Single Cell Transfection in Chick Embryos

The overall goal of this procedure is to inject single cells of chic somites or neural tubes with a plasmid encoding green fluorescent protein. This is accomplished by first pulling microinjection pipettes with an appropriate tip opening diameter. The second step of the procedure is to prepare a plasmid stock for injections.

The third step of the procedure is to prepare the embryos for injection. The fourth step of the procedure is to load the micro pipettes with plasmid DNA. The final step of the procedure is to inject the desired tissues.

Ultimately, results can be obtained that show the range of derivatives in the number of cells produced by single labeled progenitors in chosen som and neural tube subdomains through immunofluorescent microscopy. Hi, I'm Rael from the Lab of Professor Heim in the Department of Medical Neurobiology at the Hebrew University of Jerusalem. Today we'll show you a procedure for injecting single cells of chicken somites and neural tubes.

We use this procedure in the lab to study line segregation of so mite and neural tube cells. So let's get started To generate single transfected cells in chick embryos begin by pulling micro pipettes. We pull our pipettes on a standard Naga PP 83 pipette puller with a heat setting of 13.5.

The precise setting must be calibrated to achieve a pipette tip diameter of between eight and 10 microns. We use filament containing Boris silicate glass tubes with an outer diameter of one millimeter and an inner diameter of 0.75 to 0.78 millimeters. You can also use commercially available micro pipettes, lacking a filament of one millimeter outside diameter and 0.75 millimeters inner diameter.

The prepared pipettes are stored in a plastic dish with the tips protected from contact with the edge of the dish. To prepare the DNA for microinjection, the plasmid of interest is transfected into DH five alpha e coli. Using standard procedures the following day, the plasmid is purified using a Maxi prep kit for non clonal injections, we use a DNA concentration of one microgram per microliter.

For experiments where individual clones are desired, the plasmid concentration will be much lower and will need to be adjusted. For P-C-A-G-G-F-P-A concentration of 0.05 to 0.1 microgram per microliter works well. Once the optimal concentration has been determined, we recommend dilution of the entire purified plasmid stock rather than individual preparations to a predetermined concentration.

The latter might introduce some variance in success rates, probably due to variations in purity and other parameters, such as the proportion of supercoil plasmid to prepare eggs for microinjection chicken eggs that have been incubated while lying on their long axes until the desired developmental stage are swabbed with 70%ethanol and allowed to dry. Next, pierce the pointed end of the egg with surgical scissors. Using a 10 milliliter syringe and a 19 gauge needle, draw two milliliters of albumin from the egg, seal the holes with hot paraffin prior to the injection.

Use surgical scissors to open a window at the top of the eggshell to prevent debris from entering the hole. Place a piece of tape over the window and cut a hole through it. To help visualize the embryo, use a fire pulled capillary mounted on an aspirator to inject a non-toxic ink such as pelican 17 underneath the blasted derm.

Next, access the embryo by cutting and diverting the vital and or amniotic membranes using 0.14 millimeter insect pins mounted on a needle holder to load the DNA into the pulled pipette Eloqua and add a small amount of fast green powder using the capillary. Draw a minimal amount of plasmid mounted capillary of adequate diameter onto an aspirator. Then insert it into the end of the pipette opposite to the sharpened tip and load the DNA as close to the sharpened end as possible.

Next, attach the pipette onto a manual air pressure injector mounted on a micro manipulator. Apply a minimal amount of pressure if necessary to allow the plasmid solution to reach the tip of the pipette. Manipulate the pipette into the liquid medium of the egg while applying some air pressure to prevent clogging of the pipette tip.

Once the relevant tissue domain is pierced, the change in pressure is often enough to release the plasmid with visible fast green. If this does not occur, apply slight pressure to the syringe until the plasmid fast green mixture is ejected. Repeat this procedure along the axis of the embryo following the injection PBS may be added.

Finally, seal the window with sticky tape and return the eggs to the incubator. Avoid ventilation and include small containers of water in the incubator to help maintain moisture and increase yields. If the target site of the injection was superficial enough to be readily apparent, such as within the dorsal somites neural tube or ectoderm, one can measure the success rate of the experiment by performing a whole mount observation Using a fluorescent binocular.

This step is particularly helpful for evaluating whether too many or too few cells have been labeled. If further incubation is desired, add antibiotics containing PBS solution to the egg after viewing and replace the ceiling tape. In addition, one can assess the rate of success of the injection by serially sectioning the embryos and performing immunohistochemistry.

Following four hours or more, post-injection embryos are fixed and processed for paraffin sectioning. Glass mounted sections are then depa finalized and immunostain for reporter visualization amplification methods such as biotin streptavidin might be desirable. Finally, the serial sections are scanned to detect labeled cells in injected segments.

If upon initial trials multiple rather than single cells are obtained, the plasmid stock must be diluted in the process repeated until adequate results are achieved. We typically carry out a single injection of P-C-A-G-G-F-P per so mite while targeting six to 12 segments per embryo. Under this setting, a clear signal can be detected by whole mount observation, eight hours post injection in at least one segment in one embryo out of 10.

In this example, a so mite was injected with a GFP encoding plasmid, and five hours later a single fluorescent cell is observed in a whole mount view of the embryo shown here. When observed 24 hours following injection to the central so mite, a clone of fluorescent cells is observed in a whole mounted embryo two days following clonal injection. Sectioning of this segment reveals the presence of clonal progeny in both dermis and muscle, demonstrating that single progenitors generate both derivatives.

As for somites, single neuro epithelial progenitors in the dorsal neural tube can also be impaled as illustrated in this transverse section, photographed five hours after injection here. Serial section analysis revealed that clonal transfect were attained when the success rate was about 10%This means that in order to achieve clonality, the technique is by definition highly inefficient. On the other hand, under these conditions, more than 93%of the positive labeling events were found to be clonal.

We've just shown you how to inject single cells of chicken embryos when doing this procedure. It's important to remember to dilute large quantity of stock plasmid to an ideal concentration for single cell infections, and then use it throughout your experiments. So that's it.

Thanks for watching and good luck with your experiments.