The project was funded by&#160;a Research Initiative Award from the American Epilepsy Society, and an award from the Arizona Biomedical Research Commission to S.L.M., as well as a challenge grant from Research to Prevent Blindness Inc. to the Department of Ophthalmology at SUNY Downstate Health Sciences University, the New York State Empire Innovation Program, and further grants from the National Science Foundation (0726113, 0852636, &#38; 1523614), the Barrow Neurological Foundation, Mrs. Marian Rochelle, Mrs. Grace Welton, and Dignity Health SEED awards, and by federal grants from the National Science Foundation (0726113, 0852636, &#38; 1523614) and by the National Institute of Health (Awards R01EY031971 AND R01CA258021), to S.L.M and S.M.C.&#160;<span lang="EN-US" style="font-size: 12pt; line-height: 107%; font-family: Calibri, sans-serif; color: rgb(34, 34, 34); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">This work was also supported by the Office of the Assistant Secretary of Defense for Health Affairs under Award No. W81XWH-15-1-0138, to S.L.M..<span lang="EN-US" style="font-size: 12pt; line-height: 107%; font-family: Calibri, sans-serif; color: red; background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">&#160;<span lang="EN-US" style="font-size: 12pt; line-height: 107%; font-family: Calibri, sans-serif; color: rgb(34, 34, 34); background-image: initial; background-position: initial; background-size: initial; background-repeat: initial; background-attachment: initial; background-origin: initial; background-clip: initial;">L.-C. was supported by a Jos&#233; Castillejo fellowship from the Spanish Ministry of Education. We thank O. Caballero and M. Ledo for their technical advice and assistance.&#160;

