcutaneously every 24 h as long as the mouse exhibits pain-related behavior.</li>
	<li>Raise the postoperative mouse in individual housing for several weeks.</li>
</ol>
6.&#160;In vivo chronic two-photon imaging of microglia using CX3CR1-GFP mice
<ol>
	<li>After induction of anesthesia, wash the inside of the metal tube with sterile saline to remove debris. Ensure that the white alveus of the hippocampus is visible through the glass (Figure 3A) and there are no complications such as postoperative bleeding.</li>
	<li>Set the mouse under the two-photon microscope following steps 4.1 to 4.6.</li>
	<li>Check the quality and intensity of images obtained by two-photon excitation of the hippocampus (Figure 3B). Ensure that the images taken after the reduction of postoperative edema are comparable to or better than those taken on the day of the surgery (Figure 2A, see step 4.5). 
	NOTE: Image deterioration indicates the presence of tissue damage inside the brain.</li>
	<li>Ensure that microglia have already recovered their ramified morphology (Figure 3C). 
	NOTE: The imaging data of microglia indicates that at least 3-4 weeks are required for microglia to return to their basal state.</li>
	<li>Perform in vivo imaging in any layers of the CA1 for the particular purpose of the experiment.</li>
</ol>

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None of the authors have any conflicts of interest.

This method contains elaborate surgical procedures, but it can provide high-resolution images over the entire CA1 for an extended period if prepared properly. Experiments with several mice confirmed that the image quality in the chronic postoperative phase is better than that of acute postoperative imaging. The better image quality was maintained for more than 2 months. The most common cause of failure is unintentional damage to the CA1 during surgery, which is identified in the postoperative quality check (see step 4). This can be due to direct injury by aspiration, drying of the brain, excessive pressure by the glass, or toxicity from the dental cement in the final step. Another cause of failure is postoperative bleeding, which can be confirmed through the glass-bottom window within 1 week after surgery (see step 6.1). Read the protocol carefully and take all precautions to avoid such troubles.
Even with the current setup of two-photon microscope with GaAsP detectors in the laboratory, the attempt to image the dorsal CA1 through the cortex results in a significant loss of resolution due to the non-negligible scattering of light. Therefore, to obtain sufficient resolution to observe the movement of microglial processes or the formation of dendritic spines of neurons in the CA1, removing a part of the overlying cortex is inevitable, as in the present method. The only alternative way to reduce the extent of cortical removal while maintaining high image resolution is to embed a relay lens, such as a gradient refractive index (GRIN) lens. In this approach, a GRIN lens has been placed inside a guide tube implanted directly above the CA1 for chronic CA1 imaging28,29. The outer diameter of the guide tube was 1.8 mm, which is smaller than the 3.0 mm of the device in this paper, while producing high resolution (NA; 0.82) comparable to the system here (effective NA; 0.88). However, the approach using a GRIN lens has a narrow field of view for high-resolution observation and a short imaging depth limited by the WD of the GRIN lens. Therefore, imaging all hippocampal layers in a wide field of view with high resolution is a specific advantage of the method in this paper.
A limitation of this procedure is that the partial removal of the cortex and inflammation may have some detrimental effects on the hippocampus. However, because the removed cortex has no direct input to the hippocampus, the entorhinal cortex is not damaged, and the hippocampus itself is not directly injured, the overall evaluation is that the functional impairment of the hippocampus is not significant. Several previous studies have confirmed that hippocampal functions, including learning and memory, are preserved after hippocampal window implantation25,26,27,30,31,32,33,34. Therefore, the imaging approach described here is expected to faithfully report hippocampal functions after a sufficient postoperative period.
Future directions of research subjects based on this imaging technique may include synaptic pruning detected by simultaneous imaging of microglia and neurons5, learning-related microglial interaction with synapses35, microglial motility regulated by the hippocampal neural activity36, and microglial responses to peripheral inflammation or tissue damage7. This method will also help elucidate the progression of neurodegenerative diseases. For example, in Alzheimer&#39;s disease (AD), amyloid &#946; and tau proteins accumulate in parallel with neuronal cell deaths and hippocampal atrophy, leading to cognitive dysfunction; microglia have been reported to be involved in this process37,38. In vivo chronic imaging of microglia and neurons using the AD mouse models will enable us to monitor the correlation between microglial functions and neuronal pathology in the hippocampus of the AD mouse models.
This paper focuses on microglial imaging, but the same imaging procedure is applicable to the other neuronal and glial cell types in the CA1. In addition, the protocol is limited to the imaging of anesthetized mice, but the technique can also be extended to monitoring hippocampal microglia in awake mice. The motion artifacts induced by respiration, heartbeats, and body movements, especially in the deep hippocampal layers away from the glass window, may exist in experiments with awake mice. Therefore, an appropriate motion correction system may be required to capture microglial morphology and dynamics.
Discussion related to the Protocol 
It is advisable to select mice of appropriate age and genetic modifications for the expression of fluorescent probes with temporal and spatial specificity (see step 1.3). Mice at 1 month of age can be used for experiments, but mice older than 2 months are easier to handle, with their larger body size and resistance to surgical stress. In addition, the external capsule and the alveus of the hippocampus are more difficult to separate in younger mice. Therefore, removing the external capsule in young mice requires surgical skills (see steps 3.4.3-4). Assessment of the surgery and subsequent imaging will be more straightforward with mice expressing fluorescent proteins reproducibly and uniformly in the CA1 and the DG (see step 4.7). Hence, in the initial trials of the surgery, it is advisable to use transgenic mice with sparsely labeled neurons, such as Thy1-YFP or Thy1-GFP mice39, or mice with labeled microglia, such as CX3CR1-GFP mice24.
For successful surgery, the choice of anesthesia is important (see step 2.1). Different types of anesthesia can cause various degrees of intraoperative brain edema, which affect the difficulty of the surgery, the relative position of the implanted metal tube, and the extent of brain compression by the glass window. These factors may also influence the survival of hippocampal neurons after surgery.
In the aspiration process to expose the hippocampus (see step 3.4), the following points should be noted to minimize the damage to the brain and ensure successful imaging. Constantly supply sterile saline onto the tissue surface by positioning an outlet at the side of the cranial window. Prevent the tissue surface from exposure to air during aspiration. The basic principle of tissue aspiration is removing the tissue surface by a liquid flow into the suction tip. Direct contact of the suction tip to the tissue should be avoided. Adjust the negative pressure applied to the suction tube to be minimum. Aspirate the tissue step by step and confirm that bleeding is controlled in each step. When bleeding is prolonged, wait until the bleeding stops spontaneously. Keep aspiration from a short distance from the bleeding spot with a constant flow of sterile saline. Aspirate the tissue to create a cylindrical space with its size fitting to the glass-bottom metal tube (see step 1.1). Choose 23 G needles to aspirate thick pial vessels or large tissue fragments and 25 G needles to suck small tissue debris.

