This work was supported by the American Heart Association (Post-doctoral fellowship to AYS) and the National Institutes of Health (MH085499, EB003832, and OD006831 to DK). We thank Beth Friedman and Pablo Blinder for comments on the manuscript.

1. Preparing for Surgery i
<ol>
 <li>Clean the surgical tools by sonicating in a mixture of Maxizyme and Surgical Milk in an ultrasonic cleaner. Autoclave the surgical tools before each experiment.</li>
 <li>Ensure that all necessary reagents and disposables are available. A list of reagents and disposables is provided in Table 2. Reagents and disposables that come in contact with exposed tissue should be sterile, when possible.</li>
 <li>Induce anesthesia. Typical anesthetics suitable for survival studies are described in Table 1. Ensure surgical plane of anesthesia by checking for lack of toe pinch reflex. The optimal age of mouse is 3 to 6 weeks of age. The skulls of younger mice are softer and more difficult to thin. Older mice have thicker skulls that will bleed more during the thinning procedure.</li>
 <li>Secure the animal in a stereotaxic frame 1. A list of surgical equipment is provided in Table 2.</li>
 <li>Maintain body temperature at 37&deg;C using a feedback regulated rectal probe and heat pad.</li>
 <li>Apply ophthalmic ointment to the eyes to retain moisture.</li>
 <li>Shave the scalp with a small electrical razor.</li>
 <li>Clean the scalp with Betadine, followed by swabbing with 70 % (v/v) isopropyl alcohol.</li>
 <li>If desired, check that heart and breathing rates are within a normal range using a pulse oximeter. For a mouse, these numbers should center around 10 and 2 Hz, respectively.</li>
 <li>Warm an aliquot of sterile-filtered artificial cerebral spinal fluid (ACSF) to 37&deg;C (125 mM NaCl, 10 mM glucose, 10 mM HEPES, 3.1 mM CaCl2 , 1.3 mM MgCl2 , pH 7.4) (all chemicals from Sigma)2.</li>
</ol>
2. Mounting a Head Frame
<ol>
 <li>Remove the scalp over the entire dorsal skull surface with a pair of forceps and surgical scissors. Trim the skin laterally to the edges of the temporal muscles on either side of the skull and posterior to the muscles of the neck (Fig. 1A).</li>
 <li>Use a scalpel blade to remove the thin periosteum from the surface of the skull.</li>
 <li>Clean the skull with a moist cotton tipped applicator and dry the skull surface with a stream of air from a dust can. Then apply a thin layer of cyanoacrylate glue to the entire skull surface. Allow the glue to dry thoroughly. This layer of glue is required for proper adherence of the dental cement in subsequent steps. The cyanoacrylate glue does not need to be sterilized.</li>
 <li>Attach a metal connector to the skull surface, away from the area of the desired window. For anesthetized preparations, a small nut may be secured to the skull with a dab of cyanoacrylate glue, which can later be screwed into the imaging setup using a bolt (Fig. 1B, 1L and 1N). Seal the backside of the nut with tape to ensure that glue does not enter the threads during attachment to the skull. The nut and tape should be sterilized by autoclaving prior to the surgery.</li>
 <li>For awake imaging preparations, attach a more rigid custom connector with two attachment points (Fig. 1M and 1N). Drill two holes in the skull over the contralateral cortical hemisphere with a &frac12; mm drill burr, and then introduce two #000 self-tapping screws. These screws will help anchor the head mount to the skull. Apply only one full turn of the screw, in order to avoid applying pressure to the underlying cortex. Then, attach the custom metal cross bar with a small dab of cyanoacrylate glue over the cerebellum. Allow the cyanoacrylate glue to dry thoroughly. This metal connector greatly reduces the degrees of freedom and simplifies relocation of the same imaging field in longitudinal studies. A wide cross bar gives ample room for electrode placement and stimulation of vibrissae. Detailed plans for generating the custom head mount and attachment device are available online (<a href="http://physics.ucsd.edu/neurophysics/links.html">http://physics.ucsd.edu/neurophysics/links.html</a>). The screws and cross bar should be sterilized by autoclaving prior to the surgery.</li>
 <li>Cover the entire skull surface, excluding the location of the window, with a layer of dental cement (Fig. 1C and 1D). Ensure that all exposed edges of the skin are covered by cement. The components of the dental cement do not need to be sterilized.</li>
</ol>
3. The Generation of a Polished and Reinforced Thinned-skull (PoRTS) Window
<ol>
