Long Protocol for Microinjection in X. Tropicalis
1/9x Modified Ringer's solution(1/9X MR)
1/9X MR with gentamicin (100 microgram/ml)
3% Cysteine by volume in 1/9x MR, pH: 7.8-8.0.
3% Ficoll in 1/9X MR, autoclaved
10X MBS salts autoclaved (see appendix)
Instruments and Supplies:
Scissors and forceps
Mesh coated injection dishes (Fig. 2)
Tips and eppendorf 1,5 ml tubes
Pipettes (p1000 and p20)
|Steps:||At time (min)||Duration (min)|
|1: Harvest Eggs and Fertilize||0||15|
|2: Cysteine Embryos||10||5 - 7|
|3: Prepare Needles||0(during Flood interval)||10|
|4: Microinjection||stage dependent||varies|
|5: Incubate at desired temperature
Step 1: Harvest Eggs and Fertilize
Remove testes from male and place them in a 1.5 ml epi tube with 0.5 ml of 1XMBS+0.2%BSA. Manually harvest eggs into 1X MBS+0.2%BSA pre-coated 5 cm dishes. (See section for obtaining embryos and in vitro fertilization on this website). Mince testes using a microtube pestle, until mixture is cloudy. Add sperm solution to the eggs, mix with pestle thoroughly and incubate at room temperature for 3 minutes. Flood with 0.1X MBS pH 7.8-7.9 (check pH of this solution THE SAME DAY)just enough to cover the eggs. Flood time will be fertilization time (T=0). Incubate at room temperature for 10 minutes. Fertilization can be evaluated at T=2.5-3.0. An approximate measure of fertilization rate is "turning"; embryos tend to turn with the pigmented animal hemisphere up, while unfertilized eggs are impartial. One can also check the egg for a sperm-entry-point in the animal hemisphere. It will appear as a white freckle or indentation on the pigmented field (Figure 1). The dark hemisphere of fertilized eggs will contract, whereas unfertilized ones will not. This contraction will relax after a period of time, so this is only useful a useful indicator in the first 10 - 15 minutes of fertilization. Using these criteria, a rough estimate of fertilization percentage can be had by quickly scanning the dish under a stereomicroscope.
The ultimate sign of fertilization is progression to the two-cell stage. If performing time sensitive experiments, such as injections at one or two cell stage, or collecting stage specific embryos, the time to first division will be important to know. This time period depends on temperature, and can thus be controlled by changing the temperature. This can be done by changing the room temperature, or use of an incubator or a cold plate.
Figure 1 Dark sperm entry point. On an embryo with a very dark animal pole, the sperm entry point may just look like a white spot.
Step 2: Cysteine Embryos
The purpose of this step is to make embryo injection effortless (See section on de-jellying embryos on this website). The embryo is coated in a protective jelly that is difficult to bore through with a fine needle. Prepare a 3% by volume cysteine solution, using 1/9x MR as your solvent. Insure that the pH of your solution is 7.8-8.0 by adding NaOH or another base. Remove the 0.1X MBS you used to flood by gently tilting the dish. Remove as much of the 0.1X MBS as possible to avoid dilution of the cysteine solution, and replace with the cysteine solution. Place your dish on a rocker or on the tabletop and let sit for 3-5 minutes, or until done (7 min max). Gently swirl the embryos. They should become loose and separated. Wash 3 times with 0.1X MBS and two times with 3% Ficoll in 1/9x MR. Leave embryos in 3% Ficoll in 1/9x MR. If the embryos appear to clump together, repeat the cysteine step. Do not hurry through this step! Injecting embryos that are not properly de-jellied is quite frustrating.
Step 3: Prepare Needles (This can be done after flooding eggs, 10 min interval)
A 2 nanoliter (8 hash marks) injection is ideal for a one or two cell tropicalis embryo. Injections at later stages will require smaller volumes. We will prepare a needle that is large enough to deliver our volume, at a reasonable pressure, but fine enough not to rupture the embryo. Too fine of a needle will often get clogged. Use a micropipette puller to taper a small glass capillary tube (see appendix for further details). Pull necessary number of needles before starting in vitro fertilization. Back-load your injection solution into the needle using a pipetman and a thin pipet tip. We recommend the Seque/Pro Capillary tip from Bio-Rad Laboratories(Cat#223991). Place your needle on the picospritzer. Break the tip of your needle under the microscope t medium power using a pair of forceps. Lower the needle into a petri dish filled with mineral oil. At high power, apply pressure with the picospritzer. A small bubble will form in the oil, which will allow you determine the volume of your injection (see appendix for further details). Calibrate your pressure and the size of the needle accordingly until you have a 2 nanoliter (8 hash marks) injection.