Microglia are the only immune cells and tissue macrophages resident in the brain. In addition to their functions as immune cells, such as rapid response to inflammation, they have been shown to play a variety of physiological roles in the maintenance and remodeling of neural circuits, such as synaptic pruning, regulation of synaptic plasticity, and control of neuronal activity1,2,3,4,5. Physiological interactions between microglia and neurons are of increasing interest in both neural circuit development and disease neurobiology. Analyzing the physiology of microglia in vivo requires methods to observe microglia without tissue damage and inflammatory responses. However, microglia are sensitive to inflammation and dramatically change their shapes and functions in response to tissue damage6,7.
Two-photon in vivo imaging is an ideal tool for capturing the physiological dynamics of microglia8. Initial applications of in vivo two-photon imaging revealed the dynamic movement of microglial processes for environmental surveillance in the adult mouse cortex9,10. The following two-photon imaging studies further extended the in vivo monitoring of microglia for the analysis of functional and structural interactions with neurons5,11,12,13,14. However, the imaging depth of conventional two-photon microscopy has been limited to less than 1 mm from the brain surface15,16. This constraint explains the scarcity of previous studies reporting the behavior of microglia in deep brain regions, such as the hippocampus.
Recent RNA sequencing studies have revealed a considerable regional heterogeneity of microglia, raising the possibility of distinct functional roles17,18,19,20,21. Therefore, it is necessary to record microglial interactions with neurons in various brain regions, but technical difficulties have hampered such studies. In particular, microglial active involvement in synapse remodeling has been reported to be critical in the development of the hippocampal neural network and its memory-related functions22,23. However, effective methods for observing unchallenged hippocampal microglia in vivo have not been available.
This protocol explains the procedures for the chronic in vivo observation of resting microglia in the CA1 of the dorsal hippocampus using precisely controlled surgical techniques. Two-photon imaging of the CA1 area in CX3CR1-GFP mice (CX3CR1+/GFP), which express GFP in microglia24, enabled observation of the ramified microglia in all layers of the CA1 with sufficient resolution. The protocol includes several tips for successful implantation of the observation window and appropriate imaging conditions. In addition, simultaneous visualization of pyramidal neurons and microglia in the CA1 is presented as an application example. The advantages, limitations, and potentials of this technique are also discussed.

