We thank the Scientific Computing and Imaging Institute of the University of Utah for use of FluoRender software (R01GM09815). This work was supported by grants from the Glaucoma Research Foundation, Melsa M. and Frank Theodore Barr Foundation and the US National Institute of Health, (R01EY020878 and R01EY023621) to M.L.V., and (R01EY017182 and R01EY017950) to B.K.A.

In vivo imaging is performed in pathogen-free facilities using protocols approved by the Institutional Animal Care and Use Committee at the University of Utah. 
NOTE: This imaging protocol is used for reporter mice in which retinal microglia and infiltrating monocytes/macrophages express green-fluorescent protein (GFP) under the control of the fractalkine receptor locus (CX3CR1).
1. In Vivo Imaging of Retinal GFP+ Microglia by Confocal Scanning Laser Ophthalmoscopy (cSLO)
<ol>
	<li>Turn on the water-controlled heating system, set to stabilize mouse temperature between 35-37 &#186;C during procedure, and connect two heating pads.
	<ol>
		<li>Start the confocal scanning laser ophthalmoscope (cSLO) system (Figure 2A). Open the cSLO program, enter the information that will identify the individual mouse and set a corneal curvature of 2 mm (Patient Data menu).</li>
	</ol>
	</li>
	<li>Prepare for imaging. Securely fit a clean 55&#186; wide-field objective lens on its socket. Prepare the ophthalmoscope imaging platform by securing a heating pad covered with clean paper. Use clips to flatten the pad and paper and keep them from blocking the movement of the objective lens.</li>
	<li>Anesthetize the mouse by intraperitoneal injection of 1.3% 2,2,2-tribromoethanol and 0.8% tert-amyl alcohol (250 mg/kg body weight; 0.5 ml/25 g body weight) using a 30&#189; G needle fitted to a 1 ml disposable syringe. Return mouse to its cage, kept over a heating pad. 
	NOTE: The delivery of the anesthetic by injection versus inhalation allows easier positioning of the mouse for imaging and unobstructed access to the eyes, free of nose cones and tubing.</li>
	<li>Once the animal stops moving and is unresponsive to tail pinch, induce pupil dilation with Tropicamide and phenylephrine (0.5% each), by positioning the animal prone and covering both eyes with the drops for 7 min.</li>
	<li>After 5 min, place the mouse prone on the center of the ophthalmoscope platform, and fit contact lenses over both eyes, applying minimal pressure. 
	NOTE: Contact lenses are small and fragile but can be carefully handled with fingers, although forceps can be used instead40. The use of contact lenses is optional, but recommended as it minimizes eye dryness and preserves corneal transparency during imaging.</li>
	<li>After 7 min, orient the mouse with the right eye facing the objective lens and the orbit parallel to the objective lens, keeping the animal unrestrained near the edge of the platform (Figure 2B). To collect images of comparable saturation and orientation, constantly maintain the distance from eye to objective lens, as well as the alignment of the eye to the light path throughout imaging sessions. Clip long whiskers interfering with observation of the eye. 
	NOTE: For the next 8-10 min, the mouse will not move or blink, and will breath with gentle movements, which are tracked by the cSLO Eye Tracking ART (automatic real time) mode, which adjusts the scan head position based on fundus landmarks with high contrast. Past that time, mice begin panting and blinking, making imaging impractical.</li>
	<li>If imaging is performed without using contact lenses, apply PBS drops to both eyes every 2-3 min to prevent them from drying.</li>
	<li>Collect a fundus image of the retinal vasculature and optic disc, focused on the inner retina.
	<ol>
		<li>Select the infrared mode (IR) and adjust settings to 820 nm excitation, 100% laser power, 40-60% sensitivity (Figure 2C).</li>
		<li>Working in high-speed mode (12.6 frame/sec), locate the eye using the joystick to drive the ophthalmoscope. At low magnification, inspect the cornea and lens for injury or opacity, and exclude from the study eyes with defects or injury that will affect imaging the retina (Figure 3A).</li>
		<li>Locate the optic disc area (the surface of the optic nerve head, ONH) by bringing the objective closer to the eye, then center the image on it and lock this position by screwing the joystick. This alignment of the eye to the ophthalmoscope is key to obtain even focus and emission across the image.</li>
		<li>Visualize the inner planes of the retina, as well as the ONH, using the major blood vessels located on the vitreal surface of the retina as a reference focal plane (59 and 60 D), or deeper (55 D) in eyes showing excavated optic discs. The optimal focus should resolve circulating blood cells within major blood vessels, with their lumen and walls clearly distinct.</li>
