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.