The overall aim of this procedure is to orally administer materials to the intestinal lumen of zebrafish larvae. First, the larvae are anesthetized and placed onto an injection mold lined with methyl cellulose. Next, the larvae are gently submerged into the mold and aligned with the angle of the gavage needle and microm manipulator.
The needle is then inserted into a lava's mouth and maneuvered into the intestinal bulb. Once in place, the material is released into the lumen. The final step is to extract the lava from the methyl cellulose and place it in fresh media to recover from anesthesia.
Ultimately, this method can be used to deliver materials specifically to the intestinal lumen of zebrafish larvae, where transport within the intestine or in extraintestinal tissues can be studied. The main advantage of this technique over existing methods, such as immersion into water containing the material of interest, is that the amount of material delivered into the animal as well as the timing and dosage can be re rigorously controlled. This decreases experimental variability and in some cases toxicity associated with certain materials.
We first had the idea for this method when we had difficulty applying certain fluorescent probes at high enough levels to be detectable in the intestinal epithelium by microscopy. High concentrations with sometimes kill the fish and still not yield high enough fluorescent signal for imaging To begin. Pull several gavage needles using a micro pipette puller and borrow silica glass capillaries.
Move to a stereo microscope to clip the needles. First, align the tip of the needle with the end of a transparent ruler or graphical. Once aligned, use fine tip watchmaker forceps to clip the needle around one millimeter from the tip.
Examine the needle tip at 100 times.Magnification. The needle should be around 27 to 30 microns in diameter and blunt. Sharp or jagged needles should be avoided for more consistent results.
A micro forge can be used to better control the needle clip point and to fire polish the needle. Tips to remove sharp edges. Next, prepare the gavage solution.
For most applications, phenol red is added to aid in the visualization of the micro injector and placement of the gavage solution. After filling with mineral oil, mount the gavage needle onto the nano jet two Microinjection unit. Wrap a plastic 10 centimeter Petri dish with biofilm to provide a flat waterproof surface and aliquot around two microliters of the gavage solution onto the para film.
The next step is to manually backfill the gavage needle. Take care not to introduce air bubbles into the mineral oil. To calibrate the ejection volume, inject small droplets into a small Petri dish filled with mineral oil until they're consistent in size, measure the diameter of the droplets using the graphical and to calculate the droplet volume.Volume.
Consistency between needles is plotted here Before using the needle for gavage, remove any residual mineral oil by rinsing in clean zebra fish media. After warming the aros mold cover three grooves with 3%methyl cellulose. The groove should be covered just enough to hold the fish, but not so little that the methyl cellulose dries out quickly.
Next, aliquot around four milliliters of zebrafish media into the wells of a six well plate transfer zebra fish larvae for each experimental group into separate wells. To anesthetize the fish, prepare a three times trica solution in the appropriate zebra fish medium. One Welles time.
Anesthetize the fish by mixing an equal volume of trica solution into the media and swirling gently. As soon as the fish stop moving, use a wide borg glass past stir pipette to place them one by one onto the methyl cellulose with their heads on the 45 degree angle of the groove and their tails laid towards the 90 degree angle of the groove. Use a blunt dissection probe to gently press each lava into the methyl cellulose, stabilizing their position.
Next, adjust the nano jet two microinjection rig so that the needle is tilted at a shallow angle. Make sure that the extension range will be sufficient to reach past the bottom of the well. To minimize the need for future adjustments, set the microinjection unit to release 4.6 nanoliters using the slow inject setting position one hand to make minor adjustments of the plate containing the immobilized larvae and the other to control the micro manipulator of the microinjection unit.
Generally, individuals new to this method will struggle because the zebra fish are very small and easily damaged. It takes practice to identify the esophageal structure underneath the stereo microscope and to determine the appropriate amount of pressure to apply to the gavage needle to reach the intestinal lumen. First, gently maneuver the gavage needle into the mouth of an anesthetized fish.
Pass through the esophagus and slightly depress the esophageal sphincter to introduce the tip just inside the anterior intestinal wall. Once inside the anterior bulb, gently administer the material at these settings. The delivered volume should just fill the anterior bulb of the intestine and not leak out of the esophagus or clo acre.
Retract the needle quickly and smoothly. Take care not to release significant quantities of material into the esophagus. After removing the gavage needle, transfer the fish to a dish containing fresh media.
Revive the larvae from anesthesia and remove the methyl cellulose by rinsing them in fresh media several times. Transfer the larvae to a Petri dish or six well plate until needed. For subsequent analysis, micro gavage can be used to deliver diverse materials into the zebra fishing testing.
To begin, prepare the necessary gavage solutions to provide a demonstration of this method. 10 KD dextran conjugated to Texas. Red is used to analyze barrier integrity of the intestinal epithelium.
Following the steps demonstrated earlier gavage groups of 10 to 20 zebrafish larvae for each experimental condition. Repeat this on duplicate or triplicate groups if possible. After gavage, recover the larvae from anesthesia and allow them to swim freely in fresh media.
After approximately 20 minutes, re anesthetize the larvae as shown earlier. Once anesthetized position the larvae in 3%methyl cellulose on top of a 3%arose block image, the larvae with a fluorescent stereo microscope and a Texas red filter select a magnification that displays the larvae from the snout to immediately posterior to the end of the c clo acre. The fluorescent should be intense in the intestinal lumen.
However, barrier function is evaluated by the level of dextran that appears in extraintestinal tissues adjust the exposure to enable visualization of trunk fluorescence. Even if this over exposes luminal fluorescence when administered alone, dextran is retained within the lumen of the intestine. However, in larvae co gaged with EDTA, which disrupts epithelial tight junctions, dextran seen in circulation and antisemitic vessels and spaces.
In these examples, image J was used to quantitate the relative mean fluorescence in a region of interest in the zebra fish trunk and normalized to an ROI outside of the fish as an internal control. Here, results from a regular needle are compared to those from a micro forge needle. Fewer larvae in the dextrin only group exhibited extraintestinal fluorescence, suggesting that micro forge needles are less likely to damage the larvae.
Looking at one region more closely, dextran is again found restricted to the intestinal lumen without leaking into the vasculature or extraintestinal spaces. In the absence of EDTA, when co gaged with EDTA dextran is observed in extraintestinal spaces and within the lumen of the dorsal aorta and posterior cardinal vein, the blood signal here is due to blood flow. While attempting this procedure, make sure that the needle does not have jagged edges, that the fish are well positioned and secure in the Methylcellulose, and be careful and patient while learning to maneuver the needle through the larvas esophagus.
This technique can be utilized in a wide range of applications in the zebrafish that require controlled introduction or visualization of materials within the intestine. It allows for rigorous control over timing, route, and dosage of delivery into the animal, and greatly reduces variation within an experiment and between experiments.