		<li>Optimize image saturation by adjusting the dial on the touch screen control panel, until a white halo of uniform illumination spans in most of the 55&#186; fundus field around the optic disc (using slight overexposure). Next, lower the saturation to optimize contrast until blood cells moving along the major vessels can be clearly resolved. If focal dark areas persist, not due to retinal damage, realign the eye to the camera until a uniformly bright image is obtained. This adjustment is fundamental to capture reproducible sets of images in position and quality.</li>
		<li>Collect a high-resolution IR image of the central retina (1.4-1.7 mm in diameter, depending on age), averaging 30 frames in real time (4.7 frames/sec; normalized), to improve signal-to-noise ratio (Figure 2D).</li>
	</ol>
	</li>
	<li>Immediately, acquire a fluorescence image of GFP+ microglia and/or infiltrating monocytes localized to the inner planes of the retina and the ONH. 
	NOTE: Optimization of the IR fundus image of the retina determines the resolution of the fluorescence image of GFP+ cells, as well as the extent of inner retina spanned. Failure to focus the inner planes of the retina, which can be detected by poor resolution of the vasculature, will result in dimmed GFP+ cell views (Figure 3B). Failure to adjust IR saturation correctly and uniformly affects the fluorescence image (FA), introducing artifactual variations in GFP+ cell intensity and size (Figure 3C).
	<ol>
		<li>Swich to fluorescence imaging mode (FA) in the touch screen panel, using blue, 488 nm laser excitation (460-490 nm barrier filter set), and acquisition settings (100% laser power and 100-125% sensitivity), which are maintained constant for all mice (Figure 2E).</li>
		<li>Collect a single xy-point, bidimensional fluorescence image of the retina, and immediately capture a fundus image at identical position (100x scan average; 55&#186; scan angle for both images; Figure 2F).</li>
		<li>Capture a multipoint image by selecting &#8220;composite&#8221; in the control panel (Figure 2G) and moving the ophthalmoscope right and left, panning the retina across the nasal-temporal axis (Figure 2H). 
		NOTE: After scanning a single xy-point image (100x scan average), the software automatically averages new scans of the same and new areas (circumscribed with a green circle during optimal scan, or red when the scan is unfeasible), and stitches all xy-positions in real time. The resulting composite image spans 1.5-1.7 mm on the horizontal axis x 3-4 mm in the vertical axis (55&#186; x 120-124&#186; scan angle).</li>
	</ol>
	</li>
	<li>Complete imaging of each mouse within 15 min after inducing anesthesia, before blinking and motion restarts.</li>
	<li>Remove contact lenses, hydrate eyes with PBS and return the animal to its warmed cage, checking behavior until full recovery of motility.</li>
	<li>For longitudinal studies using intermittent cSLO-imaging sessions in the same individual mouse, repeat imaging no less than 3 days apart to avoid the cumulative toxicity of anesthesia, as well as corneal irritation.</li>
</ol>
2. Live Image Processing and Analysis
<ol>
	<li>Sequential image alignment:
	<ol>
		<li>Align the fundus images for a time series of the same eye using the vasculature and optic disc as landmarks. Open all images in a commercial slide-show presentation program, by dragging each image over the preceding one until their optic discs align on the xy-plane (the top image appears semi-transparent while being dragged), then rotate the top image until its major blood vessels overlap with the vasculature on the image below (focus on vessels farthest from the optic disc). Record the rotation angle for each image and save this time series for future reference, keeping track of mouse and eye identity.</li>
		<li>Align the corresponding series of single xy-point fluorescence images by applying the recorded angle to correct the x-y rotation at each time point, using a raster graphic editor program. Additionally, adjust resolution to 300 dpi (without rescaling) and background (in the RGB mode, select Levels and sample the lumen of a blood vessel as black). Save these images as TIF files, adding &#8220;aligned&#8221; to their names.</li>
	</ol>
	</li>
	<li>Microglial cell segmentation:
	<ol>