<table><tbody><tr><td>23 G blunt needles</td><td>NIPRO</td><td>02-166</td><td>Suction tips for aspiration.</td></tr><tr><td>25 G blunt needles</td><td>NIPRO</td><td>02-167</td><td>Suction tips for aspiration.</td></tr><tr><td>3 mm dermal punches (DermaPunch)</td><td>Maruho</td><td>213001610</td><td>Tools for craniotomy.</td></tr><tr><td>30 G needles</td><td>Dentronics</td><td>Disposable needle No. 30</td><td>Tools for craniotomy.</td></tr><tr><td>A femtosecond pulsed laser</td><td>Spectra-Physics</td><td>MaiTai Deep See</td><td>A Ti:Sapphire laser used at 920 nm wavelength.</td></tr><tr><td>A two-photon microscope</td><td>Nikon</td><td>A1R MP+</td><td>Microscope for the CA1 imaging.</td></tr><tr><td>AAV1-CAG-FLEX-tdTomato</td><td>Penn Vector Core</td><td></td><td>AAV for neuronal labeling.</td></tr><tr><td>AAV1-hSyn-Cre</td><td>Penn Vector Core</td><td></td><td>AAV for neuronal labeling.</td></tr><tr><td>An objective lens for two-photon imaging</td><td>Olympus</td><td>XLPLN25XSVMP</td><td>25&amp;times; objective with a long working distance, a high numerical aperture, and a correction collar</td></tr><tr><td>Aspirators</td><td>Shin-ei Industries</td><td>KS-500</td><td>Tools for aspiration.</td></tr><tr><td>Aluminum plates</td><td>Narishige</td><td>CP-1</td><td>Plates made of alminum for head fixation.</td></tr><tr><td>Chemical depilatory cream</td><td>YANAGIYA</td><td></td><td>Cream for hair removal around the surgical site.</td></tr><tr><td>Circular glass coverslips</td><td>Matsunami Glass</td><td>3&amp;phi; No.1</td><td>Coverslips bonded to stainless steel tubes.</td></tr><tr><td>CX3CR1-GFP mice</td><td>The Jackson Laboratory</td><td>008451</td><td>Transgenic mice for microglial imaging.</td></tr><tr><td>Cylindrical stainless steel tubes</td><td>MORISHITA</td><td>Custom-made</td><td>Metal tubes to be implanted. Outer diameter 3.0 mm, inner diameter 2.8 mm, height 1.7 mm.</td></tr><tr><td>Dental etching material</td><td>Sun Medical</td><td>204610461</td><td>Used to etch the skull bones.</td></tr><tr><td>Dental resin cement (Super-Bond C&amp;amp;B)</td><td>Sun Medical</td><td>204610555</td><td>Dental cement.</td></tr><tr><td>Dura pickers (Micro Points)</td><td>Fine Science Tools</td><td>10063-15</td><td>Tools for dura removal.</td></tr><tr><td>Fine forceps (Dumont #5)</td><td>Fine Science Tools</td><td>11252-20</td><td>Surgical tools.</td></tr><tr><td>Forceps</td><td>Bio Research Center</td><td>PRI13-3374</td><td>Used to disinfect the surgical site.</td></tr><tr><td>Head holding device</td><td>Narishige</td><td>MAG-2</td><td>The head holding device of mice with angle adjusters</td></tr><tr><td>Heating pad</td><td>Bio Research Center</td><td>BWT-100A and HB-10</td><td>Tools to provide thermal support for the mouse during surgery.</td></tr><tr><td>Heating pad</td><td>ALA Scientific</td><td>HEATINGPAD-1 and Hot-1</td><td>Tools to provide thermal support for the mouse on the head holding device.</td></tr><tr><td>Ketamine (Ketalar)</td><td>Daiichi Sankyo Company</td><td>S9-001665</td><td>Anesthesia during surgery.</td></tr><tr><td>Meloxicam</td><td>Tokyo Chemical Industry</td><td>M1959</td><td>Analgesia during surgery.</td></tr><tr><td>Ophthalmic ointment</td><td>Sato Pharmaceutical</td><td></td><td>Used to prevent eye dryness during surgery.</td></tr><tr><td>Povidone-iodine scrub solution</td><td>Meiji Seika Pharma</td><td>2612701Q1137</td><td>Used to disinfect the surgical site.</td></tr><tr><td>Round-tipped miniature knives</td><td>Surgistar</td><td>4769</td><td>Surgical tools.</td></tr><tr><td>Scalpel</td><td>MURANAKA MEDICAL INSTRUMENTS</td><td>450-098-67</td><td>Disposable surgical tool.</td></tr><tr><td>Stereo microscopes</td><td>Leica Microsystems</td><td>S8 APO</td><td>Microscopes for surgery.</td></tr><tr><td>Sterile saline</td><td>Otsuka Pharmaceutical Factory</td><td>3311401A2026</td><td>Washing solution during surgery.</td></tr><tr><td>Sterile waterproof pad</td><td>AS ONE</td><td>8-5945-01</td><td>Surgical platform.</td></tr><tr><td>Sulforhodamine 101</td><td>Sigma-Aldrich</td><td>S7635</td><td>A dye for in vivo vessel imaging.</td></tr><tr><td>Surgical drape</td><td>Medline Industries</td><td>MP-0606F6T</td><td>Perforated drape for surgery.</td></tr><tr><td>UV light</td><td>Toshiba</td><td>GL15</td><td>Used to cure the adhesives.</td></tr><tr><td>UV-curing optical adhesives</td><td>Thorlabs</td><td>NOA81</td><td>Adhesives for bonding coverslips to stainless steel tubes.</td></tr><tr><td>Xylazine (Celactal 2%)</td><td>Bayer</td><td></td><td>Anesthesia during surgery.</td></tr></tbody></table>

