MBL Embryology: Zoology Labs

(Henry and Martindale, 2010)

Tool Making and Handling of Marine Invertebrate Embryos and Larvae

Successful handling and culture of marine invertebrate embryos requires specific conditions and special tools. You will need to construct simple tools to facilitate manipulation of eggs and embryos of various marine invertebrate species. These tools will be used throughout the “Zoo” labs and should be prepared during this introductory lab. Be sure to have these tools ready before the first “Zoo” lab.

Topics to be covered:

1. 0.22µm Filtered Sea Water (TAs will maintain the supply of “FSW”) 2. “E-glassware”: NO detergents, heavy metals (poison to embryos!) 3. Culture conditions (sterility, water quality, temperature, etc.), Details concerning the development and culture of marine invertebrates found in local waters can be found at: http://zeus.mbl.edu/public/costello/list.php 4. Construction of mouth (transfer) pipettes with an air brake (“Braking Pipettes”) 5. Gelatin-coated “non-stick” culture dishes and transfer pipettes to raise and handle “sticky” embryos (TAs will supply the coated petri dishes) 6. Narcotizing and fixation of embryos and larvae 7. Mounting live and fixed specimens (Using Rain-X and modeling clay) 8. Removal of extra-embryonic investments (e.g., fertilization envelopes, chorions, etc.), Chemical and mechanical methods (e.g., forceps, Nitex-syringe demembranators (this topic will be covered further in future labs) 9. Forceps sharpening using a diamond sharpener (Warning! Go lightly on the sharpening. Do not shorten the forceps!) 10. Preparation of glass and tungsten dissection needles using micro- gas burners 11. Microinjection of lineage tracers, etc., Use of the microinjection apparatus

If you wish to raise embryos to larval stages, you can do this by culturing the embryos in FSW (filtered sea water) in gelatin-coated dishes. It is critical to change the water every day to avoid bacterial growth in the dish. Antibiotics, including penicillin (100units/ml) and streptomycin (200µg/ml) may also be used, but may not be required for short-term cultures. Once “sticky” embryos begin to swim, they do not need to be cultured in gelatin-coated dishes.

Recipe for Gelatin coated dishes:

Gelatin is used to prevent embryos form sticking to transfer pipettes, operating needles, and culture dishes. Make a 5x stock (this can be stored at room temp. “RT” for several months):

0.25 gm (0.5%) Knox unflavored gelatin (dissolve before adding formalin!)

250 µl formalin (0.19 % formaldehyde)

50 ml dH20

For coating dishes, dilute to 1x working strength immediately before use with dH2O. Pour several mls of solution into 35 mm Petri dishes, swirl and pour off remaining solution into next dish to be coated. A thin coat will remain in the first dish. Allow dishes to dry completely by tipping upside down, and then rinse coated dishes 3x with dH2O. Transfer pipettes and operating needles can also be coated with this 1x working solution.

General instructions regarding microinjection:

You will learn how to microinject minute quantities of fluorescent lineage tracers, morpholinos, synthetic RNAs, etc., using pressurized nitrogen gas.

The tips of glass microinjection needles are melted, or “pulled” using a Sutter P98 microelectrode puller to specific shapes. Adjustments of the microelectrode puller (the platinum filament) require great care and time; therefore, the course instructors will generally prepare these needles for you, though we will be glad to show you how the machine is operated. Please see us before using the puller. The internal bore of the glass needles contains a fine internal glass filament that facilitates backfilling with various solutions.

Fine control of the microinjection needles is accomplished using Narishige “joy-stick” hydraulic micromanipulators and the progress of injection is monitored using epi- fluorescence dissection microscopes.

Most embryos must be immobilized during injection. This may be accomplished in different ways. One technique is to engrave narrow grooves in the bottom of plastic Petri dishes. Very precise groves can be made using the sharp edge of broken glass pasture pipettes. We will demonstrate this technique in lab.

Embryos may also be electro-statically attached to glass or plastic surfaces using poly-L- lysine or protamine sulfate, but they must be removed from these treated surfaces immediately following injection to insure proper development.

Other mechanical devices may also be used to hold the embryos. Many of these approaches are specific to the embryos using used and these techniques will be introduced in subsequent labs.

Cell lineage tracing, microinjection of DiI:

Microinjection needles will be pre-pulled and loaded with DiI/Oil. These are pulled using the Sutter P98 Puller and Program #22. Please do not change the settings for that program, if you should need to use it (see notes above).

