One of the most widely used in vitro assays to model the reorganization stage of angiogenesis is the tube formation assay. The assay measures the ability of endothelial cells, plated at subconfluent densities with the appropriate extracellular matrix support, to form capillary-like structures (a.k.a tubes). Scientists typically employ this assay to determine the ability of various compounds to promote or inhibit tube formation. Compounds that are able to inhibit tube formation could be useful in various diseases, such as cancer, where tumors stimulate new blood vessel formation to receive oxygen and nutrients in order to grow beyond a relatively small size.
Upon plating, endothelial cells attach and generate mechanical forces on the surrounding extracellular support matrix to create tracks or guidance pathways that facilitate cellular migration. The resulting cords of cells will eventually form hollow lumens.
Ongoing research is focused on how specific molecules in the “matrix-integrin-cytoskeletal signaling axis” are involved in the eventual assembly into three-dimensional vascular networks.¹ Once formed, these interconnected networks are usually maintained for approximately 24 hours. Tube formation is typically quantified by measuring the number, length, or area of these capillary-like structures in two-dimensional microscope images of the culture dish.
The advantages of this assay are that it is relatively easy to set up, requires a short culture period, is quantifiable, and is amenable to high-throughput analysis. A disadvantage of this assay is the large variation of tube-forming capability among different lots of endothelial cells and support matrices, which makes the selection of these resources crucial to obtaining consistent and reliable data. While this assay is essential, users are cautioned that results should be confirmed in vivo because commercially available endothelial cells have been pre-selected for their proliferative capacity and there are no heterospecific cell interactions being represented. It has also been reported in the literature that certain non-endothelial cell types demonstrate tube formation, which suggests that tube formation by endothelial cells may not represent true differentiation of this cell type.²
All procedures should be performed in a biological safety cabinet using aseptic technique to prevent contamination.
1. Prepare a bottle of supplemented Medium 200PRF by thawing a bottle of Low Serum Growth Supplement (LSGS) and transferring the entire contents of the LSGS bottle to the bottle of Medium 200PRF.
- Note: Once Medium 200PRF has been supplemented with LSGS, the supplemented medium should be stored in the dark at 4°C and should not be frozen. When stored in the dark at 4°C, the supplemented medium is stable for 1 month.
2. Seed cryopreserved endothelial cells (HUVEC) at 2 × 105 viable cells per a 75-cm2 tissue-culture flask using LSGS-supplemented Medium 200PRF (15 mL total volume).
3. Change culture medium 24–36 hours after seeding.
4. Change the medium every other day thereafter, until the culture is approximately 80% confluent (5–6 days).
5. Thaw Gibco Geltrex in a refrigerator (4°C) overnight.
- Note: Since refrigerator temperatures may vary, thaw Geltrex in an ice bath in a refrigerator.
6. Optional step for fluorescent monitoring of tube formation using a cell-permeable dye (e.g., Invitrogen Calcein, AM): Add the dye to the endothelial cells in a 75-cm2 flask and incubate for 30 min at 37°C and 5% CO2 (protect from light). Final dye concentration should be 2 µg/mL.
7. Add 50–100 µl of Geltrex per cm2 to the growth surface and incubate coated surface for 30 minutes at 37°C to allow the gel to solidify.
- Note: 50 µL of Geltrex per cm2 is sufficient for larger well sizes (e.g., 12-well, 24-well). 100 µL per cm2 is necessary for smaller well sizes (e.g., 96-well).
8. Harvest the cells using the following procedure. Do not warm any of the following reagents prior to use. This procedure is designed for one 75‑cm2 flask. If different-sized culture vessels are used, adjust reagent volumes accordingly:
- Remove all culture medium from the flask.
- Add 12 mL of Trypsin/EDTA solution to the 75‑cm2 flask. Rock the flask to ensure the entire surface is covered.
- Immediately remove 9 mL of Trypsin/EDTA solution from the flask.
- Incubate the flask at room temperature for 1–3 minutes.
- View the culture under a microscope.
- When the cells have become completely round, rap the flask gently to dislodge the cells from the surface of the flask.
- Add 9 mL of Trypsin Neutralizer solution to the flask and transfer the detached cells to a sterile 50-mL conical tube.
- Add 9 mL of additional Trypsin Neutralizer solution to the flask and pipette the solution over the flask surface several times to remove any remaining cells.
- Add this solution to the 50-mL conical tube.
- Centrifuge the cells at 180 × g for 7 minutes, until the cells are pelleted.
- Remove the supernatant from the tube, being careful not to dislodge the cell pellet.
9. Add 4 mL of non-supplemented Medium 200PRF to the cell pellet and mix by pipetting up and down several times to ensure a homogeneous single-cell suspension.
