Our research delves into Acanthamoeba's natural behavior as it is in nature when humans and Acanthamoeba evitably cross paths. From this foundation, we can extrapolate insights into how our Acanthamoeba interacts in various environments and surfaces, whether contact lens or the cornea, enabling us to develop targeted strategies for preventing Acanthamoeba keratitis and improving contact lens hygiene practices. Historically, the Acanthamoeba field has been slow to adopt cutting-edge advancements in genetics and high-content image analysis.
However, there is now a growing trend towards the routine utilization of these tools to unravel the genetic basis of amoebic infections and to facilitate drug development via high-content throughput analysis. Numerous questions remain regarding the evolution of Acanthamoeba infections in patients. Our ongoing efforts focus on creating visual and quantifiable tools for the in vitro modeling of Acanthamoeba infections, particularly as 3D cornea models become more accessible.
These models enable us to explore how Acanthamoeba behaves during the initial Acanthamoeba keratitis infection and later with potential resurgence. To begin, fill a T75 flask with 30 milliliters of AC6 medium. Aseptically add the contents of a sample vial containing the Acanthamoeba into the flask.
Incubate the culture for three to five days at 26 to 30 degrees Celsius until the flask reaches 50%to 80%confluence. The day before the cells are required, shake and briskly strike the master culture flask twice to dislodge adherent trophozoites. Fill a new T75 flask with 30 milliliters of AC6 medium.
Then transfer two milliliters of master culture into the flask. Incubate the culture at 26 to 30 degrees Celsius for 18 to 24 hours. Prior to harvest, use a microscope set to 4x magnification to visually inspect the trophozoite population for adherence and uniformity.
Briskly strike the flask twice to dislodge adherent trophozoites. Then pour the contents into a 50-milliliter conical tube. Centrifuge the tube at 500 g for five minutes to pellet the amoeba.
After discarding the supernatant, resuspend the pellet in 2 to 10 milliliters of 1/4 Ringers solution. Vortex the resuspended sample to ensure even mixing. Then transfer 10 microliters of the sample into a disposable hemocytometer.
Count the colony-forming units per milliliter. Add Ringers solution to dilute the amoeba pellet to the required concentration. Next, seed the amoeba on the well plate.
To seed the amoeba into a sterile aluminum flow cell, slowly add four milliliters of the suspension through the sterile ports of the flow cell. Then clamp the ports close. To begin, place the strains of Acanthamoeba under a brightfield microscope.
Visualize the amoeba at 4x magnification, where the background appears light gray, and the amoeba is dark. Adjust the microscope's lighting, and focus to ensure the amoebae appear as solid, dark circles. Then configure the imaging software to record at regular intervals with a maximum gap of 30 seconds between images for accurate tracking.
Track amoebas over multiple days, recording in segments of one hour for every 12 hours over five days. If the microscope and software permits, use the XY coordinates to record multiple wells or flow cell locations in a single session. If necessary, stitch together multiple sections from a single well or flow cell for an expanded view at 4x magnification.
Open the microscope file in the imaging software. The Bio-Formats Import Options dialog box is now seen. Within the dialog box, enable Stack viewing.
Set view with to selection Hyperstack. Then check Use virtual stack under Memory management and set the Color mode to Default. Now open the Bio-Format Series Options and select the required series.
Then press OK.Navigate to Image, then select Duplicate to duplicate only the image of a single well by choosing only one C channel and one Z channel. Work exclusively with the duplicated image for further manipulation. Click on Image, followed by Type, and 8-bit to convert the image.
Then choose Process, followed by Subtract Background. Set the Rolling ball to 10. Check the Light background option, and select Sliding paraboloid.
Now navigate to Process, followed by Enhanced Contrast. Adjust Saturated pixel settings to 0.1%for individual trophozoites, or 0.3%for groups of cells and aggregates. Sequentially, click on Image, Adjust, and Threshold.
with Default greater than Black White. This will open the Binary Process Settings. Choose Default, and check Black background of binary masks.
Adjust the threshold aggressively to minimize background noise, while ensuring each amoeba is partially visible. If the software inverted the colors to make the background white and amoeba black, go to Edit, and click on Invert to switch it to a black background with white amoeba. Now sequentially, click on Process, followed by Binary and Close to connect slight gaps in the outer membrane of cells.
Fill any remaining holes by clicking on Process, Binary, and Fill Holes. Then use Shape Drawing tool, and press Edit, followed by Fill to remove any non-amoeba artifacts. If necessary, invert the image again to finalize the binary representation.
To record the size of each amoeba, click on Analyze, followed by Analyze Particles. Then set the size to 10-Infinity, the Circularity between 0 to 1, and the Show to Nothing. Check the Only display results and Summarize option to get the analysis without additional visuals.
Save the result in CSV files, then click on File, followed by Save As, and Tiff. to save the image in a Tiff format. Edit the name as required.