 <li>Ensure that drill burrs are sharp and avoid reusing them. Using a low speed on the dental drill, thin a 2 mm by 2 mm region over the somatosensory cortex with a &frac12; mm burr. Alternate between wetting the skull with ACSF and then drying the skull surface with a gentle stream of air from a gas duster; wet for cooling, and dry for thinning. This requires thinning through the cancellous layer of the skull, which may bleed, but can be controlled by flushing with ACSF (Fig. 1E). The skull begins to flex under the slight pressure of the drill when it is ~50 &mu;m, and the pial vessels should be visible through wet bone (Fig. 1F). At this thickness, small white spots within the bone will become visible for a few seconds immediately after the dry skull surface is moistened.</li>
 <li>At this point, thin the bone even further. Use a slower drill speed, i.e., 1000 rpm, which shaves the skull surface with only a slight touch. Use small controlled movements while holding the drill like a pen and only apply force in the lateral direction. The bone should be ~10 to 15 &mu;m at its final thickness (Fig. 1G). When the bone is sufficiently thin, the small white spots in the bone will no longer be visible when the dry skull surface is moistened.</li>
 <li>Polish the window region with tin oxide powder. Attach a pre-made drill bit that has been dipped in silicone aquarium sealant and withdrawn, leaving a tapered whip (inset, Fig. 1H). The silicone whip should be prepared at least one day before the surgery, and sterilized with 70% isopropanol prior to use. Place a small pinch of powder on the window along with a drop of ACSF (Fig. 1H). Agitate the slurry over the window for up to ten minutes by gently touching the tip of the moving whip to the skull surface. Flush away the tin oxide powder thoroughly from the window using ACSF and dry the bone thoroughly with a gentle stream of air. Surface irregularities and adherent bone chips left by drilling in the previous steps should be removed after polishing (Fig. 1H and 1I). The tin oxide powder does not need to be sterilized.</li>
 <li>Cut square pieces of no. 0 cover glass slightly smaller than the size of the window. Use a scribe to gently score separated horizontal and vertical lines in the cover slip using a straight edge. Then place the cover slip in a petri dish and knock the dish against a table edge to separate the glass pieces. The entire petri dish can then be autoclaved to ensure sterility.</li>
 <li>Place an appropriately sized cover glass nearby the dried window. Apply a small dab of cyanoacrylate glue over the window using the tip of a broken wooden cotton tip applicator, and quickly push the precut piece of cover glass atop the glue. Avoid creating bubbles underneath the cover glass. Gently push the cover glass against the skull surface, and hold for a few seconds (Fig. 1J and 1O). Allow the glue to dry thoroughly over 15 minutes. Excess cyanoacrylate glue can be removed from the upper surface of the cover glass with a scalpel blade after it is dried. Seal the edges of the cover glass with dental cement and form a slightly raised well to hold water for the dipping lens (Fig. 1K and 1O).</li>
</ol>
4. Recovery
<ol>
 <li>Place the animal in a warm cage following surgery. Monitor the animal periodically until it fully recovers from anesthesia.</li>
 <li>If the preparation is meant to survive for more than one day, provide buprenorphine (0.03 &mu;g per g rodent) for analgesia. We typically image the animal at least one day after the initial implantation.</li>
</ol>
5. Imaging Preparation
<ol>
 <li>Stabilize the animal on an optical breadboard for imaging, using the frame as a head support (Fig. 1L and 1M). A restraint apparatus can be made from commercially available optomechanical components from Qioptiq or ThorLabs. Our separate plate can be transported between surgical and two-photon imaging suites with the animal and all physiological monitoring devices assembled as one unit 3.</li>
 <li>If blood flow imaging is desired, inject 0.05 mL of 5% (w/v) fluorescent-dextran dye dissolved in sterile saline either through the tail vein or infraorbital vein to label the blood serum (Fig. 2A and 2C to 2H) 4-7. This must be done under general anesthesia. Infraorbital vein administration may be easier for beginners, as dextran solutions are more viscous. The tail vein can be difficult to locate in dark colored mice, and the vein tends to collapse after a failed injection. For imaging vasculature with green emission, use a fluorescein isothiocyanate dextran. For red emission, use a Texas Red dextran. With high molecular weight dextrans, the dye will remain in circulation for several hours. Alternatively, animals with endogenous fluorescent labels can be imaged directly at the appropriate two-photon excitation wavelength. The dextran solution can be frozen in aliquots for future use.</li>
