The authors would like to thank L. Belluscio for surgery filming equipment, D. Kwon for filming assistance, K. Liu for video editing assistance, and K. MacLeod for all background music. K.W. acknowledges the generous support of the NIMH Division of Intramural Research Programs and the Genes, Cognition and Psychosis Program. This work was supported by the NIMH Intramural Research Program (V.C., Y.Y., S.M. K.W.) and the NIAAA Division of Intramural Clinical and Biological Research Program (V.C., R.M.C., D.M.L.).

The experimental procedures described below were approved by the National Institute of Mental Health Animal Care and Use Committee and were in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
1. Pre-operative Preparation
<ol>
 <li>Clean all tools in a hot bead sterilizer before aseptic surgery, clean the surgery site with 70% ethanol, and put down clean drop cloths. Wear sterile gloves. Anesthetize the animal with freshly prepared 1.2% avertin solution, given at 0.02 ml/g, intraperitonially. Alternatively, anesthetize with isoflurane gas through a nose cone, 5% for induction, 1.5% for maintenance, with a passive scavenger circuit. Check the anesthesia level using tail or toe pinches to ensure full sedation. </li>
 <li>Cover the animal's eyes with sterile ophthalmic ointment, and inject freshly prepared dexamethasone (0.2 mg/kg) and carprofen (5 mg/kg) subcutaneously to prevent brain swelling and inflammation11. Shave the hair over the skull between the ears, and sterilize the skin with betadine scrub and three alternating swabs of 70% ethanol. </li>
 <li>Mount the animal in a stereotaxic surgery stage with earbars, with a water circulation heating pad below the animal set at 37 &deg;C. Regularly check the animal's anesthesia levels, and supplement the original anesthetic dose as needed. </li>
 <li>Inject 200 &mu;l of 0.5% Marcaine under the scalp skin to numb the area. Incise the skin and remove the skin flap over the skull. Remove the periosteum and dry the area with clean cotton swabs. </li>
</ol>
2. Chronic Cranial Window Surgery
<ol>
 <li>Use a high-speed dental drill with a 0.5 mm burr to gently outline a 3-5 mm diameter circle over the brain region of interest. Periodically wet the drilling site with sterile 0.9% saline and clear away bone dust with clean cotton swabs. If the bone bleeds, use gelfoam presoaked with sterile saline to blot the bleed and wait for it to stop. </li>
 <li>When the last bone layer is reached, lift and remove the bone island with fine-tipped forceps. Dura attachments to the bottom shelf of the bone may be encountered if the window crosses a bone suture. These should be gently removed as the bone flap is lifted.</li>
 <li>Roll moistened gel foam gently over the exposed dura to clean its surface and wait for any dural bleeding to stop. Critical step: do not proceed until all dural bleeding has stopped. Blood that becomes trapped inside the cranial window usually leads to an opaque window. </li>
 <li>If the region of interest is under a location where the dura is particularly thick, removing the dura above it may be necessary. If this is the case, gently separate the dura from the pia below with very fine forceps, make a small incision in the lifted dura with another pair of fine forceps, then grasp the cut edges and gently split the dura. The dura is thin but strong, so be careful not to slice the brain with the intact edges of the dura. </li>
 <li>Rinse the area with sterile ACSF or sterile saline. </li>
 <li>Lay a sterile glass coverslip (3-5 mm) over the dura or pia. Critical step: If the surrounding skull curves significantly, use Kwiksil12 adhesive to fill in spaces where the dura or pia would not make close contact with the coverslip in regions that will not be imaged. </li>
 <li>Use cyanoacrylate gel to cover the elastomer adhesive and the edges of the glass coverslip. Cover the entire exposed skull with cyanoacrylate gel, or a krazyglue and dental cement mixture. Critical step: Do not allow any cyanoacrylate gel to touch the brain surface. </li>
 <li>Embed a custom-made metal head fixation bar at the opposite end of the skull. </li>
