We thank James Perna for the graphic illustration. This work was supported by grants from the National Institute of Mental Health to Y.Z.

Approval needs to be obtained from home institutions before commencement of the surgery and imaging study. Experiments described in this manuscript were performed in accordance with the guidelines and regulations from the University of California, Santa Cruz Institutional Animal Care and Use Committee.
1. Surgery
<ol>
	<li>Autoclave all surgical instruments and sterilize the workspace with 70% alcohol thoroughly before surgery.</li>
	<li>Anesthetize the mouse by intraperitoneal (IP) injection of KX anesthetic solution (200 mg/kg ketamine and 20 mg/kg xylazine) according to the mouse&#8217;s body weight.&#160;Note: KX dosage may be adjusted according to the strain, age, and health status of the mice. Veterinary consultation to determine the optimal dose is also recommended.</li>
	<li>Perform a toe-pinch test periodically by pressing the mouse&#8217;s toes and checking for a reflexive response to monitor the anesthesia status. Make sure that the mouse is fully anesthetized before starting the surgery. Note:&#160;During prolonged imaging sessions, check the mouse&#8217;s anesthesia status periodically, administrate additional KX if necessary.</li>
	<li>Place the mouse on a heating pad to maintain the body temperature during the surgery.</li>
	<li>Shave the head of the mouse using a razor blade to expose the scalp.</li>
	<li>Sterilize the shaved area by wiping the skin with alternating alcohol pads and betadine.&#160;Remove any residual hair clippings.</li>
	<li>Gently apply eye ointment to lubricate both eyes, therefore preventing permanent damage caused by dehydration during the experiment.</li>
	<li>Make a straight incision along the midline of the scalp, and move the skin laterally towards the edges of the skull.</li>
	<li>Remove the connective tissue attached to the skull.</li>
</ol>
2. Thinned-skull Preparation
<ol>
	<li>Identify the imaging region based on stereotaxic coordinates. Note:&#160;Try to avoid large blood vessels, which block light penetration and blur the imaged structures. Vasculature is best observed when the skull is moist with sterile saline.</li>
	<li>Use the high-speed micro&#160;drill to thin a circular region of skull (0.5-1.0 mm diameter). Move the drill parallel to the skull surface, rather than holding it against the skull and pressing down. Drill until both the outer compact bone layer and the middle spongy bone layer are removed. Note:&#160;To prevent damage caused by overheating, avoid prolonged contact between the drill bit and the skull.</li>
	<li>Continue thinning the inner compact bone layer with a microsurgical blade in the center of the drill-thinned region. Hold the microsurgical blade at an angle of approximately 45&#176; to scrape the skull without pressing down against the skull until an evenly thinned small region (200-300 &#956;m diameter) with a thickness of approximately 20-30 &#956;m is obtained. Due to the auto-fluorescence of the skull, the thickness can be measured by scanning the distance between upper and lower surface of the skull under the two-photon microscope. Note: although a skull thickness of less than 20 &#956;m provides good imaging quality, it is not recommended because it makes the thinning process in subsequent re-imaging sessions difficult. To avoid over-thinning, the thickness of the skull needs to be checked periodically during the surgery (see Discussion).</li>
</ol>
3. Immobilization
<ol>
	<li>Carefully remove bone debris.</li>
	<li>Place a small drop of cyanoacrylate glue on each edge of the center opening of the sterile head plate, which has been autoclaved&#160;(see Figure 1). Hold the head plate tightly against the skull with the thinned region located in the center of the opening. Note:&#160;If the glue contaminates the thinned region by accident, remove it carefully with the microsurgical blade.</li>
	<li>Gently pull the skin from the sides of the skull to the edges of the center opening of the head plate.</li>
	<li>Wait approximately 10 min until the head plate is well attached to the skull.</li>