in vivo chronic two-photon imaging of microglia in the mouse hippocampus

Microglia, the only immune cells resident in the brain, actively participate in neural circuit maintenance by modifying synapses and neuronal excitability. Recent studies have revealed the differential gene expression and functional heterogeneity of microglia in different brain regions. The unique functions of the hippocampal neural network in learning and memory may be associated with the active roles of microglia in synapse remodeling. However, inflammatory responses induced by surgical procedures have been problematic in the two-photon microscopic analysis of hippocampal microglia. Here, a method is presented that enables the chronic observation of microglia in all layers of the hippocampal CA1 through an imaging window. This method allows the analysis of morphological changes in microglial processes for more than 1 month. Long-term and high-resolution imaging of the resting microglia requires minimally invasive surgical procedures, appropriate objective lens selection, and optimized imaging techniques. The transient inflammatory response of hippocampal microglia may prevent imaging immediately after surgery, but the microglia restore their quiescent morphology within a few weeks. Furthermore, imaging neurons simultaneously with microglia allows us to analyze the interactions of multiple cell types in the hippocampus. This technique may provide essential information about microglial function in the hippocampus.

Using this technique, the ramified and quiescent microglia can be chronically observed within all layers of the dorsal CA1, including stratum oriens (SO), stratum pyramidale (SP), stratum radiatum (SR), and stratum lacunosum-moleculare (SLM), especially 3-4 weeks after surgery when the inflammation subsides. If the surgery is performed appropriately, imaging can be performed up to several months after surgery. This section contains three topics useful for the evaluation of intravital imaging. First, sample images immediately following surgery and in the chronic phase are provided as references for appropriate surgical and imaging procedures. Second, the distinct morphology of microglia, their fluorescence intensity, and blood vessel distribution in specific hippocampal layers are described to help readers estimate the imaging depth ranging from the alveus to the DG. Third, an application example is provided that illustrates simultaneous imaging of neurons and microglia.
The representative images of microglia in CX3CR1-GFP mice immediately after surgery are shown in Figure 2. If there is no apparent damage to the CA1 and the curvature of the CA1 layers is properly flattened by the overlying coverslip, two-photon imaging depths of microglia exceed the entire CA1 layers and reach 500 &#181;m from the surgical surface, where the ML or the granule cell layer (GCL) of the DG exist (Figure 2A;&#160;see step 4.7 and Figure 4E). Microglia are slightly less ramified, especially in the layer close to the surgical surface in comparison with those in the intact hippocampus. Nevertheless, they exhibit no prominent activation, suggesting proper surgical procedures (Figure 2B). On the other hand, edema in the CA1 induced by tissue damage will limit the imaging depth to 200 &#181;m (Figure 2C). Tissue damage also induces abnormal microglial activation, such as accumulation of bulging processes toward the free tissue surface (Figure 2D; compare to the upper image of Figure 2B). These are signs of inappropriate surgery.
The representative images of microglia in the chronic phase more than 3 weeks after surgery are shown in Figure 3. Microglia can be imaged again beyond the CA1 to the deeper layers of the DG with higher fluorescent intensity and resolution and with more homogenous distribution (see step 6.3, Figure 3B). Imaging quality is better than that immediately after the surgery (Figure 2A). This difference may be explained by reduced edema and inflammation. Microglia in all layers have already restored their ramified morphology (Figure 3C), different from their postoperative appearance (Figure 2B). Time-lapse imaging of microglia in the CA1 reveals the immobile cell bodies and highly motile processes for surveillance (Video 1-2). Their motile behavior is similar to the previous report of microglial process dynamics in the cortex9.
