Name: Chronic Imaging of Mouse Visual Cortex Using a Thinned-skull Preparation (Video) | JoVE Journal
Uploaded: 2010-10-25T17:00:00.000+00:00
Duration: 11 min 12 s
Description: Learn chronic imaging of mouse visual cortex using a thinned-skull preparation to observe dendritic spine dynamics. This method enables researchers to study cortical plasticity mechanisms in vivo over time.

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5 minutes

This work was supported by Burroughs Wellcome Career Award in the Biomedical Sciences (A.K.M), The Whitehall Foundation Research Grant (A.K.M), Sloan Foundation Fellowship (A.K.M), NIH EY019277 (A.K.M), and an NEI funded, Vision Training Grant, EY013319 (E.A.K.).

<ol>
 <li>To visualize neurons in the intact brain using two-photon microscopy, a preparation where neurons are labeled with fluorescent markers is used. In the experiments presented here we use the GFP-M transgenic mouse line which labels layer 5 pyramidal cells1. Layer 5 pyramidal cells project dendritic processes into superficial layers, allowing the visualization of dendrites and dendritic spines up to a depth of 300 μm below the level of the pia. An alternate approach is to label cells using viral markers (see Lowery et al., submitted to JOVE).</li>
 <li>Sterilize the workspace using 70% ethanol and autoclave tools for aseptic surgery. Cover operating surface with clean dressing.</li>
 <li>Anesthetize mice with a Fentanyl cocktail (fentanyl 0.05 mg/kg; midazolam 5 mg/kg; metatomadin 0.5 mg/kg) followed by a quarter dose of Avertin (0.0075 mg/ml) using IACUC approved procedures. Pretreat mice with the analgesic, buprenorphine (0.5 mg/kg) administered subcutaneously (SQ). Monitor the level of anesthetic plane by testing reaction to tail/toe pinch.</li>
 <li>Maintain animal at 37.5°C using heating blanket with attached rectal thermometer.</li>
 <li>Apply ophthalmic ointment to eyes to prevent the eyes from drying out. Hood eyes with small sheet of paper to protect eyes from light source during surgery.</li>
 <li>Using scissors, remove hair from the back of head (behind ears) to the eyes.</li>
 <li>Clean the top of animal's head with sequential application of 70% ethanol, followed by betadine scrub and betadine solution.</li>
 <li>Make an L-cut incision behind the ears and up along either the right or left side of the midline (depending on which hemisphere will be imaged) to the eyes. Fold the skin over and away from surgery site. Do not remove skin as the skin will be sutured following the imaging session.</li>
 <li>Using fine forceps gently pull the skin up and away from your area of interest.</li>
 <li>Gently scrape away the periosteum from the exposed area of the skull with clean forceps or microsurgical blade. Using cotton swab, apply a small amount of 10% Ferric Chloride to the skull to completely dry the periosteum membrane and ensure that it has been removed completely. It is important that the skull is completely dry to ensure proper adhering of the glue (see 11). Using forceps or microsurgical blade, gently scrape away dried periosteum membrane.</li>
 <li> Apply a thin layer of cyanoacrylate glue around the imaging window of the stainless steel head plate (see Figure 1) and affix the plate to the skull.</li>
 <li>Using the wooden end of a cotton-swab stick, apply glue to inside seams of the imaging window; making sure to fill all gaps between the imaging plate and curvature of skull. </li>
 <li>Place a drop of glue accelerator into the window. Quickly wipe away excess glue accelerator with a cotton swab or kim wipe. The glue should be completely solidified before drilling.</li>
 <li>Begin to thin the skull in the imaging window using a dental drill affixed with a 0.7 mm stainless steel burr. Alternate drilling with occasional application of 0.9% sterile saline to avoid generating too much heat on the skull. </li>
 <li>Thin a 4x4 mm window on the skull using small strokes across the skull surface with the dental drill. Test the thinness of the skull with blunt end forceps. The skull surface will indent slightly with gentle pressure. The optimal skull thickness for imaging is between 10 and 30 μm.</li>
 <li>When the skull is thin and the window cleaned of all debris, place saline on the skull. Photograph the imaging window with a digital camera mounted to a dissecting scope. Photography and use of the brain vasculature will aid in locating the position to place the imaging plate for subsequent imaging sessions. </li>
 <li>Transport the animal to the 2-photon rig. The animal should remain on the heating blanket (with rectal probe) during the entire imaging session. Note: while under sedation, the animal's core body temperature will fluctuate greatly in the absence of a heating unit resulting in cortical damage or possibly death.</li>
 <li>A two-photon microscope with a Mai Tai laser, a 10W solid state pump for maximum power and a modified Olympus Fluoview confocal unit is used. The system is optimized for deep tissue imaging and external detectors are installed as close to the objective as possible to allow maximal detection of fluorescence from the brain. An Olympus LUMPlan fl/IR 20X/0.95NA objective lens is used. 0.9% saline is used during imaging for objective submersion.</li>
 <li>Using this microscope, and low digital zoom (x1) identify an area containing brightly labeled neurons.</li>
 <li>Using a digital camera affixed to the microscope eye piece, take a photograph of the imaging area. Alternatively, sketch out the blood vasculature pattern. This is necessary to return to the same imaging location at a subsequent time point.</li>
