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2010 Choanoflagellates, Sponges, and Animal Origins Nicole King and Stephen Fairclough

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2010 Choanoflagellates, Sponges, and Animal Origins Nicole King and Stephen Fairclough MBL Embryology -- 2010 Choanoflagellates, sponges, and animal origins Nicole King and Stephen Fairclough 1. Compare the cell morphology of choanoflagellates and sponge choanocytes • provides insights into the ancestry of animal cell biology • almost guaranteed to work (...famous last words) • provides practice with confocal microscopy OR Investigate tissue structure and embryogenesis in a sponge • provides insights into the early evolution of animal development • experiment somewhat prone to failure (e.g. one or more stains may not work) • provides practice with confocal microscopy AND 2. Investigate cell differentiation in a colony-forming choanoflagellate • fast, easy experiment -- perform during lengthy incubations 3. Observe bacterial prey capture by choanoflagellates in real time • fast, easy experiment -- perform during lengthy incubations The main players: Choanoflagellates • Monosiga brevicollis: unicellular, sequenced genome • Salpingoeca rosetta: unicellular and colonial, sequenced genome Sponges • Oscarella: thin, harbors embryos, lacks spicules, low autofluorescence • ...we'll also try to provide a calcareous sponge with a high concentration of choanocytes Protocol 1: Staining choanoflagellates and sponge choanocytes for actin, tubulin, and DNA Overview: 0. Prepare poly-L-lysine treated coverslips (we have done this for you). 1. Attach cells to coverslip. 2. Fix cells on coverslip. 3. Incubate with primary antibody, wash. 4. Incubate with secondary antibody, wash. 5. Stain with phalloidin. 6. Mount on slide with ProlongGold/DAPI. 7. Image at microscope. Reagents/Materials − 2ml choanoflagellates/ coverslip – concentration of 106-107 cells/ml − sponge tissue (to be macerated) − P20 and P200 pipettes and pipette tips − forceps − 3 Poly-L-lysine coated coverslips − 3 Glass slides − 100% EtOH in a spray/squirt bottle − Petri dish − Whatmann paper cut to fit in petri dish − Parafilm cut to fit in Petri dish − 1.5 mL eppies − 175 ul 37% formaldehyde − 30 ml 1xPBS pH 7.4 − 1500 ul Block solution (1xPBS/1%BSA/0.3%TritonX-100) − 300 µl of diluted 1˚ antibody dilution (in block solution) − 300 µl of 2˚ antibody dilution (1:500 in block solution) − 300 µl of Phalloidin solution (0.6 U Phalloidin/100µl 1xPBS) − 30ul ProlongGold + DAPI at room temperature Notes: When working with cells mounted on the coverslip , don’t let the coverslip dry at any time! When washing cells mounted on the coverslip, aspirate gently (with your pipet, not with vacuum) from the same point each time to minimize shear. You would normally want to include a secondary-only control (i.e., don’t use a primary antibody, but do everything else the same) for each sample. We have already done this for both choanoflagellate species that you are working with today and there is minimal autofluorescence, so you can skip it. For the sponge sample, be sure to include a negative control Preparing the Coverslip (coverslips have been coated in Poly-L-lysine in advance) 1. Pick up coverslip with forceps 2. Spray/squirt both sides with 100% EtOH so the coverslip is completely coated 3. Prop coverslip at an angle on a pipette tip and allow both sides to air dry completely 4. While coverslips are drying, prepare a humidified chamber by cutting a circle of Whatman paper that will line the bottom of a petri dish 5. Wet the Whatman and place it in the dish (*alternately, you can wet two Kimwipes and bunch them in the corner of the Petri dish) 6. Lay a square of parafilm on top of the Whatman that is large enough to fit all of your coverslips with at least an inch of space on all sides between each one 7. Once coverslip(s) have dried, place onto parafilm with an inch of space between each one 8. Place the lid on the dish during all incubation periods Cell Fixation and Attachment to Coverslip 9. For Monosiga: • Grow cells to a density of approximately 106-107 cells/ mL [We have done this for you.] • Spin down two tubes each containing 1mL of cells @ 500xg for 10 min; remove supernatant and re-suspend in 100µl of 4% formaldehyde (prepared by adding 125µl 37% formaldehyde stock to 1mL of 1xPBS). NOTE: cell pellet may not be visible so note the orientation at which you spin the tube. Also, be VERY, VERY gentle when re-suspending cells -- colonies will dissociate and collars/flagella will be lost. • Apply 200µl of choanoflagellates to a poly-L-lysine coated coverslip by gently pipetting sample onto the center of the coverslip so that it forms a round pool. Incubate for 15 minutes. For sponge choanocytes: • Macerate a match-head sized piece of sponge tissue in ~400µl seawater (you can do so with foreceps; gently pull the sponge apart). • Add 4% formaldehyde (prepared by adding 50µl 37% formaldehyde stock to 400 µl sea water).Transfer 200µl cell slurry onto each of two coverslips; incubate 5 minutes. 