10. Determine the concentration of cells in the non-supplemented Medium 200PRF.
11. Dilute cells in non-supplemented medium in the presence or absence of angiogenesis inducers and inhibitors. We recommend a concentration of 3.5–4.5 × 104 cells per 200 µL as a starting point and general guideline; the ideal plating density and media volume depends on cell type and should be determined experimentally. The final media volume should be ~200 µL/cm2.
- Note: For a positive inducer control, we suggest diluting cells in LSGS-supplemented Medium 200PRF. LSGS-supplemented Medium 200PRF contains 2% (v/v) FBS and bFGF (3 ng/mL). For a positive inhibitor control, we suggest diluting the cells in LSGS-supplemented media and 30 µM Suramin.
12. Gently add cells at the selected density to the gel-coated well, at a final media volume of ~200 µL/cm2.
13. Incubate the plate at 37°C, 5% CO2 overnight.
- Note: Incubation times may vary. HUVEC, for example, develop well-formed tube networks after 4–6 hours. After 24 hours, endothelial cells typically undergo apoptosis.
14. Optional Step: If cells were not pretreated with a dye before harvesting, they can be stained at the end of the incubation period after the tube network has formed using a cell-permeable dye (e.g., Calcein, AM)
- Add the dye to the cells and incubate for 30 minutes at 37°C and 5% CO2 (protect from light). Final dye concentration should be 2 µg/mL.
- Gently remove the dye-containing media with a pipette, and replace with an equivalent volume of warm Medium 200PRF. The replacement media should be identical to the media the cells were incubated in during the tube formation.
15. If a fluorescent dye was used, cells may be visualized using a fluorescence microscope. If a fluorescent dye was not used, cells may be visualized directly using a light microscope.
|Problem||Possible Causes||Recommended Solutions|
|There is no tube network formation present in the positive inducer control well.||A. The cells may not be healthy or the cells may be too old. |
B. The cell density is too high or too low.
|A. Use only healthy cells or cells from an earlier passage. |
B. The optimal seeding density is cell-type specific. Therefore, it is necessary to optimize the seeding concentration. Use a larger well size in order to increase the observable area at the center of the well. A. Make sure that the replacement media being used is identical to the media the cells were originally cultured in during the experiment. B. Always add replacement media very gently to the cells.
|The tubes at the edge of the well are out of focus.||There is a meniscus present at the edge of the well.||Use a larger well size in order to increase the observable area at the center of the well.|
|The tube network disappeared after staining.||A. The replacement media was different from the original media. |
B. The replacement media was added too fast.
|A. Make sure that the replacement media being used is identical to the media the cells were originally cultured in during the experiment. |
B. Always add replacement media very gently to the cells.
|There is high background fluorescence.||Following hydrolysis, Calcein, AM slowly leaks out of the cells.||Gently remove media and add replacement media.|
Figure 1. Induction of endothelial cell reorganization into 3D vessel structures. Human umbilical vein endothelial cells (HUVEC) (42,000 viable cells/cm2) were seeded on a 24-well polystyrene plate coated with Geltrex matrix (50 μL/cm2) using LSGS-supplemented Medium 200PRF, and incubated at 37°C and 5% CO2. At 16 hr post-seeding, 2 μg/mL of Invitrogen (A) Calcein, AM (Cat. No. C3099), (B) Calcein Blue, AM (Cat. No. C1429), or (C) CellTrace Calcein Red-Orange, AM (Cat. No. C34851) was added directly to the culture well and incubated for 20 min (37°C, 5% CO2) prior to imaging at 4x magnification. (D) A representative phase-contrast image
Figure 2. Low background of the endothelial tube formation assay. Human umbilical vein endothelial cells (HUVEC) (42,000 viable cells/cm2) were seeded on a 24-well polystyrene plate coated with Geltrex matrix (50 μL/cm2) using non-supplemented Medium 200PRF, and incubated at 37°C and 5% CO2. At 16 hr post-seeding, 2 μg/mL of (A) Calcein, AM (Cat. No. C3099), (B) Calcein Blue, AM (Cat. No. C1429), or (C) CellTrace Calcein Red-Orange, AM (Cat. No. C34851) was added directly to the culture well and incubated for 20 min (37°C, 5% CO2) prior to imaging at 4x magnification. (D) A representative phase-contrast image.
- Davis G, Senger D. (2005). Endothelial extracellular matrix: biosynthesis, remodeling, and functions during vascular morphogenesis and neovessel stabilization. Circ Res. 97(11):1093-1107.
- Donovan D et al. (2001) Comparison of three in vitro human 'angiogenesis' assays with capillaries formed in vivo. Angiogenesis. 4(2):113-121.
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LT167 updated 6-Oct-2011
For Research Use Only. Not for use in diagnostic procedures.