		<li>To identify GFP+ cells in the central retina, open in FluoRender the &#8220;aligned&#8221; TIF file corresponding to the earliest time point/age. Zoom the image to fit the screen, then define the Render view (composite, orthogonal, interpolated) and its properties (0.58 gamma, 255 saturation point, 255 luminance, 195 alpha and 1.00 shading), selecting both the RG channels as visible (hide B channel by double-clicking it in the Workspace frame; Figure 4A).</li>
		<li>To select individual cells, threshold the red channel (must be highlighted on the Workspace frame), by increasing the low threshold until the majority of cells are masked (white) with minimum overlap and cell processes (cyan), while keeping the high threshold constant (255). The low threshold value varies between 100 and 170, depending on the fluorescence intensity (Figure 4B).</li>
		<li>Save this view with the red channel visible only (Capture command) as a new TIF file, adding &#8220;cellsegm&#8221; to its original name (Figure 4C). Using a generic raster graphic editor software, invert its greyscale and adjust resolution to 300 dpi as above, and save again as RGB (Figure 4D). 
		NOTE: Delete the folders (projects) automatically created by FluoRender, and save the applied threshold for future reference.</li>
	</ol>
	</li>
	<li>Microglial soma segmentation and morphometry:
	<ol>
		<li>To identify GFP+ cell somata for area quantification, use image analysis software with automated intensity measurements and threshold capabilities, as well as threshold-based object counting and measurement. Open the &#8220;cellsegm&#8221; TIF file, and select the rescaling method (spline) and the red channel (click red tab in RGB). Improve visualization by deselecting &#8220;view channel in color&#8221; (right-click the red RGB tab), and selecting &#8220;complement colors&#8221; (Image/Adjust Image) to invert the greyscale and see the cells in grey/black over white background (Figure 4E).</li>
		<li>Calibrate the image to 1.4-1.7 mm depending on age, by drawing a horizontal line across the entire image (Calibration, Quick Calibration).</li>
		<li>Segment individual cell somata by applying intensity thresholding (Figure 4B). For this, work on the red RGB channel and open the Threshold Intensity function (Object Count; selecting &#8220;count update&#8221; command) in order to track bidimensional parameters for each thresholded region of interest (ROI) representing individual somata.</li>
		<li>Define the intensity threshold in the intensity histogram, by selecting the lowest 50-60% of the 255 levels, and refining the final threshold value by visually assessing the overlap between the threshold color mask (blue) and the cell soma perimeter (grey) in individual cells (Figure 4E).</li>
		<li>Estimate the accuracy of the cell soma masks to represent the somal dimensions by measuring the area before and after segmentation in 10 cells and accept the selected threshold range if the measurements differ less than 10% (use the Binary Toolbar and Annotated Measurements).</li>
		<li>Manually identify somal ROIs that include multiple cells and separate these elements (eliminate or separate ROIs using the Binary Toolbar controls, or the Object Catalog) while toggling on/off the binary to directly visualize the cells (Figure 4F).</li>
		<li>Classify microglial cells as ROIs with somal areas larger and smaller than 50-60 &#181;m2 to discriminate cell somata corresponding to activated microglia and deactivated microglia respectively, by using Area Restriction. 
		NOTE: Each cell somal subset is identified with a different pseudocolored binary layer, and can be further characterized for other morphological parameters (perimeter, circularity, etc.), as well as quantified within discrete retinal sectors (Figure 4G). Save the analyzed image as an ND2 file.</li>
		<li>Verify the process and branching complexity in individual activated microglial cells, by overlaying the somal mask and the single xy-point image (contrast increased 50%; Figure 4H). Manually count the processes directly extending from the cell soma or measure the diameter of the polygon encompassing the arbor (use the Binary Toolbar and Annotated Measurements). 
		NOTE: Activated cells lack processes or bear few (&#60;4) and short (&#60;2x somal diameter) ones, in contrast to non-activated cell processes, which comprise a ramified and extensive arbor (&#62; 3-10x somal diameter) surrounding their relatively small soma.</li>
		<li>Manually identify cells with somata smaller than &#60;20 &#181;m2 and/or GFP expression below the detection limit, which are therefore undetected by intensity thresholding. Use the Taxonomy command in Annotations to count manually identified elements.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