	<li>Place the head plate on two lateral blocks of the holding plate and then tighten the screws over the edges of the head plate to immobilize the mouse on the holding plate (see Figure 1). Note:&#160;Make sure that there is no skin or whiskers between the head plate and the blocks before tightening the screws. A good immobilized preparation should show no observable movement of the skull under the dissecting microscope when the back of the animal is gently patted.</li>
	<li>Rinse the exposed skull with saline to remove unpolymerized glue. Note:&#160;It is important to remove any remaining glue, as unpolymerized glue blurs images and damages microscope objectives.</li>
</ol>
4. Imaging
<ol>
	<li>Take a photo of the vasculature of the exposed skull with the thinned region as Map 1, which is used to relocate the imaged region in subsequent imaging sessions (see Figure 2).</li>
	<li>Place the mouse under the imaging microscope. Locate the thinned area under a 10X air objective using epifluorescence and move the thinnest area to the center of the view.</li>
	<li>Add a drop of saline on the top of the skull and switch to the 60X objective, which has been rinsed with sterile water. Select a region where individual dendritic spines are clearly visualized along dendrites. Identify and label the corresponding region on Map 1 by comparing vasculature between the epifluorescent view and the vasculature photo.</li>
	<li>Tune the two-photon laser wavelength according to the fluorophores. For instance, 920 nm for YFP; 890 nm for GFP; 1,000 nm for DsRed and tdTomato24.</li>
	<li>Acquire image stacks with 2 &#956;m steps along the z-axis using the 60X objective. This image stack covers an approximately 200 &#956;m x&#160;200 &#956;m area (512 x&#160;512 pixels) and is used as Map 2 for relocation during subsequent imaging sessions (see Figure 2).</li>
	<li>Acquire nine image stacks within Map 2 using 3X digital zoom. Each image stack covers an approximate area of 70 &#956;m x&#160;70 &#956;m (512 x&#160;512 pixels), with 0.7 &#956;m steps along the z-axis. Note:&#160;The intensity of the laser should be below 40 mW&#160;when measured at samples to minimize phototoxicity.</li>
</ol>
5. Recovery
<ol>
	<li>Following imaging, gently detach the head plate from the skull.</li>
	<li>Thoroughly clean the skull and the skin to remove all the remaining glue. Note:&#160;Any remaining glue on the skin will cause irritations and slow down skin healing while any remaining glue on the skull will cause erosion of the skull and angiogenesis in the newly grown bone layer, making subsequent relocation and re-imaging difficult.</li>
	<li>Rinse the skull and the skin with saline several times.</li>
	<li>Suture the scalp with sterile surgical suture.</li>
	<li>Keep the animal on a heating pad in a separate cage. Administer buprenorphine analgesic (0.1 mg/kg) subcutaneously to mitigate post-operative pain. Return the animal to the home cage after full recovery.&#160;Monitor the animal closely (check at least once daily) until the incision is healed and sutures are removed. Administer subsequent injections of buprenorphine analgesic if needed.</li>
</ol>
6. Reimaging
<ol>
	<li>Repeat Steps 1.1-1.8.</li>
	<li>Repeat Steps 3.1-3.6</li>
	<li>Locate previously imaged region by comparing the vasculature pattern to Map 1 and the dendritic branch pattern to Map 2.</li>
	<li>If reimaging is performed within 1 week, repeat Step 2.3 to remove the thin layer of newly grown bone on top of the thinned region using the microsurgical blade. If reimaging is performed after more than 1 week, repeat Steps both 2.2 and 2.3. Note:&#160;The newly grown bone layer consists of a less condensed structure compared to the original compact bone layer, leading to reduced image quality. Therefore, to acquire the same quality, it is necessary to thin the skull slightly more than the previous imaging session.</li>
	<li>Adjust the position and orientation to obtain image stacks that match previously taken image stacks under the two-photon microscope.</li>
	<li>Acquire images as in Step 4.6.</li>
	<li>Repeat Steps 5.1-5.5 after imaging.</li>
</ol>

