Fig. 1. (A) A 5-somite embryo prior to flat-mounting. (B) A 5-somite embryo after yolk removal and flat-mounting. A, anterior; P, posterior.

of buffer (warmed to 37°C and mixed well), 2 ^L of T7 polymerase. Assemble the reaction mix at room temperature and incubate at 37°C for 2 h.

3. Treat with RNase-free DNase for 20 min at 37°C in appropriate DNase buffer, check on a gel to make sure the template has been digested (see Note 17).

4. Precipitate probe with 3 M NaOAc and 100% EtOH at -20°C for at least 30 min, wash with 70% EtOH, resuspend in 10 ^L of sterile, deionized H2O. Resuspend purified template at 1 ^g/^L in RNase-free water (see Note 18).

3.3. Hemoglobin Staining With O-Dianisidine

1. Prepare the following solution: 2 mL of o-dianisidine solution; 500 ^L of 0.1 M NaOAc, pH 4.5; 2 mL of deionized H2O; 100 ^L of H2O2. We recommend adding the components in this order.

2. Add 500 ^L to live, dechorionated embryos (Subheading 3.2.1.) in glass vials after removing as much of the E3 as possible.

3. Incubate in the dark for 15 to 45 min. The samples can be examined under a dissecting microscope to monitor the reaction; it is not necessary to remove samples from the glass vials.

4. Once staining is complete, wash three times with deionized H2O. Embryos are very sticky; therefore, it is best to use a glass pipet.

5. Fix stained samples in 4% PFA for at least 1 h at room temperature.

6. If embryos are pigmented, rinse in PBST and then bleach (Subheading 3.2.2.).

7. Store embryos in PBST at 4°C or in MeOH at -20°C (see Note 19).

3.4. Observation and Photography

1. Transfer embryos from 1X PBST into 30%, 60%, and 90% glycerol/PBST.

2. For embryos older than 18 somites, it is easiest to photograph directly using a depression slide to keep embryos in the desired orientation. For embryos between 3 and 15 somites, we recommend flat-mounting to obtain representative photographs (Fig. 1).

3. Flat mounting embryos for photography.

a. Using sharp forceps, carefully remove the yolk from the embryo, leaving the embryo proper intact.

b. Using a pipet, transfer the embryo to a glass slide in a drop of glycerol, avoid placing the embryo directly in the middle of the slide.

c. Using an eyelash or fine nylon loop, carefully maneuver the embryo to the edge of the glycerol droplet such that the embryo is oriented with the dorsal side facing upwards, drag the embryo towards the middle of the slide to an area that is void of glycerol; this should help prevent the embryo from curling on itself.

d. Apply a very small amount of modeling clay to the four corners of a square cover slip by swiping the cover slip gently into the clay to scoop out a small portion.

e. Use the cover slip to gently press the embryo flat, but do not apply too much pressure, only enough to keep the embryo from curling up.

f. After the cover slip is in place, pipet a very small amount of glycerol to the edge of the coverslip to surround the embryo with glycerol. Adjust the amount of clay, the amount of glycerol, and the pressure applied until the embryo is correctly oriented and completely stationary. This process requires extensive practice and a steady hand, but in the end the results are well worth the effort.

3.5. Microinjection

Microinjection is the most widely used technique for introducing nucleic acids and nucleic acid analogues into zebrafish embryos. The technique for these experiments is identical, though the timing of injection is slightly different and will be detailed in each of the subsequent sections (Subheadings 3.6.-3.8.). Microinjection of freshly fertilized eggs has a high survival rate, is relatively nonintrusive, yields reproducible results, and with practice, it is possible to inject hundreds of embryos within an hour. Although the specific equipment used can vary, the basic requirements for microinjection are as follows: a needle puller, capillary needles, a pneumatic microinjector to modulate the amount of pressure within the needle for delivering the injection solution, a micromanipulator for controlling the injection needle, and a mold to immobilize the embryos. Injection is facilitated by temporarily immobilizing embryos in agarose molds. Various immobilization techniques are available and have been described adequately in other sources (21,22). Here, we will detail one recently developed approach that makes use of glass slides to create a simple agarose ledge that holds embryos in a specific orientation with respect to the injection needle. Very little agarose is used to create the mold, permitting clear observation of the needle, the embryos, and the subsequent injection.

3.5.1. The Ledge Mold

Simple glass cover slips are used to create an agarose ledge that will immobilize embryos in the correct orientation (see Note 20).

1. To create a cover slip mold, glue three to four coverslips together to create a "unit," and then glue one unit to another, leaving a thin recess approx 2 mm in width along the length of the unit, as shown in Fig. 2. Make two to four cover slip molds.

