Longitudinal In Vivo Imaging of Retinal Cells with Tailored Adeno-Associated Virus Serotypes via Confocal Scanning Laser Ophthalmoscopy

All experiments were conducted in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and were approved by the Institutional Animal Care and Use Committee of Capital Medical University, Beijing. Four-week-old adult male C57BL/6J mice (weighing between 15-20 g) were used for all experiments and housed in temperature-controlled rooms with a 12/12-h light/dark cycle. Standard rodent chow and water were available ad libitum. The details of the reagents and equipment used in this study are listed in the Table of Materials.

1. AAV-mediated retinal cell transfection

NOTE: For targeted transduction of RGCs, this protocol uses AAV2-hSyn-eGFP, which incorporates the hSyn promoter to drive robust expression of enhanced GFP. Müller glia is targeted using AAV9-GfaABC1D-eGFP. To achieve optimal transduction efficiency and robust transgene expression, a minimum AAV titer of 1 x 10¹² viral genomes (vg)/mL is recommended for intravitreal injections in mice¹³.

1. Adhere to stringent safety protocols when conducting AAV-mediated retinal cell transfection procedures to prevent accidental exposure and ensure a safe working environment.
2. Perform all procedures involving viral vectors within a certified biosafety cabinet (BSC). Equip personnel with appropriate personal protective equipment (PPE), including lab coats, gloves, and eye protection, throughout the duration of the procedure.
3. Dispose of all sharps and biohazard materials properly in accordance with institutional safety protocols to maintain compliance and safeguard all laboratory personnel.

2. Preparation of viral vectors and animals

1. Obtain AAV2-hSyn-eGFP or the AAV9-GfaABC1D-eGFP vectors stored at -80 °C. Thaw vectors on ice immediately before use to preserve viral integrity.
2. Anesthetize the mice with an intraperitoneal injection of pentobarbital sodium at a dosage of 50 mg/kg. Confirm the depth of anesthesia by testing the rear foot reflexes before further actions.
3. Trim the whiskers to prevent interference with the surgical area during the procedure. Disinfect the orbit with a 20% Povidone-iodine solution for 2 s, then rinse thoroughly with sterile normal saline.
4. Apply a drop of 0.5% proparacaine topically to minimize discomfort and prevent involuntary eye movements.

3. Handling and intravitreal injection of viral vectors

1. Transfer 1 µL of selected AAV solution (either AAV2-hSyn-eGFP or AAV9-GfaABC1D-eGFP) to a piece of clean, prechilled paraffin film to avoid temperature fluctuations that might affect the viral activity.
2. Aspirate the AAV solution using a Hamilton glass syringe with a 33 G beveled needle under the ophthalmic surgical microscope.
3. Place the mouse in ventral recumbency under the microscope. Tilt the head slightly to elevate the ocular side intended for injection. Adjust and optimize the magnification of the microscope to clearly identify the superior region of the mouse orbit.
4. Insert the needle 1-2 mm posterior to the limbus at the superior scleral margin of the eye into the vitreous cavity. Inject the AAV solutions slowly to avoid backflow or pressure-induced damage.
   NOTE: Injection site location: Perform the injection in the superior temporal quadrant of the eye, approximately 1 mm posterior to the limbus. Choose this location to avoid major blood vessels and minimize the risk of lens damage. Avoid injection sites too close to the limbus to prevent iris penetration or lens abrasion. Avoiding vasculature: Use an operating microscope to carefully examine the planned injection site for visible blood vessels prior to injection. If vessels are present, slightly adjust the entry point to avoid them. Insert the needle at a shallow angle, parallel to the iris, to further reduce the risk of vascular damage. Avoid repeated injections in the same area to reduce the risk of subretinal hemorrhage.
5. Keep the needle in place for 30 s to allow the vectors to disperse within the vitreous chamber and settle onto the retinal surface, thereby maximizing the chances of successful transduction.
6. Withdraw the needle slowly to minimize the risk of vitreous backflow.
7. Apply a topical antibiotic ointment to the injection site immediately after injection. Ensure that the ointment is applied uniformly to fully cover the ocular surface, preventing ocular infection and inflammation.
   NOTE: The antibiotic ointment contains 1 mg/g Dexamethasone, 3500 IU/g Neomycin Sulphate, and 6000 IU/g Polymyxin B Sulphate.
8. Place the mouse on a heating pad to maintain body temperature during recovery from anesthesia, and closely monitor them for any signs of distress or abnormalities. Once fully recovered and exhibiting no complications, return them to their cages with access to normal food and water.

4. In vivo imaging with CSLO

1. Animal preparation
   1. Select the mouse infected with AAV2-hSyn-eGFP or AAV9-GfaABC1D-eGFP after 4 weeks of intravitreal injection. Apply one drop of an eye solution containing 0.5% tropicamide and 0.5% phenylephrine to the eye surface to dilate the pupil to approximately 2 mm in diameter²².
      NOTE: A minimum of 4 weeks of AAV transduction time interval is recommended for the optimal retinal cells fluorescent labeling post intravitreal injection¹³. Additionally, in our optic nerve crush (ONC) experiments, the retinas were imaged at 4 weeks post-infection, as well as on day 7 and 14 following the crush injury. These time points were chosen to capture both the initial AAV expression and the progression of retinal changes after ONC.
   2. Cover the imaging platform with a piece of the clean pad to avoid the possible contamination of animal excrement during the operation process.
      NOTE: The custom-built imaging platform provides ample space for the animal to lie prone, ensuring stability and minimizing movement during manual restraint for optimal image acquisition.
   3. Carefully transfer the mouse onto the imaging platform and allow an acclimatization period of 2-5 min prior to imaging to minimize stress and facilitate adaptation to the imaging environment.
   4. With the assistance of a second technician, gently restrain the animal while maintaining a conscious state throughout the procedure. Provide a 10 s rest interval after 15 s of imaging to permit eye blinking and maintain corneal moisture.
      NOTE: Gently scrub the scalp skin to induce spontaneous eyelid lifting, avoiding the use of forced eyelid restraint or additional devices such as clips, thereby reducing the risk of eye injury. General anesthesia is not employed to preserve optical clarity during extended imaging sessions, as it has the potential to exacerbate transient lens opacity. Balance high-quality image acquisition with animal comfort and safety throughout the procedure. Complete the entire procedure within 5 min to ensure animal well-being and maintain imaging quality.
   5. Adjust the head posture and align the camera lens by fine-tuning the focus knob (Figure 1B) and micromanipulator (Figure 1C) to aim the region of interest (ROI) for subsequent in vivo imaging.
2. System configuration
   1. Attach a non-contact 55° wide angle lens on the camera to expand the field of view of the fundus area.
   2. Set the filter wheel to position A (angiography) to enable the acquisition of infrared (IR) and fluorescent angiography (FA) imaging mode (Figure 1A). Turn on the laser and power supply to activate the CSLO imaging system.
   3. Open the software Heidelberg Eye Explorer and create a new examination by clicking on the New Patient icon at the top of the menu bar. Enter the relevant animal information and accept the default corneal curvature of 7.7 mm in the pop-up window.
   4. Select the yellow Start button (Figure 1E) on the control panel screen to initiate the live imaging mode.
3. In vivo CSLO imaging
   1. Select the IR mode (Figure 1G) with autofluorescence at 820 nm excitation wavelength on the control panel. To achieve high-resolution imaging, the High Res. (HR) mode is activated either through the window interface or the manual control panel (Figure 1F).
      NOTE: Infrared reflectance (IR) imaging is typically the first step in CLSO ophthalmic imaging, acquired before fluorescein angiography (FA), to provide a baseline view. Even illumination, minimal artifacts, and sharp focus on major retinal vessels are crucial for the reliability of z-plane focusing and the achievement of high-quality IR images.
   2. Move the lens towards the mouse eye using the joystick with the XYZ-micromanipulator for fine adjustment. Examine the optical transparency and turn the sensitivity knob (Figure 1D) counterclockwise to decrease the brightness of the fundus image to 40%-60%, thereby avoiding overexposure of the fundus and enhancing visualization of retinal details.
   3. Adjust the focus knob and micromanipulator until the optic disc and retinal vessels are unambiguously identified on the fundus.
      NOTE: Criteria of the optimal focal plane: (1) Evenly illuminated fundus view without dark corners. (2) Uniformed illumination surrounding the optic disc without focal dark spots or distortion. (3) Clear lumen shape of the large retinal vessels with the visible movement of blood flow.
   4. Switch to the FA mode (Figure 1H) with a solid-state blue laser at an excitation wavelength of 488 nm and a barrier filter at 500 nm. Turn the sensitivity knob (Figure 1D) clockwise to increase the brightness of the fundus image to 50%-107% to illuminate the targeted retinal area.
   5. Set the ART mean value (a blue bar at the bottom left of the software interface) to at least 15 to enhance the signal-to-noise ratio. This function averages a series of B-scans acquired at the same location, improving image quality.
      NOTE: While a higher ART averaging value generally enhances image quality through averaging, it also extends scan duration. This extended duration could potentially increase the risk of corneal dehydration and animal fatigue, factors that need to be carefully considered, especially in awake animal imaging.
   6. Re-align to the inner plane of the retina, specifically targeting the GFP expressing RGCs or the Müller glial cells. Fine adjustments are made to ensure sharp illumination of the desired cell population, such as RGC soma and axons, or the Müller cell body. Imaging sensitivity is balanced to achieve optimal visualization and prevent oversaturation of the GFP-positive cells.
      NOTE: To obtain the images with comparable clarity and brightness of the GFP signal, the sensitivity acquisition should be constant for all mice in the experiment.
   7. Press down the sensitivity knob (Figure 1D) to initiate the ART mode, which generates a live mean image online. Wait until the blue bar (representing the ART Mean value) at the bottom left of the software interface has reached the set ART value (≥ 20 frames) and tap the Acquire button on the control interface to capture the fundus images.
      NOTE: For ART mode, eye-tracking is incorporated to automatically compensate for minor eye movements during in vivo imaging, enabling consistent focus on the selected retinal focal plane.
   8. Once acquired the desired images, exit the ART mode by pressing the sensitivity knob on the touch panel.
      NOTE: To prevent corneal drying and allow for spontaneous eye blinking, 10 s breaks were implemented every 15 s during the imaging process.
   9. To navigate the nasal and temporal retinal areas, move the camera head horizontally while observing the live image on the screen. Once the target region is reached, fine-tune the focus and start the imaging process as described in steps 2.3.1-2.3.8.
      NOTE: Maintain slow and controlled camera movements while ensuring consistent and bright illumination of the fundus. If dark areas appear, utilize the micromanipulator to adjust the camera position vertically and re-center the retina.
   10. To image the superior retina, gently adjust the mouse head posture to a face-up position. Moderately tilt the camera head upwards to align with the superior retinal region. Re-adjust the focus as needed and proceed with imaging.
   11. Upon completion of imaging, all desired retinal regions, exit the acquisition window, and the images are automatically saved. Apply topical lubricant on the imaged eye to maintain corneal hydration.
   12. To Initiate the examination of the contralateral eye, reposition the animal to the opposite side of the platform and repeat steps 2.3.1-2.3.11.

5. Image processing and analysis

1. Image export and preparation
   1. Open the Heidelberg Eye Explorer software and select the desired CSLO imaging session with good quality. Choose single or consecutive images of the same retinal location with consistent sensitivity settings over time.
   2. Exclude blurry images with poor focus or corneal clarity, ensuring that only images with clear visualization of retinal structures are used for analysis. Export these images in TIFF format for further processing.
2. Image alignment and cropping
   1. Group images per mouse eye based on the same retinal region.
      NOTE: The original images acquired from the CSLO system are saved in the following format: TIFF, with a dimension of 1536 x 1536 pixels, a resolution of 5.69 um/ pixel, and an ART value of ≥15.
   2. For follow-up examinations, import images into suitable image processing software. Rotate and align the images as necessary to match the orientation of the baseline image. When saving or exporting the processed images, use settings that maintain the highest possible image quality to minimize potential compression artifacts.
   3. Align the CSLO images from a time series of the same eye by rotating them to match the respective baseline image using the vasculature and optic disc as landmarks.
      NOTE: Carefully examine each aligned image, and before proceeding to crop, ensure they meet the following criteria: (1) Full retinal region capture: The image must fully encompass the intended retinal region of interest, including key landmarks like the optic disc and major blood vessels; (2) Absence of motion artifacts: The image should be free from blurring, streaking, or duplication of structures caused by eye movement during imaging; (3) Minimal distortion: The image should accurately represent the retinal structure without warping, stretching, or compression artifacts that could arise from imaging angles or lens effects. (4) Balance of image quality metrics: Assessments of background noise levels (Standard deviation of pixel intensities in structure-free areas < 5 (0-255 scale)); signal-to-noise ratio (SNR >20), illumination consistency (Coefficient of variation among six image quadrants <10%) and image uniformity (Coefficient of variation in homogeneous regions <15%).
   4. Select the rotated image and export it in TIFF format. For each time point, use the cropping tool to randomly select and crop 5-10 areas (300 x 300 pixels each) containing GFP-positive cells from the quality-approved images. Export each cropped image individually in TIFF format for further analysis.
3. Fluorescence intensity measurement
   1. Open the cropped CSLO images (8-bit grayscale, 300 x 300 pixels) in ImageJ/Fiji.
   2. To examine overall changes in fluorescent signals, measure the total fluorescence intensity per cropped image using Analyze > Measure (or Ctrl + M). Record the "Mean" value, representing the average GFP fluorescence intensity of the whole cropped image.
4. RGC quantification
   1. In ImageJ/Fiji, adjust the intensity threshold of the cropped CSLO images using Image > Adjust > Brightness/Contrast to achieve optimal visualization of individual white soma bodies against the black background.
      NOTE: Determine an individualized threshold after reviewing the batch of cropped images, or apply a consistent thresholding algorithm (e.g., Huang's method⁵ in ImageJ/Fiji). For Huang's method: (1) Convert images to 8-bit grayscale (Image > Type > 8-bit). (2) Access the thresholding tool (Image > Adjust > Threshold). (3) Select Huang as the thresholding method. (4) Preview the segmentation and click on Apply. The threshold will be automatically calculated and displayed on the image histogram.
   2. Activate the Cell Counter plugin and click on each GFP-positive RGC soma for counting. Record the total number of cells marked in the analyzed region. Repeat this process for all cropped images from each follow-up time point.
5. Data analysis
   1. Export the cell count and fluorescence intensity data to a spreadsheet or statistical software (e.g., GraphPad Prism).
   2. Perform appropriate statistical tests based on the corresponding experimental design and hypotheses. Interpret the results and conclude.