Live monitoring of microglial cell number and morphological activation during a neurodegenerative disease requires the use of non-invasive imaging methods that allow the detailed visualization of cell features. After imaging, microglial cells must be isolated (segmented) for morphometric analysis by use of multiple threshold steps to assess somal size and/or process complexity as readouts for microglia activation. In this protocol, we describe methods for live image acquisition using cSLO, and quantitative analysis of mi...

Microglia are neuroimmune cells that exclusively reside in the central nervous system (CNS) since early embryonic development and throughout adulthood. Equipped with a complex repertoire of receptors, microglial activity and regional heterogeneity are regulated by their bidirectional interplay with neighboring neurons, glia, blood-brain barrier and infiltrating neuroinflammatory cells1,2. Basal microglial functions contribute to physiological maintenance and repair, as they sample their territory for perturbations in homeostasis3,4. During CNS injury or disease, microglia are the first responders to neuronal signals that then trigger their transition to a reactive phenotype, termed &#8220;activated microglia2,5-7. Microglia activation involves an intricate cycle of gene and protein expression, which are coupled to cell soma and process resizing and remodeling6-9. Microglia activation, as well as cell redistribution and clustering, can be accompanied the local overall increase in the number of cells (termed microgliosis). This can result from cell proliferation and self-renewal, with or without recruitment of blood-derived monocytes3,4,7,10-14. In a broad range of age-dependent, chronic CNS diseases, sustained microgliosis and microglia activation parallel disease progression15-19. How microglia impact neurodegeneration remains unclear, mainly because they play both neuroprotective and deleterious roles that may have diverse contributions to disease onset and progression. Live imaging studies aimed at understanding chronic CNS disease have monitored microglial behavior in the damaged CNS of animal models and humans, and demonstrated that microglial alterations are detectable beginning at early disease stages15-17,19,20. Thus, it is critical to develop approaches to detect and monitor microglial activation in vivo.
Non-invasive detection of regional changes in brain microglia activation was established as an important in vivo indicator of neurodegenerative disease progression, using molecular imaging or bioluminescence and positron emission tomography or magnetic resonance imaging18,21,22. These highly quantitative and non-invasive molecular and nuclear imaging methods detect gliosis with regional resolution. Alternatively, two-photon confocal imaging in CX3CR1GFP/+ mice has allowed the observation of brain microglia with cellular resolution3,4,9,20,23-28. However, this approach limits long-term and repeated observation of chronic microglial alterations, given the potential risk of disturbing their behavior by even minimally invasive brain imaging procedures29. Alternatively, the retina offers optimal conditions for direct, in vivo visualization and repeated monitoring of microglia in their intact CNS niche throughout aging, following acute injury, and potentially during chronic neurodegenerative diseases. Thus, recent studies have proved the feasibility of high-resolution imaging of retinal microglia expressing GFP by adapting the confocal scanning laser ophthalmoscopy (cSLO) to image live CX3CR1GFP/+ mice. This has been used to track weekly changes in GFP+ cell numbers in individual mice for up to 10 weeks following acutely induced injury or ocular hypertension30-36.
We have extended this approach to perform long-term imaging over several months, and quantitatively track changes in microglia activation based on soma size using morphometric analysis. Somal size was defined as a useful metric of microglia activation in live imaging studies using two-photon confocal microscopy in cortical slices to perform in vivo imaging of CX3CR1-GFP+ microglia9. These and other studies also demonstrated the correlation between somal size and levels of Iba1 protein expression, which also increases with activation9,37. Thus, activated microglia can be identified in live mice, and their numbers and distribution monitored over time during CNS health and disease.
This protocol describes methods for cSLO live image acquisition and analysis to monitor microglial cell numbers, distribution and morphological activation during retinal ganglion cell (RGC) degeneration (Figure 1). Thus, this study uses: 1) a mouse model of inherited glaucoma (DBA/2J) that undergoes age-dependent optic nerve and retinal neurodegeneration and shows remarkable variability in disease progression between 5 and 10 months of age38,39, 2) monthly cSLO in vivo imaging for long-term visualization of GFP+ cells in the retina and unmyelinated optic nerve of heterozygous Cx3cr1GFP/+ DBA/2J mice aged 3-5 months, 3) live imaging analysis by segmentation and thresholding to isolate cell somata and measure their area. These strategies are applied to assess the kinetics of retinal microglial activation states during early stages of chronic glaucoma.

<table><tbody><tr><td>DBA/2J and CX3CR1-GFP/+ mice</td><td>The Jackson Laboratory, Bar Harbor, ME</td><td>000671 and 00582</td><td>Mice are bred in house, introducing new breeders every 3 - 4 generations to prevent genetic drift.</td></tr><tr><td>30&amp;frac12; G needle and 1 ml tuberculin slip-tip syringe</td><td>BD, East Rutherford, NJ</td><td>305106 and 309659</td><td></td></tr><tr><td>2,2,2-tribromoethanol, tert-Amyl alcohol 99% and phosphate buffer saline tablets</td><td>Sigma-Aldrich, St. Louis, MO</td><td>T48402, 152463 and P4417</td><td>Avertin solution (pH 7.3, sterile filtered) must be freshly made solution or stored at 4 &amp;deg;C for up to 1 week.</td></tr><tr><td>Heat therapy T/Pump</td><td>Gaymar Industries, Orchard Park, NY</td><td>TP-650</td><td></td></tr><tr><td>Cotton-tipped applicators</td><td>Fisher Scientific, Pittsburgh, PA</td><td>23-400-100</td><td></td></tr><tr><td>Tropicamide 1%</td><td>Bausch &amp;amp; Lomb, Rochester, NY</td><td>NDC 24208-585-64</td><td></td></tr><tr><td>PMMA contact lenses, 1.70 base curve, 3.2 mm total diameter, 0.40 mm thick center</td><td>Cantor &amp;amp; Nissel Ltd., Northamptonshire, UK</td><td>G003709</td><td>Lenses are rinsed with sterile PBS and stored in polypropylene boxes.</td></tr><tr><td>HRA/Spectralis confocal scanning laser ophthalmoscope and Eye Explorer software</td><td>Heidelberg Engineering GmbH, Carlsbad, CA</td><td>Version 1.7.1.0</td><td></td></tr><tr><td>PowerPoint</td><td>Microsoft, Redmond, WA</td><td>Version 14.4.3</td><td></td></tr><tr><td>Adobe Photoshop</td><td>Adobe, San Jose, CA</td><td>Version CS3</td><td></td></tr><tr><td>FluoRender</td><td>Scientific Computing and Imaging Institute, University of Utah</td><td>Version 2.13</td><td>Freeware. &lt;a href=&quot;http://www.sci.utah.edu/software/13-software/127-fluorender.html&quot; target=&quot;_blank&quot;&gt;http://www.sci.utah.edu/software/13-software/127-fluorender.html&lt;/a&gt;</td></tr><tr><td>NIS-Elements C</td><td>Nikon, Melville, NY</td><td>Version 4.30.01</td><td></td></tr></tbody></table>