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The authors declare that they have no competing financial interests.

To obtain a successful thinned-skull preparation, several steps in this protocol are crucial. 1) The thickness of the skull. The cranial bone has a sandwich structure, with two layers of high-density compact bone and a middle layer of low-density spongy bone. While the high-speed micro drill is suitable for removing the outer layers of compact bone and spongy bone, the microsurgical blade is ideal for thinning the inner layer of compact bone. As the thickness and stiffness of the skull increases during development, imaging of adult mice requires more bone to be removed in order to obtain images of good quality. The thinned area exhibits a transparent solid appearance and provides good imaging quality when the thickness of the skull is approximately 20-30 &#956;m. The skull thickness should be checked periodically during the thinning process as over-thinning makes the thinning process in subsequent reimaging sessions difficult. 2) The size of the thinned region. It is important to establish a stable thinned-skull configuration, where a cone-like cavity is achieved. The architecture of the thinned region with a proper size prevents causing damage to the cortex and helps thinning in subsequent re-imaging sessions. It is recommended to make the bottom area (approximately 200-300 &#956;m in diameter) of the thinned region smaller than the top opening (approximately 0.5-1.0 mm in diameter). 3) The stability of the images. A well-immobilized preparation helps to improve the image quality by reducing respiratory-induced movement artifacts. It is important to keep the skull dry and devoid of connective tissue and bone debris before the head plate is attached on to the skull. Extra glue can also be applied to fill the remaining gaps between the head plate and the skull. In addition, targeting an area away from big vasculature also minimizes the movements caused by blood pumping. &#160;
Investigation of the changes in the brain of living animals with high spatial and temporal resolution using two-photon microscopy requires generation of an optical window. In addition to thinned-skull preparation, fluorescently labeled structures can also be visualized by removing the skull and replacing it with a cover glass (usually 3-5 mm in diameter) that provides a clear imaging window10,13. While both thinned-skull and open-skull imaging protocols are applied broadly for in vivo imaging studies, each method has its own advantages and is suitable for different experimental designs. For instance, the open-skull preparation is useful for studies that investigate relatively large brain areas (e.g., 5 mm diameter) and require many imaging sessions with short intervals (i.e., days). Once the cranial window is successfully implanted, no additional surgery is required. However, a post-surgery period (usually approximately 1 month) is required before the first imaging session to ameliorate potential inflammatory responses caused by surgery. Re-imaging becomes impossible when the cranial window is blocked by regrowth of the bone and/or thickening of the meninges. In contrast, animals with a thinned-skull preparation can be imaged immediately after the initial surgery, making it more suitable for chronic imaging of younger animals (e.g., 2 weeks old). It is suitable for investigation with long imaging intervals (i.e. months to years), as the bone regrowth and dura thickening is not problematic. However, re-thinning of the skull is usually required for subsequent re-imaging sessions (even with 2 day intervals) due to the bone regrowth. Moreover, the newly grown bone layer consists of a less condensed structure compared to the original compact bone, greatly reducing the transparency of the thinned-skull region. Therefore, to acquire the same quality of images, it is necessary to completely remove the newly grown bone layer. And such attempts usually lead to the final thickness of the skull in the subsequent imaging thinner than the previous preparation. Since the skull thickness had been prepared to 20-30 &#956;m in the initial imaging session, leaving limited room for re-thinning, the total imaging times of thinned-skull preparation is usually less than 5 times. Recently, a new experimental approach named polished and reinforced thinned-skull (PoRTS) has been developed to combine both thinned- and open-skull preparations11,26,27.
So far, the majority of in vivo imaging in the cortex has been performed using transgenic mouse lines expressing EGFP or YFP under neuronal specific thy1 promoter. These transgenic mice express fluorescent proteins sparsely in a subset, but mixed population of cortical neurons25. Since many lines of evidence suggest that experience-dependent structural plasticity happens in selective cell types of well-defined circuits, it is important to label and image in a cell-type specific manner. To date, many approaches have been applied to achieve this goal. For example, pyramidal neurons in different cortical layers can be targeted by performing in utero electroporation at defined embryonic stages28,29. Similarly, injection of viral vectors engineered to express a fluorescent protein can also be used to label cells in different brain regions30. In addition, many lines of transgenic mice have been generated to drive the expression of Cre recombinase under cell type specific promoters31,32. Injection of viral vectors such as adeno-associated virus carrying fluorescent reporter gene following a floxed stop codon into these mice offers the possibility to target a specified class of neurons in a restricted region33. By combining these new labeling approaches with our thinned-skull preparation, changes in synaptic connections of cortical neurons can be investigated by two-photon in vivo imaging in a cell type- and/or circuit-specific manner.
With the increasing availability of transgenic animals with fluorescently labeled cell populations and rapid development of in vivo labeling techniques, similar procedures described here can also be applied to investigate other cell types (glial cells) and vasculature in the living brain. Combined with behavioral manipulations and disease models, two-photon in vivo imaging will greatly expand our understandings of the molecular, cellular, and circuit mechanisms underlying brain functions.