The protocol follows the NIH Guidelines for the Care and Use of Laboratory Animals. All procedures were approved by the Barrow Neurological Institute&#8217;s Institutional Animal Care and Use Committee.
1. Stereotaxic positioning for craniotomy
<ol>
	<li>Weigh and then anesthetize the mouse with a ketamine-xylazine (100 mg/kg&#8211;10 mg/kg i.p.) cocktail. Ensure that the animal is fully anesthetized by observing its lack of reaction to tail and/or toe pinch.</li>
	<li>Place a heating pad under the mouse and use a rectal thermometer to continuously monitor its body temperature during the initial anesthetized implantation surgery (Figure 2A).</li>
	<li>Apply ophthalmic ointment to prevent eye dryness.</li>
	<li>Stabilize the head of the animal in a rodent stereotaxic frame, outfitted with a mouse adapter. To properly orient the mouse in the stereotaxic frame, place the animal&#8217;s teeth inside the hole of the bite-bar (Figure 2A and Figure 3A, item g). Tighten the nose-clamp until it is snug (Figure 2A,B and Figure 3A, item h). The mouse&#8217;s body positioning should be as straight as possible (as in Figure 2A).
	<ol>
		<li>Further secure the animal&#8217;s head using mouse ear-bars with jaw holder cuffs (Figure 2A and Figure 3A, item i), to firmly clamp the zygomatic processes of the skull. 
		NOTE: Once secured, the mouse&#8217;s head should have no range of horizontal motion.</li>
	</ol>
	</li>
	<li>Once the mouse is adjusted to the stereotaxic frame (Figure 3B, item j), clean and sterilize the scalp by wiping it with chlor-hexedine scrub 2-3 times, each time followed by an alcohol rinse. Then, apply topical lidocaine on top of the head.</li>
	<li>Make an incision along the midline from posterior to anterior (start incision at the base of skull and complete it between the eyes; 12-15 mm) along the sagittal midline of the skull, using either fine scissors or a blade. Retract the borders of the scalp with four 28 mm bulldog serrefine clamps to expose the skull and maximize the working area (Figure 2B and Figure 3C).</li>
	<li>Use a scalpel or microspatula (Figure 3C) to scrape the periosteum to the incision&#8217;s edges. Use tissue-separating scissor or forceps (Figure 3C) to carefully retract the muscles at the back of the neck (Figure 2B).</li>
	<li>Dry the top of the skull with a cotton-tipped applicator, moistened with alcohol or 30% hydrogen peroxide, to optimize visualization of the cranial sutures. Apply saline to the skull once the connective tissue has been removed.</li>
	<li>Once the mouse&#8217;s head is stabilized in the stereotaxic apparatus, place a syringe needle tip (21 G) in the electrode holder (Figure 3B, item l), and attach the electrode holder to the stereotaxic micromanipulator, lowering the needle tip to the level of the skull (Figure 2D).</li>
	<li>Adjust the medial-lateral direction of the skull by measuring the height of the skull at any distance posterior to bregma (i.e., -2.0 mm), and two equal distances lateral from the midline on either side of the skull (i.e., &#177; 1.5 mm) (Figure 2D). Aim for a height difference across locations that is less than 0.05 mm in the dorsal-ventral direction.
	<ol>
		<li>If greater than 0.05 mm, rotate the mouse&#8217;s skull using the bite bar, or reposition the mouse&#8217;s head by loosening the ear-bars carefully, laterally shifting the head and ear-bars as needed, and then retightening the ear-bars.</li>
	</ol>
	</li>
	<li>Once the skull is positioned in the stereotaxic device, determine and record the stereotactic coordinates of the bregma and lambda skull sutures along the anteroposterior (AP), medial lateral (ML), and dorsal ventral (DV) axes.</li>
	<li>Identify the target point (right hippocampus) with the aid of a stereotaxic atlas. Then, find on the skull the target point coordinates relative to bregma (AP: -2.7 mm, ML: -2.7 mm) (Figure 2E).</li>
	<li>Using a surgical marker, mark four corners of a 1.4 mm x 2 mm imaging window on the skull with dots, centered around the target point directly above the hippocampus (Figure 2F), for subsequent craniotomy.</li>
	<li>Use the surgical marker to draw two additional dots: one over the top-left parietal bone (Figure 2G) and another over the right frontal bone (Figure 2H), to indicate the future placement of two bone screws (Figure 3A, item b) that will anchor the head-cap (Figure 3B, item m) to the skull.</li>