 <li>Obtain a low magnification stack (x20 objective, 512x512 pixel; 5 μm step) through the entire visible extent of the brain (usually spanning ~200 μm) showing the extent of dendritic ramification. Note: this low magnification image will be used to locate the imaging area at subsequent time points as both blood vessels and dendrites maintain their structure in young and adult animals.</li>
 <li>Using imaging software (Fluoview, v. 5.0, FV300), increase digital magnification 8-fold. Adjust the X,Y coordinates (pan coordinates) if necessary and collect a high magnification stack which will show detailed dendritic morphology including the locations and structure of dendritic spines. Take detailed notes regarding each stack collection, i.e. pan coordinates, the range of collection, the z-step size and the number of steps in your stack. These notes will be used when returning to this dendrite or axon at a subsequent time point.</li>
 <li>Following imaging, remove the head plate by gently separating plate from skull surface. Using forceps, remove any residual glue remaining on skull and wipe skull surface with 0.9% sterile saline. Suture the skin over the skull together using #6 suturing thread. Wipe area with 70% ethanol. To alleviate pain during post-operative recovery, administer 0.1 mg/kg Buprenorphine analgesic, SQ.</li>
 <li>Place animal in clean cage under a heat lamp until it wakes up and is mobile. Return animal to original cage.</li>
 <li>To reimage animal at a later time point (our imaging time points can be anywhere from 2 days to many months apart), remove stitches or reopen the skin on head using aseptic methods as above. Use the digital image of brain vasculature (taken on first surgery date) to locate the original imaging location. Reaffix stainless steel headplate (steps 10-12). Use a microsurgical blade to gently thin away any bone that might have re-grown between imaging time points. If necessary, thin with dental drill. Clear imaging window of debris and apply a drop of 0.9% sterile saline.</li>
 <li>Carry animal to two-photon microscope and locate original imaging field. Use the digital photograph taken during first imaging session for assistance.</li>
 <li>Open the Fluoview program and open the original low magnification image. Affix a transparency sheet to the computer screen and sketch the major blood vessel pattern using a transparency pen.</li>
 <li>Open the Live image and realign the blood vasculature to the sketched pattern on the transparency film.</li>
 <li>To reimage the 8x zoomed in structures, open up the desired collected stack from the previous imaging session. Sketch the structure (i.e. axonal, dendritic, glial) onto a transparency film, affixed to the computer screen. </li>
 <li>Zoom in 8x from the Live Image and set the pan coordinates to the first day imaging parameters. Depending on the accuracy of aligning the low magnification location, it may be necessary to also adjust the angle of the image slightly. Once the desired structure is found, match up the structure to your sketched high magnification pattern. Collect the z-stack with the identical step size and number of z-stack images for subsequent analysis. </li>
 <li>Repeat steps 29 and 30 until all the high magnification images are re-collected. </li>
 <li>When imaging is complete, repeat steps 22-24. Re-imaging can be repeated at up to 2 separate time points.</li>
 <li>Experiments on animals were performed in accordance with the guidelines and regulations set forth by the Vivarium and Division of Laboratory Animal Medicine of the University of Rochester School of Medicine and Dentistry, in full accreditation by the Association for the Assessment and Accreditation of Laboratory Animal Care, International (AAALAC) and are in compliance with state law, federal statute and NIH policy.</li>
</ol>
Representative Results
In GFP-M expressing mice, a subset of layer 5 pyramidal cells and corresponding dendritic processes can easily be visualized using 2-photon laser scanning microscopy (2PLSM)1. Here we demonstrated a thinned-skull preparation in which the skull over the desired imaging location is thinned to ~10-30 μm. Performed correctly, our protocol allowed us to clearly visualize cortical dendrites and spines in the intact visual cortex for acute or chronic analysis (Figure 2).
<img src="/files/ftp_upload/2060/2060fig1.jpg" alt="Microfluidic chip fabrication shown in four views: assembly images (A, B, C) and technical diagram (D)." data-altupdatedat="1788235205000" data-alt="Figure 1"> Figure 1. Custom head and base plate. (A) Top view of custom made head and base plate used for thinned-skull surgery preparation. (B) Side view of custom-made head and base plate. (C,D) Detailed schematic and image of head plate.
<img src="/files/ftp_upload/2060/2060fig2.jpg" alt="Blood vessel growth over 4 days, microscopy images showing vascular changes, labeled by days." data-altupdatedat="1788235205000" data-alt="Figure 2"> Figure 2. Representative results from 2PLSM following thin skull preparation. (A) Dendrites imaged in vivo in the visual cortex of a transgenic GFP-M mouse using two-photon microscopy. Image is a projection of single image obtained every 5 μm from the pia to a final depth of 175 μm. The boxed area in A represents the magnified region in (B) and (C) imaged on day 0 (D0) and four days later (D4). (D,E) Higher magnification of dendritic branch boxed in B and C demonstrating stable spines (white arrow heads), lost/retracting spines (red arrow heads) and new spines (green arrow heads). Scale bars= A. 200 μm, B,C. 20 μm, D,E. 5 μm.

