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ibidi FAQs

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Is the µ-Slide Angiogenesis available in a 96-well format?

Yes, it is. We have developed a 96-well format with the same well-in-a-well design for high throughput tube formation assays. The µ-Plate Angiogenesis 96 Well is a fully SBS (Society for Biomolecular Screening) and robotics compatible screening plate.


After completing the experiment, can the µ-Slide Angiogenesis be reused?

No, the µ-Slides Angiogenesis are disposables, and are for single-use only.


Do I have to equilibrate the µ-Slide Angiogenesis to 37°C, before starting the experiment?

No, there is no need to equilibrate the µ-Slide Angiogenesis. Slides are stored at room temperature and can be immediately filled with the gel matrix. The gas that escapes from the gel, while it is being warmed up, can be freely exchanged with the atmosphere, because of the open-well format.


Is there any recommended staining protocol for tube formation experiments?

Of course, it is possible for you to take a picture of the network in the µ-Slide Angiogenesis without staining. The phase contrast image is suitable for automated analysis at the Wimasis WimTube platform. However, if you wish to investigate a certain marker, you can then apply your normal protocol, but take care not to damage the gel matrix and network on top.

An example staining protocol can be found in Application Note 19: Tube Formation.


What would be a good positive or negative control for tube formation experiments?

A good positive control for tube formation experiments using the µ-Slide Angiogenesis would be to test your respective cell line, without using inhibitors in the medium. Endothelial cells should show a tube formation on Matrigel™ (growth factor-reduced) with a starving medium (a medium without growth factors or serum) in the upper well.

A good negative control would be to use a system of cells, consisting of gel matrix and an inhibiting substance. Be sure to use a system that will strongly inhibit tube formation, but will not affect the cell viability.


Do you have any experience with gels, other than Matrigel™, in tube formation assays?

In principle, every gel can be used in tube formation assays using the µ-Slide Angiogenesis. It is important that the cells are able to attach to the surface. Collagens, fibronectin, and laminin provide important binding motifs for cell adhesion.


We want to use growth factor-reduced Matrigel™ and serum-free media + 50 ng/ml VEGF. Is this enough to stimulate the tube formation?

Yes, it is enough to stimulate the tube formation in the µ-Slide Angiogenesis. You won’t even need to use a growth factor in the medium.


Do you use growth factor-reduced Matrigel™ for tube formation, and do you add something else into the matrix and the medium?

For tube formation assays using the µ-Slide Angiogenesis we use both growth factor-reduced Matrigel™ and non-reduced Matrigel™. So far, we have not added growth factors, or other enhancers, to the medium.


What is your experience with fetal bovine serum (FBS) concentration in tube formation experiments? Does this greatly affect the rate of tube formation and degradation?

It depends on the cell / cell line that is used in the µ-Slide Angiogenesis: The primary cells (HUVEC) showed tube formation on Matrigel, with any fetal calf serum (FCS) concentration of up to 10% in the medium. We also tested several endothelial cell lines. The result was that one line showed a much degraded tube formation, when there was an FCS concentration of 10% in the medium. The other cell lines behaved like the primary cells.


Is it really necessary to put the µ-Slide Angiogenesis into the humid chamber of the incubator for the polymerization of the gel?

The humidifying chamber is not necessary for the polymerization itself, but it minimizes the effect of evaporation. In an incubator that is shared between several people, the humidity might not be ensured to the full extent. The very small amount of gel in the µ-Slide Angiogenesis can quickly dry out, and this would lead to the formation of a meniscus.


Is the gel in tube formation assays phenol red or phenol red-free? Which one is better?

For phase contrast microscopy in tube formation assays using the µ-Slide Angiogenesis the phenol red does not disturb the picture, and due to its color, the handling is easier. When using fluorescence microscopy, the phenol red might interfere with the probe’s wavelength. In this instance, it would be better to use phenol red-free medium.


Have you tried mixing the cells in the gel, using the µ-Slide Angiogenesis?

Yes, we also cultivated cells in the gel matrix of the µ-Slide Angiogenesis. After applying Matrigel™, the cells are fine for weeks in this environment, but we did not observe tube formation. The µ-Slide Angiogenesis is a good solution for cultivating cells in a precisely defined 3D-matrix. Due to the big interface of the gel to the medium on top, the conditions in the gel can be adjusted by diffusion.


Do I need access to a video facility, in order to take the pictures of the tube formation assay every hour?

Yes, you should be able to access a microscope that allows you to consistently and precisely find the same position on the well of the µ-Slide Angiogenesis. This can be done with a microscope stage that shows the x/y-coordinates.

With an automated stage, which can save all positions of the wells (specifically, the center of each well) to a position list, you can then go to the respective positions at every time point. But, in order to conduct an imaging procedure each hour, a microscope camera that is capable of timed imaging is very handy. An incubation unit with heating capabilities is also important in most of the tube formation experiments. We can help you find a video facility in your area, which might be able to serve your needs.


Which time points are crucial in tube formation assays?

In tube formation assays, it is very important to have a good approximation on the untreated curve. For that, we recommend creating an image every hour and doing five identical experiments, in parallel. With this method, the variability of the experiment can be defined with a good approximation, and the mean and error at each time point will be correctly evaluated.

In regards to the treated experiments (both inhibitor and enhancer), we would take three experiments, in parallel for each setup, and have one untreated experiment on each µ-Slide Angiogenesis as a reference for possible calibration, or a comparability check.

The best time points for that method are as follows:

  1. at the beginning (0 hours),
  2. half way to the peak (with our example curve at 2 hours),
  3. shortly before, at, and after the peak (with our example curve at 3, 4, and 5 hours),
  4. then every 3 to 5 hours.

The experiment should run long enough for you to collect at least two points in the consolidation phase (the plateau phase at the end of the readout curve). See the chart below.


What is the right amount of cells for my tube formation experiment?

The cell number in tube formation assays using the µ-Slide Angiogenesis is very crucial to the resulting data. It is best to test several seeding densities under non-inhibited conditions, and then determine the number where most tubes are built. This will be the number where the most distinct effects of substances can be measured.


Which parameter(s) in tube formation should I evaluate?

The evaluation of image data in tube formation assays using the µ-Slide Angiogenesis gives out four parameters: cell-covered area, tube length, nodes, and loops. All of them show the same characteristic curve, over time. We decided to investigate the tube length, for each defined area in our experiments. The tube length is the most reliable parameter. It can be most easily controlled when comparing raw images to evaluated ones. It is also more stable than the values of nodes and loops. These two values show big fluctuations during the time curve. Measurements are comparable throughout all different experimental setups, when referring the tube length to a standardized area (1 mm²).