It is straightforward to specify the imaged hippocampal layers based on the distribution and the depth of neuronal cell bodies, using mice expressing fluorescent probes in the hippocampal neurons25,26,27. Without the aid of the positional information about the neuron cell bodies, the signals from fluorescent microglia alone provide some clues to the identification of hippocampal layers (Figure 3B). Microglia in the alveus have elongated cell bodies and processes which tend to extend along the axonal tracts (Figure 4A). Microglia in other layers can be distinguished by their round cell bodies and processes radiating without preferred directions. Microglia in SP are among the densely packed pyramidal neurons and can be distinguished by the lower density of microglial processes. The layers above and below SP are identified as SO and SR, respectively (Figure 4B). It is impossible to distinguish between SR and SLM based on microglial properties alone. The border between the CA1 and the DG is recognized by thick blood vessels running along the boundary (Figure 4C). These vessels can be better recognized by labeling vessels with dyes such as sulforhodamine 101 (SR101; 5 mM, 4 &#181;L/g body weight) injected intraperitoneally (Figure 4D). The fluorescence signal from microglia drops sharply in the DG compared to the overlying CA1, probably due to the difference in the refractive index of these structures. This gap in fluorescence intensity may also help to specify the border between the DG and CA1. As an application example, simultaneous imaging of GFP-positive microglia and tdTomato-labeled pyramidal neurons is shown (Figure 4E). In this experiment, CX3CR1-GFP mice received an injection of adeno-associated virus (AAV) for the expression of tdTomato in the hippocampal pyramidal neurons. Neuronal labeling will help to understand the exact layer structures of the CA1.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64104/64104fig01.jpg" /> 
Figure 1: Schematic drawings for the implantation of the observation window. (A) Cross-sectional view of the assembled glass-bottom metal tube. A circular glass coverslip adheres to the one end of the cylindrical stainless tube. (B) Cross-sectional view of the aluminum plate attached to the skull. The plate should be parallel to the marked surface of the skull so as not to interfere with the objective lens, positioned close to the plate for focusing. (C) Cross-sectional view of the implanted tube. The glass bottom should be closely attached to the alveus of the hippocampus to gently press and flatten the surface of the CA1. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64104/64104fig01large.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/64104/64104fig02.jpg" /> 
Figure 2: In vivo imaging of microglia in CX3CR1-GFP mice immediately after surgery. (A) Representative 3D reconstruction from the in vivo CA1 imaging of a CX3CR1-GFP mouse on postoperative day (POD) 0 (width: 511 &#181;m, height: 511 &#181;m, depth: 550 &#181;m). The fluorescent microglia in the ML of the DG at a depth of 500 &#181;m from the glass bottom can be imaged through the entire CA1. (B) Typical GFP-filled microglia obtained in (A) are shown as the maximum intensity projections (MIPs) of image stacks with 20 &#181;m thickness at depths of 100, 300, and 520 &#181;m from the glass bottom. Microglial morphology is detectable from the superficial CA1 to the ML of the DG, though with apparent image deterioration in the DG. Microglia especially in the layer close to the surgical surface are less ramified but exhibit no obvious activation. Scale bars, 50 &#181;m. (C) Representative 3D image reconstruction of the surgically damaged CA1. The data was acquired from a CX3CR1-GFP mouse on POD 0 (width: 399 &#181;m, height: 399 &#181;m, depth: 450 &#181;m). Microglial fluorescence is detectable only up to the depth of 200 &#181;m, in contrast with the undamaged CA1 (A) with fluorescent microglia detected at a depth of 500 &#181;m. (D) The typical image of abnormally activated microglia in (C). The MIP of GFP image stacks with 20 &#181;m thickness at a depth of 60 &#181;m shows microglial bulging processes extending toward the surgically exposed tissue surface. The overall microglial morphology is different from the image shown in (B). Scale bar, 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64104/64104fig02large.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/64104/64104fig03.jpg" /> 
