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 border="0" cellpadding="0" cellspacing="0" width="2010">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:617px;">Name</td>
			<td style="width:444px;">Company</td>
			<td style="width:208px;">Catalog Number</td>
			<td style="width:741px;">Comments</td>
		</tr>
		<tr height="60">
			<td height="60" style="height:60px;width:617px;">DBA/2J and 
			CX3CR1-GFP/+ mice</td>
			<td>The Jackson Laboratory, Bar Harbor, ME</td>
			<td>000671 and 00582</td>
			<td>Mice are breed in-house, introducing new breeders every 3 to 4 generations to prevent genetic drift.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">301/2G needle and 1 ml tuberculin slip-tip syringe</td>
			<td>BD, East Rutherford, NJ</td>
			<td>305106 and 309659</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:617px;">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&#186;C for up to 1 week.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Heat therapy T/Pump</td>
			<td>Gaymar Industries, Orchard Park, NY</td>
			<td>TP-650</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Cotton-tipped applicators</td>
			<td>Fisher Scientific, Pittsburgh, PA</td>
			<td>23-400-100</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Tropicamide 1%&#160;</td>
			<td>Bausch &#38; Lomb, Rochester, NY</td>
			<td>NDC 24208-585-64</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">PMMA contact lenses, 1.70-base curve, 3.2 mm total diameter, 0.40 mm-thick center</td>
			<td>Cantor &#38; Nissel Ltd., Northamptonshire, UK</td>
			<td>G003709</td>
			<td>Lenses are rinsed with sterile PBS and stored in polypropylene boxes.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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 height="20">
			<td height="20" style="height:20px;">PowerPoint</td>
			<td>Microsoft, Redmond, WA</td>
			<td>Version 14.4.3</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Adobe Photoshop</td>
			<td>Adobe, San Jose, CA</td>
			<td>Version CS3</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">FluoRender</td>
			<td>Scientific Computing and Imaging Institute, University of Utah</td>
			<td>Version 2.13</td>
			<td>Freeware. http://www.sci.utah.edu/software/13-software/127-fluorender.html</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">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

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

Multi-photon Imaging of Tumor Cell Invasion in an Orthotopic Mouse Model of Oral Squamous Cell Carcinoma

Loco-regional invasion of head and neck cancer is linked to metastatic risk and presents a difficult challenge in designing and implementing patient management strategies. Orthotopic mouse models of oral cancer have been developed to facilitate the study of factors that impact invasion and serve as model system for evaluating anti-tumor therapeutics. In these systems, visualization of disseminated tumor cells within oral cavity tissues has typically been conducted by either conventional histology or with in vivo bioluminescent methods. A primary drawback of these techniques is the inherent inability to accurately visualize and quantify early tumor cell invasion arising from the primary site in three dimensions. Here we describe a protocol that combines an established model for squamous cell carcinoma of the tongue (SCOT) with two-photon imaging to allow multi-vectorial visualization of lingual tumor spread. The OSC-19 head and neck tumor cell line was stably engineered to express the F-actin binding peptide LifeAct fused to the mCherry fluorescent protein (LifeAct-mCherry). Fox1nu/nu mice injected with these cells reliably form tumors that allow the tongue to be visualized by ex-vivo application of two-photon microscopy. This technique allows for the orthotopic visualization of the tumor mass and locally invading cells in excised tongues without disruption of the regional tumor microenvironment. In addition, this system allows for the quantification of tumor cell invasion by calculating distances that invaded cells move from the primary tumor site. Overall this procedure provides an enhanced model system for analyzing factors that contribute to SCOT invasion and therapeutic treatments tailored to prevent local invasion and distant metastatic spread. This method also has the potential to be ultimately combined with other imaging modalities in an in vivo setting.

A comprehensive overview of the techniques involved in generating a mouse model of oral cancer and quantitative monitoring of tumor invasion within the tongue through multi-photon microscopy of labeled cells is presented. This system can serve as a useful platform for the molecular assessment and drug efficacy of anti-invasive compounds.

Loco-regional invasion of head and neck cancer is linked to metastatic risk and presents a difficult challenge in designing and implementing patient ...