	<li>Use the surgical marker to draw three more dots indicating where the epidural EEG recording electrodes will be inserted: one dot positioned 1 mm to either side of the midline, and one additional dot on the midline positioned 1 mm behind Lambda (Figure 2E, Figure 4A and Figure 5B).</li>
</ol>
2. Head-Cap installation
<ol>
	<li>Using a 0.7 mm diameter burr (Figure 3C), carefully drill two small holes in the skull, at the dot positions indicating the bone screws placement (see step 1.15 above). Then, take two bone screws (stainless steel-slotted, length: 4 mm; diameter: 0.85 mm) previously disinfected with ethanol 70-80%, and screw them to the skull for additional head-cap stability (Figure 2I-J and Figure 3A, item b).
	<ol>
		<li>Ensure that the screw tips do not protrude beyond the bottom of the bone into the skull cavity, risking damage to the brain. Note that one of the two bone screws will be anterior to the two head-cap screws, and the other posterior to them (Figure 2G-J).</li>
	</ol>
	</li>
	<li>Squirt saline over the skull to cool it down and clean it. Then dry the skull completely and apply a thin layer of cyanoacrylate around the screws.</li>
	<li>Use a custom alignment piece (Figure 3A, item c) clamped to the microdrive mounted to the stereotaxic device (Figure 2F-J and Figure 3A, item d), so that the position of the head-cap is aligned along the midline, with the anterior ridge overlying bregma. This custom stainless-steel piece is L-shaped (angled at 90&#176;), with two holes separated by 4 mm (Figure 2F-J and Figure 3A, item c).
	<ol>
		<li>Use the stereotaxic manipulator to position the L-shaped alignment piece, with two Fillister Head Slotted drive screws attached (M1.6 x 0.35 Metric Coarse, 12 mm Length; Figure 3A, item a) over the bregma, until the base of the screws lies flat on the skull, being careful to not exert pressure on the skull (Figure 2I-J).</li>
	</ol>
	</li>
	<li>Apply cyanoacrylate followed by dental cement to attach the base of the screws to the skull. Be careful to not obstruct the imaging window reference marks over the hippocampus (Figure 2K).</li>
</ol>
3. Imaging window craniotomy surgery
<ol>
	<li>At each of the positions of the craniotomy reference marks that indicate the corners of the imaging window (see step 1.14 above), drill a 0.7 mm diameter burr hole, followed by gently delineating the periphery of the 1.4 mm x 2 mm craniotomy window with the drill, with iteratively deeper cuts (Figure 2L).</li>
	<li>Take care to not to drill too deeply or with too much force on the skull, which is thin and fragile, having a thickness of approximately 300 &#181;m. Once the bone flap is complete, pry the loose flap up with the tip of a scalpel, being mindful to not damage the underlying dura mater.</li>
	<li>Cover the open window with bone wax (Figure 2M).</li>
	<li>Using a 0.9 mm diameter burr, drill three holes for future placement of the epidural EEG electrodes (see step 1.15 above) taking care to not to cut through the dura mater (Figure 2E,L).</li>
	<li>Cover the three holes with bone wax.</li>
	<li>Gently apply cyanoacrylate around the edges of the bone-wax covered imaging window and EEG holes, being careful to not seal the craniotomies (Figure 2L).</li>
	<li>Build a wall of dental cement that encompasses the imaging window and EEG holes and binds it to the previously cemented head-cap (Figure 2M and Figure 3B, item m).</li>
	<li>Let the cement dry for at least 10 minutes.</li>
	<li>Close any loose wound margins with simple interrupted sutures, using 4-0 or 5-0 black braided silk (Figure 2N and Figure 3C).</li>
</ol>
4. Post-surgery
<ol>
	<li>Use a cotton-tipped applicator to apply lidocaine ointment around the exposed skin to deaden sensation around the wound&#8217;s edge.</li>
	<li>Use a sterile cotton-tipped applicator to apply Neosporin to the exposed skin.</li>
	<li>Remove the animal from the stereotaxic frame (Figure 2N).</li>
	<li>Inject ketoprofen (5 mg/kg) IP or IM for post-surgical analgesia.</li>
	<li>Observe the animal until it is alert and moving (this will usually occur within minutes).</li>
	<li>Place the animal in a warm cage and continue to monitor its recovery until it drinks or eats and is ready to transfer to animal housing.</li>
	<li>Check on the animal at least once daily for approximately one week after the surgery, watching for any signs of listless behavior and/or inadequate eating/drinking (Figure 2O).</li>
</ol>
5. Dye injection and EEG recording
<ol>
	<li>Wait a minimum of one day after the head-cap implantation surgery to initiate the recordings.</li>
	<li>Place the mouse in a foam-restraining device molded to its body, within the stereotaxic frame (Figure 3B, items n &#38; j).</li>