The mammalian cortex participates in many brain functions, from sensory perception and movement control to abstract information processing and cognition. Various cortical functions build upon different neural circuits, which are made up of different types of neurons communicating and exchanging information at individual synapses. The structure and function of synapses are consistently being modified in response to experiences and pathologies. In the mature brain, synaptic plasticity takes the form of both strength changes and addition/removal of synapses, playing important roles in formation and maintenance of a functional neural circuitry. Dendritic spines are the postsynaptic components of the majority of excitatory synapses in the mammalian brain. The constant turnover and morphological changes of spines are believed to serve as a good indicator of modifications in synaptic connections1-7.
Two-photon laser scanning microscopy offers deep penetration through thick, opaque preparations and low phototoxicity, which makes it suitable for live imaging in the intact brain8. In combination with fluorescent labeling, two-photon imaging provides a powerful tool to peek into the living brain and follow structural reorganization at individual synapses with high spatial and temporal resolution. Various methods have been used to prepare mice for live imaging9-13. Here, we describe a thinned-skull preparation of in vivo two-photon imaging to investigate the structural plasticity of postsynaptic dendritic spines in the mouse cortex. Using this approach, our recent studies have depicted a dynamic picture of dendritic spine changes in response to motor skill learning With increasing availability of transgenic animals with fluorescently labeled neuronal subsets and rapid development of in vivo labeling techniques, similar procedures described here can also be applied to investigate other cell types and cortical regions, combined with other manipulations, as well as used in disease models16-23.

<table><tbody><tr><td>Ketamine</td><td>Bioniche Pharma</td><td>67457-034-10</td><td>Mixed with xylazine for anesthesia</td></tr><tr><td>Xylazine</td><td>Lloyd laboratories</td><td>139-236</td><td>Mixed with ketamine for anesthesia</td></tr><tr><td>Saline</td><td>Hospira</td><td>0409-7983-09</td><td>0.9% NaCl for injection and imaging</td></tr><tr><td>Razor blades</td><td>Electron microscopy sciences</td><td>72000</td><td>Double-edge stainless steel razor blades</td></tr><tr><td>Alcohol pads</td><td>Fisher Scientific</td><td>06-669-62</td><td>Sterile alcohol prep pads</td></tr><tr><td>Eye ointment</td><td>Henry Schein</td><td>102-9470</td><td>Petrolatum ophthalmic ointment sterile ocular lubricant</td></tr><tr><td>High-speed micro drill</td><td>Fine Science Tools</td><td>18000-17</td><td>The high-speed micro drill is suitable for thinning the outer layer of compact bone and targeting a small area</td></tr><tr><td>Micro drill steel burrs</td><td>Fine Science Tools</td><td>19007-14</td><td>1.4 mm diameter</td></tr><tr><td>Microsurgical blade</td><td>Surgistar</td><td>6961</td><td>The microsurgical blade is suitable for thinning the inner layer of compact bone and middler layer of spongy bone</td></tr><tr><td>Cyanoacrylate glue</td><td>Fisher Scientific</td><td>NC9062131</td><td>Fix the head plate onto the skull</td></tr><tr><td>Suture</td><td>Havard Apparatus</td><td>510461</td><td>Non-absorbale, sterile silk suture, 6-0 monofilament</td></tr><tr><td>Dissecting microscope</td><td>Olympus</td><td></td><td>SZ61</td></tr><tr><td>CCD camera</td><td>Infinity</td><td></td><td></td></tr><tr><td>Two-photon microscope</td><td>Prairie Technologies</td><td></td><td>Ultima IV</td></tr><tr><td>10X objective</td><td>Olympus</td><td></td><td>NA 0.30, air</td></tr><tr><td>60X objective</td><td>Olympus</td><td></td><td>NA 1.1, IR permeable, water immersion</td></tr><tr><td>Ti-sapphire laser</td><td>Spectra-Physics</td><td></td><td>Mai Tai HP</td></tr></tbody></table>

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.

