This work was supported by grants from the Research to Prevent Blindness Foundation (Career Development Award to P.R.W. and an unrestricted grant to the Department of Ophthalmology and Visual Sciences at Washington University School of Medicine in St. Louis), National Glaucoma Research (a program of BrightFocus Foundation), and the McDonnell Center for Cellular and Molecular Neurobiology. Z.W. is supported by an Institutional National Research Service Award T32 EY013360. This work was also supported by the Hope Center Viral Vectors Core at Washington University School of Medicine.

<ol>
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The authors have nothing to disclose.

The two-photon imaging procedure described herein enables longitudinal in vivo imaging of the mouse retina. Repeatable images of the same region of retina can be obtained for a continuous period of up to 6 or more h under isoflurane. The mouse can also be imaged on different days using cellular and vascular landmarks to locate the same imaging area (Figure 3). The use of a clear gel immersion combined with cover glass for this purpose has previously been applied to a range of procedures, including visualization of retina for subretinal injection, laser-induced retinal injury models, and fundus imaging20,21,22.
The anatomy of the eye presents unique challenges to in vivo imaging, as the high optical power of the mouse cornea and lens impedes direct imaging though the pupil without correction. Several other in vivo imaging methods rely on the use of a plano-concave contact lens for correction of the anterior optics of the mouse eye7,17,18,19.&#160;With only optical correction at the cornea, the high optical power of the mouse lens results in an inevitable amount of parallax, particularly of structures in the peripheral scan field, manifesting as stretching and translational movement in the X-Y dimension at different Z-planes. To minimize image parallax-related distortion in X and Y dimensions, it is crucial that the mouse eye is oriented such that the tangent plane to the retina at the imaging area is perpendicular to the microscope light path. The setup described here is conducive to precise manipulation of the angle of the eye to achieve this alignment. An adjustable mouse head holder that permits rotation along two axes allows for easy manual adjustments of the angle of the eye as the experimenter scrolls through the Z-dimension to minimize parallax. This tilting also circumvents the field stop effect of the pupil to permit imaging greater areas of the retina. The restraint of the head holder also greatly reduces motion artifacts caused by respiration.
Care must be taken to maintain clarity of the mouse eye, as image quality will deteriorate with opacification during continuous imaging. Frequent reapplication of lubricant gel during imaging, and ointment application after each imaging session help to prevent the eye from drying and developing opacities. Some corneal opacities will spontaneously resolve after 24-48 h. The use of clear gel and cover glass as described in this protocol provides for similar image quality and aberration correction as a contact lens7, while allowing for easier adjustments of the eye angle without the need to realign the cover glass. Additionally, the gel provides continual hydration to the eye, making it possible to perform acute imaging sessions of up to several hours. Finally, since the cover glass does not contact the cornea, it causes minimal irritation to the eye that may reduce optical clarity for repeat imaging sessions.
A limitation of this approach is the fact that optical aberrations are not entirely corrected. While this severely diminishes axial resolution due to the heavy parallax, quantitative measurements of the soma can be obtained in single-image planes. It should be noted that as fluorescence signal intensity of retinal neurons is dependent on sample alignment with this method, excitation and emission ratiometric based sensors are more appropriate for experiments comparing samples chronically across different imaging sessions. An approach to correct optical aberrations at the system level is adaptive optics, which allows for subcellular resolution in the retina8,9,14,21. However, adaptive optics requires highly specialized equipment and extensive expertise to implement.
Alternative approaches to two-photon in vivo&#160;retinal imaging are confocal microscopy or ophthalmoscopy6. The approach presented here should be readily translatable to widefield or&#160;confocal microscopy. Single photon imaging is perhaps more robust and poses less risk of damaging the retina due to high energy of the two-photon laser necessary to achieve efficient two-photon effect through the cornea and lens of the eye. To avoid two-photon laser damage, the threshold for maximal laser power should be empirically determined by examining wholemount retinas after completion of imaging experiments and immunostaining for cell types in the layers imaged. In the system presented here, RGCs were labelled with the pan-RGC marker, Rbpms, and densities were normal up to 45 mW imaging power, whereas 55 mW caused a significant loss of RGCs (not shown).
A drawback of single-photon imaging is the fact that this approach will very heavily stimulate the native visual circuits of the retina compared to two-photon imaging23. Previous experiments using retinal wholemounts or eyecup preparations have shown that two-photon laser scanning elicits circuit activation that is largely transient24. Here, imaging of RGC activity with the Ca2+ sensor Twitch2b shows that the onset of laser scanning induces Ca2+ elevations, which return to baseline over the course of 5-20 s in most RGCs (Figure 8). Given that the laser power in this protocol is in the range of previous experiments reporting in vivo&#160;retinal light response8, the currently described method is likely amenable to recordings of circuit activity in the retina.&#160;Such considerations are important for experiments that may be influenced by circuit activity.
This protocol demonstrates in vivo imaging of two types of retinal neurons, RGCs and amacrine cells. Similar labeling of other major cell types can be achieved, including horizontal cells (Cx57-Cre25), bipolar cells (Chx10-Cre26; mGluR6-GFP27), cone photoreceptors (S- or M-opsin-Cre28), rod photoreceptors (Nrl-Cre29), M&#252;ller glia (Foxg1-Cre26), and pericytes (NG2-DsRed9). Transgenic mice are also available to label discrete subsets of RGCs (e.g., KCNG4-Cre for &#945;RGCs30; OPN4-Cre for ipRGCs31; JAM-B-CreER for J-RGCs32) and amacrine cells (e.g., ChAT-Cre for starburst amacrine cells26 and neuropeptide promoter drivers for various amacrine cell subtypes 3,34). Viral vectors can be used to target specific cell populations in lieu of transgenic mice. Intravitreal injections of AAV2 with a ubiquitous CAG promoter element almost exclusively label RGCs, amacrine cells and horizontal cells25. Pairing the modified AAV2.7m8-Y444F capsid with an engineered mGluR6 promoter construct allows for broad labeling of ON bipolar cells35. Subretinal injections of AAV lead to an enrichment of photoreceptors, with serotype AAV2/5 having the highest transduction efficiency36. Shh10, a modified AAV6 capsid protein, paired with glial fibrillary acidic protein promoter elements has been demonstrated specific for M&#252;ller glia37.
The ability to observe cells in the central nervous system with a completely non-invasive approach can be used to study both basic properties of neural circuits8, as well as mechanisms of neurodegeneration3,4,5,6,38. Many blinding diseases target cellular populations in the retina, and in vivo imaging approaches in mice have been used to study optic nerve injury1,3,4, macular degeneration13, stroke5, glaucoma2,6, and uveitis7. Furthermore, many central nervous system neurodegenerative conditions manifest in the retina including Alzheimer&#39;s disease39, multiple sclerosis40, and Parkinson&#39;s disease41. Therefore, this readily accessible technique for in vivo imaging of the retina can be applied as a tool to study a broad set of neurodegenerative conditions.

In vivo visualization of the central nervous system generally requires invasive procedures like skull thinning and installation of glass windows or optical relay lenses. The retina is the only structure in the nervous system that can be directly observed without the need for invasive preparation as it natively receives light from the environment. The ease of optical access to the retina makes it an attractive model system for studying the central nervous system.
Live fluorescent imaging of the retina in mice has been used to track RGC death in models of glaucoma1,2, optic nerve injury1,3,4, and stroke5, as well as changes in microglial activation6,7,8 and vasculature9 in degenerative conditions. Intrinsic signals can also be used to visualize photoreceptors10,11,12 and retinal pigment epithelial cells13. Many approaches to in vivo imaging of the retina use either highly specialized devices specifically designed for ophthalmological purposes6 or highly modified optical systems to correct for the native aberrations of the cornea and lens8,9,11,12,13,14.
The present protocol demonstrates an approach to in vivo imaging of fluorescent signals in the retina at cellular resolution, utilizing a basic method of partially correcting for the anterior optics of the mouse eye. This strategy requires very minor adaptations to multiphoton microscope setups that are commonly used for in vivo imaging of the brain. As this approach is straightforward to set up, and the mice are under little stress, it is conducive to perform time-lapse experiments over both acute and chronic durations. Additionally, genetic and organic dye-based procedures that label individual retinal components, including RGCs, amacrine cells, microglia, and vasculature, are compatible with this imaging technique and enable in vivo observation of cell types and structures critical for retinal function. These tools can be adapted to label most other neuronal cell types as well as glial and vascular components of the retina.

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		<tr>
			<td>#1.5 coverslip</td>
			<td>ThermoFisher</td>
			<td>152440</td>
			<td>Richard-Allan #1.5 24 mm x 40 mm</td>
		</tr>
		<tr>
			<td>50 mL glass syringe</td>
			<td>Hamilton</td>
			<td>80950</td>
			<td>22G cemented needle</td>
		</tr>
		<tr>
			<td>Adeno-associated virus (AAV2)</td>
			<td>Hope Center Viral Core</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Anesthesia Air Pump</td>
			<td>RWD Life Science</td>
			<td>R510-30</td>
			<td></td>
		</tr>
		<tr>
			<td>Atropine</td>
			<td>Sigma</td>
			<td>A0132</td>
			<td>For pupil dilator solution</td>
		</tr>
		<tr>
			<td>Basic Small Animal Anesthesia Device</td>
			<td>RWD Life Science</td>
			<td>R500IE</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass capillary</td>
			<td>Sutter</td>
			<td>B150-86-10</td>
			<td>Outside diameter 1.50 mm, inside diameter 0.86 mm, length 10 cm</td>
		</tr>
		<tr>
			<td>CFP/YFP filter cube</td>
			<td>Chroma</td>
			<td>custom</td>
			<td>480/40, 505 long pass, 535/30</td>
		</tr>
		<tr>
			<td>ChromoFlex - Two channel PMT detection unit</td>
			<td>Scientifica</td>
			<td>S-MPLG-1002</td>
			<td></td>
		</tr>
		<tr>
			<td>Circulating heating pump</td>
			<td>Braintree Scientific</td>
			<td>tp-700</td>
			<td>Set to 37 &#176;C</td>
		</tr>
		<tr>
			<td>Cling film</td>
			<td>VWR</td>
			<td>10713-916</td>
			<td></td>
		</tr>
		<tr>
			<td>Compact Filter Holder</td>
			<td>ThorLabs</td>
			<td>DH1</td>
			<td>Holds coverslip over mouse eye</td>
		</tr>
		<tr>
			<td>Cx3cr1-GFP transgenic mice (B6.129P2(Cg)-Cx3cr1tm1Litt/J)</td>
			<td>The Jackson Laboratory</td>
			<td>005582</td>
			<td></td>
		</tr>
		<tr>
			<td>DAQ controller chassis</td>
			<td>National Instruments</td>
			<td>PXIe-1073</td>
			<td></td>
		</tr>
		<tr>
			<td>Data acquisition device</td>
			<td>National Instruments</td>
			<td>BNC-2090A</td>
			<td></td>
		</tr>
		<tr>
			<td>Evans Blue dye</td>
			<td>Fisher Scientific</td>
			<td>AAA1677409</td>
			<td></td>
		</tr>
		<tr>
			<td>FPGA module with digitizer</td>
			<td>National Instruments</td>
			<td>NI-5734</td>
			<td></td>
		</tr>
		<tr>
			<td>Gas Evacuation Apparatus</td>
			<td>RWD Life Science</td>
			<td>R546W</td>
			<td></td>
		</tr>
		<tr>
			<td>GenTeal Severe lubricant eye gel&#160;&#160;</td>
			<td>Alcon</td>
			<td>(from local pharmacy)</td>
			<td>For use during imaging</td>
		</tr>
		<tr>
			<td>GFP/Red filter cube</td>
			<td>Chroma</td>
			<td>custom</td>
			<td>535/30, 560 long pass, 605/70</td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>McKesson Medical and Surgical</td>
			<td>190147</td>
			<td></td>
		</tr>
		<tr>
			<td>HyperScope Launch Optics for use with Pockels Cell</td>
			<td>Scientifica</td>
			<td>S-MP-101080</td>
			<td></td>
		</tr>
		<tr>
			<td>HyperScope Main module</td>
			<td>Scientifica</td>
			<td>S-MP-100466</td>
			<td></td>
		</tr>
		<tr>
			<td>HyperScope Scan Path</td>
			<td>Scientifica</td>
			<td>S-MP-100406</td>
			<td></td>
		</tr>
		<tr>
			<td>HyperScope X galvo Module</td>
			<td>Scientifica</td>
			<td>MP-100443</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ Fiji software</td>
			<td></td>
			<td>Freeware</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Patterson Veterinary</td>
			<td>NDC 14043-704-06</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane gas filter cannister (active scavenging)</td>
			<td>RWD Life Science</td>
			<td>R510-31</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane gas filter cannister (passive scavenging)</td>
			<td>RWD Life Science</td>
			<td>R510-31S</td>
			<td></td>
		</tr>
		<tr>
			<td>ketamine HCl (100 mg/mL)</td>
			<td>Vedco</td>
			<td>NDC 50989-161-06</td>
			<td></td>
		</tr>
		<tr>
			<td>M32 to M26 adapter</td>
			<td>ThorLabs</td>
			<td>M32M26S</td>
			<td></td>
		</tr>
		<tr>
			<td>MaiTai GUI software</td>
			<td>Spectra-Physics</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>MATLAB software</td>
			<td>MathWorks</td>
			<td>NA</td>
			<td>R2015b</td>
		</tr>
		<tr>
			<td>meloxicam (5 mg/mL)</td>
			<td>Boehringer Ingelheim</td>
			<td>NDC 0010-6013-01</td>
			<td>Analgesic</td>
		</tr>
		<tr>
			<td>Micorscope Objective</td>
			<td>Edmund Optics</td>
			<td>46-404</td>
			<td>Mitutoyo WE715042319</td>
		</tr>
		<tr>
			<td>micropipette puller</td>
			<td>Sutter</td>
			<td>Flaming/Brown Model P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Mineral oil</td>
			<td>Fisher</td>
			<td>BP2629-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini bulldog hemostatic clamp</td>
			<td>Fine Science Tools</td>
			<td>18053-28</td>
			<td></td>
		</tr>
		<tr>
			<td>Miniature EVA Tubing 0.02&#34; ID, 0.06&#34; OD</td>
			<td>McMaster Carr</td>
			<td>1883T1</td>
			<td></td>
		</tr>
		<tr>
			<td>Miniature EVA Tubing 0.05&#34; ID, 0.09&#34; OD</td>
			<td>McMaster Carr</td>
			<td>1883T4</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse head holder</td>
			<td>Narishige</td>
			<td>SGM-4</td>
			<td></td>
		</tr>
		<tr>
			<td>No. 5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Optic Posts 1/2&#34;</td>
			<td>ThorLabs</td>
			<td>TR3-P5</td>
			<td></td>
		</tr>
		<tr>
			<td>Optical power meter kit</td>
			<td>ThorLabs</td>
			<td>PM100D</td>
			<td></td>
		</tr>
		<tr>
			<td>pE-300 Ultra LLG Deivery</td>
			<td>Scientifica</td>
			<td>COO-LED3ULLGs</td>
			<td></td>
		</tr>
		<tr>
			<td>Phenylephrine hydrochloride</td>
			<td>Sigma</td>
			<td>P6126</td>
			<td>For pupil dilator solution</td>
		</tr>
		<tr>
			<td>Pockels cell</td>
			<td>Conoptics</td>
			<td>350-80-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Pockels cell amplifier</td>
			<td>Conoptics</td>
			<td>Model 302RM</td>
			<td></td>
		</tr>
		<tr>
			<td>Proparacaine hydrochloride</td>
			<td>Sigma</td>
			<td>1571001</td>
			<td>For eye immobilization</td>
		</tr>
		<tr>
			<td>Red &#38; Far Red short pass filter Cube</td>
			<td>Chroma</td>
			<td>custom</td>
			<td>560 short pass</td>
		</tr>
		<tr>
			<td>Rotating 1/2&#34; post clamp</td>
			<td>ThorLabs</td>
			<td>SWC</td>
			<td></td>
		</tr>
		<tr>
			<td>ScanImage package</td>
			<td>Vidrio Technologies</td>
			<td>Freeware</td>
			<td>Image acquisition software; Version 5.4.0 (2018); requires MATLAB</td>
		</tr>
		<tr>
			<td>sodium chloride solution, sterile (0.9%)</td>
			<td>Fresenius Kabi</td>
			<td>NDC 63323-186-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>S9 E</td>
			<td></td>
		</tr>
		<tr>
			<td>Tabletop centrifuge</td>
			<td>Oxford</td>
			<td>Benchmate C8</td>
			<td></td>
		</tr>
		<tr>
			<td>Terramycin oxytetracycline/polymyxin B antibiotic ophthalmic ointment</td>
			<td>Zoetisus</td>
			<td>NA</td>
			<td>For use after intravitreal injection</td>
		</tr>
		<tr>
			<td>ThermoRack cooling system</td>
			<td>Solid State Cooling Systems</td>
			<td>ThermoRack 401</td>
			<td>Set to 20 &#176;C</td>
		</tr>
		<tr>
			<td>Ultrafast Ti:Sapphire laser</td>
			<td>Spectra-Physics</td>
			<td>Mai Tai DeepSee</td>
			<td></td>
		</tr>
		<tr>
			<td>Vgat-Cre transgenic mice (Slc32a1tm2(cre)Lowl/J)</td>
			<td>The Jackson Laboratory</td>
			<td>016962</td>
			<td></td>
		</tr>
		<tr>
			<td>VGlut2-Cre transgenic mice (Slc17a6tm2(cre)Lowl/J)</td>
			<td>The Jackson Laboratory</td>
			<td>016963</td>
			<td></td>
		</tr>
		<tr>
			<td>VivoScope for In Vivo Imaging</td>
			<td>Scientifica</td>
			<td>S-MPVS-1200-00P</td>
			<td></td>
		</tr>
		<tr>
			<td>White petrolatum-mineral oil lubricant eye ointment</td>
			<td>Stye</td>
			<td>NA</td>
			<td>For use after imaging</td>
		</tr>
		<tr>
			<td>xylazine HCl (20 mg/mL)</td>
			<td>Akorn</td>
			<td>NDC 59399-110-20</td>
			<td></td>
		</tr>
	</tbody>
</table>

transpupillary two-photon in vivo imaging of the mouse retina

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

Various transgenic, viral vector, or inorganic dye-labeling approaches can be used to specifically visualize several retinal cells types and structures in vivo using a simple adaptation of a basic multiphoton microscope. To visualize RGCs and amacrine cells, VGlut2-Cre and VGat-Cre transgenic mice, respectively, were given an intravitreal injection of a Cre-dependent AAV expression construct encoding Twitch2b, a cytoplasmic fluorescence resonance energy transfer (FRET)-based Ca2+ sensor that contains cyan and yellow fluorescent proteins (CFP and YFP, respectively) and the Ca2+ binding domain of troponin15. In VGlut2-Cre mice, RGC somas are clearly discernable, and fascicles of axons are often apparent (Figure 3).
It should be noted that the trajectory of axons and the negative image of the vasculature makes it very straightforward to identify the optic nerve head in VGlut2-Cre mice, which is useful as a landmark in chronic imaging experiments (Figure 3). Although amacrine cells appear less bright than RGCs, possibly due to their smaller soma size and/or less efficient AAV transduction, their somas are still readily apparent in the inner nuclear layer. In contrast to RGCs, amacrine cell neurites are more often observed in the inner plexiform layers (Figure 4). Retinal microglia can be imaged in the Cx3cr1-GFP transgenic mouse line6. Microglia associate with the vasculature, making it possible to find the same region in time-lapse imaging experiments.
This approach can be used to track the dynamics of fine microglia processes, a procedure that has better spatial resolution in single-plane images, or if maximum-intensity projections are prepared focusing on individual cells (Figure 5). The poor axial resolution caused by optical aberration through the mouse lens precludes examination of fine details in the z-dimension. To determine whether this imaging technique can observe degenerative changes in cellular ultrastructure, 1 &#181;L of 50 mM N-methyl-D-aspartate (NMDA) was injected into the vitreous to induce excitotoxic lesion. One day after injection, the microglia demonstrated short processes or amoeboid morphology (Figure 5) in accordance with previous reports16. It should be noted that cells in the Cx3cr1-GFP transgenic line exhibited more uniform and complete fluorescent protein expression across the cellular cohort than in experiments with AAV-mediated delivery of fluorescent protein expression cassettes. The benefits of varied and sparse versus complete and uniform labeling should be considered when designing experiments.
To label retinal vasculature as previously described8, mice were injected intraperitoneally with 200 &#181;L of Evans blue dye (20 mg/mL in sterile saline) 30-60 min before imaging. This led to strong labeling of blood vessels emanating from the optic nerve head (Figure 6). Surprisingly, the fluorescent signal of a single injection persisted for at least seven days. Two distinct methods were used to estimate the dimensions of the in vivo images. First, the same retinal regions were imaged in vivo and after fixation in flattened retinal wholemounts using confocal microscopy (Figure 7). Random cell pairs were selected from four different in vivo&#160;samples, and the true distance between cell pairs was measured in confocal scans and matched with in vivo pixel distance to obtain an average pixel size of 0.99 &#181;m with 1x digital magnification. Using similar methods correlating in vivo images with confocal wholemount scans has revealed that a single head position allows for imaging over a roughly 650 mm2 patch of retina.
Repositioning of the head holder along one torsional axis can allow access to a linear region of the retina 2.2 mm in length (not shown). Further, 1 or 2 &#181;m diameter fluorescent microspheres were injected into the eyes of mice and their diameter was measured as full-width half-maximum of line scans from in vivo images with 10x digital zoom. This gave a slightly larger pixel size estimate, but with more variance (Figure 7). Overall, confocal imaging of wholemount samples after completion of in vivo experiments is the most consistent method to assign scale to individual images, as variance in corneal and lens properties may alter image scale from sample to sample.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61970/61970fig1v2.jpg" /> 
Figure 1: Light path schematic. The basic components of the two-photon microscope used in this protocol consist of a Pockels Cell to modulate laser power, a lens pair to reduce the laser beam diameter to match the back aperture of the microscope objective, and a pair of galvo scan mirrors for beam steering. A pair of steering mirrors is present before each major optical component. The focus is controlled by a motor that drives the objective mount. The emission light path can be customized for different fluorophores by changing out dichroic and barrier filters. A general setup for cyan/yellow/red imaging is displayed in which a short pass dichroic mirror directs red light to the first PMT, and a long pass dichroic mirror paired with appropriate band pass filters is used to separate cyan and yellow emissions. Abbreviation: PMT = photomultiplier tube. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61970/61970fig2v2.jpg" /> 
Figure 2: Positioning mice for in vivo imaging. To position the mouse with the pupil on axis with the light path, the anesthetized mouse is first restrained in a head holder, the head is rotated and angled, a large drop of lubricant eye gel is placed on the eye, and the mouse is placed on the stage. A coverslip is mounted in the coverslip holder perpendicular to the light path and lowered down towards the eye. The coverslip should not contact the cornea or mouse head (left), which will be evident if the coverslip is deflected. However, the coverslip should also be close enough to avoid waisting of the droplet (right), because this will have a demagnifying effect on the sample. After applying the gel immersion and securing the coverslip, the stage should be moved in place directly under the microscope objective. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61970/61970fig3v3.jpg" /> 
Figure 3: Imaging retinal ganglion cells. For image display, maximum intensity projections with the z-planes containing cells of interest are created, and resultant images are median-filtered to remove PMT shot noise. Two examples of retinal ganglion cells labelled by injecting AAV-EF1&#945;-FLEX-Twitch2b into VGlut2-Cre transgenic mice are shown, specifically the CFP signal. Images were acquired at sessions four days apart, and vascular landmarks were used to return to the same region near the optic nerve head. The optic nerve head is oriented towards the bottom of the image. Although both samples show some variance in orientation (regions with decreased intensity are indicated with arrows), most cells are present at both time points. Scale bar = approximately 50 &#181;m. Abbreviations: PMT = photomultiplier tube; AAV = adeno-associated virus; EF1&#945; = elongation factor-1alpha; FLEX = flip-excision; VGlut2 = vesicular glutamate transporter 2; CFP = cyan fluorescent protein. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig3v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61970/61970fig5v2.jpg" /> 
Figure 4: Imaging amacrine cells. Amacrine cells were labeled by injecting AAV-EF1&#945;-FLEX-Twitch2b into VGat-Cre transgenic mice. The CFP signal of Twitch 2b is specifically shown. Small maximum-intensity projections focused on the depths of the inner nuclear layer indicate amacrine cell somas, while focusing on the inner plexiform resolves amacrine cell neurites (arrow). The optic nerve head is oriented towards the right of the image. Scale bar = approximately 50 &#181;m. Abbreviations: AAV = adeno-associated virus; EF1&#945; = elongation factor-1alpha; FLEX = flip-excision; VGat = vesicular gamma aminobutyric acid transporter; CFP = cyan fluorescent protein; INL = inner nuclear layer; IPL = inner plexiform layer. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61970/61970fig4v2.jpg" /> 
Figure 5: Imaging microglia. The transgenic mouse line Cx3cr1-GFP was used to label microglia. A maximum-intensity projection of the full scan volume shows many microglia, some with fine process detail that can be resolved. Note that cells towards the lower left of the field have less distortion in the maximum projection than those towards the upper right due to parallax in this region. Maximum-intensity projections containing only the cell of interest significantly reduces this parallax (center, boxed in corresponding colors). Furthermore, this imaging strategy can document the dynamics of fine microglia process remodeling (lower panels). Comparatively, many microglia can be seen with short processes or amoeboid morphology one day after an excitotoxic lesion by intravitreal injection of 50 mM NMDA (right). Scale bar = approximately 50 &#181;m. Abbreviations: GFP = green fluorescent protein; Cx3cr1 = Cx3 chemokine receptor 1; NMDA = N-methyl-D-aspartate. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61970/61970fig6v2.jpg" /> 
Figure 6: Labelling vascular landmarks. Mice were injected with 200 &#181;L of 20 mg/mL Evans blue intraperitoneally 30-60 min prior to the first imaging session. Full thickness maximum-intensity projections demonstrate lasting fluorescence in the retinal vasculature that persisted for at least seven days. Scale bar = approximately 50 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig6v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/61970/61970fig7v2.jpg" /> 
Figure 7: Image dimensions. Retinal ganglion cells labelled by injecting AAV-EF1&#945;-FLEX-Twitch2b into VGlut2-Cre transgenic mice were imaged in vivo, and the same region was then imaged by confocal laser scanning microscopy after fixation and wholemount preparation of the retina. Yellow fluorescent protein channel is shown for both. Colored arrow pairs indicate the same cell in both preparations (upper panels). Single-plane image of 2 &#181;m diameter fluorescent microspheres injected intravitreally and imaged in vivo&#160;(lower left panel). Microspheres did not settle and thus were in constant motion making measurement of axial resolution impossible. Pixel sizes calculated from full-width half-maximum measurements of in vivo fluorescent microspheres or correlative confocal measurements taken from 2-4 retinas per group (lower right). Scale bar = 50 &#181;m. Abbreviations: AAV = adeno-associated virus; EF1&#945; = elongation factor-1alpha; FLEX = flip-excision; VGlut2 = vesicular glutamate transporter 2. <a href="https://www.jove.com/files/ftp_upload/61970/61970fig7v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/61970/61970fig8v2.jpg" /> 
Figure 8: Calcium activity induced by two-photon scanning. Retinal ganglion cells labelled by injecting AAV-EF1&#945;-FLEX-Twitch2b into VGlut2-Cre transgenic mice, YFP is pseudocolored magenta and CFP green, imaged in a single plane as a time series at 4.22 Hz. All RGCs had a similar starting YFP/CFP ratio. Most responded with an increase in FRET ratio (excluding the orange cell), and one maintained a high YFP/CFP ratio throughout the time series (yellow cell). YFP/CFP ratios were normalized to the first frame average, and colored circles match with colored traces. Asterisks indicate time points with representative images displayed on left. Scale bar = 20 &#181;m. Abbreviations: AAV = adeno-associated virus; EF1&#945; = elongation factor-1alpha; FLEX = flip-excision; VGlut2 = vesicular glutamate transporter 2; YFP = yellow fluorescent protein; CFP = cyan fluorescent protein; RGCs = retinal ganglion cells; FRET = fluorescence resonance energy transfer.&#160;<a href="https://www.jove.com/files/ftp_upload/61970/61970fig8v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina at JoVE.com

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

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

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

transpupillary-two-photon-in-vivo-imaging-of-the-mouse-retina

Research

JoVE Journal

Neuroscience

4.0K Views.  Washington University School of Medicine. In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

Video: Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has significantly broadened our knowledge of physiological and pathological processes in numerous tissues in vivo 4-7. In studies of the central nervous system (CNS), there has been a broad application of in vivo imaging in the brain, which has produced a plethora of novel and often unexpected findings about the behavior of cells such as neurons, astrocytes, microglia, under physiological or pathological conditions 8-17. However, mostly technical complications have limited the implementation of in vivo imaging in studies of the living mouse spinal cord. In particular, the anatomical proximity of the spinal cord to the lungs and heart generates significant movement artifact that makes imaging the living spinal cord a challenging task.
We developed a novel method that overcomes the inherent limitations of spinal cord imaging by stabilizing the spinal column, reducing respiratory-induced movements and thereby facilitating the use of two-photon microscopy to image the mouse spinal cord in vivo. This is achieved by combining a customized spinal stabilization device with a method of deep anesthesia, resulting in a significant reduction of respiratory-induced movements. This video protocol shows how to expose a small area of the living spinal cord that can be maintained under stable physiological conditions over extended periods of time by keeping tissue injury and bleeding to a minimum. Representative raw images acquired in vivo detail in high resolution the close relationship between microglia and the vasculature. A timelapse sequence shows the dynamic behavior of microglial processes in the living mouse spinal cord. Moreover, a continuous scan of the same z-frame demonstrates the outstanding stability that this method can achieve to generate stacks of images and/or timelapse movies that do not require image alignment post-acquisition. Finally, we show how this method can be used to revisit and reimage the same area of the spinal cord at later timepoints, allowing for longitudinal studies of ongoing physiological or pathological processes in vivo.

In vivo Imaging of the Mouse Spinal Cord Using Two-photon Microscopy

A minimally invasive protocol to stabilize the mouse spinal column and perform repetitive in vivo spinal cord imaging using two-photon microscopy is described. This method combines a spinal stabilization device and an anesthetic regimen to minimize respiratory-induced movements and produce raw imaging data that require no alignment or other post-processing.

In vivo imaging using two-photon microscopy 1 in mice that have been genetically engineered to express fluorescent proteins in specific cell types 2-3 has ...

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

In Vivo Two-Photon Microscopy of Single Nerve Endings in Skin

Nerve endings in skin are involved in physiological processes such as sensing1 as well as in pathological processes such as neuropathic pain2. Their close-to-surface positioning facilitates microscopic imaging of skin nerve endings in living intact animal. Using multiphoton microscopy, it is possible to obtain fine images overcoming the problem of strong light scattering of the skin tissue. Reporter transgenic mice that express EYFP under the control of Thy-1 promoter in neurons (including periphery sensory neurons) are well suited for the longitudinal studies of individual nerve endings over extended periods of time up to several months or even life-long. Furthermore, using the same femtosecond laser as for the imaging, it is possible to produce highly selective lesions of nerve fibers for the studies of the nerve fiber restructuring. Here, we present a simple and reliable protocol for longitudinal multiphoton in vivo imaging and laser-based microsurgery on mouse skin nerve endings.

In Vivo Two-Photon Microscopy of Single Nerve Endings in Skin

The protocol for dynamic longitudinal imaging and selective laser lesion of nerve endings in reporter transgenic mice is presented.&#160;

Nerve endings in skin are involved in physiological processes such as sensing1 as well as in pathological processes such as neuropathic pain2. Their ...

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as addition/removal of synapses, occur in an experience-dependent manner, providing the structural foundation of neuronal plasticity. As postsynaptic components of the most excitatory synapses in the cortex, dendritic spines are considered to be a good proxy of synapses. Taking advantages of mouse genetics and fluorescent labeling techniques, individual neurons and their synaptic structures can be labeled in the intact brain. Here we introduce a transcranial imaging protocol using two-photon laser scanning microscopy to follow fluorescently labeled postsynaptic dendritic spines over time in vivo. This protocol utilizes a thinned-skull preparation, which keeps the skull intact and avoids inflammatory effects caused by exposure of the meninges and the cortex. Therefore, images can be acquired immediately after surgery is performed. The experimental procedure can be performed repetitively over various time intervals ranging from hours to years. The application of this preparation can also be expanded to investigate different cortical regions and layers, as well as other cell types, under physiological and pathological conditions.

Two-Photon in vivo Imaging of Dendritic Spines in the Mouse Cortex Using a Thinned-skull Preparation

Time-lapse imaging in the living animal provides valuable information on structural reorganization in the intact brain. Here, we introduce a thinned-skull preparation that allows transcranial imaging of fluorescently labeled synaptic structures in the living mouse cortex by two-photon microscopy.

In the mammalian cortex, neurons form extremely complicated networks and exchange information at synapses. Changes in synaptic strength, as well as ...

Two-photon Imaging of Cellular Dynamics in the Mouse Spinal Cord

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for live-cell imaging in the murine spinal cord. This technique is uniquely suited to analyze neural precursor cell (NPC) dynamics following transplantation into spinal cords undergoing neuroinflammatory demyelinating disorders. NPCs migrate to sites of axonal damage, proliferate, differentiate into oligodendrocytes, and participate in direct remyelination. NPCs are thereby a promising therapeutic treatment to ameliorate chronic demyelinating diseases. Because transplanted NPCs migrate to the damaged areas on the ventral side of the spinal cord, traditional intravital 2P imaging is impossible, and only information on static interactions was previously available using histochemical staining approaches. Although this method was generated to image transplanted NPCs in the ventral spinal cord, it can be applied to numerous studies of transplanted and endogenous cells throughout the entire spinal cord. In this article, we demonstrate the preparation and imaging of a spinal cord with enhanced yellow fluorescent protein-expressing axons and enhanced green fluorescent protein-expressing transplanted NPCs.

A new ex vivo preparation for imaging the mouse spinal cord. This protocol allows for two-photon imaging of live cellular interactions throughout the spinal cord.

Two-photon (2P) microscopy is utilized to reveal cellular dynamics and interactions deep within living, intact tissues. Here, we present a method for ...

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon microscopy in Thy1-GFP transgenic mouse line. This approach creates a possibility to investigate the correlation of behavioural manipulations with changes in neuronal morphology in vivo.
The cranial window implantation procedure was considered to be limited only to the easily accessible cortex regions such as the barrel field. Our approach allows visualization of neurons in the highly vascularized RSC. RSC is an important element of the brain circuit responsible for spatial memory, previously deemed to be problematic for in vivo two-photon imaging.
The cranial window implantation over the RSC is combined with an injection of mCherry-expressing recombinant adeno-associated virus (rAAVmCherry) into the dorsal hippocampus. The expressed mCherry spreads out to axonal projections from the hippocampus to RSC, enabling the visualization of changes in both presynaptic axonal boutons and postsynaptic dendritic spines in the cortex. 
This technique allows long-term monitoring of experience-dependent structural plasticity in RSC.

Simultaneous Two-photon In Vivo Imaging of Synaptic Inputs and Postsynaptic Targets in the Mouse Retrosplenial Cortex

This video shows the craniotomy procedure that allows chronic imaging of neurons in mouse retrosplenial cortex using in vivo two photon microscopy in Thy1-GFP transgenic line. This approach is combined with injection of mCherry-expressing adeno-associated virus into dorsal hippocampus. These techniques allow long-term monitoring of experience-dependent structural plasticity in RSC.

This video shows the craniotomy procedure that allows chronic imaging of neurons in the mouse retrosplenial cortex (RSC) using in vivo two-photon ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

In Vivo Vascular Injury Readouts in Mouse Retina to Promote Reproducibility

Advancements in ophthalmic imaging tools offer an unprecedented level of access to researchers working with animal models of neurovascular injury. To properly leverage this greater translatability, there is a need to devise reproducible methods of drawing quantitative data from these images. Optical coherence tomography (OCT) imaging can resolve retinal histology at micrometer resolution and reveal functional differences in vascular blood flow. Here, we delineate noninvasive vascular readouts that we use to characterize pathological damage post vascular insult in an optimized mouse model of retinal vein occlusion (RVO). These readouts include live imaging analysis of retinal morphology, disorganization of retinal inner layers (DRIL) measure of capillary ischemia, and fluorescein angiography measures of retinal edema and vascular density. These techniques correspond directly to those used to examine patients with retinal disease in the clinic. Standardizing these methods enables direct and reproducible comparison of animal models with clinical phenotypes of ophthalmic disease, increasing the translational power of vascular injury models.

In Vivo Vascular Injury Readouts in Mouse Retina to Promote Reproducibility

Here, we present three data analysis protocols for fluorescein angiography (FA) and optical coherence tomography (OCT) images in the study of Retinal Vein Occlusion (RVO).

Advancements in ophthalmic imaging tools offer an unprecedented level of access to researchers working with animal models of neurovascular injury. To ...