 <li>Gently clean the window surface with a moist cotton tipped applicator.</li>
 <li>For prolonged imaging of anesthetized preparations, inject sterile lactated Ringer's solution intraperitoneally at a volume of 3 &mu;L per gram every 2 hours to maintain body fluids and energy requirements.</li>
 <li>When imaging animals in the awake state, limit head restraint to only a few hours at a time to reduce stress levels. Return the animal to the home cage between imaging sessions for food and water.</li>
</ol>
6. Representative Results
A successful window will allow imaging depths up to 250 &mu;m below the pial surface for several months. This method has been used to study in vivo capillary blood flow 4, 8, microglial activation 8, 9, and dendritic structure within the cortical parenchyma 8. In one example, we use two-photon imaging to show the cortical vasculature of an anesthetized Thy1-yellow fluorescent protein (YFP) mouse, after the blood serum is labeled by intravenous injection of Texas Red dextran (Fig. 2A). Dural vessels are often visible slightly above the cortical surface in the dura mater (Fig. 2C, arrow). Large pial arterioles and venules lie on the cortical surface (Fig 2D). Penetrating vessels branch from this surface network and dive into the cortex where they ramify into a dense capillary bed that feeds the cortical tissue (Fig. 2E to 2H). Dendritic arbors of deep YFP expressing cortical neurons, a signal endogenous to this mouse line, can be imaged concurrently in a second channel 10 (Fig. 2B to 2H). The second harmonic signal of the bone was collected in a third channel, and can be used to gauge the thickness of the thinned skull after collection of image stacks (Fig. 2A to 2C).
Cortical vascular dynamics are profoundly affected by anesthetics 11. In a second example, we show a video of spontaneous vasoactivity collected by two-photon microscopy from a habituated awake mouse. Prominent vasomotor oscillations in the lumen diameter are seen with a pial arteriole, but not with a neighboring venule. This basal range of vasoactivity is diminished with urethane anesthesia 4. To quantify spontaneous and evoked changes in blood flow, we use adapted line scanning techniques to capture both the vascular diameter and red blood cell velocity of individual vessels. Detailed resources on quantitative blood flow imaging using two-photon microscopy are available 3, 12.
<img alt="Figure 1" src="/files/ftp_upload/3742/3742fig1.jpg" /> 
Figure 1. Procedure for a PoRTS window. (A to K) Images of sequential steps in the procedure for generating a PoRTS window. See text for detailed instructions. &beta; = bregma and &lambda; = lambda. (L) Bolt and nut system for securing the head during imaging of anesthetized preparations. (M) Custom machined cross bar head mount for awake preparations. In this example, a connector was also implanted for repeated electrocorticogram recordings. (N) Schematic diagram showing dorsal view of the head mount and position of various components. The nut used in panel L is meant as alternative to the cross bar using in panel M. Two #000 self-tapping screws are added with the cross bar mount for added stability with awake imaging preparations. (O) Schematic diagram showing cross section of a PoRTS window.
<img alt="Figure 2" src="/files/ftp_upload/3742/3742fig2.jpg" /> 
Figure 2. Two-photon imaging of vasculature and neuronal structure in mouse cortex. All images were collected through a PoRTS window in a Thy1-YFP mouse at 2 days after window implantation 10. Maximal projection over 150 &mu;m of tissue in the coronal orientation showing the thinned skull in relation to the vasculature (A) and dendrites (B). The bone (blue) was detected by collecting the second harmonic fluorescence at 450 nm emission with 900 nm excitation 8. The vasculature (red) was labeled by intravenously injected 70 kDa Texas Red dextran 6. The dendritic fields of neurons (green) are endogenous to the Thy1-YFP transgenic mouse line. (C-H) Maximal projections over 50 &mu;m of tissue in the horizontal orientation at different depths below the pia. Data is from the same image stack shown in Panels A and B. Dural vessels may be visible just above the cortical surface (arrow in C).
Abbreviations
ACSF = artificial cerebral spinal fluid 
PoRTS = polished and reinforced thinned skull window 
YFP = yellow fluorescent protein
iEnsure that the procedures described are approved by your local Institutional Animal Care and Use Committee.

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Nothing to disclose.

Two-photon imaging through a PoRTS window requires transmission through the thinned bone and the dura, which attenuates the laser light and adds optical aberrations at greater depths 8. However, despite this drawback, imaging depths up to 250 &mu;m below the pial surface can be achieved with 900 nm excitation. Greater imaging depths may in principle be possible with longer excitation wavelengths 13. A major advantage of this method is the absence of cortical inflammation that might exist transiently...

<table border="1">
<tbody>
 <tr>
 <td> Agent</td>
 <td>Route of delivery</td>
 <td>Dose for mouse</td>
 <td>Duration</td>
 <td>Notes</td>
 <td>Source</td>
 <td> Ref Ref</td>
 </tr>
 <tr>
 <td>Pentobarbital (Nembutal) </td>
 <td>IP</td>
 <td>90 &mu;g/g</td>
 <td>15-60 min</td>
 <td>Narrow safety margin. Work up to proper dose of anesthesia slowly. Supplement 10 % of induction dose as required.</td>
 <td>036093; Butler Schein</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Ketamine (Ketaset)
 mixed with
 Xylazine (Anased)</td>
 <td>IP</td>
 <td>60 &mu;g/g (K)
 
 10 &mu;g/g (X) (mix in same syringe)</td>
 <td>20-30 min</td>
 <td>Xylazine is co-injected as a muscle relaxant and analgesic. Supplement only Ketamine at 50% of induction dose as required.</td>
 <td>(K) 010177, (X) 033198; Butler Schein</td>
 <td>7</td>
 </tr>
 <tr>
 <td>Isoflurane (Isothesia) </td>
 <td>Inhalation</td>
 <td>4% mean alveolar concentration (MAC) for induction;
 1-2% MAC for maintenance</td>
 <td>4-6 h. </td>
 <td>Provided in mixture of 70% oxygen and 30% nitrous oxide. Prolonged anesthesia leads to slow recovery.</td>
 <td>029403; Butler Schein</td>
 <td>26</td>
 </tr>
 </tbody>
 </table>
 Table 1. Suggested anesthetic agents for survival studies.
 <table border="1">
 <tbody>
 <tr>
 <td>ITEM</td>
 <td>COMPANY</td>
 <td>CATALOG # / MODEL</td>
 </tr>
 <tr>
 <td>Betadine </td>
 <td>Butler Schein</td>
 <td>6906950</td>
 </tr>
 <tr>
 <td>Buprenorphine (Buprenex) </td>
 <td>Butler Schein</td>
 <td>031919</td>
 </tr>
 <tr>
 <td>Fluorescein isothiocyanate dextran, 2 MDa molecular weight</td>
 <td>Sigma</td>
 <td>FD2000S</td>
 </tr>
 <tr>
 <td>Isopropyl alcohol</td>
 <td>Fisher</td>
 <td>AC42383-0010</td>
 </tr>
 <tr>
 <td>Lactated Ringer's Solution</td>
 <td>Butler Schein</td>
 <td>009846;</td>
 </tr>
 <tr>
 <td>Lidocaine solution, 2 % (v/v) </td>
 <td>Butler Schein</td>
 <td>002468</td>
 </tr>
 <tr>
 <td>Saline </td>
 <td>Butler Schein</td>
 <td>009861</td>
 </tr>
 <tr>
 <td>Surgical Milk</td>
 <td>Butler Schein</td>
 <td>014325</td>
 </tr>
 <tr>
 <td>Texas Red dextran, 70 kDa molecular weight</td>
 <td>Invitrogen</td>
 <td>D1864</td>
 </tr>
 <tr>
 <td>Maxizyme</td>
 <td>Butler Schein</td>
 <td>035646</td>
 </tr>
 <tr>
 <td>DISPOSABLES</td>
 </tr>
 <tr>
 <td>Carbide burrs, 1/2 mm tip size</td>
 <td> Fine Science Tools</td>
 <td>19007-05</td>
 </tr>
 <tr>
 <td>Cottoned tip applicators </td>
 <td>Fisher Scientific</td>
 <td>23-400-100</td>
 </tr>
 <tr>
 <td>Cover Glass, no.&nbsp;0 thickness </td>
 <td>Thomas Scientific</td>
 <td>6661B40</td>
 </tr>
 <tr>
 <td>Cyanoacrylate glue </td>
 <td>ND Industries</td>
 <td>31428 H04308</td>
 </tr>
 <tr>
 <td>Gas duster</td>
 <td>Newegg</td>
 <td>N82E16848043429</td>
 </tr>
 <tr>
 <td>Grip cement powder </td>
 <td>Dentsply</td>
 <td>675571</td>
 </tr>
 <tr>
 <td>Grip cement solvent</td>
 <td>Dentsply</td>
 <td>675572</td>
 </tr>
 <tr>
 <td>Insulin syringe, 0.3 mL volume with 29.5 gauge needle </td>
 <td>Butler Schein</td>
 <td>018384</td>
 </tr>
 <tr>
 <td>Nut and bolt to secure the head </td>
 <td>Digikey</td>
 <td>Nut, H723-ND; bolt, R2-56X1/4-ND</td>
 </tr>
 <tr>
 <td>Opthalmic ointment </td>
 <td>Butler Schein</td>
 <td>039886</td>
 </tr>
 <tr>
 <td>Scalpel blades </td>
 <td>Fisher Scientific</td>
 <td>12-460-448</td>
 </tr>
 <tr>
 <td>Screws, self-tapping #000 </td>
 <td>J.I. Morris Company</td>
 <td>FF000CE125</td>
 </tr>
 <tr>
 <td>Silicone aquarium sealant</td>
 <td>Perfecto Manufacturing</td>
 <td>31001</td>
 </tr>
 <tr>
 <td>Tin oxide powder</td>
 <td>Mama's Minerals</td>
 <td>EQT-TINOX</td>
 </tr>
 <tr>
 <td>EQUIPMENT</td>
 </tr>
 <tr>
 <td>Glass scribe</td>
 <td>Fisher Scientific</td>
 <td>08-675</td>
 </tr>
 <tr>
 <td>Dissecting microscope </td>
 <td>Carl Zeiss</td>
 <td>OPMI-1 FC</td>
 </tr>
 <tr>
 <td>Electric powered drill</td>
 <td> Foredom or Osada</td>
 <td>K.1020 (Foredom) or EXL-M40 (Osada)</td>
 </tr>
 <tr>
 <td>Electrical razor</td>
 <td>Wahl</td>
 <td>Series 8900</td>
 </tr>
 <tr>
 <td>Forceps, Dumont no. 55 </td>
 <td>Fine Science Tools</td>
 <td>11255-20</td>
 </tr>
 <tr>
 <td>Feedback regulated heat pad</td>
 <td>FHC</td>
 <td>40-90-8 (rectal thermistor, 40-90-5D-02; heat pad, 40-90-2-07)</td>
 </tr>
 <tr>
 <td>Isoflurane vaporizer </td>
 <td>Ohmeda</td>
 <td>IsoTec4</td>
 </tr>
 <tr>
 <td>Pulse oximeter</td>
 <td>Starr Life Sciences</td>
 <td>MouseOx</td>
 </tr>
 <tr>
 <td>Screwdriver, miniature </td>
 <td>Garret Wade</td>
 <td>26B09.01</td>
 </tr>
 <tr>
 <td>Stereotaxic frame</td>
 <td>Kopf Instruments</td>
 <td>Model 900 (with mouse anesthesia mask and non-rupture ear bars)</td>
 </tr>
 <tr>
 <td>Surgical scissors, blunt end </td>
 <td>Fine Science Tools</td>
 <td>14078-10</td>
 </tr>
 <tr>
 <td>Ultrasonic cleaner </td>
 <td>Fisher Scientific</td>
 <td>15-335-30</td>
 </tr>
 </tbody>
 </table>
 Table 2. List of specific reagents, disposables and equipment.

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.

Watch this Scientific Journal Video about A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain at JoVE.com

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

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 ...

a-polished-reinforced-thinned-skull-window-for-long-term-imaging

Research

JoVE Journal

Neuroscience

26.8K 조회수.  University of California, San Diego. 우리는 mm에 걸쳐하고 뇌의 염증없이 몇 달 동안 안정 마우스 두개골에 이미지 창을 형성하는 방법을 제시한다. 이 방법은 잘 두 광자 현미경을 사용하여 혈류, 세포 역학 및 세포 / 혈관 구조의 종방향 연구에 가장 적합합니다.

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

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 ...

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, inflammatory and metabolic conditions, using in vivo fluorescence microscopy. A closed cranial window is placed over the left parieto-occipital cortex of the mice. Local microcirculation is recorded in real time through the window using epi and fluorescence illumination, and measurements of vessels diameters and red blood cell (RBC) velocities are performed. RBC velocity is measured using real-time cross-correlation and/or fluorescent-labeled erythrocytes. Leukocyte and platelet adherence to pial vessels and assessment of perfusion and vascular leakage are made with the help of fluorescence-labeled markers such as Albumin-FITC and anti-CD45-TxR antibodies. Microcirculation can be repeatedly video-recorded over several days. We used for the first time the close window brain intravital microscopy to study the pial microcirculation to follow dynamic changes during the course of Plasmodium berghei ANKA infection in mice and show that expression of CM is associated with microcirculatory dysfunctions characterized by vasoconstriction, profound decrease in blood flow and eventually vascular collapse.

Intravital microscopy to follow temporal and spatial hemodynamic and inflammatory events in the pial microcirculation.

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, ...

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Extinction Training During the Reconsolidation Window Prevents Recovery of Fear

Fear is maladaptive when it persists long after circumstances have become safe. It is therefore crucial to develop an approach that persistently prevents the return of fear. Pavlovian fear-conditioning paradigms are commonly employed to create a controlled, novel fear association in the laboratory. After pairing an innocuous stimulus (conditioned stimulus, CS) with an aversive outcome (unconditioned stimulus, US) we can elicit a fear response (conditioned response, or CR) by presenting just the stimulus alone1,2 . Once fear is acquired, it can be diminished using extinction training, whereby the conditioned stimulus is repeatedly presented without the aversive outcome until fear is no longer expressed3. This inhibitory learning creates a new, safe representation for the CS, which competes for expression with the original fear memory4. Although extinction is effective at inhibiting fear, it is not permanent. Fear can spontaneously recover with the passage of time. Exposure to stress or returning to the context of initial learning can also cause fear to resurface3,4.
Our protocol addresses the transient nature of extinction by targeting the reconsolidation window to modify emotional memory in a more permanent manner. Ample evidence suggests that reactivating a consolidated memory returns it to a labile state, during which the memory is again susceptible to interference5-9. This window of opportunity appears to open shortly after reactivation and close approximately 6hrs later5,11,16, although this may vary depending on the strength and age of the memory15. By allowing new information to incorporate into the original memory trace, this memory may be updated as it reconsolidates10,11. Studies involving non-human animals have successfully blocked the expression of fear memory by introducing pharmacological manipulations within the reconsolidation window, however, most agents used are either toxic to humans or show equivocal effects when used in human studies12-14. Our protocol addresses these challenges by offering an effective, yet non-invasive, behavioral manipulation that is safe for humans.
By prompting fear memory retrieval prior to extinction, we essentially trigger the reconsolidation process, allowing new safety information (i.e., extinction) to be incorporated while the fear memory is still susceptible to interference. A recent study employing this behavioral manipulation in rats has successfully blocked fear memory using these temporal parameters11. Additional studies in humans have demonstrated that introducing new information after the retrieval of previously consolidated motor16, episodic17, or declarative18 memories leads to interference with the original memory trace14. We outline below a novel protocol used to block fear recovery in humans.