<ol><li>Feng, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Imaging+neuronal+subsets+in+transgenic+mice+expressing+multiple+spectral+variants+of+GFP%5BTitle%5D)+AND+%22Neuron%22%5BJournal%5D)">Imaging neuronal subsets in transgenic mice expressing multiple spectral variants of GFP.</a> Neuron. 28 (1), 41-51 (2000).</li>
<li>Holtmaat, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((high-resolution+imaging+in+the+mouse+neocortex+through+a+chronic+cranial+window%5BTitle%5D)+AND+%22Nat+Protoc%22%5BJournal%5D)">high-resolution imaging in the mouse neocortex through a chronic cranial window.</a> Nat Protoc. 4 (8), 1128-1144 (2009).</li>
<li>Majewska, A. K., Newton, J. R., Sur, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Remodeling+of+synaptic+structure+in+sensory+cortical+areas+in+vivo%5BTitle%5D)+AND+%22J+Neurosci%22%5BJournal%5D)">Remodeling of synaptic structure in sensory cortical areas in vivo.</a> J Neurosci. 26 (11), 3021-3029 (2006).</li>
<li>Mostany, R., Portera-Cailliau, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+Craniotomy+Surgery+Procedure+for+Chronic+Brain+Imaging%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">A Craniotomy Surgery Procedure for Chronic Brain Imaging.</a> J Vis Exp. , (2008).</li>
<li>Yang, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Thinned-skull+cranial+window+technique+for+long-term+imaging+of+the+cortex+in+live+mice%5BTitle%5D)+AND+%22Nat+Protocols%22%5BJournal%5D)">Thinned-skull cranial window technique for long-term imaging of the cortex in live mice.</a> Nat Protocols. 5 (2), 201-208 (2010).</li>
<li>Xu, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Choice+of+cranial+window+type+for+in+vivo+imaging+affects+dendritic+spine+turnover+in+the+cortex%5BTitle%5D)+AND+%22Nat+Neurosci%22%5BJournal%5D)">Choice of cranial window type for in vivo imaging affects dendritic spine turnover in the cortex.</a> Nat Neurosci. 10 (5), 549-551 (2007).</li>
</ol>

No conflicts of interest declared.

