An Assay for Permeability of the Zebrafish Embryonic Neuroepithelium

1. Preparing for Microinjection

1. Prepare microinjection needles by pulling capillary tubes using Sutter instruments needle puller.
2. Load microinjection needle with fluorescent dye (FITC-Dextran).
3. Mount needle on micromanipulator and microinjection apparatus.
4. Carefully break microinjection needle using forceps to roughly 2 μm in width, however, this will vary depending on your microinjector setup. For our microinjection needles, this corresponds to the first region of the needle from the tip that does not bend.
5. Measure drop size in oil, adjusting injection time and pressure, so that each injection delivers 1 nl. Example settings for Harvard Apparatus microinjector are: P balance = 1.4 psi, P out = 1.4 psi, P inject = 22.9 psi, P clear = 67.8 psi with an injection time of .4 to .7 sec. The diameter of our needle using these settings is roughly 2 μm. However, settings will be microinjector specific, and vary according to needle diameter.

2. Preparing the Embryos

1. Coat 2 dishes with 1% agarose in water for each condition, poke holes into agarose with a 1-200 μl pipette tip, and remove agarose plugs. Fill dishes with embryo media.
2. Using forceps, dechorionate embryos that are 18 hpf or older under a stereomicroscope. Embryos are staged according to Kimmel et al.¹¹ .
3. Transfer dechorionated embryos into first agarose coated dish.
4. To anesthetize embryos, add tricaine (0.1 mg/ml) to the dish until embryos stop moving (made according to Westerfield¹²).

3. Injecting the Brain Ventricles

1. Orient embryos so you are looking at their dorsal side by putting the tail of the embryo into the hole. If your micromanipulator is on the right, then move the embryo so that the forebrain is to the left and hindbrain to the right.
2. Position needle at widest point of hindbrain ventricle.
3. Carefully pierce roof plate of hindbrain ventricle being sure not to go through the depth of the brain into the yolk (Figure 1A).
4. Inject 1-2 nl of fluorescent dye into the ventricles making sure the dye fills the whole length of the brain ventricles.
5. Transfer embryos to the second agarose coated dish filled with embryo media and re-anesthetize as described in 2.4.
6. IMMEDIATELY start imaging, as described in section 4 in order to get a time zero image.

4. Imaging

1. Orient embryos with their tail in the hole as described in 3.1.
2. Use a dissecting microscope with both transmitted and fluorescent light to take a brightfield dorsal image. Keep the magnification constant between imaging of different embryos. This allows for direct comparison of analyses performed using Image J (5.2-6).
3. WITHOUT MOVING the embryo, microscope or dish, take a corresponding fluorescent image.
4. Repeat for each embryo at desired time points.

5. Quantification of Dye Movement

1. Merge brightfield and fluorescent images in Photoshop as previously described by Gutzman and Sive¹⁰.
2. Measure the distance the dye front moves in Image J software available at http://rsbweb.nih.gov/ij/.
3. Open merged file in Image J and use line tool to draw a line from the forebrain hinge-point to dye front at a 10-20 ° angle from neuroepithelium (Figure 1A). This region was chosen because it is the first and most noticeable site of dye leaking out of the wild type neuroepithelium.
4. Select measurement tool to calculate length of line.
5. Repeat for each time-point.
6. Calculate net distance the dye front moved over time by subtracting distance at t=0 from other time points.
7. Plot on graph.

6. Representative Results

An example of results obtained in a neuroepithelial permeability assay using wild type embryos is shown in Figure 1B-D. To accurately differentiate permeability, it is useful to test dyes with different molecular weightsto identify a size that is only slightly leaky in wild type or control embryos (Figure 2). This allows for identification of genetic mutants or environmental conditions that either increase or decrease permeability (Figure 1D, green and red lines respectively). For the 24 hpf zebrafish neuroepithelium, 70 kDa FITC Dextran leaks slowly over 2 hr, whereas 2,000 kDa does not and 10 kDa almost immediately leaks out. Therefore 70 kDa is the ideal molecular weight to identify conditions that both increase and decrease neuroepithelial permeability.

If the needle misses the ventricular lumen, fluorescence will appear outside the brain at t=0 (for an example see Gutzman and Sive, 2009¹⁰). These embryos should be discarded since the injected dye was not initially contained within the brain and no clear conclusion regarding movement of the dye and permeability of the neuropeithelium can be made.

Finally, if embryos have small ventricles or un-inflated brain ventricles, pre-injection of ventricles with a saline solution can be done prior to injection of the fluorescent dye. This inflates the ventricles making subsequent visualization of the ventricles easier when injecting with the fluorescent dye. Proper controls must be performed to determine whether injection of saline disrupts normal neural tube development.

Figure 1. Time course of different molecular weight dyes. (A) Experimental Diagram. First, fluorescent dye is injected into the ventricles. X = position of needle for injection. Next dorsal images are captured over time. Finally, the distance moved by the dye front from the forebrain hinge-point is measured (represented by a red line). (B-C) Merged brightfield and fluorescent dorsal images at 22 hpf (t = 0 min, B) and 24 hpf (t =120 min, C). White line indicates distance of the dye front from forebrain ventricle. (D) Hypothetical sample permeability data. Blue = wild type or controls, red = sample with decreased permeability relative to control, and green = sample with increased permeability relative to control.

Figure 2. Measurement of neuroepithelial permeability to different molecular weight dyes. (A-E) Dorsal merged brightfield and fluorescent images of 22 hpf wild type embryos at t=0 min after injection with FITC-dextran of the following molecular weights: 10 kDa (A), 40 kDa (B), 70 kDa (C), 500 kDa (D) and 2,000 kDa (E). (A’-E’) Same embryo as in (A-E) at t=120 min at 24 hpf. Anterior to left. F= forebrain, M= midbrain, H= hindbrain. Asterisk = ear.