This work was supported by National Institutes of Health (NIH) grants (R01 EY024963 and EY028100) to J.C. Z.W. was supported by a Knights Templar Eye Foundation Career Starter Grant. The hyaloid isolation procedure described in this study was adapted with modification from protocols generously shared by Drs. Richard Lang, Toshihide Kurihara, and Lois Smith, to whom the authors are thankful.

All animals were treated in accordance with the Association for Research in Vision and Ophthalmology (ARVO) Statement for the Use of Animals in Ophthalmic and Vision Research for animal experiments, following the Guidelines of the National Institutes of Health (NIH) regarding the care and use of animals for experimental procedures and the regulations set forth by the Institutional Animal Care and Use Committee (IACUC) at Boston Children&#8217;s Hospital. Lrp5-/- mice (stock no. 005823; Jackson Laboratory) and its wild-type (WT) control C57BL/6J mice (stock no. 000664; Jackson Laboratory) were used for this study.
1. Part I: In vivo imaging of hyaloid vessels using a rodent retinal imaging system
<ol>
	<li>Preparation of the mice anesthesia and pupil dilation
	<ol>
		<li>Select the mice.
		<ol>
			<li>Use mice that are older than P12 (after they have opened their eyes) for retinal imaging; both sexes are suitable.</li>
			<li>As desired, image mutant mice with delayed hyaloid regression (and their WT controls) throughout adulthood. WT mice older than 3 weeks may not exhibit much detectable remaining hyaloid vessels, hence P12&#8211;P21 is ideal for imaging remaining visible WT hyaloid.</li>
		</ol>
		</li>
		<li>Prepare a ketamine/xylazine mixture for anesthesia.
		<ol>
			<li>Take 2.3 mL of ketamine stock solution (100 mg/mL) and 0.7 mL of xylazine stock solution (20 mg/mL) and add 20 mL of sterile saline (0.9% sodium chloride) to make a working solution of a ketamine/xylazine mixture (10 mg/mL ketamine and 0.6 mg/mL xylazine).</li>
		</ol>
		</li>
		<li>Anesthetize the mice by intraperitoneally injecting ketamine/xylazine mixture in a volume of 8x the mouse&#8217;s body weight (e.g., a 20 g mouse needs 160 &#956;L of ketamine/xylazine working solution mixture to achieve a working dose of ketamine [80 mg/kg body weight] and xylazine [4.8 mg/kg body weight] mixture).</li>
		<li>Apply one drop of pupil-dilating drug solution (see Table of Materials) to each eye to dilate the mouse&#8217;s pupils immediately following anesthesia.</li>
		<li>Assess the level of anesthesia by pedal reflex (firm hindlimb toe pinch). Wait until the mouse is sufficiently anesthetized (no pedal reflex) and its pupils are widely dilated.</li>
	</ol>
	</li>
	<li>Optical coherence tomography imaging of hyaloid vessels
	<ol>
		<li>Moisturize the mouse&#8217;s corneas with artificial tears, and then, place the mouse on the positioning stage.</li>
		<li>Gently contact the mouse&#8217;s cornea with the lens of the OCT probe. Adjust the mouse and probe so that the optic nerve head is in the center of the visual field in the fundus image, to guide the OCT imaging. 
		NOTE: For more details on the general setup of a rodent retinal imaging system (see Table of Materials) and adjusting the mouse&#8217;s position, please refer to Gong et al.28.</li>
		<li>Adjust the focus to achieve the optimal images of OCT.</li>
		<li>Adjust the angle of the indicated line in the OCT imaging software (see Table of Materials) to reveal persistent hyaloid vessels and, then, take images.</li>
	</ol>
	</li>
	<li>Fundus fluorescein angiography imaging of hyaloid vessels
	<ol>
		<li>Prepare a fluorescein solution: add 9 mL of sterile phosphate-buffered saline (PBS) to 1 mL of fluorescein solution (stock concentration: 100 mg/mL). The concentration of the final working solution is 10 mg/mL.</li>
		<li>Intraperitoneally inject the fluorescein working solution (5 &#956;L/g body weight).</li>
		<li>Moisturize the mouse&#8217;s cornea with artificial tears, and then, place the mouse on the positioning stage.</li>
		<li>Position the lens of the Micron IV retinal imaging microscope to make direct gentle contact with the mouse cornea. Adjust the alignment slightly to position the optic nerve head in the center of the view field.</li>
		<li>Change the microscope filter to a green fluorescent channel.</li>
		<li>Focus on persistent hyaloid vessels to take images.</li>
		<li>Take multiple images after 1 min, 3 min, 5 min, and 10 min (no more than 10 min) post-injection to determine the best time point (signal-to-background ratio) for observing hyaloid vessels. Complete the FFA procedure within 10 min, after which the fluorescein may become too diffused and make vessels invisible.</li>
	</ol>
	</li>
	<li>Recovery of the mice from anesthesia
	<ol>
		<li>After the procedures, keep the mice on a warm heating pad.</li>
		<li>Wait until the mice are mobile again to return them to the mouse cage.</li>
	</ol>
	</li>
</ol>
2. Part II: Ex vivo visualization of hyaloid vessels
<ol>
	<li>Preparation of fixed mouse eyes
	<ol>
		<li>Select mice of the right age. 
		NOTE: Neonatal mice are typically sacrificed at P8 for hyaloid dissection. Both sexes are suitable. Mutant mice with a delayed hyaloid regression (and their respective WT controls) may be dissected and analyzed throughout adulthood.</li>
		<li>Euthanize the mice by exposure to CO2.</li>
		<li>Enucleate the mouse&#8217;s eyes by blunt dissection.</li>
		<li>Open the eyelids wide with microsurgery forceps to allow access to the eyeball. Place curved microsurgery forceps under the globe in the orbit to grasp the optic nerve without squeezing the eyeball. Gently pull and remove the eyeball with the forceps.</li>
		<li>Alternatively, dissect the eyeball by using microsurgery scissors to carefully cut parallel to the globe from the four sides toward the back of orbit and separating the globe from surrounding connective tissues.</li>
		<li>Immerse the eyes in 4% paraformaldehyde in PBS buffer for 30 min at room temperature for fixation.</li>
		<li>Transfer the fixed eyeballs to ice-cold PBS buffer. 
		NOTE: The eyeballs may be stored in PBS at 4 &#176;C for up to 1 week.</li>
	</ol>
	</li>
	<li>Embedding of hyaloid vessels with gelatin injection
	<ol>
		<li>Prepare a 5% (w/v) gelatin solution.
		<ol>
			<li>Weigh out 50 mg of gelatin.</li>
			<li>Dissolve the gelatin in 1 mL of distilled water.</li>
			<li>Incubate the gelatin solution in a 37 &#176;C water bath until fully dissolved. Keep the solution in 37 &#176;C water until use. 
			NOTE: A larger batch of solution may be prepared and stored at 4 &#176;C as aliquots. Every time before use, the solution needs to be warmed in 37 &#176;C water bath to achieve a clear consistency.</li>
		</ol>
		</li>
		<li>Under a dissection microscope, inject 50 &#956;L of gelatin solution into the vitreous body at the limbus; repeat the injection 3x at different sites to make a total of four injections, evenly spaced around the limbus (Figure 2A).</li>
		<li>Incubate the eyeballs in a 4 &#176;C refrigerator or on ice for 30 min to solidify the injected gelatin in the vitreous space.</li>
	</ol>
	</li>
	<li>Dissection and isolation of hyaloid vessels 
	NOTE: See Figure 2B-E.
	<ol>
		<li>Place the eyeballs in a Petri dish containing PBS to keep the tissue from drying. Make an incision with microsurgery scissors at the limbus and remove the cornea.</li>
		<li>Snip and remove the optic nerve.</li>
		<li>Under a dissection microscope, use two pairs of forceps to peel off and discard the sclera, choroid, and retinal pigment epithelium (RPE) layers and remove the iris.</li>
		<li>With the optic-nerve side of the retina facing up and the lens side down, inject 50 &#956;L of PBS just beneath the retina cup to allow accumulation of the PBS buffer between the gelatinized vitreous body and the retina.</li>
		<li>Gently peel off and remove the retina cup and ciliary body from the vitreous body with microsurgery forceps.</li>
		<li>Using a transfer pipette, transfer the gelatin cup containing the hyaloid tissue immersed in PBS to a microscope slide</li>
		<li>Turn the rest of the tissue (lens/hyaloid) over, so the lens side is facing up.</li>
		<li>Lift the lens and gently loosen the connection between the lens/TVL and VHP, and then, cut the HA with microsurgery scissors to remove the lens-TVL. Keep the VHP-part of the hyaloid cup for the flat mount.</li>
	</ol>
	</li>
	<li>Flat mounting and staining of hyaloid vessels 
	<ol>
		<li>Gently rinse the hyaloid cup with PBS on the slide to remove all dissection debris.</li>
		<li>Make sure the hyaloid cup is floating on the slide in adequate PBS solution.</li>
		<li>Gently arrange and adjust the position of the gelatinized hyaloid cup with microsurgery forceps, with the rim of the cup facing down on the slide, to achieve optimal appearance after melting.</li>
		<li>Place the slide with the hyaloid cup immersed in PBS onto a slide warmer at 37 &#176;C, wait until the gelatin melts and the hyaloid flattens, and remove the slide when it is barely dry (do not over-dry).</li>
		<li>(Optional) Immunostain the hyaloid vessels
		<ol>
			<li>Make blocking and penetrating buffer (e.g., 5% bovine serum albumin [BSA] and 0.1% Triton X-100 in PBS buffer). Add 50 &#956;L of blocking buffer onto a flat-mounted hyaloid isolation and incubate it at room temperature for 30 min.</li>
			<li>Apply 50 &#956;L of primary antibody (e.g., CD31) (1:100 dilution in blocking buffer) onto a flat mount of hyaloid isolation, and then, incubate it in a wet box at 4 &#176;C overnight.</li>
			<li>Gently rinse 3x, for 5 min each time, with PBS.</li>
			<li>Apply 50 &#956;L of secondary antibody (e.g., goat anti-rabbit secondary antibody-Alexa 488, 1:100 dilution) onto the hyaloid flat mount and incubate it at room temperature for 1 h.</li>
			<li>Gently rinse 3x, for 5 min each time, with PBS.</li>
		</ol>
		</li>
		<li>Add a drop of anti-fade mounting medium with DAPI (4&#8217;, 6-diamidino-2-phenylindole) to stain the nuclei in the hyaloid vessels.</li>
		<li>Gently place a coverslip on the hyaloid to complete the flat mount.</li>
	</ol>
	</li>
	<li>Imaging and quantification of flat-mounted hyaloid vessels
	<ol>
		<li>Image the DAPI staining of hyaloid vessels with a fluorescent microscope and make sure the central HA is visible (exclude the samples without the HA).</li>
		<li>Identify vessel branches directly derived from the HA and manually count the number of vessel branches for quantification.</li>
	</ol>
	</li>
	<li>Visualization of hyaloid vessels in the cross section of eyes 
	<ol>
		<li>Embed fixed eyeballs into optimal cutting temperature compound in a suitable tissue mold, and then, freeze the embedded eyes on dry ice or store them in a -20 &#176;C freezer.</li>
		<li>Use a cryostat microtome to make a cross section of the embedded eyes into sections of 12 &#956;m thickness at -20 &#176;C.</li>
		<li>Collect the eye cross sections onto a microscope slide room temperature by gently touching the tissue sections, which will adhere to the slide.</li>
		<li>Air-dry to dehydrate the tissue sections for ~30 min, which can then be stored in a -20 &#176;C freezer until ready for staining.</li>
		<li>Rinse the slide 1x with PBS to remove the remaining optimal cutting temperature compound on the slide.</li>
		<li>(Optional) Immerse the slide into an appropriate fixative buffer (e.g., 4% paraformaldehyde in PBS) for 15 min at room temperature for additional fixation, if needed.</li>
		<li>Permeabilize the tissue with 0.1% Triton X-100 in PBS for 30 min at room temperature.</li>
		<li>Prepare isolectin-IB4 staining buffer for blood vessel staining.</li>
		<li>Dissolve and reconstitute isolectin-IB4 (500 &#956;g) powder with 50 mL of PBS in a 50 mL centrifuge tube.</li>
		<li>Slowly add 50 &#956;L of 1 M CaCl2 solution, drop by drop, into the 50 mL of PBS/isolectin-IB4 mixture. This step needs to be performed slowly to avoid precipitation. The final concentration of CaCl2 is 1 &#956;M. The presence of Ca2+ is required for isolectin-IB4 staining.</li>
		<li>Apply 30 &#956;L of isolectin-IB4 staining buffer onto the eye cross sections on the slide and incubate it in a wet box at 4 &#176;C overnight.</li>
		<li>Rinse 3x, for 5 min each time, with PBS.</li>
		<li>Add a drop of anti-fade mounting medium with DAPI to stain the nuclei in the sections.</li>
		<li>Gently place a coverslip on the tissue sections to complete the flat mount.</li>
		<li>Check the tissue under a microscope to visualize the presence of hyaloid vessels within the eyecup.</li>
	</ol>
	</li>
</ol>

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<li>Lang, R. A., Bishop, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Macrophages+are+required+for+cell+death+and+tissue+remodeling+in+the+developing+mouse+eye%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">Macrophages are required for cell death and tissue remodeling in the developing mouse eye.</a> Cell. 74 (3), 453-462 (1993).</li>
<li>Riazifar, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Phenotypic+and+functional+characterization+of+Bst%2B%2F-+mouse+retina%5BTitle%5D)+AND+%22Disease+Models+%26+Mechanisms%22%5BJournal%5D)">Phenotypic and functional characterization of Bst+/- mouse retina.</a> Disease Models & Mechanisms. 8 (8), 969-976 (2015).</li>
</ol>

The authors have nothing to disclose.

Techniques to assess and characterize hyaloid vessels are intuitive and necessary procedures to observe the hyaloid vessel regression in animal models, to allow studies on the mechanisms underlying the vascular regression during development. While the in vivo retinal imaging allows the longitudinal observation of hyaloid regression in the same animal, access to a rodent fundus imaging system for OCT and FFA may be a limiting factor. In addition, in vivo imaging in live mice is not feasible before they open their eyes. Th...

The blood supply in the eye is essential to ensure the normal development of the retina and surrounding ocular tissues and to equip a proper visual function. There are three vascular beds in the eye: the retinal vasculature, the choroid, and a transient embryonic circulatory network of hyaloid vessels. The development of the ocular vasculature requires spatial and temporal coordination throughout embryogenesis and tissue maturation. Among the three vascular beds, the hyaloid vasculature is the first functional blood supply system to provide nutrition and oxygen to the newly formed embryonic lens and the developing retina. Hyaloid vessels regress at the same time that the retina vasculatures develop and mature1. The regression of hyaloid vasculature is pivotal to allow a clear visual pathway for the development of visual function; hence, this vascular regression process is as important as the growth of retinal vasculature. Impaired hyaloid regression may lead to eye diseases. Moreover, the regression of hyaloid vessels provides a model system to investigate the cellular and molecular mechanisms involved in the regulation of vascular regression, which may have implications for the angiogenic regulation in other organs as well.
The hyaloid vasculature, derived from the hyaloid artery (HA), is composed of vasa hyaloidea propria (VHP), tunica vasculosa lentis (TVL), and pupillary membrane (PM). It provides nourishment to the developing retina, the primary vitreous, and the lens during embryonic development2. Arising from the HA, VHP branches anteriorly through the vitreous to the lens. The TVL cups the posterior surface of the lens capsule, and anastomoses to the PM, which connects to the anterior ciliary arteries, covering the anterior surface of the lens2,3, resulting in the formation of a network of vessels in the PM3,4,5. Interestingly, there are no veins in the hyaloid vasculature, and the system makes use of choroidal veins to accomplish venous drainage.
In the human embryo, the hyaloid vasculature is nearly complete at approximately the ninth week of gestation and starts to regress when the first retinal vessels appear, during the fourth month of gestation2. Beginning with atrophy of the VHP, regression of the capillary networks of the TVL, the PM, and lastly, the HA occurs subsequently2,3. Meanwhile, the primary vitreous retracts and the secondary vitreous starts to form, composed of the extracellular matrix components, including collagen fibers. By the sixth month of gestation, the primary vitreous is reduced to a small transparent canal extending from the optic nerve disc to the lens, called the Cloquet&#8217;s canal or hyaloid canal, and the secondary vitreous becomes the main component of the posterior segment2,3. The hyaloid circulation vanishes mostly at 35 to 36 weeks of gestation, just before birth3.
Unlike humans, in whom hyaloid vasculature is completely regressed at birth, the mouse hyaloid vascular system starts to regress after birth. As the mouse retina is born avascular and retinal vessels develop postnatally, hyaloid vessels regress concurrently from postnatal day (P) 4 and are mostly completely regressed by P216 (Figure 1). The PM disappears first between P10 and P12, and the VHP disappears between P12 and P16, while a small number of TVL and HA cells remain even at P16, and by P21 the hyaloid vascular system regression is almost complete6. In the meantime, retinal vasculature begins developing after birth. The superficial layer of vascular plexus fully extends to the peripheral retina at P7&#8211;P8, the deep layer (located in the outer plexiform layer) develops from P7&#8211;P12, and finally, the intermediate plexus in the inner plexiform layer develops between P12 and P157. As the retinal vasculature develops, it gradually replaces the function of concomitantly regressing hyaloid vessels, providing nutrition and oxygen to the developing eye. The postnatal occurrence of hyaloid vessel regression in mice provides an easily accessible experimental model to observe and study the hyaloid vasculature, as well as the molecular basis governing vascular regression processes under both physiological and pathological conditions8.
Failure of hyaloid regression can be seen in diseases such as PHPV, which is a rare congenital developmental anomaly of the eye resulting from a failed or incomplete regression of the embryological, primary vitreous and hyaloid vasculature9. The mechanisms regulating the regression process of hyaloid vasculature are complicated and broadly studied. One major molecular pathway essential for the normal regression of hyaloid vessels is the Wnt signaling pathway10, as genetic mutations in this pathway affecting both Wnt ligand and receptors have been linked with PHPV in humans9. Experimental studies identified a Wnt ligand, Wnt7b, which is produced by macrophages around hyaloid vessels in the developing eye to mediate this regression process. Wnt7b activates Wnt signaling by binding with the receptors frizzled4 (FZD4)/LRP5 in adjacent endothelial cells to initiate cell apoptosis, leading to the regression of hyaloid vessels10. As a result, Wnt7b-deficient mice show a persistence of hyaloid vessels10. Similarly, a nonconventional Wnt ligand, Norrin (encoded by the Ndp gene), also binds to FZD4/LRP5 to induce the hyaloid vessel regression during development. Ndpy/-, Lrp5-/-, and Fzd4-/- mice all display postponed hyaloid vessel regression, supporting a critical regulatory role of Wnt signaling11,12,13,14,15,16. Moreover, another Wnt coreceptor LRP6 overlaps with LRP5 in their function on modulating the Wnt signaling pathway in hyaloid vascular endothelial cells17. Other factors that may also contribute to hyaloid regression include the hypoxia-inducible factor18,19, vascular endothelial growth factor20,21, collagen-1822,23, Arf24, angiopoietin-225, and bone morphogenetic protein-426. In this paper, we use Lrp5-/- mice as a model of persistent hyaloid vessels to demonstrate the techniques of assessing and characterizing hyaloid vasculature through both in vivo and ex vivo methods.
The visualization of hyaloid vasculature in vivo and ex vivo is essential for studying the mechanisms of hyaloid vessel regression. Current methods to observe hyaloid vasculature mainly focus on visualizing and analyzing the VHP and HA, through OCT and FFA images, eye cross sections, and the hyaloid flat mount. OCT and FFA are powerful in vivo imaging tools, allowing longitudinal observation in live animals after they have opened their eyes. Moreover, isolated hyaloid flat mount provides visualization of the whole hyaloid vasculature and a means to achieve an accurate quantification of the vessel numbers. Yet the delicate and fragile nature of hyaloid vessels and the resulting technical difficulties of its isolation may have limited its use in eye research somewhat10,17,27. In this paper, we provide a detailed protocol of the visualization of hyaloid vessels, combining both in vivo live retinal imaging and ex vivo isolated hyaloid flat mount to enhance the feasibility of these techniques. This protocol has been adapted with modification and expansion from previous publications on the in vivo method of live fundus and OCT imaging28 and the ex vivo method of isolated hyaloid flat mount11.

<table><tbody><tr><td>AK-Fluor (fluorescein injection, USP)</td><td>Akorn</td><td>17478-253-10</td><td></td></tr><tr><td>Anti-CD31 antibody</td><td>Abcam</td><td>ab28364</td><td></td></tr><tr><td>Antifade mounting medium</td><td>Thermo Fisher</td><td>S2828</td><td></td></tr><tr><td>Antifade Mounting Medium with DAPI</td><td>Vector Laboratories</td><td>H-1200</td><td></td></tr><tr><td>Artificial tear eyedrop</td><td>Systane</td><td>N/A</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Sigma-Aldrich</td><td>A2058</td><td></td></tr><tr><td>C57BL/6J mice</td><td>The Jackson Laboratory</td><td>Stock NO: 000664</td><td></td></tr><tr><td>Calcium chloride (CaCl2)</td><td>Sigma-Aldrich</td><td>C1016</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM3050S</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM3050 S</td><td></td></tr><tr><td>Cyclopentolate hydrochloride and phenylephrine hydrochloride eyedrop</td><td>Cyclomydril</td><td>N/A</td><td></td></tr><tr><td>Gelatin&amp;nbsp;</td><td>Sigma-Aldrich</td><td>G9382</td><td></td></tr><tr><td>Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 488</td><td>ThermoFisher Scientific</td><td>A-11008</td><td></td></tr><tr><td>Heating board</td><td>Lab-Line Instruments Inc.</td><td>N/A</td><td></td></tr><tr><td>Isolectin GS-IB4, 594 conjugate</td><td>ThermoFisher Scientific</td><td>I21413</td><td></td></tr><tr><td>Ketamine hydrochloride injection</td><td>KetaVed</td><td>NDC 50989-996-06</td><td></td></tr><tr><td>&lt;em&gt;Lrp5-/-&lt;/em&gt; mice</td><td>The Jackson Laboratory</td><td>Stock NO. 005823</td><td>Developed by Deltagen Inc., San Mateo, CA</td></tr><tr><td>Micron IV and OCT</td><td>Phoenix Research Labs</td><td>N/A</td><td>Imaging software: InSight</td></tr><tr><td>Microscope</td><td>Zeiss</td><td>discovery v8</td><td></td></tr><tr><td>Microsurgery forceps</td><td>Scanlan International</td><td>4004-05</td><td></td></tr><tr><td>Microsurgery scissors</td><td>Scanlan International</td><td>6006-44</td><td></td></tr><tr><td>Optimal cutting temperature compound</td><td>Tissue-Tek</td><td>4583</td><td></td></tr><tr><td>Optimal cutting temperature compound</td><td>Agar Scientific</td><td>AGR1180</td><td></td></tr><tr><td>Paraformaldehyde (16%)</td><td>Electron&amp;nbsp;Microscopy&amp;nbsp;Sciences</td><td>15710</td><td></td></tr><tr><td>Peel-A-Way disposable embedding molds (tissue molds)</td><td>Fisher Scientific</td><td>12-20</td><td></td></tr><tr><td>Phosphate-buffered saline (PBS) buffer (10X)</td><td>Teknova</td><td>P0496</td><td></td></tr><tr><td>Slide cover glass</td><td>Premiere</td><td>94-2222-10</td><td></td></tr><tr><td>Superfrost microscope slides&amp;nbsp;</td><td>Fisherbrand</td><td>12-550-15</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>X100</td><td></td></tr><tr><td>Xylazine sterile solution</td><td>Akorn: AnaSed</td><td>NDC: 59399-110-20</td><td></td></tr></tbody></table>

assessment and characterization of hyaloid vessels in mice

In the eye, the embryonic hyaloid vessels nourish the developing lens and retina and regress when the retinal vessels develop. Persistent or failed regression of hyaloid vessels can be seen in diseases such as persistent hyperplastic primary vitreous (PHPV), leading to an obstructed light path and impaired visual function. Understanding the mechanisms underlying the hyaloid vessel regression may lead to new molecular insights into the vascular regression process and potential new ways to manage diseases with persistent hyaloid vessels. Here we describe the procedures for imaging hyaloid in live mice with optical coherence tomography (OCT) and fundus fluorescein angiography (FFA) and a detailed technical protocol of isolating and flat-mounting hyaloid ex vivo for quantitative analysis. Low-density lipoprotein receptor-related protein 5 (LRP5) knockout mice were used as an experimental model of persistent hyaloid vessels, to illustrate the techniques. Together, these techniques may facilitate a thorough assessment of hyaloid vessels as an experimental model of vascular regression and studies on the mechanism of persistent hyaloid vessels.

In vivo imaging of hyaloid vessels in live mice 
Figure 3A reveals cross-sectional views of OCT images for the retina and hyaloid tissues in 3-month-old WT and Lrp5-/- mice, an animal model with persistent hyaloid. The WT eye shows the absence of hyaloid tissue, whereas the Lrp5-/- eye shows two persistent hyaloid vessels derived from the optic nerve head. Figure 3B d...

Watch this Scientific Journal Video about Assessment and Characterization of Hyaloid Vessels in Mice at JoVE.com

Assessment and Characterization of Hyaloid Vessels in Mice

This protocol describes both in vivo and ex vivo methods to fully visualize and characterize hyaloid vessels, a model of vascular regression in mouse eyes, using optical coherence tomography and fundus fluorescein angiography for the live imaging and ex vivo isolation and subsequent flat mount of hyaloid for quantitative analysis.

In the eye, the embryonic hyaloid vessels nourish the developing lens and retina and regress when the retinal vessels develop. Persistent or failed ...

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Research

JoVE Journal

Medicine

9.2K Views.  Harvard Medical School. This protocol describes both in vivo and ex vivo methods to fully visualize and characterize hyaloid vessels, a model of vascular regression in mouse eyes, using optical coherence tomography and fundus fluorescein angiography for the live imaging and ex vivo isolation and subsequent flat mount of hyaloid for quantitative analysis.

Video: Assessment and Characterization of Hyaloid Vessels in Mice

Assessment of Maternal Vascular Remodeling During Pregnancy in the Mouse Uterus

The placenta mediates the exchange of factors such as gases and nutrients between mother and fetus and has specific demands for supply of blood from the maternal circulation. The maternal uterine vasculature needs to adapt to this temporary demand and the success of this arterial remodeling process has implications for fetal growth. Cells of the maternal immune system, especially natural killer (NK) cells, play a critical role in this process. Here we describe a method to assess the degree of remodeling of maternal spiral arteries during mouse pregnancy. Hematoxylin and eosin-stained tissue sections are scanned and the size of the vessels analysed. As a complementary validation method, we also present a qualitative assessment for the success of the remodeling process by immunohistochemical detection of smooth muscle actin (SMA), which normally disappears from within the arterial vascular media at mid-gestation. Together, these methods enable determination of an important parameter of the pregnancy phenotype. These results can be combined with other endpoints of mouse pregnancy to provide insight into the mechanisms underlying pregnancy-related complications.

This protocol describes a technique to assess changes in the maternal vasculature during pregnancy in mice. Using stereological methods, remodeling of the decidual spiral arteries is assessed quantitatively and the results confirmed qualitatively using immunohistochemistry.

The placenta mediates the exchange of factors such as gases and nutrients between mother and fetus and has specific demands for supply of blood from the ...

Developmental Biology

A Method for Labeling Vasculature in Embryonic Mice

The establishment of a functional blood vessel network is an essential part of organogenesis, and is required for optimal organ function. For example, in the thymus proper vasculature formation and patterning is essential for thymocyte entry into the organ and mature T-cell exit to the periphery. The spatial arrangement of blood vessels in the thymus is dependent upon signals from the local microenvironment, namely thymic epithelial cells (TEC). Several recent reports suggest that disruption of these signals results in thymus blood vessel defects 1,2. Previous studies have described techniques used to label the neonatal and adult thymus vasculature 1,2. We demonstrate here a technique for labeling blood vessels in the embryonic thymus. This method combines the use of FITC-dextran or Griffonia (Bandeiraea) Simplicifolia Lectin I (GSL 1 - isolectin B4) facial vein injections and CD31 antibody staining to identify thymus vascular structures and PDGFR-&beta; to label thymic perivascular mesenchyme 3-5. The option of using cryosections or vibratome sections is also provided. This protocol can be used to identify thymus vascular defects, which is critical for defining the roles of TEC-derived molecules in thymus blood vessel formation. As the method labels the entire vasculature, it can also be used to analyze the vascular networks in multiple organs and tissues throughout the embryo including skin and heart 6-10.

This article describes a method for labeling embryonic skin and thymus blood vessels.

The establishment of a functional blood vessel network is an essential part of organogenesis, and is required for optimal organ function. For example, in ...

Immunology and Infection

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Determining Bile Duct Density in the Mouse Liver

Mouse is broadly used as a model organism to study biliary diseases. To evaluate the development and function of the biliary system, various techniques are used, including serum chemistry, histological analysis, and immunostaining for specific markers. Although these techniques can provide important information about the biliary system, they often do not present a full picture of bile duct (BD) developmental defects across the whole liver. This is in part due to the robust ability of the mouse liver to drain the bile even in animals with significant impairment in biliary development. Here we present a simple method to calculate the average number of BDs associated with each portal vein (PV) in sections covering all lobes of mutant/transgenic mice. In this method, livers are mounted and sectioned in a stereotypic manner to facilitate comparison among various genotypes and experimental conditions. BDs are identified via light microscopy of cytokeratin-stained cholangiocytes, and then counted and divided by the total number of PVs present in liver section. As an example, we show how this method can clearly distinguish between wild-type mice and a mouse model of Alagille syndrome. The method presented here cannot substitute for techniques that visualize the three-dimensional structure of the biliary tree. However, it offers an easy and direct way to quantitatively assess BD development and the degree of ductular reaction formation in mice.

We present a rather simple and sensitive method for accurate quantification of bile duct density in the mouse liver. This method can aid in determining the effects of genetic and environmental modifiers and the effectiveness of potential therapies in mouse models of biliary diseases.

Mouse is broadly used as a model organism to study biliary diseases. To evaluate the development and function of the biliary system, various techniques ...

Visualizing Leukocyte Rolling and Adhesion in Angiotensin II-Infused Mice: Techniques and Pitfalls

Epifluorescence intravital video microscopy (IVM) of blood vessels is an established method to evaluate the activation of immune cells and their ability to role and adhere to the endothelial layer. Visualization of circulating cells by injection of fluorescent dyes or fluorophore-coupled antibodies is commonly used. Alternatively, fluorescent reporter mice can be used. Interactions of leukocytes, in particular lysozyme M+ (LysM+) monocytes, with the vessel wall play pivotal roles in promoting vascular dysfunction and arterial hypertension. We here present the technique to visualize and quantify leukocyte rolling and adhesion in carotid arteries in angiotensin II (AngII)-induced hypertension in mice by IVM.
The implantation of a catheter damages the vascular wall and leads to altered blood cell responses. We compared different injection techniques and administration routes to visualize leukocytes in a LysMCre+IRG+ mouse with widespread expression of red fluorescent protein and conditional expression of green fluorescent protein in LysM+ cells. To study LysM+ cell activation, we used AngII infused mice in which rolling and adhesion of leukocytes to the endothelium is increased. We either injected acridine orange using a jugular catheter or directly though the tail vein and compared the amount of rolling and adhering cells. We found that jugular catheter implantation per se increased the number of rolling and adhering LysM+ cells in sham-infused LysMCre+IRG+ mice compared to controls. This activation was augmented in AngII-infused mice. Interestingly, injecting acridine orange directly through the tail vein did not increase LysM+ cell adhesion or rolling in sham-infused mice. We thereby demonstrated the importance of transgenic reporter mice expressing fluorescent proteins to not interfere with in vivo processes during experimentation. Furthermore, tail vein injection of fluorescent tracers might be a possible alternative to jugular catheter injections.

This manuscript describes the use of transgenic reporter mice and different administration routes of fluorescent dyes in angiotensin II-induced hypertension using intravital video microscopy of blood vessels to evaluate the activation of immune cells and their ability to roll and adhere to the endothelium.

Epifluorescence intravital video microscopy (IVM) of blood vessels is an established method to evaluate the activation of immune cells and their ability ...

Assessing Therapeutic Angiogenesis in a Murine Model of Hindlimb Ischemia

Critical limb ischemia (CLI) is a serious condition that entails a high risk of lower limb amputation. Despite revascularization being the gold-standard therapy, a considerable number of CLI patients are not suited for either surgical or endovascular revascularization. Angiogenic therapies are emerging as an option for these patients but are currently still under investigation. Before application in humans, those therapies must be tested in animal models and its mechanisms must be clearly understood. An animal model of hindlimb ischemia (HLI) has been developed by the ligation and excision of the distal external iliac and femoral arteries and veins in mice. A comprehensive panel of tests was assembled to assess the effects of ischemia and putative angiogenic therapies at functional, histologic and molecular levels. Laser Doppler was used for the flow measurement and functional assessment of perfusion. Tissue response was evaluated by the analysis of capillary density after staining with the anti-CD31 antibody on histological sections of gastrocnemius muscle and by measurement of collateral vessel density after diaphonization. Expression of angiogenic genes was quantified by RT-PCR targeting selected angiogenic factors exclusively in endothelial cells (ECs) after laser capture microdissection from mice gastrocnemius muscles. These methods were sensitive in identifying differences between ischemic and non-ischemic limbs and between treated and non-treated limbs. This protocol provides a reproducible model of CLI and a framework for testing angiogenic therapies.

Here, a critical hindlimb ischemia experimental model is presented followed by a battery of functional, histologic and molecular tests&#160;to assess the effectiveness of angiogenic therapies.&#8203;

Critical limb ischemia (CLI) is a serious condition that entails a high risk of lower limb amputation. Despite revascularization being the gold-standard ...

Monitoring Functionality and Morphology of Vasculature Recruited by Factors Secreted by Fast-growing Tumor-generating Cells

The subcutaneous matrigel plug assay in mice is a method of choice for the in vivo evaluation of pro- and anti-angiogenic factors. In this method, desired factors are introduced into cold-liquid ECM-mimic gel which, after subcutaneous injection, solidifies to form an environment mimicking the cancer milieu. This matrix permits the penetration of host cells, such as endothelial cells, and therefore, the formation of vasculature.
Herein we propose a new modified matrigel plug assay, which can be exploited to illustrate the angiogenic potential of a pool of factors secreted by cancer cells, as opposed to a specific factor (e.g., bFGF and VEGF) or agent. The plug containing ECM-mimic gel is utilized to introduce the host (i.e., mouse) with a pool of factors secreted to the C.M. of fast-growing tumor-generating glioblastoma cells. We have previously described an extensive comparison of the angiogenic potential of U-87 MG human glioblastoma and its dormant-derived clone, in this system model, showing induced angiogenesis in the U-87 MG parental cells. The C.M. is prepared by filtering collected media from confluent tissue culture plates of either cell line following 48 hr incubation. Hence, it contains only factors secreted by the cells, without the cells themselves. Described here is the combination of two imaging modalities, microbubbles contrast-enhanced ultrasound imaging and intravital fibered-confocal endomicroscopy, for an accurate, real-time characterization of the extent, morphology and functionality of newly-formed blood vessels within the plugs.

We describe a matrigel plug assay to illustrate angiogenic potential of a pool of factors secreted by cancer cells using two complementary imaging modalities, ultrasound and endomicroscopy. The matrigel, an extracellular matrix (ECM)-mimic gel, is utilized to introduce the host (mouse) with angiogenic factors secreted to the conditioned media (C.M.).&#160;

The subcutaneous matrigel plug assay in mice is a method of choice for the in vivo evaluation of pro- and anti-angiogenic factors. In this method, desired ...

Biology

Quantification of Vascular Parameters in Whole Mount Retinas of Mice with Non-Proliferative and Proliferative Retinopathies

Retinopathies are a heterogeneous group of diseases that affect the neurosensory tissue of the eye. They are characterized by neurodegeneration, gliosis&#160;and a progressive change in vascular function and structure. Although the onset of the retinopathies is characterized by subtle disturbances in visual perception, the modifications in the vascular plexus are the first signs detected by clinicians. The absence or presence of neovascularization determines whether the retinopathy is classified as either non-proliferative (NPDR) or proliferative (PDR). In this sense, several animal models tried to mimic specific vascular features of each stage to determine the underlying mechanisms involved in endothelium alterations, neuronal death and other events taking place in the retina. In this article, we will provide a complete description of the procedures required for the measurement of retinal vascular parameters in adults and early birth mice at postnatal day (P)17. We will detail the protocols to carry out retinal vascular staining with Isolectin GSA-IB4 in whole mounts for later microscopic visualization. Key steps for image processing with Image J Fiji software are also provided, therefore, the readers will be able to measure vessel density, diameter and tortuosity, vascular branching, as well as avascular and neovascular areas. These tools are highly helpful to evaluate and quantify vascular alterations in both non-proliferative and proliferative retinopathies.

This article describes a well-established and reproducible lectin stain assay for the whole mount retinal preparations and the protocols required for the quantitative measurement of vascular parameters frequently altered in proliferative and non-proliferative retinopathies.

Retinopathies are a heterogeneous group of diseases that affect the neurosensory tissue of the eye. They are characterized by neurodegeneration, ...

Neuroscience

Sample Preparation for Computed Tomography-based Three-dimensional Visualization of Murine Hind-limb Vessels

Blood vessels are complex networks with tree-like structures, and vascular networks are essential for maintaining both circulation and maintaining organ function. Clarifying the mechanism of blood vessel formation is therefore extremely useful for elucidating developmental processes and pathological mechanisms. Murine hind-limb vessels are often used as a model for physiological and pathological angiogenesis. Evaluation is mainly performed via a two-dimensional method using tissue sections. However, methods for evaluating three-dimensional (3D) vascular morphology are particularly limited. This paper introduces a method for visualizing murine hind-limbs using computed tomography (CT). Radiation-opaque resin is injected through the descending aorta, and whole vessels are filled with dye. By adjusting the time of dye injection, arterial-specific filling is also possible, and samples can be obtained with any micro-X-ray CT device. This contrast method provides a basic technique for the 3D evaluation of murine blood vessels in the lower extremities. Furthermore, this method can be used to visualize all blood vessels below the diaphragm and evaluate blood vessels in the abdominal organs.

Here, we describe a visualization and quantification method for murine hind-limb vessels using micro-X-ray computed tomography.

Blood vessels are complex networks with tree-like structures, and vascular networks are essential for maintaining both circulation and maintaining organ ...

Dolichoectasia: Vascular Dementia and Alzheimer&#39;s Disease Models

A Visual Approach for Inducing Dolichoectasia in Mice to Model Large Vessel-Mediated Cerebrovascular Dysfunction

The blood-brain (BBB) is a crucial system that regulates selective brain circulation with the periphery, as an example, allowing necessary nutrients to enter and expel excessive amino acids or toxins from the brain. To model how the BBB can be compromised in diseases like vascular dementia (VaD) or Alzheimer's disease (AD), researchers developed novel methods to model vessel dilatation. A compromised BBB in these disease states can be detrimental and result in the dysregulation of the BBB leading to untoward and pathological consequences impacting brain function. We were able to modify an existing technique that enabled us to inject directly into the Cisterna magna (CM) to induce dilatation of blood vessels using elastase, and disrupt the tight junctions (TJ) of the BBB. With this method, we were able to see various metrics of success over previous techniques, including consistent blood vessel dilatation, reduced mortality or improved recovery, and improving the fill/opacifying agent, a silicone rubber compound, delivery for labeling blood vessels for dilatation analysis. This modified minimally invasive method has had promising results, with a 19%-32% increase in sustained dilatation of large blood vessels in mice from 2 weeks to 3 months post-injection. This improvement contrasts with previous studies, which showed increased dilatation only at the 2 week mark. Additional data suggests sustained expansion even after 9.5 months. This increase was confirmed by comparing the diameter of blood vessels of the elastase and the vehicle-injected group. Overall, this technique is valuable for studying pathological disorders that affect the central nervous system (CNS) using animal models.

Author Spotlight: Modeling Vascular Contributions to Alzheimer's Disease in Transgenic Mice

We demonstrate chemically inducing large blood vessel dilatation in mice as a model for investigating cerebrovascular dysfunction, which can be used for vascular dementia and Alzheimer's disease modeling. We also demonstrate visualizing the vasculature by injecting silicone rubber compound and providing clear visual guidance for measuring changes in blood vessel size.

The blood-brain (BBB) is a crucial system that regulates selective brain circulation with the periphery, as an example, allowing necessary nutrients to ...