High Throughput Microinjections of Sea Urchin Zygotes

Summary

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.

Abstract

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.

Introduction

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 urchins^3,4, 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 conducted^10-12.

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 model^15,16. 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 process^18-21, cell biological processes^22-24, 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.

Protocol

1. Preparation of Protamine Sulfate (PS) Coated Dishes

1. Prepare 1% solution of protamine sulfate (PS) by adding 0.5 g of PS to 50 ml of deionized, distilled water (ddH₂O) in a 50 ml conical tube. Shake well at high speed on a bench shaker at room temperature for 1-2 hr to ensure complete dissolution of PS. This solution can be stored at 4 °C for at least 3 months (make sure to completely dissolve gel-like precipitate before each use)³.
2. Take a sleeve of 60 mm x 15 mm polystyrene Petri dishes and lay out both lids and bottoms on the bench.
3. Pour 1% PS solution in each dish (both bottoms and lids can be used) just enough to cover the surface, leave for at least 2 min. The leftover PS solution can be reused many times within 3 months when stored at 4 °C.
4. Place PS-treated dishes in a beaker filled with distilled water (dH₂O). Leave the beaker under running dH₂O for at least 10 min.
5. PS-coated dishes can be used immediately or air dry for storage. Cover them to prevent dust accumulation. They can be stored at room temperature for 1 month³.

2. Obtain Sea Urchin Gametes and Immobilize the Eggs on a PS-coated Dish for Microinjection

1. Spawn sea urchins by shaking or intracoelomic injection of up to 1 ml of 0.5 % KCl to induce contraction of smooth muscles to release gametes³ (Figure 2).
2. If the animal is a male, as indicated by the white color sperm, collect sperm dry from the surface of an animal into a 1.5 ml tube. Dry sperm can be stored at 4 °C for a week without significant loss of biological function.
3. If the animal is a female, as indicated by the yellow color due to the egg yolk content, collect its gametes by immersing the gonopores in a beaker full of sea water. The eggs will be released from gonopores and settle down by gravity.
4. Filter the eggs from debris such as animal spines using 80 μm Nylon filter mesh. The best results are achieved if the eggs are used within 5-7 hr after spawning.
5. Dejelly the eggs:
   1. Prepare 100 ml of acidic sea water for approximately 1 ml of egg. Use 0.5 M citric acid to adjust pH of sea water from pH 8.0 to 5.1-5.2. Too low of a pH will decrease fertilization and too high of a pH will not allow eggs to attach to the PS coated plates.
   2. Add eggs (not more than 1 ml) to acidic sea water and allow the eggs to settle down on ice for 10 min. Alternatively, the removal of the jelly coat can be facilitated by gentle swirling of the eggs during the treatment. One can effectively dejelly S. purpuratus eggs in 3-4 min this way.
   3. Carefully pour off acidic sea water, add fresh sea water and let the eggs settle down on ice again. Dejellied eggs can be stored on ice for up to 6 hr without fertilization problems

3. Row the Eggs

1. Prepare 1 M stock solution of 3-aminotriazole (3-AT, MW = 84.08) by dissolving 0.84 g of 3-AT in 10 ml of ddH₂O. This solution can be stored at 4 °C for up to 6 months.
2. Prepare 1 mM working solution of 3-AT by adding 50 μl of 1 M stock solution to 50 ml of prechilled sea water. Keep the solution on ice.
3. Prepare mouth pipette (Figure 3) for gentle handling of the eggs:
   1. Pipette tip is manually pulled from glass micropipettes (100 x OD 1.09 mm). Hold the micropipette with both hands over the open flame. As the glass begins to melt, carefully pull the ends of the pipette in opposing directions.
   2. Connect one pulled pipette to a plastic tubing (ID 1.14 mm), seal tight with Parafilm. The length of the tubing should be approximately 50-70 cm.
   3. Insert a sterile filtered P20 or P200 tip at the end of the tube opposite to the glass tubing to be used as the mouth piece.
   4. Clip the end of the glass micropipette with scissors to adjust the diameter of the tip. The optimal diameter slightly exceeds the diameter of the egg (80 μm).
4. Pour   4 ml of 1 mM 3-AT sea water into each PS-coated dish (bottom or lid).
5. Take the mouth pipette, put the P20/P200 tip in the mouth and secure it with the teeth. To aspirate eggs, first aspirate a small amount of sea water. By having a column of liquid prior to loading the eggs, one will have maximal control over the mouth pipetting technique.
6. Position the micropipette tip near the eggs and gently aspirate in several hundreds of eggs in the pipette. Confine the eggs within the glass micropipette.
7. Row dejellied eggs in a straight line on a dish by gently blowing them out of the mouth pipette while moving the tip in a straight line. Row the eggs right before the injections, because fertilization problems may occur due to prolonged exposure to 3-AT sea water³. Moreover, cationic adhesives like PS can be toxic to eggs, and it is not advisable to expose embryos to these reagents for long periods, especially if the fertilization envelopes have been removed.
8. Make a scratch on the dish near the line of rowed eggs using a glass pipette. This scratch will be important to break the injection needle to adjust the flow of solution as needed. Alternatively, the scratch can be made before pouring 3-AT sea water to the dish.