Chronic imaging using two-photon microscopy is becoming an increasingly popular technique to study morphological changes triggered during plasticity2-5. Here we demonstrate a thinned-skull preparation to follow identified dendritic spines in the intact mouse brain on different imaging days. In this protocol, the skull is left intact, causing minimal damage to the cortex and resulting in very low levels of neuroinflammation which may alter brain function6. This allows the animal to be imaged immedia...

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chronic imaging of mouse visual cortex using a thinned-skull preparation

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 ...

Dendritic spines are membrane protrusions from a neuron's dendrite that receive synaptic input. Lengthening, shrinking appearance or disappearance of spines occurs in response to changes in neural network function that result from specific patterns of neural activity. This phenomenon is known as cortical plasticity to observe morphological changes in dendritic spines.In live animals, an imaging plate is affixed to a mouse's skull. An area of the skull is surgically thinned and images are required at low and high magnification. The areas are then re-image at later time points.The results obtained show that dendritic spine morphology can be clearly visualized using a thin skull preparation on multiple time points. And that spine morphology is highly plastic, showing changes in structure and also the appearance and disappearance of dendritic spines. Hi, I'm Emily Kelly in the laboratory of Anaya Maka in the Department of Neurobiology and Anatomy at the University of Rochester.Today we'll show you procedure for chronic imaging of the mouse vial cortex following a fins called preparation. We use this procedure in our laboratory to study dendritic spine turnover. So let's get started.Begin the procedure by sterilizing the tools for aseptic surgery. Then sterilize the workspace using 70%ethanol and cover the operating surface with clean dressing. Monitor the depth of anesthesia in a fentanyl cocktail anesthetized mouse by testing its response to a toe pinch the mouse shown here is A-G-F-P-M transgenic mouse in which GFP is expressed.In layer five parametal cells that project dendritic processes into superficial layers. To maintain a body temperature of 37.5 degrees Celsius, place the animal on a heating blanket and monitor it with an attached rectal thermometer. Then apply ophthalmic ointment to the eyes to prevent them from drying out using scissors.Remove the hair from the back of the mouse's head from behind the ears to the eyes. Then clean the top of the animal's head with an application of 70%ethanol, followed by Betadine scrub and Betadine solution. Make an LCU incision behind the ears and up along either the right or left side of the midline, depending on which hemisphere will be imaged to the eyes, fold the skin over and away from the surgery site.Do not remove the skin since it will be sutured later using fine forceps. Gently pull the skin up and away from the area of interest. Using clean forceps, gently scrape away the periosteum from the exposed area of the skull.Apply a small amount of 10%fair chloride to the skull to completely dry the periosteum membrane and ensure that it has been removed completely. It is important that the skull is completely dry. To ensure proper adhering of the glue, gently scrape away dried periosteum membrane with a microsurgical blade.Next, apply a thin layer of Sano acrylic glue around the imaging window of the stainless steel head plate and affix the plate to the skull. With the wooden end of a cotton swab stick, apply glue to the inside seams of the imaging window, making sure to fill all the gaps between the imaging plate and the curvature of the skull. Place a drop of the glue accelerator into the window.Then quickly wipe away the excess. Allow the glue to dry once the glue has completely dried. Using a dental drill affixed with a 0.7 millimeter stainless steel burr, begin to thin the skull in the imaging window.Alternate drilling with occasional application of 0.9%sterile saline. To avoid generating too much heat on the skull. Thin a four by four millimeter window on the skull using small strokes across the skull surface.With the dental drill. Test the thinness of the skull with blunt end forceps. The skull surface will indent slightly with gentle pressure.The optimal skull thickness for imaging is between 10 and 30 microns. Once the skull is thinned, clean the window of all debris and place saline on the skull. Photograph the imaging window with a digital camera mounted to a dissecting scope.The brain vasculature in this photograph will aid in locating the position to place the imaging plate for subsequent imaging sessions. Transport the animal to the two photon rig. The animal should remain on the heating blanket and body temperature should be monitored continuously throughout the imaging session.A two photon microscope with a mi tai laser. A 10 watt solid state pump for maximum power and a modified Olympus flow view confocal unit are used. The system is optimized for deep tissue imaging and external detectors are installed as close to the objective as possible to allow for maximum detection of fluorescence from the brain, add a drop of saline to the imaging window using low digital zoom.Identify an area containing brightly labeled neurons using a digital camera fixed to the microscope. Ips, take a photograph of the imaging area. This is