	<li>Secure the head-cap (Figure 3B-I, item m) to a custom mounting bar which is attached to the stereotaxic frame (Figure 3A,B, items e &#38; j, and Figure 4A). The long section of item e (Figure 3A,B) should have a length between 9.4 &#8211; 13 mm. Fix item f (a mounting plate with dimensions 1.5 cm x 0.5 cm, with two holes separated 4 mm from center to center) to item e (Figure 3A,B, items e, f and k). This alignment system will position the awake mouse securely to the stereotaxic frame throughout the imaging sessions (Figure 4).</li>
	<li>Insert the epidural EEG recording electrodes (Figure 5A), placing the ground electrodes (white wires) within the posterior central hole, and the recording electrodes (red wires) in the anterior left and right holes (Figure 2E, Figure 4E and Figure 5E). Position each of the wires at an L-shape between the skull and the dura mater (Figure 5C) to optimize electrical contact.</li>
	<li>Inject the tail vein with green fluorescein lysine-fixable dextran (0.2 mL/25 g body wt. of Fluorescein 5% w/w), to visualize blood flow within the hippocampus.
	<ol>
		<li>To enhance dilation of the central vein of the mouse&#8217;s tail previous to injection, use one or more of the following strategies: a) Apply 70% ethanol to the central vein using a cotton-tipped applicator; b) Dip the tail into warm water (35-40 &#176;C) for 1-3 minutes; c) Expose the tail to a heating lamp for a few minutes.</li>
		<li>Secure the mouse&#8217;s tail with two fingers and locate the central vein, which lies immediately below the skin. Insert the needle (ultrafine insulin syringe with integrated needle 30G), with the bevel up, into the vein, approximately halfway to two thirds from the base of the tail. Keep the needle parallel to the tail during the injection.</li>
		<li>Inject the dye slowly and steadily, taking care to avoid any changes in the position of the needle or tail. 
		NOTE: There should be no resistance upon depressing the syringe plunger.</li>
		<li>After completing the injection, withdraw the needle and apply gentle pressure to the puncture site to seal the vessel.</li>
		<li>Allow clotting to occur.</li>
	</ol>
	</li>
</ol>
6. In vivo fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE)
<ol>
	<li>Directly following the tail vein injection (step 5.5.3 above), clamp the 300 &#181;m beveled fiber-optic bundle (containing 5000&#8211;7000 3 &#181;m fibers in each bundle) in the downward orientation to the mobile arm of the robotic stereotaxic drive.</li>
	<li>Remove the bone wax from the craniotomy.</li>
	<li>Remove the dura, using the bent tip of a syringe needle.</li>
	<li>Using the robotic positioning system, first move the fiber objective to the appropriate anterior-posterior and left-right stereotaxic location, with the tip zeroed in the z-axis on the surface of the cortex.</li>
	<li>Initiate the EEG recording (Figure 5D).</li>
	<li>Initiate confocal laser-scanning of the back plane of the fiber-objective, followed directly by descending the objective slowly in the z-axis using the robotic microdrive&#8217;s z-controls, until it reaches the target depth within the brain (Figure 4C). View the imaging results in the microscopic software while descending.
	<ol>
		<li>If the fiber tip is within the target X-Y-Z recording region, and two or more vessels enter the imaging window&#8217;s FOV, stop the descent and initiate video recordings. Obtain live full-field time-lapse movies of the blood flow at 11.7 Hz using Cellvizio Lab&#8217;s imaging software. (Higher speeds may be obtained with a smaller FOV). Recordings may take place for 4-5 hours at a time, in consecutive days if needed.</li>
	</ol>
	</li>
	<li>Using a 10 mL syringe with beveled silicon tubing fixed to the tip of the needle, feed the animal during the recordings by periodically providing paste made with mice pellets ground with water.</li>
	<li>At the end of the imaging session, terminate the recordings.</li>
	<li>Gently remove the EEG leads and clean them with Tergazyme.</li>
	<li>Slowly remove the fiber-optic bundle from the animal&#8217;s brain by reversing the fiber along the z-axis of entry.</li>
	<li>Release the animal from the head post and body restraints.</li>
	<li>If appropriate, follow the necessary euthanasia and tissue processing protocols to visualize blood vessels and to perform immunohistochemistry.</li>
	<li>If planning to record from the same animal in a future session, cover the imaging window with bone wax or silastic.</li>
</ol>