in vivo dynamics of retinal microglial activation during neurodegeneration: confocal ophthalmoscopic imaging and cell morphometry in mouse glaucoma

Microglia, which are CNS-resident neuroimmune cells, transform their morphology and size in response to CNS damage, switching to an activated state with distinct functions and gene expression profiles. The roles of microglial activation in health, injury and disease remain incompletely understood due to their dynamic and complex regulation in response to changes in their microenvironment. Thus, it is critical to non-invasively monitor and analyze changes in microglial activation over time in the intact organism. In vivo studies of microglial activation have been delayed by technical limitations to tracking microglial behavior without altering the CNS environment. This has been particularly challenging during chronic neurodegeneration, where long-term changes must be tracked. The retina, a CNS organ amenable to non-invasive live imaging, offers a powerful system to visualize and characterize the dynamics of microglia activation during chronic disorders.
This protocol outlines methods for long-term, in vivo imaging of retinal microglia, using confocal ophthalmoscopy (cSLO) and CX3CR1GFP/+ reporter mice, to visualize microglia with cellular resolution. Also, we describe methods to quantify monthly changes in cell activation and density in large cell subsets (200-300 cells per retina). We confirm the use of somal area as a useful metric for live tracking of microglial activation in the retina by applying automated threshold-based morphometric analysis of in vivo images. We use these live image acquisition and analyses strategies to monitor the dynamic changes in microglial activation and microgliosis during early stages of retinal neurodegeneration in a mouse model of chronic glaucoma. This approach should be useful to investigate the contributions of microglia to neuronal and axonal decline in chronic CNS disorders that affect the retina and optic nerve.

Our recent in vivo studies used these live image acquisition and analysis methods to visualize and track the kinetics and patterns of ONH and retinal microglial changes during early stages of chronic glaucoma and their relationship to late neurodegeneration59. Here we illustrate a cSLO image acquisition protocol to visualize microglial cells across a large area of the central retina in individual young heterozygous Cx3CR1-GFP DBA/2J retinas (Figure 5). Based on the high cell resolutio...

Watch this Scientific Journal Video about In Vivo Dynamics of Retinal Microglial Activation During Neurodegeneration: Confocal Ophthalmoscopic Imaging and Cell Morphometry in Mouse Glaucoma at JoVE.co

In Vivo Dynamics of Retinal Microglial Activation During Neurodegeneration: Confocal Ophthalmoscopic Imaging and Cell Morphometry in Mouse Glaucoma

Microglia activation and microgliosis are key responses to chronic neurodegeneration. Here, we present methods for in vivo, long-term visualization of retinal CX3CR1-GFP+ microglial cells by confocal ophthalmoscopy, and for threshold and morphometric analyses to identify and quantify their activation. We monitor microglial changes during early stages of age-related glaucoma.

In Vivo Dynamics of Retinal Microglial Activation During Neurodegeneration: Confocal Ophthalmoscopic Imaging and Cell Morphometry in Mouse Glaucoma

Microglia, which are CNS-resident neuroimmune cells, transform their morphology and size in response to CNS damage, switching to an activated state with ...

in-vivo-dynamics-retinal-microglial-activation-during

Universidad de Cartagena UDEC

Research

JoVE Journal

Medicine

10.5K Views.  University of Utah. Microglia activation and microgliosis are key responses to chronic neurodegeneration. Here, we present methods for in vivo, long-term visualization of retinal CX3CR1-GFP+ microglial cells by confocal ophthalmoscopy, and for threshold and morphometric analyses to identify and quantify their activation. We monitor microglial changes during early stages of age-related glaucoma.