In vivo Bioluminescence Imaging of Tumor Hypoxia Dynamics of Breast Cancer Brain Metastasis in a Mouse Model

It is well recognized that tumor hypoxia plays an important role in promoting malignant progression and affecting therapeutic response negatively. There is little knowledge about in situ, in vivo, tumor hypoxia during intracranial development of malignant brain tumors because of lack of efficient means to monitor it in these deep-seated orthotopic tumors. Bioluminescence imaging (BLI), based on the detection of light emitted by living cells expressing a luciferase gene, has been rapidly adopted for cancer research, in particular, to evaluate tumor growth or tumor size changes in response to treatment in preclinical animal studies. Moreover, by expressing a reporter gene under the control of a promoter sequence, the specific gene expression can be monitored non-invasively by BLI. Under hypoxic stress, signaling responses are mediated mainly via the hypoxia inducible factor-1&alpha; (HIF-1&alpha;) to drive transcription of various genes. Therefore, we have used a HIF-1&alpha; reporter construct, 5HRE-ODD-luc, stably transfected into human breast cancer MDA-MB231 cells (MDA-MB231/5HRE-ODD-luc). In vitro HIF-1&alpha; bioluminescence assay is performed by incubating the transfected cells in a hypoxic chamber (0.1% O2) for 24 hr before BLI, while the cells in normoxia (21% O2) serve as a control. Significantly higher photon flux observed for the cells under hypoxia suggests an increased HIF-1&alpha; binding to its promoter (HRE elements), as compared to those in normoxia. Cells are injected directly into the mouse brain to establish a breast cancer brain metastasis model. In vivo bioluminescence imaging of tumor hypoxia dynamics is initiated 2 wks after implantation and repeated once a week. BLI reveals increasing light signals from the brain as the tumor progresses, indicating increased intracranial tumor hypoxia. Histological and immunohistochemical studies are used to confirm the in vivo imaging results. Here, we will introduce approaches of in vitro HIF-1&alpha; bioluminescence assay, surgical establishment of a breast cancer brain metastasis in a nude mouse and application of in vivo bioluminescence imaging to monitor intracranial tumor hypoxia.

In vivo Bioluminescence Imaging of Tumor Hypoxia Dynamics of Breast Cancer Brain Metastasis in a Mouse Model

Bioluminescence imaging of hypoxia inducible factor-1&alpha; activity is applied to monitor intracranial tumor hypoxia development in a breast cancer brain metastasis mouse model.

It is well recognized that tumor hypoxia plays an important role in promoting malignant progression and affecting therapeutic response negatively. There ...

In vivo Imaging of Tumor Angiogenesis using Fluorescence Confocal Videomicroscopy

Fibered confocal fluorescence in vivo imaging with a fiber optic bundle uses the same principle as fluorescent confocal microscopy. It can excite fluorescent in situ elements through the optical fibers, and then record some of the emitted photons, via the same optical fibers. The light source is a laser that sends the exciting light through an element within the fiber bundle and as it scans over the sample, recreates an image pixel by pixel. As this scan is very fast, by combining it with dedicated image processing software, images in real time with a frequency of 12 frames/sec can be obtained.
We developed a technique to quantitatively characterize capillary morphology and function, using a confocal fluorescence videomicroscopy device. The first step in our experiment was to record 5 sec movies in the four quadrants of the tumor to visualize the capillary network. All movies were processed using software (ImageCell, Mauna Kea Technology, Paris France) that performs an automated segmentation of vessels around a chosen diameter (10 &mu;m in our case). Thus, we could quantify the 'functional capillary density', which is the ratio between the total vessel area and the total area of the image. This parameter was a surrogate marker for microvascular density, usually measured using pathology tools.
The second step was to record movies of the tumor over 20 min to quantify leakage of the macromolecular contrast agent through the capillary wall into the interstitium. By measuring the ratio of signal intensity in the interstitium over that in the vessels, an 'index leakage' was obtained, acting as a surrogate marker for capillary permeability.

In vivo Imaging of Tumor Angiogenesis using Fluorescence Confocal Videomicroscopy

In this paper, we present a method to analyze tumor microvessels in vivo using dynamic contrast-enhanced fluorescence videomicroscopy. Two quantitative parameters were acquired: functional capillary density reflecting the vascularity of the tumor, and index leakage reflecting the leakiness of the endothelial wall.

Fibered confocal fluorescence in vivo imaging with a fiber optic bundle uses the same principle as fluorescent confocal microscopy. It can excite ...

In vivo 19F MRI for Cell Tracking

In vivo 19F MRI allows quantitative cell tracking without the use of ionizing radiation. It is a noninvasive technique that can be applied to humans. Here, we describe a general protocol for cell labeling, imaging, and image processing. The technique is applicable to various cell types and animal models, although here we focus on a typical mouse model for tracking murine immune cells. The most important issues for cell labeling are described, as these are relevant to all models. Similarly, key imaging parameters are listed, although the details will vary depending on the MRI system and the individual setup. Finally, we include an image processing protocol for quantification. Variations for this, and other parts of the protocol, are assessed in the Discussion section. Based on the detailed procedure described here, the user will need to adapt the protocol for each specific cell type, cell label, animal model, and imaging setup. Note that the protocol can also be adapted for human use, as long as clinical restrictions are met.

