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Microinjection Protocol


Pre-injection Checklist:

Reagents:
1/9x Modified Ringer's solution (1/9x MR)
1/20x MR with gentamicin (100 microgram/ml)
3% Cysteine by volume in 1/9x MR
1/9xMR+3% Ficoll
10% Calf serum by volume in Leibovitz L15 media (L15+10%CS)

Instruments:
Micropipette puller
Picospritzer Microinjector
Scissors and forceps
Mesh coated injection dish
Eppendorf tube


Post Fertilization

Steps:
At time (min)
Duration (min)
1. Flood eggs (1/9xMR+3%Ficoll)
0
10
2: Cysteine Embryos
30
as needed (10-20 min)
3: Prepare Needles
35
10
4: 2 cell
60-80
30


Step 1: Harvest Eggs and Fertilize

Manually harvest eggs into L15+10%CS pre-coated dishes. (See section for obtaining embryos and in vitro fertilization on this website.) Mince testes in 500 microliters of L15+10%CS. We mince the testis in 1.5 ml eppendorf tubes using a microtube pestle. Pipet sperm solution on to the eggs and incubate at room temperature for 2 minutes. Flood with 1/9x MR+3% Ficoll (see below). Flood time will be fertilization time (T=0). Fertilization can be evaluated at T=25-30. An approximate measure of fertilization rate is "turning": embryos tend to turn with the pigmented animal hemisphere up, while unfertilized eggs are impartial. However "turning" is not as reliable for X. tropicalis as it appears to be for X. laevis in our experience. Fertilized eggs can be recognized by a sperm-entry-point (SEP) in the animal hemisphere and contraction of the animal hemisphere. SEPs will appear as a dark freckle or indentation on the pigmented field. In embryos with very pigmented animal poles, the SEP often appears to be a white spot, but don't confuse it with the germinal vesicle!


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.

In our experience, trop embryos can be difficult to microinject for two reasons: first, the embryos can be very leaky when punctured with the needle and they develop mild gastrulation defects (failure to close the blastopore) due to aggressive removal of the jelly coat. (Uninjected completely de-jellied embryos will fail to fully gastrulate while only partially de-jellied embryos will gastrulate without problems). In order to minimize the leakiness, we try to keep the embryos in a Ficoll solution as much as possible. However, once the wound has healed we then transfer the embryos to a low salt solution which seems to improve gastrulation.


Step 2: Cysteine Embryos

The purpose of this step is to make the injection of the embryo easier by complete removal of the jelly coat. The embryo is coated in a projective jelly that is difficult to bore through with a fine needle. To de-jelly the embryos, prepare a 3% cysteine solution, using 1/9x MR. Insure that the pH of your solution is 7.8-8.0 by adding NaOH or another base. Free base cysteine rather than cysteine-HCl can be used which requires less NaOH to bring to neutral pH. This appears to improve the firmness of the eggs (E. Amaya). Remove as much of the 1/9x MR+Ficoll as possible to avoid dilution of your cysteine solution, and replace with the cysteine solution. Place your dish on the tabletop and let sit for 5-20 minutes. Gently stir the embryos. Avoid vigorous swirling or the embryos will develop secondary axes. The embryos should become loose and separated. Wash 3-6 times with 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. Once the embryos are adequately de-jellied, transfer them back to 1/9xMR+3% Ficoll.

Step 3: Prepare Needles

A 2 nanoliter injection is ideal for a one or two cell tropicalis embryo. Injections at later stages will require smaller volumes. We prepare needles that are large enough to deliver our volume, at a reasonable pressure, and fine enough not to rupture our embryo. Too fine a needle will often get clogged. Use a micropipette puller to taper a small glass capillary tube. Back-load your solution to be injected into the needle using a pipetman and a thin pipet tip. We recommend the Seque/Pro Capillary tip from Bio-Rad Laboratories. Place your needle on the picospritzer. Under the microscope using a pair of scissors or a forceps break the tip of your needle. Lower the needle into a petri dish filled with mineral oil and apply pressure with the picospritzer. A small bubble will form in the oil, which will allow you to determine the volume of your injection. Calibrate your pressure and the size of the needle accordingly until you have a 2 nanoliter injection.