Video: In Vivo Dynamics of Retinal Microglial Activation During Neurodegeneration: Confocal Ophthalmoscopic Imaging and Cell Morphometry in Mouse Glaucoma

A Laser-induced Mouse Model of Chronic Ocular Hypertension to Characterize Visual Defects

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model of ocular hypertension to mimic human high-tension glaucoma. Here laser illumination is applied to the corneal limbus to photocoagulate the aqueous outflow, inducing angle closure. The changes of IOP are monitored using a rebound tonometer before and after the laser treatment. An optomotor behavioral test is used to measure corresponding changes in visual capacity. The representative result from one mouse which developed sustained IOP elevation after laser illumination is shown. A decreased visual acuity and contrast sensitivity is observed in this ocular hypertensive mouse. Together, our study introduces a valuable model system to investigate neuronal degeneration and the underlying molecular mechanisms in glaucomatous mice.

Chronic ocular hypertension is induced using laser photocoagulation of the trabecular meshwork in mouse eyes. The intraocular pressure (IOP) is elevated for several months after laser treatment. The decrease of visual acuity and contrast sensitivity of experimental animals are monitored using the optomotor test.

Glaucoma, frequently associated with elevated intraocular pressure (IOP), is one of the leading causes of blindness. We sought to establish a mouse model ...

In vivo Imaging of Optic Nerve Fiber Integrity by Contrast-Enhanced MRI in Mice

The rodent visual system encompasses retinal ganglion cells and their axons that form the optic nerve to enter thalamic and midbrain centers, and postsynaptic projections to the visual cortex. Based on its distinct anatomical structure and convenient accessibility, it has become the favored structure for studies on neuronal survival, axonal regeneration, and synaptic plasticity. Recent advancements in MR imaging have enabled the in vivo visualization of the retino-tectal part of this projection using manganese mediated contrast enhancement (MEMRI). Here, we present a MEMRI protocol for illustration of the visual projection in mice, by which resolutions of (200 &#181;m)3 can be achieved using common 3 Tesla scanners. We demonstrate how intravitreal injection of a single dosage of 15 nmol MnCl2 leads to a saturated enhancement of the intact projection within 24 hr. With exception of the retina, changes in signal intensity are independent of coincided visual stimulation or physiological aging. We further apply this technique to longitudinally monitor axonal degeneration in response to acute optic nerve injury, a paradigm by which Mn2+ transport completely arrests at the lesion site. Conversely, active Mn2+ transport is quantitatively proportionate to the viability, number, and electrical activity of axon fibers. For such an analysis, we exemplify Mn2+ transport kinetics along the visual path in a transgenic mouse model (NF-&#954;B p50KO) displaying spontaneous atrophy of sensory, including visual, projections. In these mice, MEMRI indicates reduced but not delayed Mn2+ transport as compared to wild type mice, thus revealing signs of structural and/or functional impairments by NF-&#954;B mutations.
In summary, MEMRI conveniently bridges in vivo assays and post mortem histology for the characterization of nerve fiber integrity and activity. It is highly useful for longitudinal studies on axonal degeneration and regeneration, and investigations of mutant mice for genuine or inducible phenotypes.

In vivo Imaging of Optic Nerve Fiber Integrity by Contrast-Enhanced MRI in Mice

This video illustrates a method, using a clinical 3 T scanner, for contrast-enhanced MR imaging of the na&#239;ve mouse visual projection and for repetitive and longitudinal in vivo studies of optic nerve degeneration associated with acute optic nerve crush injury and chronic optic nerve degeneration in knock-out mice (p50KO).

The rodent visual system encompasses retinal ganglion cells and their axons that form the optic nerve to enter thalamic and midbrain centers, and ...

Neuroscience

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, including phagocytosis of damaged synapses or cells, debris, and/or invading pathogens. Impaired phagocytic function has been implicated in the pathogenesis of diseases such as Alzheimer's and age-related macular degeneration, where amyloid-&#946; plaque and drusen accumulate, respectively. Despite its importance, microglial phagocytosis has been challenging to assess in vivo. Here, we describe a simple, yet robust, technique for precisely monitoring and quantifying the in vivo phagocytic potential of retinal microglia. Previous methods have relied on immunohistochemical staining and imaging techniques. Our method uses flow cytometry to measure microglial uptake of fluorescently labeled particles after intravitreal delivery to the eye in live rodents. This method replaces conventional practices that involve laborious tissue sectioning, immunostaining, and imaging, allowing for more precise quantification of microglia phagocytic function in just under six hours. This procedure can also be adapted to test how various compounds alter microglial phagocytosis in physiological settings. While this technique was developed in the eye, its use is not limited to vision research.

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglial phagocytosis is critical for the maintenance of tissue homeostasis and inadequate phagocytic function has been implicated in pathology. However, assessing microglia function in vivo is technically challenging. We have developed a simple but robust technique for precisely monitoring and quantifying the phagocytic potential of microglia in a physiological setting.

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, ...

