Figure 1.
Schematic illustration of the choroid sprouting assay.
Eyes are first enucleated from the mice and an initial incision is made along the corneal limbus. A micro-scissor is then used to cut 0.5 mm posterior to the corneal limbus in order to remove the cornea/iris complex and the lens. An incision perpendicular to the corneal limbus towards the optic nerve is made. After peeling off the retina from RPE/choroid/sclera complex, the central and peripheral regions of the complex are separated and further cut into approximately 1 mm×1 mm pieces and embedded in matrigel. The microvascular sprouts from the choroid tissue is visualized under a microscope.
Figure 2.
Mouse and rat central and peripheral choroid sprouting: intra- and inter-animal variability.
(a) Five replicates from 5 animals were compared for the intra-animal (from the same eye of the same animal) variability and inter-animal variability of choroid sprouting. Scale bar: 500 µm. (b) The sprouting from the peripheral mouse choroid is more consistent than central mouse choroid (n = 5) (c) Flow cytometry analysis of choroid sprouting cell populations. About 60% of the cell population from the choroid sprouts stained positive for both CD-31 and isolectin, indicating ECs/macrophages. 36% of the cell population is isolectin-positive but did not stain for CD-31. (d) The extending growth cone resembles vascular tube formation in vivo and stains positively with isolectin GS (arrow head) surrounded by chondroitin sulfate proteoglycan neuron-glial antigen 2 (NG2) positive pericytes (arrow). Scale bar: 10 µm. (e) Real time-PCR analysis of choroid sprouts and aortic ring sprouts indicates that the expression of endothelial marker VE-cadherin is not significantly different between the two assays when normalized to CD31 expression (n = 12) p = 0.97; unpaired T-test. (f) The expression of NG2 by real time PCR is higher in choroid sprouts compared to aortic ring sprouts (n = 8) *** p<0.0001; unpaired t-test. (g) The rate of choroid sprouting is not correlated to the size of the choroid tissue embedded within the rage of 0.2 mm2 to 0.8 mm2.
Table 1.
Choroid Sprouting Standardization and Optimization
Figure 3.
Computerized and manual choroid assay quantification methods yield reproducible and comparable normalized results.
(a) The percentage change between control and treatment groups was highly reproducible between independent experiments run at different times; (n = 6–12) ** p = 0.0042; 2-way ANOVA with Bonferroni correction; (b) Representative photos demonstrate manual and computerized quantification of choroid endothelial sprouts. Scale bar: 500 µm. (c) A step-by-step demonstration of SWIFT-Choroid method quantifying the area of the sprouts. (c1) The area of the choroid tissue was first selected by the magic wand tool from the ImageJ software. (c2) Step 1 of the macro was then used to delete the choroid tissue from the image and calculate the choroid area in pixels. (c3) The excess area of the image was then removed using the polygon selection tool and (c4) the threshold function and the second step of the macro was followed to calculate the sprouting area in pixels. (d) The SWIFT-Choroid method calculates sprouting area without extracellular space. The absolute value of the calculated area is 2.27±0.26 mm2 for manual (n = 11) versus 0.87±0.09 mm2 (n = 11) *** p<0.0001; unpaired t-test. (e) However, the differences between control (n = 11) and treated groups normalized to respective controls (n = 12) were similar for each method: 46.6±8.1% using manual quantification and 46.9±8.1% using SWIFT-Choroid. *** p<0.0001 between treatment group and control group, p = 0.986 between quantification methods; 2-way ANOVA.
Figure 4.
RPE cells promote choroidal endothelial sprouting in endothelial-selective media.
Three different cell culture media (CSC, EGM-2 and DMEM) were compared for standardizing choroidal tissue response. (A) Complete CSC medium with growth factor Boost promotes rapid sprouting. An intact RPE layer on choroid further accelerates sprouting (n = 5–12) p<0.0001 compared to choroid without RPE. (B) RPE cells also potentiate choroid sprouting in EGM-2 medium (n = 6 for each time point) p<0.0001, (C) However, in DMEM there is no difference in sprouting rate (n = 6–18, p = 0.1) regardless of the presence of RPE. The sprouting in DMEM medium contained RPE-like contamination (arrows) and the sprouts did not form growth cones as shown in CSC or EGM-2 medium. All comparisons are 2-way ANOVA with Bonferroni correction. Scale bar: 500 µm.
Figure 5.
The age and strain of the animal affects the rate of choroidal sprouting in explant culture.
(A&B) Choroid with or without RPE from P8 animals sprouts significantly faster than that from P240 aging animals (n = 6–18, p<0.0001 with RPE; n = 6–12, p = 0.0002 without RPE). (C) The choroid explants from 129S6/SvEvTac mice (n = 10) grow significantly faster than choroid explants from C57BL/6J mice (n = 12) at day 7 p = 0.0017. All comparisons are 2-way ANOVA with Bonferroni corrections.
Figure 6.
Choroid sprouting assay responds dose-dependently to pro- and anti-angiogenic factors.
(A) Choroid sprouting increases with increasing doses of VEGF (n = 8 for each time point). After 4 days of incubation, 250 ng/mL of VEGF increased the sprouting compared to vehicle treated control *** p<0.001. (B) After treating with 4-HDHA, the sprouting rate decreased as the concentration of 4-HDHA increases. At 15 µM 4-HDHA, the existing sprouts regressed over time (n = 4–8 for each treatment group) p<0.0001 between different 4-HDHA doses. All comparisons are 2-way ANOVA with Bonferroni correction.
Table 2.
Suggested Conditions for Choroid Microvascular Sprouting Assay