Figure 3: In vivo imaging of microglia in CX3CR1-GFP mice in the chronic phase. (A) The representative appearance of the implanted window on the right dorsal CA1 in the chronic phase. Through the glass bottom, the white fibers of the alveus are clearly observed without postoperative bleeding. The DHC and the LV are partly visible in the caudomedial area and in the rostrolateral edge, respectively. Scale bar, 1 mm. (B) Representative 3D reconstruction from the in vivo chronic CA1 imaging of a CX3CR1-GFP mouse on POD 24 (width: 511 &#181;m, height: 511 &#181;m, depth: 670 &#181;m). Compared to the images immediately after surgery (Figure 2A), microglia show a more homogeneous distribution and can be observed more clearly in the deeper layers of the DG because of the reduction of edema and inflammation. (C) Typical images of microglia from (B) with fully restored ramified morphology are shown as the MIPs of GFP image stacks with 20 &#181;m thickness at depths of 100, 280, and 500 &#181;m from the surgical surface. Scale bars, 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64104/64104fig03large.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/64104/64104fig04.jpg" /> 
Figure 4: Clues to identifying each layer of the hippocampus during in vivo imaging and an application example of this method. (A) The typical image of microglia in the alveus of CX3CR1-GFP mice in the chronic phase. Microglial cell bodies and processes extending along the axonal tracts are shown in the MIP of GFP image stacks with 10 &#181;m thickness at a depth of 15 &#181;m. Scale bar, 50 &#181;m. (B) Representative images of microglia in SO, SP, and SR in the chronic phase shown as the MIP of GFP image stacks with 5 &#181;m thickness. The local density of microglial processes is lower in SP than in surrounding SO or SR due to the densely packed pyramidal neurons in SP. Scale bars, 100 &#181;m. (C) The thick blood vessels running along the boundary between the CA1 and the DG can be recognized only by the microglial fluorescence. The average intensity projection of tiled GFP image stacks with 90 &#181;m thickness in the chronic phase at a depth of about 500 &#181;m is shown. The void of the GFP signal corresponds to the thick vessels. Scale bar, 500 &#181;m. (D) (Upper) The image of the same field of view as C after intraperitoneal injection of SR101. Scale bar, 500 &#181;m. (Lower) 3D reconstruction of the tiling images after SR101 injection (width: 2.60 mm, height: 1.73 mm, depth: 0.69 mm). The thick vessels running along the border between the CA1 and the DG can be easily recognized by the intravascular SR101 fluorescence. (E) (Left) 3D reconstruction of the CA1 and the DG (width: 255 &#181;m, height: 255 &#181;m, depth: 730 &#181;m) and (right) the MIP of GFP and tdTomato fluorescence in SP made from image stacks with 20 &#181;m thickness. At 1 week before surgery, 1.5 &#181;L of a mixture of AAV1-CAG-FLEX-tdTomato (5.0 x 1012 vg/mL) and AAV1-hSyn-Cre (1.0 x 109 vg/mL) was injected into the hippocampus of a CX3CR1-GFP mouse to label the neurons sparsely. Imaging was performed on POD 0. SP and GCL are easily recognized by the position of neuronal cell bodies. Scale bar, 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64104/64104fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Time-lapse imaging of SO microglia in the chronic phase. The MIP of GFP image stacks with 20 &#181;m thickness in the time-lapse imaging of SO microglia, animated so that 28 min play in 1 sec. Scale bar, 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64104/Video1.zip" target="_blank">Please click here to download this Video.</a>
Video 2: Time-lapse imaging of SR microglia in the chronic phase. The MIP of GFP image stacks with 20 &#181;m thickness in the time-lapse imaging of SR microglia, animated so that 28 min play in 1 sec. Scale bar, 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64104/Video2.zip" target="_blank">Please click here to download this Video.</a>