Step 4: Microinjection
We use a mesh-bottomed dish to holds the embryos (figures 2 and 3), (See appendix for instructions on making dishes). Add a small amount of 3% Ficoll in 1/9x MR to the dish. Under the microscope, select embryos for injection and transfer them (50- 100 embryos depending of the experiment) to the mesh-bottomed dish. Try to do this promptly before the contraction discussed in Step 1 relaxes, or selecting fertilized embryos may become difficult. If the rate of fertilization is lower (10 - ~70%), select embryos for injection by picking fertilized embryos individually. I the fertilization rate is high, you may choose to simply suck up more embryos than you need with your pipette, transfer them to the dish, inject everything, and sort out the unfertilized ones later. This method saves some time before cell division, giving a bit more injection time.Arranging the embryos in rows makes it easy to keep track of which ones have been injected. Avoid placing embryos too close to the wall of the dish because the angle of the needle will cause it to break when it contacts the dish wall. Using a flexible pipette tip, gently nudge and rotate embryos into the isolated wells of the mesh until the are all aligned and ready to be injected. Carefully draw out almost all the 1/9x MR without disturbing the embryos. Once the liquid is drawn out, the embryos will be fixed in position in the wells and will not move when you inject. Inject in an organized manner-row by row or column by column. At the end of each row or column check the needle is not clogged by injecting above the embryos(try injecting in the solution in the dish and/or increase the air pressure to try to unclog the needle. Ultimately you may need to re-break and re-calibrate your needle, or start over with a new needle and solution). After all embryos are injected, fill the injection dish with 3% Ficoll in 1/9x MR and dislodge embryos by agitating gently. Label dishes covers. After 1 hour, transfer the injected embryos to a LABELED (cover and bottom) petri dish containing 1/9x MR and incubate at desire temperature (22-28ºC). Be sure to remove any dead or deformed embryos at this point.
Figure 2: Injection dish
Figure 3: Embryos sitting on mesh in injection dish
Step 5: Store embryos and collect at appropriate stages
Grow embryos until the desired stage is reached. See Raising Tadpoles section on this website for more instructions.
While X. tropicalis can be successfully raised in a more limited temperature range than X. laevis, temperature can be manipulated to facilitate collecting embryos at the desired stage. Andrea Wills has generated the following table as a guide:
Table to Calculate Time from IVF to Collection of Desired Embryonic Stage
* hours at room temperature—in practice this varies from about 21-24 degrees C.
^ I have tried growing embryos up at 19-20oC, which slows them down by a couple hours and makes these stages a bit more manageable.
** fertilized and injected at room temp., then shifted to 28 after injections (usually about 2 hrs after fertilization)
^^ remember, injections can't take place till about half an hour after fertilization, and then you have to stick around another hour or so to put them in 1/9 MR +gent, so you can't go home till about 1.5 - 2 hours after fertilization.
1x Modified Ringer's Solution Recipe:
0.1 M NaCl
1.8 mM KCl
2.0 mM CaCl2
1.0 mM MgCl2
5.0 mM Hepes-NaOH, pH 7.6 or 300 mg/I NaHCO3
Buffers (From Early Development of Xenopus Laevis, Laboratory Manual, Sive, Grainger, Harland, Cold Spring Harbor Laboratory Press)
1X MBS (Modified Barth's Saline), pH 7.8
Two solutions: 0.1 M CaCl2 and 10X MBS salts.
0.1 M CaCl2= Dissolve 11.1 g/liter, Autoclave and store aliquots at -20ºC or 4ºC. Usually do 200 ml: 2.22 g.
10X MBS salts, pH 7.8
For 1 L:
NaCl 51.43 g (880 mM)
KCl 0.7455 g (10mM)
MgSO4 1.204 g (10mM)
HEPES (pH 7.8) 50 ml Hepes 1M (50 mM)
NaHCO3 2.1g (25 mM)
Dissolve in dH2O, adjust final pH to 7.8 with NaOH, adjust volume and autoclave. Store at RT.
Prepare the final MBS solution by mixing 100 ml of 10X salt solution with 7 ml of 0.1 M CaCl2, and adjust the volume to 1 liter with distilled water, adjust pH to 7.8. This is MBS 1X.
1X MBS + 0.2% BSA
For 200 ml: dissolve 0.4 g of BSA in 180 ml of 1X MBS solution. Adjust volume. Check pH 7.8. Filter sterilized.
Dilute 1X MBS 10 times in distilled water, check pH: 7.8-8.0 Important: check pH the day of fertilization.
We use the Picospritzer II from Parker Hannifin Corporation (www.parker.com). For our picospritzer we try to use a duration around 300 milliseconds. To determine the volume of your injection measure the distance between units on your reticle at a certain magnification. By knowing the distance between units one can determine the diameter of the bubble that forms when you inject into mineral oil. Once you know the diameter you can determine the volume of your injection by using the equation for the volume of a sphere V= 4πr3/3. Post a list of the volumes associated with different sized bubbles beside the injection bench for ease of use.
We employ a type Z-1 microinjector from the Narishige Group.
We utilize Sutter Instrument Company's model p87 micropipette puller to fabricate our needles.Our needles are pulled using program 1 under the following settings: heat 360, pull 180, velocity 80, time 60, and pressure 500. For our needles we use borosilicate glass capillaries, item number TW100F-4, from World Precision Instruments.
Mesh coated injection dish:
You will need the following supplies to make a mesh coated injection
dish: a 35 mm Falcon petri dish or any other polystyrene dish, a 500 or 800 micrometer polypropylene mesh, and methylene chloride . We use Spectrum Laboratories' Spectra/Mesh. To make a dish cut out an appropriately sized piece of mesh, take the Falcon dish and add methylene chloride to its center and then place the mesh on top. The methylene chloride will melt the polystyrene but not the polypropylene mesh.
Special thanks to L. Zimmerman, E. Amaya, P. Meade, N. Hirsch, and S. Borland (Indiana U-Axolotl Colony) for help with developing this protocol.
This page contributed by Timothy Grammer and updated by Joanna Yeh.
Updated by Maura Lane 02/10/17