 <li>Return the animal to a warm recovery chamber, after an intraperitonial injection of ketoprofen, 5 mg/kg, for pain management. Continue the analgesic for two more days post-operatively. </li>
 <li>After two weeks of post-operative recovery, anesthetize the animal with isoflurane (5% for induction, 1.5% for maintenance), mount it in a custom-made microscope stage with a head-fixation frame, and check the cranial window for optical clarity under illumination with blue light. Brain surface blood vessel clarity is highly indicative of cranial window quality. If the blood vessel edges are sharply defined, the window is likely to be useable. The two-week post-operative recovery time is recommended to allow sufficient time for the animal to recover from surgery. Cranial windows typically clear and improve during this time, and remain optically transparent for additional weeks to months until regrowth of skull or thickening of the dura degrades window quality. </li>
</ol>
3. Behavioral Protocol and Laser Scanning Two-photon Microscopy
<ol>
 <li>Animals with clear cranial windows will be exposed to an environmental stimulus or subjected to a behavioral training session, and then imaged afterwards at the time of maximal Arc-GFP expression. Before beginning a behavioral protocol, the animal should be kept in a consistent home cage environment to minimize day-to-day variation in baseline Arc-GFP expression levels. </li>
 <li>Begin behavioral training or environmental stimulation paradigms, depending on the brain region of interest. For example, animals can be exposed to different visual environments over consecutive days, and imaged each day after visual stimulation to identify stimulus-specific responses in the visual cortex10. </li>
 <li>Arc-GFP fluorescence in neurons typically reaches its peak level two hours after stimulation. The experimental timeline may be optimized to facilitate the detection of Arc-GFP expression in neurons at their peak fluorescence levels. </li>
 <li>After a behavioral training session or environmental stimulation is completed, anesthetize the animal with isoflurane (5% for induction, 1.5% for maintenance) and mount the animal in the custom-made microscope stage with head-fixation frame under the two-photon microscope. </li>
 <li>The animal's head position is fixed to the head-fixation frame that connects directly to the stage, using the implanted metal bar on the skull. Isoflurane and oxygen are continuously supplied to the mouse through a nose cone. Body temperature is maintained using a heating pad.</li>
 <li>Ensure that the microscope detectors are protected from ambient light by conducting two-photon laser scanning in a dark room. Use a 20x or 25x (1.05 numerical aperture) water immersion lens for imaging. First, under epi-fluorescence illumination, acquire an image of the surface blood vessel patterns over the brain region of interest with a CCD camera for future image alignment. </li>
 <li>Start two-photon laser scanning to acquire a 3-D image stack. An Olympus FV1000MPE multi-photon microscope is used in our setup. The excitation wave-length of the two-photon laser is set at 920 nm, and the power of the laser emitted from the objective is set at approximately 50 mW. Emitted fluorescence is simultaneously detected in green and red channels (dichroic mirror at 570 nm, barrier filters at 495-540 nm and 570-625 nm), using an external two-channel photomutiplier detection system placed close to the specimen. Arc-GFP fluorescence only appears in the green channel, whereas tissue auto-fluorescence appears in both channels13, 14. </li>
 <li>Typical image stacks have dimensions of approximately 320x320x100 &mu;m (width x length x depth), a horizontal resolution of 0.5 &mu;m/pixel, and a vertical resolution of 3 &mu;m/pixel.</li>
 <li>After acquiring the image stack, return the animal to its home cage. Do not disturb the animal until the next behavioral and imaging session. Repeat the behavior and imaging procedure across days as desired. Use the previously acquired brain surface blood vessel image to orient back to the same imaging location. </li>
</ol>

