Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.
Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.
Efficient and reproducible treatment delivery is one of the main methodological challenges for researchers. Several methods have been established to transiently deliver treatment solutions into the eggs, embryos, and cells. These methods include electroporation (based on a generating transient pores in the membrane using short electrical pulses)1,2, lipofection (delivery through the fusion of treatment-containing liposomes with the membrane)1, microparticle bombardment1 (DNA is precipitated on the micron-sized metal particles that are then used to penetrate the cells at high velocity), and transduction (virus is used as a delivery vehicle of transgenes). At the moment, microinjection is the only approach that holds the advantage of delivering any solution with 100% efficiency with minimal reagents. Moreover, a single injection solution can be composed of a complex cocktail of treatments. This technique has been used to successfully microinject the eggs and embryos from numerous species such as sea urchins3,4, zebra fish5, mouse6, frog7, and cattle8 as well as single cells in the tissue culture9. Single blastomeres injections at later developmental stages have also been conducted10-12.
The current methods of microinjections are based on the pressure-injection method that was initially described by Hiramoto10; however, great progress has been made towards optimization of this process. Excellent microinjection techniques have been described elsewhere11, and here we describe one of the specific methods that is currently used to microinject sea urchin (Strongylocentrotus purpuratus) newly fertilized eggs. For over a century, sea urchins have been a valuable experimental model15,16. Sea urchins are evolutionarily closely related to chordates (including us) and analysis of their genome revealed that they contain all the major gene families as the human17. They produce a large number of synchronously developing transparent embryos that can be easily manipulated. Using sea urchin as a model organism, the sea urchin community has contributed to our understanding of the fertilization process18-21, cell biological processes22-24, and the gene regulatory networks (GRNs)25-28.
Microinjection into sea urchin zygotes requires several steps. First, the eggs need to be immobilized prior to injections (described below). Microinjection dishes are coated with protamine sulfate (PS), which creates a positively-charged surface to which the negatively charged eggs can adhere3. The eggs are dejellied by incubation in acidic sea water (pH 5.15) for 10 min, followed by two washes in natural sea water or artificial sea water (pH 8.0). The dejellied eggs are carefully rowed in a straight line in the middle of PS-coated dish in the presence of 1 mM 3-aminotriazole (3-AT), which is required to inhibit the activity of ovoperoxidase that is secreted from the cortical granules of the egg as a result of fertilization29. This step is important to prevent hardening of the fertilization envelope and to facilitate microinjection needle entry. As an alternative to 1 mM 3-AT, 10 mM paraminobenzoic acid (PABA) can be used. The injection solution is loaded into a microinjection needle using specialized microloading pipette tip and mounted on a holder attached to micromanipulator and pressure unit (Figure 1). Each needle can be used to microinject individual zygotes in multiple experiments on separate days. Microinjection can be performed for 10-15 min until the zygotes harden. The zygotes are then washed with the sea water and cultured at 15 °C. When the embryos reach hatching blastula stage, they release hatching enzyme that digests components of the fertilization envelope30 and allow them to naturally detach from the PS-coated dish. If necessary, the embryos can be gently detached from the dish using a mouth pipette or Pasteur pipette by gently blowing sea water onto the embryos. The described method enables efficient and reliable microinjection of 100-400 newly fertilized eggs on a single dish, providing a high-throughput method for downstream analyses.
1. Preparation of Protamine Sulfate (PS) Coated Dishes
2. Obtain Sea Urchin Gametes and Immobilize the Eggs on a PS-coated Dish for Microinjection
3. Row the Eggs
4. Microinjection of the Sea Urchin Zygotes
GFP and mCherry reporter constructs were in vitro transcribed and microinjected into the newly fertilized eggs. Embryos were incubated at 15 °C for 24 hr (until the blastula stage) and imaged using Zeiss Observer Z1 microscope. Injection of reporter constructs did not lead to any developmental defects (Figure 6). For quantification of fluorescent signals, image acquisition was performed at low magnification (100X) to maximally capture fluorescent pixels (Figures 6D-F). Fluorescent signals were quantified using Axiovision 4.8.2.0. The standard errors for the intensity of fluorescent signals within the population of 100-200 blastulae did not exceed 1.5% (data not shown).