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We have nothing to disclose.

We developed a head-cap restraint system for simultaneous electrophysiological and fiber-optic pCLE experiments in awake mice, reducing potential response contamination due to anesthetic drugs. The head-cap and mounting apparatus are straightforward to construct and are reusable for chronic awake-behaving imaging experiments. We checked the quality of the recordings against the gold-standard for in vivo microscopic blood flow imaging, TPLSM.
Proficient surgical skills are necessary to implemen...

In contrast with other microscopic imaging methods1,2,3,4,5,6,7,8, in vivo fiber-optic-based confocal microscopy allows the measurement of blood flow dynamics in any brain region, at any depth, at high speed (up to 240 Hz depending on field-of-view size9). A fiber-optic probe enables in vivo confocal laser scanning imaging at 3 &#181;m resolution because the tip of the probe (a lens-less objective made up of a bundle of 5000-6000 3 &#181;m diameter individual fibers) can be positioned with a microelectrode&#8217;s accuracy, within 15 &#181;m of the fluorescent target of interest. As with in vivo two-photon imaging, fluorophores must be previously introduced into the imaging target. For example, fluorescein dextran (or quantum dots) may be injected into the vasculature, or genetically-encoded fluorescent proteins can be transfected into cells, or fluorescent dyes such as Oregon Green BAPTA-1 can be bulk-loaded into cells, prior to imaging.
Recent research using these techniques has found that mural cell motor activity leading to ictal capillary vasospasms&#8212;sudden constrictions that occur at the position of the mural cells during seizures9&#8212;can contribute to neurodegeneration in the ictal hippocampus9. Whereas previous imaging studies showed in vitro and in vivo pericyte constrictions connected to drug applications6,7,10,11,12, Leal-Campanario et al. found the first evidence of in vivo spontaneous capillary constrictions in the murine brain. To establish relevance to human temporal lobe epilepsy, they studied male (P30-40 old) knockout (KO) Kv1.1 (kcna1-null) mice14,15 (JAX stock #003532), a genetic model of human episodic ataxia type 115. Pericytes drove both pathological and physiological hippocampal mural vasoconstrictions9 in the spontaneously epileptic animals and their wild-type (WT) littermates. These observations were replicated in WT animals rendered epileptic with kainic-acid, thereby indicating their generalization to other forms of epilepsy. Leal-Campanario et al moreover determined, using novel stereological microscopy approaches, that apoptotic&#8212;but not healthy&#8212;neurons in epileptic animals were spatially coupled to the hippocampal microvasculature. Because excitotoxicity has no known spatial association to the vasculature, this result indicated that abnormal capillary vasospasmic ischemia-induced hypoxia contributes to neurodegeneration in epilepsy. Figure 1 shows a schematic of the general setup.

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in vivo fiber-coupled pre-clinical confocal laser-scanning endomicroscopy (pcle) of hippocampal capillaries in awake mice

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application to elucidate capillary blood flow effects during seizures, driven by mural cells. In vitro and in vivo cortical imaging have shown that capillary constrictions driven by pericytes can result from functional local neural activity, as well as from drug application, in healthy animals. Here, a protocol is presented on how to use pCLE to determine the role of microvascular dynamics in neural degeneration in epilepsy, at any tissue depth (specifically in the hippocampus). We describe a head restraint technique that has been adapted to record pCLE in awake animals, to address potential side-effects of anesthetics on neural activity. Using these methods, electrophysiological and imaging recordings can be conducted over several hours in deep neural structures of the brain.

We developed these methods to assess whether abnormal pericyte-driven capillary vasospasms in the hippocampus&#8212;occurring as the result of seizures&#8212;could cause frank hypoxia that contributes to cell death in the ictal focus9,13.
The development of the head-cap and its proper installation afforded high-stability in the recordings, allowing simultaneous recording of EEG and blood flow deep in the hippocampus of wild-type and ep...