The injection needles are essentially closed at the tips and need to be opened by gently tapping them against the bottom of the injection dish. This procedure will be demonstrated to you. Injections are performed in gelatin-coated dishes using demembranated embryos. A series of small scratches will be made in the bottom of these dishes using a broken, jagged-edged Pasteur pipette (“scratch dishes”), which will be used to help immobilize the embryos for injection.

Preparation of DiI

Stock solution: 100 mg/ml DiI in 100% EtOH

Store at RT, 4 oC or –20 oC away form strong light.

To use, dilute DiIC18 (3) 1:20 in Soybean Oil. Heating with sonication or vortexing may be required to completely dissolve the DiI. Store at 4 oC or –20 oC away from strong light.

Backfill microinjection needles well before use to allow DiI/oil to reach the tip, using a Hamilton syringe equipped with a long 26g needle. The viscosity of the oil slows backfilling. Thus, be sure to prepare these 24 hours in advance.

The DiI may come out of solution during storage. It can be re-dissolved by heating with sonication or vortexing before use. Usually the presence of a few small crystals in the injection needles is not a problem.

DiI fluorescence is often quenched after fixation. Observations are generally made while specimens are alive. However, one can preserve fluorescence by adding 50 mM EDTA (pH 8.0) to the fixative (0.1% to 4% formaldehyde in FSW) (Henry et al., 2001).

Blastomeres can also be labeled with fluorescent DiI directly through the vitelline envelope via iontophoresis (Henry et al., 2001, see also Sweet et al., 2004). Hoder and Ettensohn (1998) described this technique, and the construction of an inexpensive device, which is powered by a 9 volt battery. Quicker labeling is achieved by lowering the value of the current-limiting resistor to 100,000 or 10,000 ohms. Make up the DiI in EtOH at a concentration of 20 mg/ml. If the DiI EtOH solution is too concentrated you will not be able to label the cells.

1. Glass microelectrodes are pulled using a standard mechanical puller, such as the Sutter P97. The tips should not be too long and flexible (about 4-5mm). Microelectrodes are back-filled with diI-EtOH as described by Hoder and Ettenshon (1998) and Henry et al., 2001; 25mg/ml diI, cat. no. D-282, Molecular Probes Inc., Eugene, OR dissolved in 100% EtOH). These “needles” should effectively be “closed”. If the tip breaks, diI will crystallize and no labeling will take place. The needle is held using standard neurobiology microelectrode holders. The positive platinum electrode wire is threaded into the back end of the glass microelectrode so that it contacts the diI-EtOH solution. The negative reference electrode (platinum wire) is immersed in filtered sea water (FSW) in a small plastic petri dish (but not in direct contact with the embryos). Using a micromanipulator, firm contact is established between the tip of the needle and the surface of the embryo that overlies the desired blastomere. Pressure is applied to the embryo such that the surface is visibly indented, otherwise diI will not reach the cell. It is not necessary to actually penetrate the envelope. To prevent the embryos from rolling away, a small groove is engraved in the bottom of the plastic petri dish using the sharp edge of a broken Pasture pipette.

2. Current is applied until the desired amount of fluorescence labeling is achieved (15 sec. to a few min.). Labeling is monitored with an epifluorescence dissecting microscope. Labeling will not take place, if the needle becomes coated with extra-cellular matrix or cell debris. One must then clean the needle by wiping the tip gently against the bottom of the operating dish, or replace it with a fresh one.

Cell lineage tracing, microinjection of Dextran lineage tracers:

Alternatively, one can microinject Dextran lineage tracers (e.g., 10,000MW Rhodamine Green, or Fluor Ruby, Molecular Probes, Inc.). Use a working stock of 50mg/ml in 0.5M KCL. Such aqueous dyes can be co-injected with RNA/DNA for molecular studies.

Demembranation:

Different techniques are used to remove extracellular investments and you will be introduced to these in subsequent labs. For instance, some annelid embryos can be demembranated using a Ca++ chelator. Place the unfertilized eggs in a fresh mixture of 1:1 0.25M NaCitrate and 1.0M Sucrose. Place approximately 1-2ml in a polypropylene tube and place a few hundred eggs in a tiny volume of FSW (approx 10µl) in the solution. After 45 sec., pour the eggs into a coated dish containing FSW, as the denuded embryos will be very sticky. Wash the eggs two times in FSW and fertilize.