necessary to return to the same imaging location at a subsequent time point.Obtain a low magnification stack through the entire visible extent of the brain showing the extent of dendritic ramification. This will be used to locate the imaging area at subsequent time points as both blood vessels and dendrites. Maintain their structure in young and adult animals using imaging software.Increase the digital magnification eightfold, adjust the XY coordinates if necessary, and collect a high magnification stack, which will show detailed dendritic morphology including the locations and structure of dendritic spines. Take detailed notes regarding each stack collection, including pan coordinates, the range of collection, the Z step size, and the number of steps in the stack. These notes will be used when returning to this dendri or axon at a subsequent time point.Following imaging, remove the head plate by gently separating it from the skull surface. Using forceps. Remove any residual glue and wipe the skull with 0.9%sterile saline.Suture the skin over the skull together using number six suture thread. Wipe the area with 70%ethanol following the imaging procedure. Place the animal in a clean cage under a heat lamp until it wakes up and is mobile.Then return the animal to its original cage to image the animal again at a later time point anywhere from two days to many months later, remove the stitches and reopen the skin on the head. Using aseptic methods, use the digital image of the brain vasculature taken during the first surgery to locate the original imaging location. Then re-fix the head plate as before.Use a microsurgical blade to gently thin away any bone that might have regrown between imaging time points if necessary. Thin with a dental drill, clear the imaging window of debris and apply a drop of 0.9%sterile saline. Carry the animal to a two photon microscope and locate the original imaging area using the digital fluorescent image taken during the previous session.Start the flow view program. Open the original low magnification image, affix a transparency sheet to the computer screen and sketch the major blood vessel pattern using a transparency pen. Next, open the live image and realign the blood vasculature to the sketched pattern on the transparency film.Remove the transparency to to re-image the eight x zoomed in structures. Open up the desired collected stack from the previous imaging session. Sketch the structure onto a transparency film affixed to the computer screen.Zoom in eight x from the live image and align the image with the transparency. Depending on the accuracy of aligning the low magnification location, it may be necessary to also adjust the angle of the image slightly set the pan coordinates to the first day's Imaging parameters, including step size and number of images collect the Zack re-imaging can be performed twice. The number of times the scalp is opened should be limited to avoid disrupting the integrity of the skull, which will result in poor imaging resolution in GFPM expressing mice.A subset of layer five parametal cells and corresponding dendritic processes were visualized using in vivo two photon laser scanning microscopy here, GFP labeled visual cortex dendrites are shown. This image is a projection of a single image obtained every five microns from the PIA to a final depth of 175 microns. The boxed region is shown at higher magnification here on day zero, and again on day four here, higher magnification of a dendritic branch image from day zero and day four compared.These images demonstrate stable spines indicated by the white arrow heads lost retracting spines, which are marked by red arrowhead and new spines indicated by the green arrowhead. We've just shown you how to perform chronic two photon imaging in mouse visual cortex using a thin skull preparation. When doing this procedure, it's important to remember to take special care during the thinning process to avoid damage to the skull and underlying dura as this will result in poor imaging resolution and take detailed notes.So that's it. Thanks for watching and good luck with your experiments.

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Research

JoVE Journal

Neuroscience

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Acute brain trauma is a severe injury that has no adequate treatment to date. Multiphoton microscopy allows studying longitudinally the process of acute brain trauma development and probing therapeutical strategies in rodents. Two models of acute brain trauma studied with in vivo two-photon imaging of brain are demonstrated in this protocol.

Acute Brain Trauma in Mice Followed By Longitudinal Two-photon Imaging

This manuscript describes a procedure to track the remodeling of the cerebrovasculature during amyloid plaque accumulation in vivo using longitudinal two-photon microscopy. A thinned-skull preparation enables the visualization of fluorescent dyes to assess the progression of cerebrovascular damage in a mouse model of Alzheimer's disease.

Longitudinal In Vivo Imaging of the Cerebrovasculature: Relevance to CNS Diseases

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

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

Learn chronic imaging of mouse visual cortex using a thinned-skull preparation to observe dendritic spine dynamics. This method enables researchers to study cortical plasticity mechanisms in vivo over time.

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

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