In YFP-H line mice25, yellow fluorescent protein expresses in a subset of layer V pyramidal neurons, which project their apical dendrites to the superficial layers in the cortex. Through the thinned-skull preparation, the fluorescently labeled dendritic segments can be repetitively imaged under two-photon microscope over various imaging intervals, ranging from hours to months. Here we show an example of a four-time imaging of the same dendrites over 8 days in the motor cortex of a 1&#160;month old mouse, where individual spines as well as filopodia can be clearly visualized along the dendrite. Usually, the depth of image stacks is approximately 100-200 &#956;m from the pial surface. Various analyses can be performed based on these images. For instance, the spine formation, elimination and turnover can be quantified by comparing images from different sessions. Spine density can be calculated by dividing the number of spines by the length of the dendritic segment. Changes of spine motility and morphology can also be analyzed.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51520/51520fig1highres.jpg" src="/files/ftp_upload/51520/51520fig1.jpg" /> 
Figure 1.&#160;Custom-made immobilization plates of thinned-skull preparation for two-photon in vivo imaging. (A) A photograph of the head plate, which is made of two or three razor blades glued together, with sharp edges covered by tapes. (B) A photograph of the holding plate, which consists of 1 stainless steel plate, 2 stainless steel blocks, 2 screws, and 2 spacers.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51520/51520fig2highres.jpg" width="500" /> 
Figure 2.&#160;Transcranial two-photon imaging through a thinned-skull preparation in the mouse motor cortex, showing dynamics of dendritic spines over eight days. (A) A CCD image of the vasculature pattern with the thinned skull area (Map 1). The black box indicates the region where two-photon in vivo images were acquired. (B) A low-magnification maximum projection of dendritic branches in the motor cortex of a 1&#160;month old mouse (Map 2). (C) Repetitive images of the same dendritic segment reveal newly formed spines (arrowheads), eliminated spines (arrows), and filopodia (stars) on day 0, 2, 4, and 8. The left panel is a higher-magnification view of the dendritic segment shown in the boxed region in (B).&#160; Scale bars: 500 &#956;m (A), 20 &#956;m (B), and 2 &#956;m (C).

Watch this Scientific Journal Video about Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation at JoVE.com

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.

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 ...

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Nanyang Technological University

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JoVE Journal

Neuroscience

17.8K Views.  University of California, Santa Cruz. 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.

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

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 ...

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, direction5-7, ocular dominance8,9 and binocular disparity9. Orientation selectivity is the most widely studied feature and a continuous map with a quasi-periodic layout for preferred orientation is present across the entire primary visual cortex10,11. Integrating the synaptic, cellular and network contributions that lead to stimulus selective responses in these functional maps requires the hybridization of imaging techniques that span sub-micron to millimeter spatial scales. With conventional intrinsic signal optical imaging, the overall layout of functional maps across the entire surface of the visual cortex can be determined12. The development of in vivo two-photon microscopy using calcium sensitive dyes enables one to determine the synaptic input arriving at individual dendritic spines13 or record activity simultaneously from hundreds of individual neuronal cell bodies6,14. Consequently, combining intrinsic signal imaging with the sub-micron spatial resolution of two-photon microscopy offers the possibility of determining exactly which dendritic segments and cells contribute to the micro-domain of any functional map in the neocortex. Here we demonstrate a high-yield method for rapidly obtaining a cortical orientation map and targeting a specific micro-domain in this functional map for labeling neurons with fluorescent dyes in a non-rodent mammal. With the same microscope used for two-photon imaging, we first generate an orientation map using intrinsic signal optical imaging. Then we show how to target a micro-domain of interest using a micropipette loaded with dye to either label a population of neuronal cell bodies or label a single neuron such that dendrites, spines and axons are visible in vivo. Our refinements over previous methods facilitate an examination of neuronal structure-function relationships with sub-cellular resolution in the framework of neocortical functional architectures.

A method is described for labeling neurons with fluorescent dyes in predetermined functional micro-domains of the neocortex. First, intrinsic signal optical imaging is used to obtain a functional map. Then two-photon microscopy is used to label and image neurons within a micro-domain of the map.

In the primary visual cortex of non-rodent mammals, neurons are clustered according to their preference for stimulus features such as orientation1-4, ...

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 Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

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 ...

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 ...

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

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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

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

Targeted Labeling of Neurons in a Specific Functional Micro-domain of the Neocortex by Combining Intrinsic Signal and Two-photon Imaging

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 Imaging of Cellular Dynamics in the Mouse Spinal Cord

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

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