Imaging KV allows for easy analysis of cilia structure and function. It can be initially seen from the 5 somite stage (ss), but is most easily visualized between the 8 and 12 ss as it increases in size over this developmental window ( Figure 2 and note 2).

To assess the age of the fish embryos, roll the embryo on to its side and count along the number of fully-formed somites, starting from the head to the tail ( Figure 1). At the 10 ss the tail will begin to bud off from the yolk and this can be used as a secondary measure ( Figure 2).

Throughout this procedure you should use glass vials and pipettes ( note 4).

At the end of the staining procedure, embryos can be stored for up to two weeks at 4°C in PBS, or long-term at 4°C in 2% PFA in PBS. Staining of embryos does not appear to be significantly diminished by longer-term storage.

Once zebrafish embryos have been stained, the easiest way to visualize KV is by mounting the tail only in relief slides. Many tails can be imaged on a single slide, making slide preparation, imaging and storage easier.

Imaging can be undertaken with any fluorescent microscope that is able to remove out-of-focus light. While the KV can be seen with a traditional global excitation and capture system, there is too much noise resulting from the surrounding tissue. A confocal microscope offers the best image quality, but optical sectioning using structured illumination also gives good results. The staining and mounting method described are suitable for both upright and inverted microscopy.

Following capture, images can be analysed for relevant parameters using FIJI (ImageJ) software. To measure the length of cilia use the segmented line tool on a maximum intensity projection of a z-stack. KV volume can be calculated using the area delineated by aPCK; to obtain a good approximation of the average radius, use the oval tool to outline the KV and measure the area. Note that although the KV may appear as an ellipse in the section, it should be modelled in 3D as a sphere. Using the area of the KV in 2D, calculate the average radius ( r = A / π ) and subsequently the volume ( V = 4 3 π r 3 ).

Light microscopy images were acquired using a Leica MZ16F stereo microscope and Leica Application Suite V4.2. The presence of fluorescently labelled acetylated-α tubulin was confirmed in flat-mounted embryos using a Leica MZ16F stereo microscope with a Leica external light source for fluorescence excitation EL6000 with a red fluorescence protein filter.

Fluorescent microscopy images were captured using an Axio Imager Z1 fluorescent microscope (Zeiss), using an apotome for optical sectioning and a Colibri light source and filter sets for DAPI (blue), Alexa Fluor488 (green) and Alexa Fluor 594 (Red) with a 20X air objective.

Zebrafish were maintained in a Home Office aquarium facility at a temperature of 28°C. All zebrafish procedures were performed under Home Office UK license regulations.

     1. Preparation of paraformaldehyde

Paraformaldehyde is dangerous in its powdered form and should be handled with care using a fume hood and mask. To successfully get PFA into solution it is necessary to warm it to 60°C and add sodium hydroxide to raise the pH. At the correct temperature and pH (pH 7), the powder will go into solution rapidly. After cooling check the pH is correct. You can freeze aliquots (e.g. 10–20 ml) of PFA for later use. Once thawed, keep at 4°C for up to three days.

     2. Fixing zebrafish at the 10 ss

At 28.5°C, embryos reach the 10 ss approximately 14 hpf, making collection difficult (as this is typically very late at night, assuming fish lay eggs in the morning in a standard aquarium with unmodified night/day cycles). To facilitate collection of the 10 ss embryos, it is possible to slow the development of the embryos down by placing them at a slightly lower temperature. At around 24°C the embryos will not reach the 10 ss until approximately 21 hpf (i.e. the following morning).

     3. Embryo fixation

It is usual to remove the chorion before fixation of fish embryos. At early stages, however, the embryos are very fragile and also the yolk tends to stick to almost anything when rehydrated. Thus it is best to leave the chorion on when undertaking the rehydration procedure, prior to immunostaining.

     4. Immunostaining in glass

Embryos should be transferred using glass pipettes; they will stick to plastic and disintegrate easily. Immunostaining should be carried out in glass vials only. Using 2-ml glass vials allows for staining in a small volume of liquid (≤200 µl) which helps to reduce reagent costs and reduces the storage space required. Typical volumes are 2 ml for washes and 200 μl of antibody solution.

     5. Embryo rehydration

Embryos are very fragile at this stage and should be handled with care. When rehydrating, only remove 50% of the solution they are in and replace with the next one (e.g. remove half of the 100% methanol, and replace with half of 75% methanol to give 87.5%, then remove most of this and place into 75% methanol such that the steps are of roughly 12.5%). Mixing methanol with water leads to an exothermic reaction so embryos should be kept on ice throughout to maintain their low temperature.

     6. Primary antibodies

Any antibody which specifically detects ciliary proteins can be used for analysis of KV, provided that it cross-reacts with zebrafish antigen. Staining of the intra-flagellar transport (IFT) machinery, the axoneme, or the ciliary membrane are all possible with this technique and allow for analysis of both structural and functional defects within zebrafish cilia. For simple detection of cilia structure the best antibody is anti-acetylated-α tubulin (Sigma-Aldrich; catalogue number T6793) at 1:500 for the ciliary axoneme. To visualise KV epithelium, counterstain with anti-aPKC at 1:500.

     7. Generation of imaging chamber

While it is possible to purchase slides with wells of varying sizes, most commercially available slides have a circular bottom making it difficult to get a uniform distance from the cover slip to the sample. As a result only one or two samples can be placed in each. To generate a uniformly flat imaging surface, PVC tape can be used to convert any flat slide to a relief slide in which you can create an imaging chamber to fit your own size requirements. For KV visualization, one layer of tape provides ample depth to prevent the sample being crushed, while keeping the sample close to the cover slip. Detailed information and a visual demonstration of imaging chamber generation has been reported previously ¹⁹ .

     8. Embryo dissection for KV visualization

To get a good image of KV it is best to remove the tail tip from the rest of the embryo and lay it flat. Using two pairs of forceps, pinch two-thirds of the way along the back of the embryo, to separate the posterior part of the embryo from the anterior, and between the head and tail to detach the tail from the yolk sac. Remove as much yolk from the remaining tissue as possible.

     9. Removal of PBS from the relief slide

Due to the small size of the dissected zebrafish tail tip, it can be difficult to remove the PBS from the imaging chamber without disturbing the samples. To facilitate this process, use a razor blade to cut off a medium-sized (20–200 µl) plastic micropipette tip at 45°. The cut tip can be placed flat on to the microscope slide allowing for aspiration of PBS without disturbing the samples.

     10. Mounting medium

Mounting medium with an anti-fading agent is essential for the successful imaging of immunofluorescence-stained KV. Some mounting media is available in a hard-setting format e.g. Vectashield H-1500, but it is preferable to use a non-setting medium such as Vectashield H-1200.