Figure 1. The microinjection setup. (A) A PS-coated dish with immobilized eggs is located on the stage of (B) an inverted microscope. Microinjection needle is attached to (C) a needle holder which is connected to a (D) pressure unit. Movement in the x-, y-, and z-dimensions is directed with using (E) the micromanipulator or (F) the coarse manipulator. (G) A screw driver wrapped with paper towels is used to gently tap the microscope stage to induce slight trembling to facilitate needle entry into the newly fertilized egg.
Figure 2. Spawning of the sea urchins. (A) Sea urchin is induced to shed by intracoelomic injection of 0.5% KCl. The needle points to the peristomial membrane surrounding the mouth of the animal, the only soft part of the animal. (B) Female sea urchin is placed with its gonapores immersed in the sea water in a plastic beaker to collect yellow eggs. (C) White sperm is released from gonapores of the male sea urchin.
Figure 3. Mouth pipette. A mouth pipette consists of three parts: (A) Glass micropipette needle, (B) plastic tubing, and (C) a sterile filtered P20 or P200 tip. The glass micropipette is inserted into the plastic tubing which is connected to the P20 or P200 mouth piece.
Figure 4. Microinjection needle. (A) The microinjection needle is pulled using a needle puller with a specific setting. The setting of the needle puller needs to be empirically determined. Using a Narishege PC-10 needle puller, we use 72.8 °C for heat setting 1 and 83.7 °C for heat setting 2. Using the scratch on the plate we break the tip of the needle to facilitate solution flow. (B) The good needle should have a length of 500-600 μm from a width of 50 μm at the shoulder to 5 μm at the tip.
Figure 5. Injection of the newly fertilized sea urchin eggs. Dejellied eggs were rowed on a PS-coated dish and fertilized. Newly fertilized eggs were injected one by one while moving the microscope stage along the line of zygotes (from top to bottom). (A) Injection bolus is clearly seen to be forming inside the newly fertilized egg at the tip of the needle. (B) Transparent fertilization envelope is formed upon fertilization. (C) Sperm appear as small black dots in the background. Scale bar is 50 μm. Click here to view larger image.
Figure 6. Microinjected zygotes developed into blastulae without developmental or morphological defects. Newly fertilized eggs were injected with fluorescent markers which allow them to be clearly distinguished from the uninjected embryos (shown by arrows). (A-C) Embryos imaged at 400X magnification. (D-F) Embryos are imaged at 100X magnification to capture most of the emitted fluorescence signals for reliable quantification. After 24 hr post fertilization, blastulae were collected and treated with 2x sea water for 2 min to immobilize the swimming embryos3, followed by 1x sea water wash. Images were acquired with Zeiss Observer Z1 microscope and AxioCam monochromic camera. (A,D) DIC images of the blastulae. (B,E) Overlay images of embryos injected with in vitro transcribed mCherry in fluorescence and DIC channels. (C,F) Overlay images of embryos injected with in vitro transcribed GFP in fluorescence and DIC channels. Scale bar is 50 μm. Click here to view larger image.
Microinjection is a powerful technique for delivering various treatments such as DNA, mRNA, recombinant proteins, loss of function and gain of function reagents, dyes and their combinations into eggs, embryos, and cells of various organisms1-7. However, several considerations should be kept in mind when designing a microinjection experiment.
It is critically important to consider the solubility of the delivered treatment and the injection volume. If the microinjected solution tends to form even a slight precipitate, the microinjecting needle would be clogged. Generally it is recommended to centrifuge the solution for more than 15 min before loading into the needle to make sure any precipitate is removed from the solution3. The other approach is to pass the solution through the Millipore spin columns. If it is not possible to avoid clogging, it may be possible to unclog the needle by gentle breaking the tip again, but it becomes hard to control the flow of the solution if the tip is too wide. The large amount of the injected solution can be toxic to the embryo; moreover, the wide tip introduces greater variations among the volumes of solution injected into individual embryos. It is also important that the amount of injected volume into the newly fertilized eggs is adjusted. The injected volume can be calibrated by pulling the needles at various settings followed by injecting the solution into oil. This allows for measuring the diameter (and thus volume) of the droplet32. We find that an acceptable amount of injection solution in the newly fertilized egg should not exceed 1/5 diameter of the zygote as shown in Figure 5.
The main advice is to load the needle right before the injection, to work fast, and to frequently use the cleaning button function of the injector which triggers application of the maximum available pressure to the outlet in order to flush out the blocked injection needle.
Sometimes the needle may clog from the outside, when the sperm and debris from the previously injected eggs stick to the tip of the needle. In that case, it is useful to raise the needle all the way up from the 3-AT water in the injecting dish. Usually the tension caused by the liquid-air interface is enough to remove the debris. Alternatively, gently brushing the injection needle against the scratch on the injection plate may help also. In the extreme case, it is possible to wipe the tip of the needle with the dripping wet Kimwipe. However, such wiping requires extreme caution and practice.
Fluorescent dyes such as 10,000 MW Texas Red lysine charged dextran (2 mg/ml) produce a strong signal that can be detected and used for reliable normalization of the injected solution. When mRNA is to be injected, it is recommended to coinject with mCherry or GFP mRNAs along with the mRNA of interest to ensure that no mRNA degradation took place. In that case visualization of the fluorescent signal is dependent on the translation of mCherry or GFP. If the use of dyes or fluorescent constructs is not possible, it is recommended to row a small amount of eggs on every individual dish to make sure that all the newly fertilized eggs on that dish are injected.
In conclusion, microinjection is a valuable tool for research in genetics, molecular, cellular, and developmental biology. Using this technique, the sea urchin community has made significant contribution to our understanding of cell and developmental biology such as cell fate specification, gene function, gene regulation and biochemical pathways underlying embryonic development16.
The authors have nothing to disclose.
We thank Santiago Suarez for critical reading of the manuscript and Betty Cowgill for aid in photography. We also thank the anonymous reviewers for their critical feedback. This work is supported by the University of Delaware Research Fund.
Glass pasteur pipettes | VWR | 14673-043 | |
Inverted microscope Axiovert 40C | Zeiss | 4109431007990000 | Injection microscope |
Microloader tips | Eppendorf | 5242 956.003 | Load injection solution |
Nylon filter mesh 80 μm | Amazon.com | 03-80-37 | Filter eggs to get rid of debris |
P20 or P200 Aerosol Barrier Pipette Tips | Fisher Scientific | 02707432 or 02707430 | Part of a mouth pipette |
Parafilm | Fisher Scientific | 13 374 12 | Part of a mouth pipette |
Polyethylene tubing | Intramedic | PE-160 | Part of a mouth pipette |
Protamine sulfate | MP Biomedicals, LLC | 194729 | Attach dejellied eggs to injection dishes |
Sea urchins S. purpuratus | Pt. Loma Marine Invertebrate Lab | N/A | |
Sea water | any pet store | Instant Ocean | |
Sterile 60 mm x 15 mm Polystyrene Petri Dish | Fisher Scientific | 0875713A | Injection dishes |
Three-Axis Coarse Positioning Micromanipulator MMN-1 | Narishige | 9124 | Manipulate injection needle |
Three-Axis Joystick Type Oil Hydraulic Fine Micromanipulator MMO-202ND | Narishige | 9212 | Manipulate injection needle |
Transfer pipettes | Fisher Scientific | 13-711-9AM | |
Vertical needle puller | Narishige | PC-10 | Pull injection needles |