In vivo 19F MRI for Cell Tracking

We describe a general protocol for in vivo cell tracking using MRI in a mouse model with ex vivo labeled cells. A typical protocol for cell labeling, image acquisition processing and quantification is included.

In vivo 19F MRI allows quantitative cell tracking without the use of ionizing radiation. It is a noninvasive technique that can be applied to humans. ...

Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Bone turns over continuously and is highly regenerative following injury. Osteogenic stem/progenitor cells have long been hypothesized to exist, but in vivo demonstration of such cells has only recently been attained. Here, in vivo imaging techniques to investigate the role of endogenous osteogenic stem/progenitor cells (OSPCs) and their progeny in bone repair are provided. Using osteo-lineage cell tracing models and intravital imaging of induced microfractures in calvarial bone, OSPCs can be directly observed during the first few days after injury, in which critical events in the early repair process occur. Injury sites can be sequentially imaged revealing that OSPCs relocate to the injury, increase in number and differentiate into bone forming osteoblasts. These methods offer a means of investigating the role of stem cell-intrinsic and extrinsic molecular regulators for bone regeneration and repair.

Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Quantitative measurement of bone progenitor function in fracture healing requires high resolution serial imaging technology. Here, protocols are provided for using intravital microscopy and osteo-lineage tracking to sequentially image and quantify the migration, proliferation and differentiation of endogenous osteogenic stem/progenitor cells in the process of repairing bone fracture.

Bone turns over continuously and is highly regenerative following injury. Osteogenic stem/progenitor cells have long been hypothesized to exist, but in ...

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. Despite the discovery of key regulators of uterine contractility, this process is still not fully understood. Transgenic mice provide an ideal model in which to study parturition. Previously, the only method to study uterine contractility in the mouse was ex vivo isometric tension recordings, which are suboptimal for several reasons. The uterus must be removed from its physiological environment, a limited time course of investigation is possible, and the mice must be sacrificed. The recent development of radiometric telemetry has allowed for longitudinal, real-time measurements of in vivo intrauterine pressure in mice. Here, the implantation of an intrauterine telemeter to measure pressure changes in the mouse uterus from mid-pregnancy until delivery is described. By comparing differences in pressures between wild type and transgenic mice, the physiological impact of a gene of interest can be elucidated. This technique should expedite the development of therapeutics used to treat myometrial disorders during pregnancy, including preterm labor.

Intrauterine Telemetry to Measure Mouse Contractile Pressure In Vivo

This manuscript provides a protocol for implanting telemeters in the mouse for the purpose of measuring intrauterine pressures during pregnancy.

A complex integration of molecular and electrical signals is needed to transform a quiescent uterus into a contractile organ at the end of pregnancy. ...

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo: A Model of Single Vessel Stroke

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to image cellular events in an intact animal. Additionally, the ability to induce physiological disease states such as stroke in vivo increases its utility. The technique described herein allows for physiological assessment of cellular responses within the CNS following a stroke and can be adapted for other pathological conditions being studied. The technique presented uses laser excitation of the photosensitive dye Rose Bengal in vivo to induce a focal ischemic event in a single blood vessel. 
The video protocol demonstrates the preparation of a thin-skulled cranial window over the somatosensory cortex in a mouse for the induction of a Rose Bengal photothrombotic event keeping injury to the underlying dura matter and brain at a minimum. Surgical preparation is initially performed under a dissecting microscope with a custom-made surgical/imaging platform, which is then transferred to a confocal microscope equipped with an inverted objective adaptor. Representative images acquired utilizing this protocol are presented as well as time-lapse sequences of stroke induction. This technique is powerful in that the same area can be imaged repeatedly on subsequent days facilitating longitudinal in vivo studies of pathological processes following stroke.

Rose Bengal Photothrombosis by Confocal Optical Imaging In Vivo:  A Model of Single Vessel Stroke

Here, we describe a semi-invasive optical microscopy approach for the induction of a Rose Bengal photothrombotic clot in the somatosensory cortex of a mouse in vivo. The technical aspects of the imaging procedure are described from induction of a photothrombotic event to application and data collection.

In vivo imaging techniques have increased in utilization due to recent advances in imaging dyes and optical technologies, allowing for the ability to ...

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Glaucoma is a disease of the central nervous system affecting retinal ganglion cells (RGCs). RGC axons making up the optic nerve carry visual input to the brain for visual perception. Damage to RGCs and their axons leads to vision loss and/or blindness. Although the specific cause of glaucoma is unknown, the primary risk factor for the disease is an elevated intraocular pressure. Glaucoma-inducing procedures in animal models are a valuable tool to researchers studying the mechanism of RGC death. Such information can lead to the development of effective neuroprotective treatments that could aid in the prevention of vision loss. The protocol in this paper describes a method of inducing glaucoma - like conditions in an in vivo rat model where 50 &#181;l of 2 M hypertonic saline is injected into the episcleral venous plexus. Blanching of the vessels indicates successful injection. This procedure causes loss of RGCs to simulate glaucoma. One month following injection, animals are sacrificed and eyes are removed. Next, the cornea, lens, and vitreous are removed to make an eyecup. The retina is then peeled from the back of the eye and pinned onto sylgard dishes using cactus needles. At this point, neurons in the retina can be stained for analysis. Results from this lab show that approximately 25% of RGCs are lost within one month of the procedure when compared to internal controls. This procedure allows for quantitative analysis of retinal ganglion cell death in an in vivo rat glaucoma model.

Glaucoma-inducing Procedure in an In Vivo Rat Model and Whole-mount Retina Preparation

Glaucoma is characterized by damage to retinal ganglion cells. Inducing glaucoma in animal models can provide insight into the study of this disease. Here, we outline a procedure that induces loss of RGCs in an in vivo rat model and demonstrates the preparation of whole-mount retinas for analysis.

Glaucoma is a disease of the central nervous system affecting retinal ganglion cells (RGCs). RGC axons making up the optic nerve carry visual input to the ...

Depletion of Mouse Cells from Human Tumor Xenografts Significantly Improves Downstream Analysis of Target Cells

The use of in vitro cell line models for cancer research has been a useful tool. However, it has been shown that these models fail to reliably mimic patient tumors in different assays1. Human tumor xenografts represent the gold standard with respect to tumor biology, drug discovery, and metastasis research 2-4. Tumor xenografts can be derived from different types of material like tumor cell lines, tumor tissue from primary patient tumors4 or serially transplanted tumors. When propagated in vivo, xenografted tissue is infiltrated and vascularized by cells of mouse origin. Multiple factors such as the tumor entity, the origin of xenografted material, growth rate and region of transplantation influence the composition and the amount of mouse cells present in tumor xenografts. However, even when these factors are kept constant, the degree of mouse cell contamination is highly variable.
Contaminating mouse cells significantly impair downstream analyses of human tumor xenografts. As mouse fibroblasts show high plating efficacies and proliferation rates, they tend to overgrow cultures of human tumor cells, especially slowly proliferating subpopulations. Mouse cell derived DNA, mRNA, and protein components can bias downstream gene expression analysis, next-generation sequencing, as well as proteome analysis 5. To overcome these limitations, we have developed a fast and easy method to isolate untouched human tumor cells from xenografted tumor tissue. This procedure is based on the comprehensive depletion of cells of mouse origin by combining automated tissue dissociation with the benchtop tissue dissociator and magnetic cell sorting. Here, we demonstrate that human target cells can be can be obtained with purities higher than 96% within less than 20 min independent of the tumor type.

Human tumor xenografts are vascularized and infiltrated by cells of mouse origin during the growth phase in vivo. To circumvent the bias caused by these contaminating cells during downstream analysis, we developed a method allowing for comprehensive depletion of all mouse cells by magnetic cell sorting.

The use of in vitro cell line models for cancer research has been a useful tool. However, it has been shown that these models fail to reliably mimic ...

Fluorescent Dye Labeling of Erythrocytes and Leukocytes for Studying the Flow Dynamics in Mouse Retinal Circulation

The retinal and choroidal blood flow dynamics may provide insight into the pathophysiology and sequelae of various ocular diseases, such as glaucoma, diabetic retinopathy, age-related macular degeneration (AMD) and other ocular inflammatory conditions. It may also help to monitor the therapeutic responses in the eye. The proper labeling of the blood cells, coupled with live-cell imaging of the labeled cells, allows for the investigation of the flow dynamics in the retinal and choroidal circulation. Here, we describe the standardized protocols of 1.5% indocyanine green (ICG) and 1% sodium fluorescein labeling of mice erythrocytes and leukocytes, respectively. Scanning laser ophthalmoscopy (SLO) was applied to visualize the labeled cells in the retinal circulation of C57BL/6J mice (wild type). Both methods demonstrated distinct fluorescently labeled cells in the mouse retinal circulation. These labeling methods can have wider applications in various ocular disease models.

Live-cell imaging of the labeled blood cells in ocular circulation can provide information about inflammation and ischemia in diabetic retinopathy and age-related macular degeneration. A protocol to label blood cells and image the labeled cells in the retinal circulation is described.

The retinal and choroidal blood flow dynamics may provide insight into the pathophysiology and sequelae of various ocular diseases, such as glaucoma, ...