Watch this Scientific Journal Video about In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus at JoVE.com

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

This paper describes a method for chronic in vivo observation of the resting microglia in the mouse hippocampal CA1 using precisely controlled surgery and two-photon microscopy.

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

Microglia, the only immune cells resident in the brain, actively participate in neural circuit maintenance by modifying synapses and neuronal ...

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Nanyang Technological University

Research

JoVE Journal

Neuroscience

3.8K Views.  The University of Tokyo. This paper describes a method for chronic in vivo observation of the resting microglia in the mouse hippocampal CA1 using precisely controlled surgery and two-photon microscopy.

Video: In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or thinned-skull 3 preparations. While the open-skull technique is very versatile, it is not optimal for studying microglia because it is invasive and can cause microglial activation. Even though the thinned-skull approach is minimally invasive, the repeated re-thinning of skull required for chronic imaging increases the risks of tissue injury and microglial activation and allows for a limited number of imaging sessions. Here we present a chronic thin-skull window method for monitoring murine microglia in vivo over an extended period of time using two-photon microscopy. We demonstrate how to prepare a stable, accessible, thinned-skull cortical window (TSCW) with an apposed glass coverslip that remains translucent over the course of three weeks of intermittent observation. This TSCW preparation is far more immunologically inert with respect to microglial activation than open craniotomy or repeated skull thinning and allows an arbitrary number of imaging sessions during a time period of weeks. We prepare TSCW in CX3CR1 GFP/+ mice 4 to visualize microglia with enhanced green fluorescent protein to &le;150 &mu;m beneath the pial surface. We also show that this preparation can be used in conjunction with stereotactic brain injections of the HIV-1 neurotoxic protein Tat, adjacent to the TSCW, which is capable of inducing durable microgliosis. Therefore, this method is extremely useful for examining changes in microglial morphology and motility over time in the living brain in models of HIV Associated Neurocognitive Disorder (HAND) and other neurodegenerative diseases with a neuroinflammatory component.

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

We describe a method for repeatedly visualizing murine microglia and circulating monocytes in vivo over hours, days or weeks using transcranial two-photon microscopy. We demonstrate how to prepare a thinned-skull window that allows intermittent observation of quiescent microglia that can be activated by adjacent stereotactic injection of the HIV-1 regulatory protein Tat.

Traditionally in neuroscience, in vivo two photon imaging of the murine central nervous system has either involved the use of open-skull1,2 or ...

Chronic Imaging of Mouse Visual Cortex Using a Thinned-skull Preparation

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The technique presented here allows the imaging of the same area of the brain at several time points (chronic imaging) with microscopic resolution allowing the tracking of dendritic spines which are the small structures that represent the majority of postsynaptic excitatory sites in the CNS. The ability to clearly resolve fine cortical structures over several time points has many advantages, specifically in the study of brain plasticity in which morphological changes at synapses and circuit remodeling may help explain underlying mechanisms. In this video and supplementary material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation. The thinned-skull preparation is a minimally invasive approach, which avoids potential damage to the dura and/or cortex, thus reducing the onset of an inflammatory response. When this protocol is performed correctly, it is possible to clearly monitor changes in dendritic spine characteristics in the intact brain over a prolonged period of time.