Immunology and Infection

A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice

The use of rodent models of glaucoma has been essential to understand the molecular mechanisms that underlie the pathophysiology of this multifactorial neurodegenerative disease. With the advent of numerous transgenic mouse lines, there is increasing interest in inducible murine models of ocular hypertension. Here, we present an occlusion model of glaucoma based on the injection of magnetic microbeads into the anterior chamber of the eye using a modified microneedle with a facetted bevel. The magnetic microbeads are attracted to the iridocorneal angle using a handheld magnet to block the drainage of aqueous humour from the anterior chamber. This disruption in aqueous dynamics results in a steady elevation of intraocular pressure, which subsequently leads to the loss of retinal ganglion cells, as observed in human glaucoma patients. The microbead occlusion model presented in this manuscript is simple compared to other inducible models of glaucoma and also highly effective and reproducible. Importantly, the modifications presented here minimize common issues that often arise in occlusion models. First, the use of a bevelled glass microneedle prevents backflow of microbeads and ensures that minimal damage occurs to the cornea during the injection, thus reducing injury-related effects. Second, the use of magnetic microbeads ensures the ability to attract most beads to the iridocorneal angle, effectively reducing the number of beads floating in the anterior chamber avoiding contact with other structures (e.g., iris, lens). Lastly, the use of a handheld magnet allows flexibility when handling the small mouse eye to efficiently direct the magnetic microbeads and ensure that there is little reflux of the microbeads from the eye when the microneedle is withdrawn. In summary, the microbead occlusion mouse model presented here is a powerful investigative tool to study neurodegenerative changes that occur during the onset and progression of glaucoma.

Here, we present a protocol to induce ocular hypertension in the murine eye that results in the loss of retinal ganglion cells as observed in glaucoma. Magnetic microbeads are injected into the anterior chamber and attracted to the iridocorneal angle using a magnet to block the outflow of aqueous humour.

The use of rodent models of glaucoma has been essential to understand the molecular mechanisms that underlie the pathophysiology of this multifactorial ...

In Vivo Imaging of Cx3cr1gfp/gfp Reporter Mice with Spectral-domain Optical Coherence Tomography and Scanning Laser Ophthalmoscopy

Spectral domain optical coherence tomography (SD-OCT) and scanning laser ophthalmoscopy (SLO) are extensively used in experimental ophthalmology. In the present protocol, mice expressing green fluorescent protein (gfp) under the promoter of Cx3cr1 (BALB/c-Cx3cr1gfp/gfp) were used to image microglia cells in vivo in the retina. Microglia are resident macrophages of the retina and have been implicated in several retinal diseases1,2,3,4,5,6. This protocol provides a detailed approach for generation of retinal B-scans, with SD-OCT, and imaging of microglia cell distribution in Cx3cr1gfp/gfp mice with SLO in vivo, using an ophthalmic imaging platform system. The protocol can be used in several reporter mouse lines. However, there are some limitations to the protocol presented here. First, both SLO and SD-OCT, when used in the high-resolution mode, collect data with high axial resolution but the lateral resolution is lower (3.5 &#181;m and 6 &#181;m, respectively). Moreover, the focus and saturation level in SLO is highly dependent on parameter selection and correct alignment of the eye. Additionally, using devices designed for human patients in mice is challenging due to the higher total optical power of the mouse eye compared to the human eye; this can lead to lateral magnification inaccuracies7, which are also dependent on the magnification by the mouse lens among others. However, despite that the axial scan position is dependent upon lateral magnification, the axial SD-OCT measurements are accurate8.

In Vivo Imaging of Cx3cr1gfp/gfp Reporter Mice with Spectral-domain Optical Coherence Tomography and Scanning Laser Ophthalmoscopy

This protocol describes how high-resolution imaging techniques such as spectral domain optical coherence tomography and scanning laser ophthalmoscopy can be utilized in small rodents, using an ophthalmic imaging platform system, to obtain information on retinal thickness and microglial cell distribution, respectively.

Spectral domain optical coherence tomography (SD-OCT) and scanning laser ophthalmoscopy (SLO) are extensively used in experimental ophthalmology. In the ...

Digestion of Whole Mouse Eyes for Multi-Parameter Flow Cytometric Analysis of Mononuclear Phagocytes

The innate immune system plays important roles in ocular pathophysiology including uveitis, diabetic retinopathy, and age-related macular degeneration. Innate immune cells, specifically mononuclear phagocytes, express overlapping cell surface markers, which makes identifying these populations a challenge. Multi-parameter flow cytometry allows for the simultaneous, quantitative analysis of multiple cell surface markers in order to differentiate monocytes, macrophages, microglia, and dendritic cells in mouse eyes. This protocol describes the enucleation of whole mouse eyes, ocular dissection, digestion into a single cell suspension, and staining of the single cell suspension for myeloid cell markers. Additionally, we explain the proper methods for determining voltages using single color controls and for delineating positive gates using fluorescence minus one controls. The major limitation of multi-parameter flow cytometry is the absence of tissue architecture. This limitation can be overcome by multi-parameter flow cytometry of individual ocular compartments or complimentary immunofluorescence staining. However, immunofluorescence is limited by its lack of quantitative analysis and reduced number of fluorophores on most microscopes. We describe the use of multi-parametric flow cytometry to provide highly quantitative analysis of mononuclear phagocytes in laser-induced choroidal neovascularization. Additionally, multi-parameter flow cytometry can be used for the identification of macrophage subsets, fate mapping, and cell sorting for transcriptomic or proteomic studies.

This protocol provides a method to digest whole eyes into a single cell suspension for the purpose of multi-parameter flow cytometric analysis in order to identify specific ocular mononuclear phagocytic populations, including monocytes, microglia, macrophages, and dendritic cells.

The innate immune system plays important roles in ocular pathophysiology including uveitis, diabetic retinopathy, and age-related macular degeneration. ...

Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. Although studying these cells in isolation in vitro or together ex vivo provides useful physiological information; salient features of neural cell physiology will be missed in such contexts. Therefore, there is a need for studying neural cells in their native in vivo environment. The protocol detailed here describes repetitive in vivo two-photon imaging of neural cells in the rodent cortex as a tool to visualize and study specific cells over extended periods of time from hours to months. We describe in detail the use of the grossly stable brain vasculature as a coarse map or fluorescently labeled dendrites as a fine map of select brain regions of interest. Using these maps as a visual key, we show how neural cells can be precisely relocated for subsequent repetitive in vivo imaging. Using examples of in vivo imaging of fluorescently-labeled microglia, neurons, and NG2+ cells, this protocol demonstrates the ability of this technique to allow repetitive visualization of cellular dynamics in the same brain location over extended time periods, that can further aid in understanding the structural and functional responses of these cells in normal physiology or following pathological insults. Where necessary, this approach can be coupled to functional imaging of neural cells, e.g., with calcium imaging. This approach is especially a powerful technique to visualize the physical interaction between different cell types of the CNS in vivo when genetic mouse models or specific dyes with distinct fluorescent tags to label the cells of interest are available.

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.

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. ...

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

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

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

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

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 ...

Mouse Retina OCT Fibergram Alignment with Confocal Images

Alignment of Visible-Light Optical Coherence Tomography Fibergrams with Confocal Images of the Same Mouse Retina

In recent years, in vivo retinal imaging, which provides non-invasive, real-time, and longitudinal information about biological systems and processes, has been increasingly applied to obtain an objective assessment of neural damage in eye diseases. Ex vivo confocal imaging of the same retina is often necessary to validate the in vivo findings especially in animal research. In this study, we demonstrated a method for aligning an ex vivo confocal image of the mouse retina with its in vivo images. A new clinical-ready imaging technology called visible light optical coherence tomography fibergraphy (vis-OCTF) was applied to acquire in vivo images of the mouse retina. We then performed the confocal imaging of the same retina as the &#34;gold standard&#34; to validate the in vivo vis-OCTF images. This study not only enables further investigation of the molecular and cellular mechanisms but also establishes a foundation for a sensitive and objective evaluation of neural damage in vivo.

Author Spotlight: Advancements in In Vivo and Ex Vivo Retinal Imaging for Improved Glaucoma Diagnosis and Treatment 

The present protocol outlines the steps for aligning in vivo visible-light optical coherence tomography fibergraphy (vis-OCTF) images with ex vivo confocal images of the same mouse retina for the purpose of verifying the observed retinal ganglion cell axon bundle morphology in the in vivo images.

In recent years, in vivo retinal imaging, which provides non-invasive, real-time, and longitudinal information about biological systems and processes, has ...

In Vivo Dynamics of Retinal Microglial Activation During Neurodegeneration: Confocal Ophthalmoscopic Imaging and Cell Morphometry in Mouse Glaucoma

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Chapters in this video

A Laser-induced Mouse Model of Chronic Ocular Hypertension to Characterize Visual Defects

In vivo Imaging of Optic Nerve Fiber Integrity by Contrast-Enhanced MRI in Mice

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice

In Vivo Imaging of Cx3cr1^gfp/gfp Reporter Mice with Spectral-domain Optical Coherence Tomography and Scanning Laser Ophthalmoscopy

Digestion of Whole Mouse Eyes for Multi-Parameter Flow Cytometric Analysis of Mononuclear Phagocytes

Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

Simultaneous Imaging of Microglial Dynamics and Neuronal Activity in Awake Mice

Alignment of Visible-Light Optical Coherence Tomography Fibergrams with Confocal Images of the Same Mouse Retina