Conditioned fear can be diminished through an inhibitory process called extinction, but can resurface under conditions such as the passage of time or exposure to stress. Our protocol presents a novel way of preventing fear recovery by introducing extinction during the reconsolidation window (the re-storage phase of a reactivated memory).

Fear is maladaptive when it persists long after circumstances have become safe. It is therefore crucial to develop an approach that persistently prevents ...

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

Optical coherence tomography (OCT) is a biomedical imaging technique with high spatial-temporal resolution. With its minimally invasive approach OCT has been used extensively in ophthalmology, dermatology, and gastroenterology1-3. Using a thinned-skull cortical window (TSCW), we employ spectral-domain OCT (SD-OCT) modality as a tool to image the cortex in vivo. Commonly, an opened-skull has been used for neuro-imaging as it provides more versatility, however, a TSCW approach is less invasive and is an effective mean for long term imaging in neuropathology studies. Here, we present a method of creating a TSCW in a mouse model for in vivo OCT imaging of the cerebral cortex.

Thinned-skull Cortical Window Technique for In Vivo Optical Coherence Tomography Imaging

We present a method of creating a thinned-skull cortical window (TSCW) in a mouse model for in vivo OCT imaging of the cerebral cortex.

Optical coherence tomography (OCT) is a biomedical imaging technique  with high spatial-temporal resolution. With its minimally invasive approach OCT  has ...

Improved Preparation and Preservation of Hippocampal Mouse Slices for a Very Stable and Reproducible Recording of Long-term Potentiation

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory encoding. LTP elicited in the CA1 region of acute hippocampal slices has been extensively studied. However the molecular mechanisms underlying the maintenance phase of this phenomenon are still poorly understood. This could be partly due to the various experimental conditions used by different laboratories. Indeed, the maintenance phase of LTP is strongly dependent on external parameters like oxygenation, temperature and humidity. It is also dependent on internal parameters like orientation of the slicing plane and slice viability after dissection.
The optimization of all these parameters enables the induction of a very reproducible and very stable long-term potentiation. This methodology offers the possibility to further explore the molecular mechanisms involved in the stable increase in synaptic strength in hippocampal slices. It also highlights the importance of experimental conditions in in vitro investigation of neurophysiological phenomena.

This paper presents a complete methodology to prepare and preserve in vitro acute hippocampal slices from adult mice. This protocol allows the recording of very stable long-lasting long-term potentiation (LTP) for more than 8 hours with a success rate of 95%.

Long-term potentiation (LTP) is a type of synaptic plasticity characterized by an increase in synaptic strength and believed to be involved in memory ...

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 ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure the activity of neurons during various behaviors. To correlate neural activity with behavior, the animal should not be immobilized but should be able to move. Many behavioral changes occur during long time scales and require recording over many hours of behavior. This also makes it necessary to culture the worms in the presence of food. How can worms be cultured and their neural activity imaged over long time scales? Agarose Microchamber Imaging (AMI) was previously developed to culture and observe small larvae and has now been adapted to study all life stages from early L1 until the adult stage of C. elegans. AMI can be performed on various life stages of C. elegans. Long-term calcium imaging is achieved without immobilizing the animals by using short externally triggered exposures combined with an electron multiplying charge-coupled device (EMCCD) camera recording. Zooming out or scanning can scale up this method to image up to 40 worms in parallel. Thus, a method is described to image behavior and neural activity over long time scales in all life stages of C. elegans.

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Imaging behavior and neural activity over long time scales without immobilization of the animal is a prerequisite to understand behavior. Agarose microfluidic chambers imaging (AMI) can be used to image neural activity and behavior for all life stages of Caenorhabditis elegans.

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure ...

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Subtitles

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