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F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mapping+behaviorally+relevant+neural+circuits+with+immediate-early+gene+expression.">Mapping behaviorally relevant neural circuits with immediate-early gene expression.</a> Current opinion in neurobiology. 15 (5), 599-606 (2005).</li><li>Barth, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+circuits+and+systems+using+transgenic+reporters+of+neural+activity.">Visualizing circuits and systems using transgenic reporters of neural activity.</a> Curr. Opin. Neurobiol. 17 (5), 567-5671 (2007).</li><li>Lyford, G. 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No conflicts of interest declared.

The in vivo imaging method described here enables repeated examination of Arc gene expression changes in the same sets of neurons over multiple days in the live animal. It is an efficient and versatile method to obtain information about neural plasticity-related molecular dynamics in individual neurons in response to various behavioral experiences. Standard histochemical methods such as in situ hybridization and immunostaining can achieve single-cell resolution3, but lack the ability to tra...

<table border="1">
<tbody>
 <tr>
 <td>Name 			of the Equipment</td>
 <td>Company</td>
 <td>Catalogue 			number</td>
 <td>Comments 			(optional)</td>
 </tr>
 <tr>
 <td>FV1000 			multi-photon laser scanning microscope</td>
 <td>Olympus</td>
 <td>FV1000MPE</td>
 <td>Imaging</td>
 </tr>
 <tr>
 <td>Dissection 			microscope</td>
 <td>Omano</td>
 <td>555V107</td>
 <td>Surgery</td>
 </tr>
 <tr>
 <td>Stereotaxis 			surgery stage for mice</td>
 <td>Harvard 			Apparatus</td>
 <td>726335</td>
 <td>Surgery</td>
 </tr>
 <tr>
 <td>20X 			or 25X water immersion objective</td>
 <td>Olympus</td>
 <td>XLPL25XWMP</td>
 <td>Imaging</td>
 </tr>
 <tr>
 <td>Microscope 			stage with head-fixation frame</td>
 <td>Custom 			made</td>
 <td>N/A</td>
 <td>Imaging</td>
 </tr>
 <tr>
 <td>Fine 			forceps</td>
 <td>Fine 			Science Tools</td>
 <td>11251-20</td>
 <td>Surgery</td>
 </tr>
 <tr>
 <td>Dental 			drill burr</td>
 <td>Fine 			Science Tools</td>
 <td>19007-05</td>
 <td>Surgery</td>
 </tr>
 <tr>
 <td>CCD 			camera</td>
 <td>QImaging</td>
 <td>QICAM 			12-bit</td>
 <td>Imaging</td>
 </tr>
 </tbody>
</table>

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.

This protocol describes a method to track experience-dependent molecular changes in individual cortical neurons in live animals. A chronic cranial window is first created over a cortical region of interest in a mouse carrying a fluorescent reporter of gene expression. Two-photon microscopy can then be coupled with various behavioral paradigms to observe behaviorally induced molecular changes in individual neurons and track such changes in the same sets of neurons over multiple days (Figure 1).
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Watch this Scientific Journal Video about In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons at JoVE.com

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.

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 ...

in-vivo-two-photon-imaging-experience-dependent-molecular-changes

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21.5K Views.  National Institute of Mental Health. 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.

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

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 ...

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 Two-Photon Microscopy of Single Nerve Endings in Skin

Nerve endings in skin are involved in physiological processes such as sensing1 as well as in pathological processes such as neuropathic pain2. Their close-to-surface positioning facilitates microscopic imaging of skin nerve endings in living intact animal. Using multiphoton microscopy, it is possible to obtain fine images overcoming the problem of strong light scattering of the skin tissue. Reporter transgenic mice that express EYFP under the control of Thy-1 promoter in neurons (including periphery sensory neurons) are well suited for the longitudinal studies of individual nerve endings over extended periods of time up to several months or even life-long. Furthermore, using the same femtosecond laser as for the imaging, it is possible to produce highly selective lesions of nerve fibers for the studies of the nerve fiber restructuring. Here, we present a simple and reliable protocol for longitudinal multiphoton in vivo imaging and laser-based microsurgery on mouse skin nerve endings.

In Vivo Two-Photon Microscopy of Single Nerve Endings in Skin

The protocol for dynamic longitudinal imaging and selective laser lesion of nerve endings in reporter transgenic mice is presented.&#160;

Nerve endings in skin are involved in physiological processes such as sensing1 as well as in pathological processes such as neuropathic pain2. Their ...

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 ...

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 ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent and cause visual impairment and blindness. Understanding how such diseases progress is critical to formulating new treatments. In vivo&#160;microscopy in animal models of disease is a powerful tool for understanding neurodegeneration and has led to important progress towards treatments of conditions ranging from Alzheimer&#39;s disease to stroke. Given that the retina is the only central nervous system structure inherently accessible by optical approaches, it naturally lends itself towards in vivo imaging. However, the native optics of the lens and cornea present some challenges for effective imaging access.
This protocol outlines methods for in vivo two-photon imaging of cellular cohorts and structures in the mouse retina at cellular resolution, applicable for both acute- and chronic-duration imaging experiments. It presents examples of retinal ganglion cell (RGC), amacrine cell, microglial, and vascular imaging using a suite of labeling techniques including adeno-associated virus (AAV) vectors, transgenic mice, and inorganic dyes. Importantly, these techniques extend to all cell types of the retina, and suggested methods for accessing other cellular populations of interest are described. Also detailed are example strategies for manual image postprocessing for display and quantification. These techniques are directly applicable to studies of retinal function in health and disease.

In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent ...