Currently one of the keys to the creation of transgenic or otherwise genetically modified insects is the precise delivery of a cocktail of DNA, RNA and/or proteins needed to affect the desired modification of germ cells.
While having the right equipment is essential, this does not mean the equipment needs to be complex or expensive.
There is an anecdote shared among photographers of an accomplished chef looking at the work of a prominent photographer and after remarking on the stunning quality of the images asked, “What kind of camera do you use?” The photographer, slightly miffed, turned to the chef and says, “I think the food you prepare is exquisite, what type of stove do you use?”
Skilled hands can be more important than fancy equipment.
So it is, more or less, with creating genetically modified insects.
A recent description by Watanabe et al (2014) of their protocol for creating genetically modified crickets illustrates this point very well. Watanabe et al used TALENs to knockout a gene by targeted mutagenesis and used embryo microinjection to deliver the TALENs to presumptive germ cells. The microinjection setup used to deliver the TALENs was an inverted compound microscope in combination with a simple oil-filled glass syringe, some tubing and a glass microinjection needle. Not exactly a ‘high-tech’ arrangement for embryo microinjection. It was quite effective however.
At the University of Maryland’s Insect Transformation Facility where we have created upwards of 500 transgenic insect lines of a variety of species over the last few years we have a number of embryo microinjection stations equipped with a variety of technologies.
My preferred insect embryo microinjection setup is a little different from Watanbe et al’s (2014). I use a stereo-zoom dissecting-microscope equipped high-aperture objective and ocular lenses, a precision mechanical stage, an injection fluidics controller such as Femtojet® (Eppendorf) or Picopump (World Precision Instruments) and quartz-glass needles attached to a remote, hydraulically controlled micromanipulator, all sitting on a compressed-air vibration-dampening table.
I prefer to use a stereo-zoom dissecting-microscope for insect embryo microinjections for a number of reasons. Objects in view are not inverted, the needle is easily accessible, you have a large working distance and depth of field and you can easily adjust magnification to suit your particular needs. In addition, the embryos and needle are situated in a comfortable position relative to your eyes and hands. Overall, I find these features make it easier and quicker for me to get the needle into focus and into position for injection as well as allowing me to rapidly change needles and get back to injecting. Importantly, the system allows me to operate while being comfortable, allowing the Insect Transformation Facility to operate more efficiently.
I find using inverted or traditional compound microscopes somewhat less convenient. Reduced working distances and depth of field make keeping track of the needle more challenging. You tend to see the needle only when it is in the focal plane of the embryo.
Also, with inverted microscopes objects appear inverted and I find positioning the needle a somewhat tricky maneuver without breaking the needle on something as you get it into position. Of course with an inverted compound microscope you have lots of space to move and position the injection slide/coverslip, and injection needle, and that is nice.
The ability to change a needle and get back into position and focus is critical in terms of saving time, but not absolutely necessary for performing high quality microinjections.
Having said all of that, inverted compound microscopes have been used since the early days of insect transformation when Rubin and Spradling first reported making transgenic Drosophila with P-elements in 1982, and as Watanabe et al. (2014) demonstrate, it is a perfectly effective insect microinjection configuration, just not my favorite.
There are many devices for fluidics control from oil filled syringes, “home-built” fluidic devices, and high-end pneumatic microinjection controllers such as Femtojet® (Eppendorf) and Picopump (World Precision Instruments).
The Femtojet® and Picopump (and others) are pneumatic controllers allowing users to precisely control the head pressure being applied to the injection solution in the glass injection needle at all times. For example, one can provide constant pressure to counteract pressure inside the embryo and this can help prevent needle-clogs. Also, one can apply precisely controlled pulses of pressure that can facilitate controlling the volume of liquid delivered.
Watanabe et al (2014) used an oil-filled syringe to control fluid delivery from the glass injection needle. With practice I find these types of systems can be quite effective, but in my hands the rate at which I can inject insect eggs is reduced, as is the rate at which I can change needles, and for some species of insects this can be quite often. Replacing needles when using a pneumatic injection-control system is relatively quick and easy.
Microinjection technologies are used for may applications in biology, with delivering genetic technologies to developing insect embryos being only one. Good equipment can be helpful and there is currently a lot of room for improvements and innovations, but one thing remains constant. Skilled and practiced hands are still required.
The University of Maryland’s Insect Transformation Facility is a not-for-profit service provider specializing in the genetic modification of arthropods. The UM-ITF provides insect embryo microinjection services, transgenic insect production services with experience with over a dozen species as well as training services to researchers world-wide.
Watanabe T, Noji S, Mito T (2014) Gene knockout by targeted mutagenesis in a hemimetabolous insect, the two-spotted cricket Gryllus bimaculatus, using TALENs. Methods 69: 17-21 DOI: 10.1016/j.ymeth.2014.05.006