Embryo Microinjections: High Tech, Low Tech ?

Rob Harrell

Rob Harrell, Manager          University of Maryland’s Insect Transformation Facility,       Producing genetically modified insects for the research community.

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.

Oil-filled syringe for microinjecting cricket eggs

Oil-filled syringe for microinjecting cricket eggs (black arrow).  Image from Watanabe et al (2014)

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.

UM-ITF Microinjection Station

One of the University of Maryland’s Insect Transformation Facility’s insect embryo microinjection stations.

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.


Genetic Technology Delivery by Embryo Microinjection

Genetic Technology Delivery by Embryo Microinjection

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



  1. Thanks very much for that interesting post Rob. I totally agree that a fancy setup is not required for insect microinjections, although might be helpful to achieve a higher throughput. In my previous lab I had occasionally injected with a Femtojet setup and was at first shocked to find that the injections in my new lab are done with an air filled 20 ml syringe. However, with a little practice the results are very much comparable.
    I would be interested what your thoughts on the injection needle are? In my mind breaking a good needle is probably the most important factor that determines the success of an injection. Currently, I am pulling glass needles on an laser equipped needle puller, fill the closed needles with the injection solution and then break the needle tip under oil on the edge of a glass cover slide. Unfortunately, how the needle breaks is pretty much random and I might have to try several needles until finding a good one. In my previous lab needles were broken on a needle grinder, which seems to improve the needle openings. However, the needle grinder is an expensive piece of equipment. How do you open your needles and what is your experience with different ways of doing it?

    Many thanks for sharing your insights!

  2. As Fillip Port points out the process of opening needles is one of the critical factors in successful embryo microinjections because the less damage the needle causes to the embryo during injection the more likely the injected embryo is to survive and if embryos do not survive you are not going to produce transgenic lines. He also points out that the process for opening these needles produces pretty random results, some are sharp and some are completely dull. I would love to find a method for opening needles that removes this randomness. There are many ways to open a needle, again it all depends on which way works the best for you. In the past when I was doing mostly injections in Drosophila melanogaster and the Caribbean Fruit Fly Anastrepha suspense using borosilicate glass capillaries I beveled every needle I used. These beveled needles worked well and the injections had very good survival and good transformation rates. You could almost inject the entire day with one needle. Before I started to bevel the needles I opened them the way I was taught, bring the needle tip close to the edge of the slide until it is barely touching then gently move the slide until the needle is open. This method does not work very well for me, I find that opening the needle in this fashion more often than not produces blunt needles. It’s probably just me since I know this method works for other people. The reason I think the needles come out blunt from this method is you are brushing the glass tip against a very hard unforgiving object, the glass slide. I believe it breaks the tip into a flat end making it dull rather than an asymmetric break which makes the needle slip into the embryo more easily. In the Insect Transformation Facility (ITF) we inject mainly mosquitoes and use quartz glass capillaries. I have tried to bevel the quartz needles, but in my experience it does not seem to make them any sharper than opening them by brushing them against something. The way I open needles currently, and I would be interested in how others are opening their needles, is I find something on the double stick tape that is under the oil, like a fiber of some kind, the smaller the better, I gently lower the needle on to the fiber so the needle is perpendicular to the fiber just barely touching it along the needle taper, I slowly draw the needle back across the fiber. I repeat this a few times until air starts to pass through the tip forming small bubbles. When opening a needle I always use the same pressure to start, 40 psi, the max on the system I have currently set up. When starting with the same pressure each time, I know that the smaller the bubbles at opening, the sharper the needle. If I cannot find a fiber, I use one of the eggs and draw the needle back across it until the needle is open. This technique works well for me, but as I said I am interested to hear how others are opening their needles.

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