2. Place two molds in a Petri dish as shown in Fig. 2 such that the recess faces up and is closest to the top of the Petri dish.

3. Using a transfer pipet, carefully add 1% molten agarose to cover only the recess of the cover slip mold and the surrounding area, as shown in gray in Fig. 2. Only a small amount of agarose is needed as most of the recess will fill by capillary action.

4. After the agarose has cooled, carefully cut along the edge of the recess with a fresh razor blade, separating the agarose from the glass cover slip mold. Extract the cover slip mold from the agarose with the aid of blunt forceps, leaving behind a delicate ledge of agarose.

5. Fill the dish with E3 containing Methylene blue and store at 4°C (see Note 21).

Top of I'l ln dish

Top of I'l ln dish

Fig. 2. (A) A cover slip unit assembled by gluing two sets of three to four cover slips together, forming a recess that will be filled with molten agarose/E3 as shown in (B) to create a ledge for immobilizing embryos.

3.5.2. Needle Preparation

1. Using a suitable pipet/needle puller, pull needles that narrow quickly to a tip. Microinjection needles should be thick enough to easily penetrate the chorion but not so thick as to damage the embryo. Needles that are pulled too long and fine will bend when in contact with the chorion and are more prone to blockages.

2. Break the tip of the needle with fine forceps creating an oblique angle to facilitate penetration. For reproducible results, use a microscope equipped with a micrometer. A large opening will damage embryos, while a small opening is more likely to become blocked. If the needle becomes blocked, use the forceps to re-open the needle by removing the very tip to enlarge the needle. Adjustment of injection pressure or time may be required to maintain a constant volume.

3. Load the needle using a pipet fitted with a microloader pipet tip. Place the tip as far into the back of the capillary as possible and slowly expel the solution as the tip is withdrawn. The filaments facilitate flow of the solution towards the needle tip. Several hundred embryos can be injected with 2 ^L of solution. Insert the needle into the microelectrode; the fit should be snug (see Note 22).

3.5.3. Embryo Collection and Microinjection

Postinjection survival rate is influenced by the quality of zebrafish embryos obtained. Adult fish can be kept together overnight, or they can be kept separate and put together in the morning, in which case it usually takes around 10 min for mating to occur. Immediately after collection, it is difficult to ascertain whether eggs are unfertilized or of low-quality. Therefore some researchers prefer to mix clutches prior to injection, ensuring that not too much time is wasted on injecting embryos that will not develop.

1. Carefully align the embryos under the ledge as shown in Fig. 3 such that the cell is facing outwards, we recommend using a 24-gage wire that has been lightly filed to remove sharp edges. The chorion should remain intact with gentle pressure as it is quite durable.

Fig. 3. (A) Embryos are gently pushed under an agarose ledge for injection, which accommodates almost 50 embryos. (B) The microinjection needle glides through the chorion, the blastomere, and the injection solution is expelled into the cell, right at the margin of yolk and cytoplasm in a volume approx one-fifth the volume of the cell.

Fig. 3. (A) Embryos are gently pushed under an agarose ledge for injection, which accommodates almost 50 embryos. (B) The microinjection needle glides through the chorion, the blastomere, and the injection solution is expelled into the cell, right at the margin of yolk and cytoplasm in a volume approx one-fifth the volume of the cell.

2. After the embryos are aligned, set up the micromanipulator so that the needle is in line with the embryos in the center of the microscope stage. It helps to orient the needle at an angle (around 45°) with respect to the cell.

3. Using a single fluid motion, drive the needle through the chorion and into the cytoplasm of the embryo, expel the injection solution, extract the needle, and repeat. When using the ledge mold it is possible to see the injection location and volume without the aid of a dye, but at first it might be helpful to use Phenol Red as a marker for the solution (see Note 23; Fig. 3).

4. The injection volume should be one fifth that of the embryo, which is approx 1 nL at the one-cell stage. The injection pressure and injection time should be adjusted accordingly to maintain a consistent injection volume. To calibrate the volume, inject into a drop of mineral oil, measuring the diameter of the resulting droplets and convert to volume; adjust injection time, pressure, and needle diameter accordingly; see Note 24).

3.5.4. Postinjection Care

1. After all embryos have been injected, carefully remove the embryos from the mold using gentle pressure from the 24-gage wire; embryos with punctured chorions are very fragile, so exercise caution. The chorions will recover within a few hours.

2. After 4 to 5 h have passed, remove any embryos that are unfertilized, necrotic, or irreparably damaged, transfer to a new dish with fresh E3, and allow development to progress. The survival of resulting embryos is increased by maintaining sanitary conditions (21,22). Embryos that will be raised in a fish facility should be carefully monitored over the course of their development into juveniles and adults.

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