In this video and supplemental material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation.

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The ...

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution . The method consists in implanting a cranial imaging window, injecting a fluorescent calcium indicator dye that can be taken up by large numbers of neurons and finally recording the activity of neurons with time lapse calcium imaging using an in-vivo two photon microscope. Co-injection of astrocyte-specific dyes allows one to differentiate neurons from astrocytes. The technique can be performed in mice expressing fluorescent molecules in specific subpopulations of neurons to better understand the network interactions of different groups of cells.

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution .

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons .  ...

Biology

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Longitudinal Two-Photon Imaging of Dorsal Hippocampal CA1 in Live Mice

Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from subcellular to network levels and at temporal scales from milliseconds to weeks. In addition, two-photon imaging can be combined with a variety of behavioral tasks to explore the causal relationships between brain function and behavior. However, in mammals, limited penetration and scattering of light have limited two-photon intravital imaging mostly to superficial brain regions, thus precluding longitudinal investigation of deep-brain areas such as the hippocampus. The hippocampus is involved in spatial navigation and episodic memory and is a long-standing model used to study cellular as well as cognitive processes important for learning and recall, both in health and disease. Here, a preparation that enables chronic optical access to the dorsal hippocampus in living mice is detailed. This preparation can be combined with two-photon optical imaging at cellular and subcellular resolution in head fixed, anesthetized live mice over several weeks. These techniques enable repeated imaging of neuronal structure or activity-evoked plasticity in tens to hundreds of neurons in the dorsal hippocampal CA1. Furthermore, this chronic preparation can be used in combination with other techniques such as micro-endoscopy, head-mounted wide field microscopy or three-photon microscopy, thus greatly expanding the toolbox to study cellular and network processes involved in learning and memory.

This method describes a chronic preparation that allows optical access to the hippocampus of living mice. This preparation can be used to perform longitudinal optical imaging of neuronal structural plasticity and activity-evoked cellular plasticity over a period of several weeks.

Two-photon microscopy is a fundamental tool for neuroscience as it permits investigation of the brain of live animals at spatial scales ranging from ...

Behavior

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. Although studying these cells in isolation in vitro or together ex vivo provides useful physiological information; salient features of neural cell physiology will be missed in such contexts. Therefore, there is a need for studying neural cells in their native in vivo environment. The protocol detailed here describes repetitive in vivo two-photon imaging of neural cells in the rodent cortex as a tool to visualize and study specific cells over extended periods of time from hours to months. We describe in detail the use of the grossly stable brain vasculature as a coarse map or fluorescently labeled dendrites as a fine map of select brain regions of interest. Using these maps as a visual key, we show how neural cells can be precisely relocated for subsequent repetitive in vivo imaging. Using examples of in vivo imaging of fluorescently-labeled microglia, neurons, and NG2+ cells, this protocol demonstrates the ability of this technique to allow repetitive visualization of cellular dynamics in the same brain location over extended time periods, that can further aid in understanding the structural and functional responses of these cells in normal physiology or following pathological insults. Where necessary, this approach can be coupled to functional imaging of neural cells, e.g., with calcium imaging. This approach is especially a powerful technique to visualize the physical interaction between different cell types of the CNS in vivo when genetic mouse models or specific dyes with distinct fluorescent tags to label the cells of interest are available.

This protocol describes a chronic cranial window implantation technique that can be used for longitudinal imaging of neuro-glio-vascular structures, interactions, and function in both healthy and diseased conditions. It serves as a complementary alternative to the transcranial imaging approach that, while often preferred, possesses some critical limitations.

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. ...

Immunology and Infection

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.

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 ...

In Vivo Chronic Two-Photon Imaging of Microglia in the Mouse Hippocampus

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Rozdziały w tym wideo

A Thin-skull Window Technique for Chronic Two-photon In vivo Imaging of Murine Microglia in Models of Neuroinflammation

Chronic Imaging of Mouse Visual Cortex Using a Thinned-skull Preparation

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Longitudinal Two-Photon Imaging of Dorsal Hippocampal CA1 in Live Mice

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice