Angiogenesis is a hallmark process involved in embryology and a disease development, including tumor growth and vasoproliferative eye disease. In this project we aim to present a comprehensive framework of in vitro techniques that can be used to study angiogenesis on a molecular level in a very controlled setting, and to screen for potential fewer product options. Translating in vitro to in vivo and ultimately to the clinical setting is challenging due to the risk of false positive or false negative results.
Selecting the right angiogenesis assay is therefore crucial, and knowing its limitations. This study aims to offer guidelines and how to interpret and how to establish two common angiogenesis assays. Recently we compared a 2D scratch wound migration assay with a 3D spheroid sprouting assay.
Where the 2D assay is easily scalable, the 3D assay captures angiogenesis in greater detail, including tip stalk cell formation, matrix interaction, and the glycolytic switch. This data offer scientists guidelines on which assay to choose next in their projects. Our laboratory will continue to characterize key aspects of the presented in vitro assays by directly comparing them to in vivo settings in order to facilitate translations of in vitro results to clinical applications.
To begin, seed 100 microliters of HUVEC cells into the wells of a 96-well plate. Place the plate in a cell culture incubator at 37 degrees Celsius under 5%carbon dioxide for six hours to facilitate cell settling. Next, replace the medium in each well with 100 microliters of endothelial cell growth basal medium containing 2%FBS, and incubate again.
The following day, position the plate in a 96-Well Woundmaker Tool. Press down the lever of the device to generate a scratch. Carefully lift off the lid of the Woundmaker Tool before releasing the lever to prevent double scratching.
Wash the wells two times with 200 microliters of FBS supplemented endothelial cell growth basal medium. After confirming total debris removal, pipette 100 microliters of the prepared stimulation or control solutions into each well. Use a live cell imaging microscope to acquire images every hour for 24 hours.
The positive control showed successful migration in the original scratched area. Technical issues were seen as patchy cell growth, double scratching, or waves in the scratch borders. A 25%increase in relative wound distance was considered optimal.
Incorrect separation of the positive and negative controls indicated low dynamic range. To begin, add 30 milliliters of 5%potassium hydroxide in methanol to a 50 milliliter tube. Incubate 100 cover slips with a diameter of 12 millimeters in the tube for 30 minutes under constant agitation.
Next, wash the cover slips in demineralized water for 30 minutes. Once washing is complete, transfer the cover slips into 70%isopropyl solution for storage. To coat the cover slips, first lean them individually against the wall of a square Petri dish.
When the isopropyl alcohol has evaporated, transfer them into a 24-well plate. Pipette one milliliter of collagen in PBS into each well and incubate. Next, wash the cover slips about four to five times for five minutes each with one milliliter of PBS.
Pipette cultured HUVECs into the wells with the coated cover slips. After culturing the cells for a desired length of time, wash the cells in one milliliter of PBS for five minutes. Fix the cells in one milliliter of 2%paraformaldehyde for 20 minutes at room temperature.
Now, rinse the cells in one milliliter of PBS for three to four wash cycles of five minutes each. Finally, pipette one milliliter of blocking buffer into the well plate before staining. To begin, add HUVECs into a 10 milliliter Falcon tube containing eight milliliters of endothelial growth medium.
Add two milliliters of methylcell stock solution into the tube and shake well. Next, transfer 25 microliters of the mixture onto the inverted inner surface of a large, square Petri dish. Gently rotate the lid by 180 degrees and place it on the bottom of a cell culture vessel and incubate overnight.
The next day, add 2.3 milliliters of Rat Tail Collagen Type 1 to a vial containing 0.28 milliliters of 10 x medium 199. Keep this mixture on ice and mix until it is evenly yellow. Titrate the mixture against sodium hydroxide until it turns orange.
Then add 50 microliters of HEPES buffer to the final product. Collect the hanging cultured cells into a 50 milliliter falcon tube with 20 milliliters of PBS, then centrifuge. After discarding the supernatant, add 0.1 milliliters of FBS and 0.4 milliliters of endothelial basal medium to the spheroid pellet.
Gently tap the tube to resuspend the pellet. Now pipette two milliliters of methylcell stock solution and mix well. Then add two milliliters of the prepared collagen mixture to the resuspended spheroids.
Dispense 0.5 milliliters of the resulting mixture into the wells of a 24-well plate and incubate. Next, pipette 100 microliters of diluted recombinant Human Vascular Endothelial Growth Factor into the wells of the plate. Incubate the cell culture at 37 degrees Celsius under 5%carbon dioxide supplementation for 12 hours.
The next day, with an inverted microscope, capture the images of the spheroids that are not in contact with the well rim with each other or show signs of damage. The spheroids stimulated with only basal medium or recombinant Human Vascular Endothelial Growth Factor were clearly distinguishable. Spheroids in low-quality gels appeared to drop to the ground and dispersed.
The negative control showed a moderate baseline sprouting rate while the Human Vascular Endothelial Growth Factor doubled or tripled the relative sprouting length. The number of sprouts showed a similar dynamic range. To begin, fix the sprouted spheroid HUVEC gels in one milliliter of 4%paraformaldehyde for one hour.
Wash the gels gently in one milliliter of PBS about three to four times. Detach the gels fully from the surface of each well of the plate. Then transfer them into new 24-well plates.
Add one milliliter of blocking solution to the well plate, then incubate on an orbital shaker for one hour at room temperature. After incubation, pipette 500 microliters of the primary antibody solution into the wells before incubation. The next day, wash the gels gently with PBS for five wash cycles of five minutes each.
Add 500 microliters of the corresponding secondary antibody diluted with fluid and FTSE in blocking buffer. Incubate the plate overnight at four degrees Celsius on an orbital shaker. After washing the gels in PBS, transfer them onto microscope slides.
Add two drops DAPI containing mounting medium onto a cover slip and place it over each gel. Seal the dried gels with nail polish. Place them in the dark before storing at four degrees celsius until further analysis.