Watch this Scientific Journal Video about In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice at JoVE.com

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice

Whereas multiphoton imaging is only effective at limited depths from the tissue&#8217;s surface, it is possible to achieve 3 &#181;m resolution imaging at any depth via pCLE. Here, we present a protocol to conduct pCLE imaging to measure microvascular dynamics in the hippocampus of ictal and wild-type mice.

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy (pCLE) of Hippocampal Capillaries in Awake Mice

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application ...

in-vivo-fiber-coupled-pre-clinical-confocal-laser-scanning

Research

JoVE Journal

Neuroscience

849 Views.  Universidad Pablo de Olavide. Dit protocol stelde ons in staat om de confocale laserscanning glasvezel-endomicroscoop te gebruiken om gedurende enkele uren gelijktijdige beeldvorming en elektrofysiologische opnames in vivo uit te voeren van neurale structuren op elke diepte van de hersenen in het wakkere of relatief verdoofde dier. We ontwikkelden een pijnloze fixatiemethode die diepe beeldvorming van de hersenen bij wakkere dieren mogelijk maakt met behulp van methoden zoals die eerder werden gebruikt voor elektrofysiolo...

Video: In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy pCLE of Hippocampal Capillaries in Awake Mice

Flat-floored Air-lifted Platform: A New Method for Combining Behavior with Microscopy or Electrophysiology on Awake Freely Moving Rodents

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a living animal&#8217;s brain. Moreover, the lengthy recovery from anesthesia limits the frequency of repeated recording/imaging episodes in longitudinal studies. Hence, new methods that would allow stable recordings from non-anesthetized behaving mice are expected to advance the fields of cellular and cognitive neurosciences. Existing solutions range from mere physical restraint to more sophisticated approaches, such as linear and spherical treadmills used in combination with computer-generated virtual reality. Here, a novel method is described where a head-fixed mouse can move around an air-lifted mobile homecage and explore its environment under stress-free conditions. This method allows researchers to perform behavioral tests (e.g., learning, habituation or novel object recognition) simultaneously with two-photon microscopic imaging and/or patch-clamp recordings, all combined in a single experiment. This video-article describes the use of the awake animal head fixation device (mobile homecage), demonstrates the procedures of animal habituation, and exemplifies a number of possible applications of the method.

This method creates a tangible, familiar environment for the mouse to navigate and explore during microscopic imaging or single-cell electrophysiological recordings, which require firm fixation of the animal&#8217;s head.

It is widely acknowledged that the use of general anesthetics can undermine the relevance of electrophysiological or microscopical data obtained from a ...

Behavior

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 ...

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and magnetic resonance imaging (MRI) have enabled in vivo quantification of CSF flow but they are limited by reduced imaging volumes and low spatial resolution, respectively. Recent work has found that CSF enters the brain parenchyma through a network of perivascular spaces surrounding the pial and penetrating arteries of the rodent cortex. This perivascular entry of CSF is a primary driver of the glymphatic system, a pathway implicated in the clearance of toxic metabolic solutes (e.g., amyloid-&#946;). Here, we illustrate a new macroscopic imaging technique that allows real-time, mesoscopic imaging of fluorescent CSF tracers through the intact skull of live mice. This minimally-invasive method facilitates a multitude of experimental designs and enables single or repeated testing of CSF dynamics. Macroscopes have high spatial and temporal resolution and their large gantry and working distance allow for imaging while performing tasks on behavioral devices. This imaging approach has been validated using two-photon imaging and fluorescence measurements obtained from this technique strongly correlate with ex vivo fluorescence and quantification of radio-labeled tracers. In this protocol, we describe how transcranial macroscopic imaging can be used to evaluate glymphatic transport in live mice, offering an accessible alternative to more costly imaging modalities.

Transcranial optical imaging allows wide-field imaging of cerebrospinal fluid transport in the cortex of live mice through an intact skull.

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and ...