Step 4: Microinjection

We place embryos in a mesh-bottomed dish which holds the embryos (figures 2 and 3). (See appendix for instructions on how to make dishes.) Pre-coat your mesh-bottomed dish with L15 and then fill with 1/9x MR+3% Ficoll. Under the microscope, select embryos for injection and transfer them to a mesh-bottomed dish. Fill the dish from the center out. Avoid placing embryos too close to the wall of the dish. 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 the 1/9x MR+Ficoll without disturbing the embryos. Inject in an organized manner-row by row or column by column. This will allow you to keep track of what you injected with minimal effort. At the end of each row or column check that the needle is not clogged.

Fig 2. Injection dish
Fig 3. Embryos sitting on mesh in injection dish

Alternatively, one can use an agarose coated dish. The agarose coated dishes are much more gentle on embryos especially when they are especially soft and squishy. One difficulty with the agarose coated dish is that the embryos don't line up in rows and columns and instead form a large clump which can make keeping track of which embryo has been injected from those that need to be injected difficult. Nevertheless, some users find agarose coated dishes much easier.

After all embryos are injected fill the injection dish with 1/9x MR+2.5% Ficoll and dislodge embryos by agitating. Transfer the injected embryos to an agarose-coated petri dish containing 1/9xMR+3% Ficoll. Some injected embryos may need to be discarded due to leakage of cytoplasm, which will become apparent shortly after injecting. After 30 min to a few hours (before gastrulation begins), transfer the healthy embryos to an agarose-coated dish with 1/20xMR+gent. We raise our embryos in agarose covered dishes (E. Amaya) which greatly prevents the embryos from sticking to the dishes. We don't use agarose-coated dishes during the fertilization steps because we found that the cysteine solution/de-jellying process would sometimes breakdown or loosen the agarose coating.

Selection of firm embryos for injection appears to be critical. We ovulate and collect eggs from several females for each experiment, keeping the individual clutches of eggs separate during the fertilization process. Only the firmest clutch of eggs is chosen for injection. Selecting only the firm eggs has greatly enhanced our success rate. Paul Meade's lab recommends this also. We are currently selecting females in our colony for firm, well-developing eggs and cull frogs that fail to do so, in order to generate a colony of good egg producers for microinjection experiments.

In our experience, completely de-jellying the eggs is critical for easy microinjection. However, with the jelly coat completely removed, the vitelline envelope can become loose and the eggs can become soft especially in Ficoll. However, injection without Ficoll leads to significant amounts of leakage from the injection site so Ficoll is critical. If the embryos are allowed to grow up in Ficoll, we find that a sizable percentage (20-30%) often develop blastopore closure defects. We believe this effect is because complete removal of the jelly coat inevitably results in loosening of the vitelline envelope. The envelope appears to be important for giving some support for gastrulation movements. Therefore it is important to remove the embryos from Ficoll into a low salt solution (1/20xMR+gent). We suspect that at this low salt, water enters the embryo and adds to its rigidity which improves its support for gastrulation. Although we can only speculate on the reasons, we have had a significant improvement in microinjection success with this protocol.


Step 5: Store embryos and collect at appropriate stages

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

 

Stage

Approx. Time at 22*

IVF time

Collect Time

Boost time

Approx.

Time at 28**

IVF time

Collect Time

Boost time

9

7 hrs

1pm

8pm

9am

6 hrs

1pm

7pm

9am

10

8 hrs

1pm

9pm

9am

6.5 hrs

1pm

7:30pm

9am

11

9.5 hrs

1pm

10:30pm

9am

7.5 hrs

1pm

8:30pm

9am

12

11 hrs

Noon

11pm

8am

8.5-9 hrs

1pm

10pm

9am

13^

13 hrs

7pm

8am

3pm

10.5 hrs

noon

10:30pm

8am

14^

14 hrs

7pm

9am

3pm

11-12 hrs

Noon

11pm-12am

8am

15

16 hrs

5pm

9am

1pm

--

 

 

 

16

18 hrs

3pm

9am

11am

--

 

 

 

17

20 hrs

1pm

9am

9am

--

 

 

 

18

21 hrs

1pm

10am

9am

--


 

 

19-20

22-23 hrs

1pm

noon

9am

--

 

 

 

21-23

24-25 hrs

1pm

1pm

9am

--

 

 

 

24-26

26-28 hrs

1pm

afternoon

9am

--

 

 

 

* 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 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 2 hours after fertilization.



Appendix

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

Picospritzer:
We use the Picospritzer II from General Valve Corporation.

Microinjector:
We employ a type Z-1 microinjector from the Narishige Group.


Micropipette puller:
We utilize Sutter Instrument Company's model p87 micropipette puller to fabricate our needles.

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