4. Microinjection of the Sea Urchin Zygotes

1. Turn on microinjection station. Here we describe the use of FemtoJet injection system.
2. Pull injection capillaries using a needle puller. An example of a good needle is depicted in Figure 4. Note: For RNA injections, injection capillaries should be pretreated to avoid RNAase contamination³¹:
   1. Place injection capillaries into 50 ml conical tubes and add 50 ml of filtered methanol/HCl (9:1). Shake well and leave overnight for no longer than 10 hr.
   2. Pour off the methanol/HCl and rinse the capillaries 2x with filtered methanol in the same tube.
   3. Drain methanol as much as possible and lyophilize the capillaries overnight to ensure complete removal of methanol.
3. Carefully mount the needle capillary on a needle loading holder and load it with <0.5 μl of a sample solution using microloader tips. Sample solutions usually contain 20% glycerol, depending on the injected material³.
4. Place the dish with rowed eggs on the microscope stage (eggs should be aligned vertically when viewed through eyepieces) and focus on the eggs.
5. Carefully mount the loaded needle onto the needle microinjecting holder and adjust the position of the needle to have a perfectly focused tip in the middle of the field.
6. Depress the button to apply the maximum pressure for needle cleaning. Solution should come out of the needle. Adjust the pressure if necessary: injection pressure should be in the range of 90-360 hPa, with the compensation pressure at approximately 1/3 of the injection pressure.
7. Adjust the solution flow by injecting into unfertilized egg. Using a joystick micromanipulator, direct the tip of the injection needle to the unfertilized egg. To facilitate needle entry, create a slight trembling by gentle tapping of the stage with a hard object such as a screw driver. An injection bolus forming inside the egg should be seen (Figure 5).
8. If the injection bolus is not visible, slightly raise the tip of the needle above the eggs, locate the scratch on the dish and adjust it to be positioned in the middle of the field. Gently tap the needle tip to the scratch mark to break the tip of the injection needle. Adjust the injection and compensation pressures to ensure that the injection bolus does not exceed 1/5 of the diameter of the egg (1-2 pm).
9. Once the injection bolus is calibrated, fertilize the eggs by adding diluted sperm (1:1,000) or 1-2 μl of sperm. Mix well carefully to prevent dislodging the row of eggs.
10. Start injections immediately after the fertilization envelope becomes visible (Figure 5). Move along the line of rowed eggs using the stage controller and inject as many zygotes as possible by poking them with the needle controlled by joystick micromanipulator.
11. After 10-15 min, the newly fertilized eggs will become hardened and cannot be injected any longer. Remove the dish from the stage and carefully aspirate out sperm in 3-AT sea water using a plastic Pasteur pipette.
12. Do not let the zygotes dry. Quickly add fresh prechilled sea water to cover the dish. Incubate the embryos at 15 °C.

Results

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). Fluo...

Discussion

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 organisms^1-7. 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 ...

Materials

Name	Company	Catalog Number	Comments
Glass Pasteur pipettes	VWR	14673-043
Inverted microscope  Axiovert 40 °C	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