Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

The overall goal of this procedure is to enable time-lapse analysis of diverse developmental processes using a fluorescent dissecting microscope with simple strategies for realignment after drifting in any direction. This method can help answer key questions in the developmental biology field because it allows direct observation of tissues and organ development in living embryos over several hours. The main advantage of this method is that it's affordable and easy to use alternative to more complex techniques normally used for time-lapsed recording of developing tissues and organs, such as laser scanning or lighted microscopy.

Though this method can provide insights into spacial and temporal alterations that occur during development of sweeper fish embryo and larvae, it can also be applied to investigate early developmental processes and other model organisms. Furthermore, it is suitable to observe dynamic processes in adult organisms, such as root healing and regeneration. In preparation, identify highly reproductive adult fish from the transgenic GFP expressing line.

On the afternoon before the plant micro-injection, set up five male-female pairs of reproductive fish in five breeding containers. In the containers, keep the male and female separated by a removable sieve overnight. On the following morning, just after the lights come on, put a male and female together above the sieve, which will prevent them from eating up their eggs.

Now, observe the fish while they spawn. Once the pair produces about 50 eggs, separate them. Then, transfer the eggs using a Pasteur pipette to a Petrie dish filled with embryo water for a first round of injection.

Repeat this process for each mated pair. If additional eggs are needed, the same mating pairs can be brought together and separated several times. For the micro-injection, advanced preparation of dishes and injection needles is done in a fairly standard manner.

Details can be found in the text protocol. Begin with loading a prepared injection needle. Using a micro-loader tip, backfill the needle with two microliters of MO working solution, and then attach the needle in a micro-manipulator.

Then, the needle tip can be opened. While viewing it with the dissection microscope, use tweezers to pinch the end open, or push the tip against a rigid surface. Then, calibrate the injection volume by injecting the MO solution into a drop of mineral oil on a graticule.

Adjust the injection to make 75 micron drops which will correspond to about 200 picoliters of solution. Next, prepare the embryos to be injected. Arrange the approximately 21 cell-stage embryos along the groove of the micro-injection dish with the vegetal pole oriented toward the micro-injection needle's approach, which is at a 45 degree angle from the groove.

Now, inject the embryos. With the needle, perforate the chorion and the yolk, then inject the loaded Morpholino solution into the yolk. This works for embryos at the one or two cell stage.

After injecting all the embryos, use a wide bore Pasteur to collect them. Transfer the injected embryos into Petri dishes filled with embryo water, and incubate them at 28.5 degrees Celsius. Later in the afternoon, siphon out dead embryos and replace the embryo water.

The next morning, after the embryos have gone through gastrulation, inhibit their melanization by replacing their water with embryo water containing a trace amount of PTU. Be careful, PTU will disrupt proper development prior to gastrulation. Later, at the desired developmental stage, dechorionate the embryos with sharp tweezers.

Using the microscope, grab the chorion with one pair of tweezers, and try to grab the other side of the chorion with a second pair of tweezers. Then, carefully pull the tweezers apart without disrupting the embryo. Then, anesthetize the embryos with Tricaine, and proceed with embedding the embryos and agarose on a 15 micron relocation grid placed in a cover glass bottomed micro-dish.

Then, load one milliliter aliquots of melted agarose into 1.5 milliliter reaction tubes. Put the tubes in heat blocks set to 36 degrees Celsius, and wait for them to cool down. Then, using a wide bore pipette, transfer an embryo to the micro-dish.

Siphon off the excess embryo water and overlay the embryo with agarose. While the agarose is still liquid, use two small pipette tips to orient the embryos. Position the structure of interest, in this case, the pronephros, as close as possible to the glass bottom, and in close vicinity to the grid.

The grid should not overlay the structure of interest. It is advantageous to embed up to four embryos in different micro-dishes, and plan to image the best one. Proper embedding needs some practice.

While the agarose is still liquid, the embryo has to be oriented in a desired position. If it's a pronephros close to the glass bottom, and in a suitable distance to the relocation grid. Check the agarose routinely.

Once it is hard enough to hold the embryos, let it settle for another two to three minutes. Then, overlay the embedded embryo with embryo water containing trace amounts of PTU and tricaine. Decant or siphon off the excess embryo water and lock on the lid to prevent evaporation.

The result shouldn't have any air bubbles. The embryo can now be imaged with the glass bottom facing the objective lens. Using the preparation to make lapse images, for example, by taking Z-stack images periodically over several hours, it is necessary to correct for focal drift.

If possible, apply an auto-focus strategy. Toggle the auto-focus option in the software control. For the reference channel, choose the transmitted light bright field channel to minimize photo toxicity.

It is also essential to choose a sufficiently large specimen dependent search range for the auto-focus to work properly. Before collecting images, position a particular point of the imprinted grid close to the structure of interest in the cross-hairs of the software, then save this position using a screenshot. Then, just before each imaging time interval is over, reference the screenshot and readjust the position of the stage and the focus if auto-focus is not in use.

The presented method was used to analyze kidney development in WT1A morphened embryos of a transgenic zebra fish line. During early nephrogenous, this line chose green fluorescence in the intermediate mesoderm where the kidney progenitors arise, and later on during development in the forming tubules and nephron primordia. Time-lapse imaging revealed normal nephrogenesis in the control Morpholino injected embryos.

At 20 hours post-fertilization, the developing pronephric tubules were visible. At their anterior tips, a spherical accumulation of cells representing the forming nephron primordia could be detected. During the next hours, tubules and nephron primordia grew, and the primodia started to fuse at the midline.

In contrast, nephrogenesis was severely disrupted in the mutants. Although GFP positive tubular structures were visible at 20 hours post-fertilization, they were diffused and underdeveloped. Time-lapse imaging showed nephron primorgia never formed, and a striking number of fluorescent cells located outside the developing pronephrons migrated ventrally, leaving the pronephric field.

While attempting this protocol, it's important to remember that the absence of additional equipment, such as temperature control or anti-evaporation tables requires regular checks for drifts and readjustments. Following this procedure, image embryos can be processed to perform other methods, like immunohistochemistry, incetrohypertization, or real-time PCR in order to identify specific components of cells, tissues, or to assess gene expression.