Vitelline envelopes can also be removed by mechanical means by hand, using forceps, or with a Nitex screen with openings slightly smaller than that of the egg’s surrounding envelope. The Nitex is fused to the end of small plastic syringe using heat. This is accomplished by first cutting off the end of the syringe and melting the plastic in a microburner. The softened plastic is quickly fused to the Nitex screen. Excess screen is trimmed away after the plastic has solidified (cooled). Back-load the embryos or eggs into the syringe and using the plunger, slowly push the eggs through the Nitex screen into a gelatin coated dish containing FSW. Demembranated eggs and embryos are very sticky so use both coated dishes and coated transfer pipettes. Demembranation can also be performed prior to fertilization in some species. Success is greater if the oocytes are nicely rounded, otherwise wait until after fertilization when the eggs usually assume a nice rounded shape.

Blastomere Isolation:

One can examine the developmental properties of isolated blastomeres or partial embryos by removing specific cells.

A). One method involves the use of fine glass or tungsten needles to separate the blastomeres. Due to the presence of the vitelline envelope in many species, these must be removed. Various chemical and mechanical techniques can be used, which are specific for different embryos (see examples below).

1. Glass needles are pulled by hand using a gas micro-burner (Hamburger, 1960). Alternatively, sharpened tungsten needles may be used. Two techniques are recommended for sharpening tungsten wire (diameter of 0.004 inches). One method involves electrolytic sharpening in a saturated solution of either sodium hydroxide, or potassium or sodium nitrite. This technique is well described in the papers by Hubel (1957) and Conrad et al. (1993). The second technique involves burning the tungsten using a small microburner (Dossell, 1958; Hamburger, 1960). These needles can be mounted in small needle holders or melted glass rods.

2. To ablate specific blastomeres, firm pressure is applied to the desired cell using the tip of the micro-needle. In some cases this can be done through the vitelline envelope. The needle will penetrate the cell and lyse the blastomere. The presence of residual cell debris may or may not interfere with the development of the remaining live cells (Henry et al., 2001). To avoid killing sister cells, one must wait until each division is complete. The operated embryos should be transferred to a clean dish containing fresh FSW.

B. An alternative method can be used to separate embryonic cells, which involves the use of Calcium-low or calcium-free sea water. This treatment can also remove certain vitelline envelops (e.g., some annelids).

1. Approximately 45 mins. prior to first cleavage, (or the cleavage stage of interest) the fertilized eggs are placed in calcium-low or calcium-free sea water. To remove any residual calcium, the eggs are washed five times in calcium-free sea water. One ccan use Rulon’s (1941) modification of Herbst’s calcium-free sea water (see also Cavanaugh, 1956). After completion of the desired cleavage stage and successful cell separation, the embryos are washed three times in FSW. In some species this can be done in the presence of loose-fitting vitelline envelopes, though some blastomeres may not be entirely separated or they may refuse at later stages of development.

C). Early and late cleavage stage blastomeres/embryos may be separated by a third method using fine nylon ligatures. Certain brands dental floss provide an excellent source for very fine nylon ligatures, which are approximately 20µm in diameter. (POH, “Personal Oral Hygene,” un-waxed dental floss, Oral Health Products, Inc, P.O. Box 470623, Tulsa, OK 74147). Filaments can also be teased from nylon stockings.

The embryos are placed in a small petri dish containing FSW. It is helpful to pre-cut short pieces (approx. 2mm) and pre-tie single overhand knots in preparation for applying the ligatures. The loop is placed around the embryo and tightened using watchmaker’s forceps. The filaments should be kept short, and one must be careful not to allow the nylon thread to contact the air-water interface, as the surface tension may damage the embryo. It is sometimes helpful to use a glass or tungsten needle to pre-crease the embryo to aid in the proper placement of the ligature. The ligature is left in place until the desired stage of development is reached.

Narcotizing larvae

To “anesthetize” larvae and prevent muscular contraction, they can be placed into a 1:1 mixture of 6.5% MgCl2 (made up in dH2O) and FSW. The Mg++ competes with Ca++ at neuromuscular junctions and prevents muscle contraction. This rarely helps reduce motion created by cilia. Many other methods are also available.

BODIPY FL Phallacidin stain for filamentous actin (muscles)

Stock = 200 units/ml BODIPY FL Phallacidin in 100% MeOH (Molecular Probes, Inc.)

Store at –20oC.

1. Fix larvae in 2-4% formaldehyde in filtered sea water for 10-15’ at RT. DO NOT allow larvae to be exposed to EtOH or MeOH!

2. Wash fixed larvae in FSW with 0.2% Triton X-100 2 to 3 times.

3. * Make up working solution of BIODIPY FL phallacidin as a 1:500 dilution in FSW with 0.2 % Triton X 100.

4. Transfer fixed larvae to BODIPY FL phallacidin working solution and stain for 20’- 2hr at RT (depending upon species) or overnight at 4oC.

5. Rinse 3 times in FSW with 0.2% Triton X-100.

6. Mount stained larvae in 70% glycerol in PBS, and view fluorescence staining in the larvae.

*Alternatively, one can place the required volume of the BIODIPY FL Phallacidin stock as a drop in the bottom of a 3 spot depression dish. Once the methanol has evaporated, add the required volume of SW with 0.2% Triton X-100 to the dish and mix. Finally, add the fixed embryos to the staining solution.

Hoescht 33342 nuclear staining

Stock = 10 mg/ml Hoescht 33342 in dH2O

Store at 4oC.

1. Fix embryos and larvae in 3.7% formaldehyde in FSW for 10’- 1 hr at RT.

2. Wash larvae in FSW or PBS 2-3 times.

3. * Make up working solution of Hoescht 33342 as a 1:10,000 dilution in FSW, dH2O, or PBS.

4. Transfer fixed larvae to Hoescht 33342 working solution and stain for 1 hr at RT or 12- 16 hr at 4oC.

5. Rinse 3 times in FSW, PBS or dH20.

6. Mount stained larvae in 70% glycerin in FSW or PBS and view fluorescence staining in the larvae.

*Alternatively, Hoescht 33342 can be added directly to the fixative or even to the glycerol mounting media.

*Hoescht 33342 is a vital stain and can be added to FSW to stain the nuclei of living embryos and larvae, however, viewing will damage DNA and ultimately lead to abnormal development.

CsCl2 Induction of metamorphosis of cnidarian planula larvae

A number of planula larvae can be induced to undergo metamorphosis by treatment with

CsCl2. Make a stock solution of 0.4 M CsCl2 in DIH20. Dilute to 40 mM working strength in FSW. Leave the larvae in CsCl2 continuously and after a few hours the larvae should settle to the bottom and undergo metamorphosis. Wash out CsCl2 after 3-6 hr (they will not develop normally if left in CsCl2 for extended periods of time. This treatment works especially well for Hydractinia.

References

Cavanaugh, G. M. (1956). “Formulae and Methods VI of the Marine Biological Laboratory Chemical Room.” 84p. Marine Biological Laboratory, Woods Hole, MA.

Conrad, G. W., Bee, J. A., Roche, S. M. and Teillet, M. A. (1993). Fabrication of microscaples by electrolysis of tungsten wire in a meniscus. J. Neurosci. Methods 50: 123-127.

Dossel, W. E. (1958). Preparation of tungsten micro-needles for use in embryology research. Lab. Invest. 7: 171-173.

Hamburger, V. (1960). “A Manual of Experimental Embryology” revised edition, Univ. of Chicago Press.. 221p.

Henry, J. Q., Tagawa, K. and Martindale, M. Q. (2001). Deuterostome evolution: Early development in the enteropneust hemichordate Ptychodera flava. Evolution & Development. 3: 375-390.

Hoder, P. G. and Ettensohn, C. A. (1998). The dynamics and regulation of mesenchymal cell fusion in the sea urchin embryo. Dev. Biol. 199: 111-163.

Hubel, D. H. (1957). Tungsten microelectrode for recording from single units. Science 125: 549-550.

Rulon, O. (1941). Modification of development in the sand dollar by NaCNS and Ca- free sea water. Phys. Zool. 14: 305-315.

Sweet, H., and Amemiya, S., with Ransick, A., Minokawa, T., McClay, D., Wikramanayake, A., Kuraishi, R., Kiyomoto, B. M., Nishida, H. and Henry, J. J., as contributing authors. (2004). “Methods in Cell Biology: Experimental Analysis of the Development of Sea Urchins and Other Non-Vertebrate Deuterostomes.” Chapter 11, “Blastomere Isolation and Transplantation,” Academic Press, San Diego, CA.

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