Craniotomy Procedure for Visualizing Neuronal Activities in Hippocampus of Behaving Mice

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit functions in systems neuroscience. However, high absorbance and scattering of light in mammalian tissue limit intravital imaging mostly to superficial brain regions, leaving deep-brain areas, such as the hippocampus, out of reach for optical microscopy. In this video, we show the preparation and implantation of the custom-made imaging window to enable chronic in vivo imaging of the dorsal hippocampal CA1 region in head-fixed behaving mice. The custom-made window is supplemented with an infusion cannula that allows targeted delivery of viral vectors and drugs to the imaging area. By combining this preparation with wide-field imaging, we performed a long-term recording of neuronal activity using a fluorescent calcium indicator from large subsets of neurons in behaving mice over several weeks. We also demonstrated the applicability of this preparation for voltage imaging with single-spike resolution. High-performance genetically encoded indicators of neuronal activity and scientific CMOS cameras allowed the recurrent visualization of subcellular morphological details of single neurons at high temporal resolution. We also discuss the advantages and potential limitations of the described method and its compatibility with other imaging techniques.

This article demonstrates the preparation of a custom-made imaging window supplemented with infusion cannula and its implantation onto the CA1 region of the hippocampus in mice.

Imaging neuronal activities at single-cell resolution in awake behaving animals is a very powerful approach for the investigation of neural circuit ...

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their activity in vivo in behaving mice. This paper describes a method for the implantation of a chronic cranial window to allow for the longitudinal imaging of brain cells in awake, head-restrained mice. In combination with genetic strategies and viral injections, it is possible to label specific cells and regions of interest with structural or physiological markers. This protocol demonstrates how to combine viral injections to label neurons in the vicinity of GCaMP6-expressing astrocytes in the cortex for simultaneous imaging of both cells through a cranial window. Multiphoton imaging of the same cells can be performed for days, weeks, or months in awake, behaving animals. This approach provides researchers with a method for viewing cellular dynamics in real time and can be applied to answer a number of questions in neuroscience.

Implantation of a Cranial Window for Repeated In Vivo Imaging in Awake Mice

Presented here is a protocol for the implantation of a chronic cranial window for the longitudinal imaging of brain cells in awake, head-restrained mice.

To fully understand the cellular physiology of neurons and glia in behaving animals, it is necessary to visualize their morphology and record their ...

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.

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.

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

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 ...

Real-Time Imaging for Neuron Activity in the Dentate Gyrus Cells of Mice 

In Vivo Calcium Imaging of Granule Cells in the Dentate Gyrus of Hippocampus in Mice

Real-time approaches are typically needed in studies of learning and memory, and in vivo calcium imaging provides the possibility to investigate neuronal activity in awake animals during behavior tasks. Since the hippocampus is closely associated with episodic and spatial memory, it has become an essential brain region in&#160;this field&#39;s research. In recent research, engram cells and place cells were studied by recording the neural activities in the hippocampal CA1 region using the miniature microscope in mice while performing behavioral tasks including open-field and linear track. Although the dentate gyrus is another important region in the hippocampus, it has rarely been studied with in vivo imaging due to its greater depth and difficulty for imaging. In this protocol, we present in detail a calcium imaging process, including how to inject the virus, implant a GRIN (Gradient-index) lens, and attach a base plate for imaging the dentate gyrus of the hippocampus. We further describe how to preprocess the calcium imaging data using MATLAB. Additionally, studies of other deep brain regions that require imaging may benefit from this method.

Author Spotlight: Investigating Neural Activity of Dentate Gyrus Granule Cells with Miniature Microscope

The dentate gyrus of the hippocampus carries out essential and distinct functions in learning and memory. This protocol describes a set of robust and efficient procedures for in vivo calcium imaging of granule cells in the dentate gyrus in freely moving mice.

Real-time approaches are typically needed in studies of learning and memory, and in vivo calcium imaging provides the possibility to investigate neuronal ...

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in an Awake Mouse

In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy (pCLE) of Hippocampal Capillaries in Awake Mice

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