10. Very gently aspirate liquid off of each coverslip by placing pipette tip at one corner of the coverslip and letting it touch the edge of the pool of liquid. Be very gentle to avoid disrupting the microvillar collar, but work quickly enough that the coverslip doesn’t dry. 11. Wash 4 times by gently pipetting 100µl 1xPBS onto the coverslip and subsequently aspirate off using the same corner of the coverslip. Remember, don’t let the coverslip dry at any time! Stain Cells and Mount on Slide 12. ** Apply 100 µl block solution (make fresh each day) to each coverslip. Incubate 30 minutes. 13 **Aspirate block solution and apply 100 µl of diluted 1˚ antibody to choanoflagellates and one sponge choanocyte sample. For the second sponge choanocyte sample, apply 100 ul block. Incubate 30 minutes at room temperature. 14. Aspirate remaining fluid, wash 4 times with block solution. 15. **Apply 100 µl of diluted α-mouse 2˚ antibody and incubate 30 minutes, room temperature, in a DARK DRAWER 16. Aspirate remaining fluid, wash 4 times with block solution 17. Apply 100 µl of Phalloidin solution and incubate for 15 minutes in a DARK DRAWER 18. Aspirate remaining fluid, wash 2 times with 1xPBS 19. Let coverslip dry almost completely in a DARK DRAWER (about 5 or 10 minutes) 20. Add 10µl of room temperature ProlongGold + DAPI to a clean, dry slide and place coverslip face down on slide – be sure to place coverslip slowly and gently. Don’t move the coverslip after placing it on the slide to avoid damaging cells. 21. Allow time for ProlongGold to equilibrate (about 30 minutes) **Can stop for a while at these points; for antibodies, can leave at 4˚ overnight Protocol 2: Staining sponge tissue for whole mount Overview 1. Collect sponge tissue from sea table in laboratory 2. Fix tissue in paraformaldehyde (we have done this for you) 3. Incubate with primary antibody, wash. 4. Incubate with secondary antibody, wash. 5. Stain with phalloidin. 6. Mount on slide with ProlongGold/DAPI. 7. Image at microscope. Reagents/Materials − P20 and P200 pipettes and pipette tips − 2 Poly-L-lysine coated coverslips − 2 Glass slides − 100% EtOH in a spray/squirt bottle − Petri dish − Whatmann paper cut to fit in petri dish − Parafilm cut to fit in Petri dish − sponge tissue − 1.5 mL eppies − 4 mls 4% paraformaldehyde − 10ml 1xPBS pH 7.4 − 500ul Block solution (1xPBS/1%BSA/0.3%TritonX-100) − 400 µl of diluted 1˚ antibody dilution (in block solution) − 400 µl of 2˚ antibody dilution (1:500 in block solution) − 400 µl of Phalloidin solution (0.6 U Phalloidin/100µl 1xPBS) − Glycerol − Glycerol + DAPI Notes: Living sponges will be held in the sea tables next to the laboratory. When you are ready I will orient you as to which species are available. Try not to handle the sponges - it is bad for them and you. Also, do not expose them to air -- this generally kills them. All incubation and wash steps should take place on a rocker or nutator. A rocker is preferred because it is gentler. Most sponges exhibit autofluorescence that can be highly variable between individuals, fixation conditions, etc. Therefore, you will want to include a secondary-only control. Prepare samples 1. in seawater, excise a piece of tissue about the size of a match-head. Be careful not to compress or damage the tissue unnecessarily. Tissue can be neatly removed from Oscarella by pipetting with a transfer pipette (usually - some individuals are tougher and require a razor or scissors). Other sponges will be a bit trickier to handle. Fix Tissue 2. balancing the tissue on the top of your foreceps (i.e., don’t squeeze it), transfer up to 2 pieces into 1ml of 4% paraformaldehyde (note: this is not the same fixative that you used in Protocol 1). Fix two samples, one for staining with primary and secondary, and a second sample as a negative control (stain only with secondary antibody). 3. Incubate at room temp for 1-2 hours (in a perfect world fixation would occur overnight at 4C; I will announce if I have tissue already fixed in this manner, cooking show style) 4. rinse the tissue once in 1mL 1xPBS 5. wash the tissue 3 x 5minutes in 1ml 1xPBS Stain and Mount Tissue on Slide 6. **Incubate in 1mL block solution for at least 30 minutes 7. **transfer one piece of tissue to a 0.5mL tube containing 200µl of primary antibody diluted in block solution. The second piece of tissue should be incubated in 200µl block solution without primary antibody. Note: if the tissue seems a bit large at this point, you can trim it further so that it takes up less than 20% of the volume of the primary antibody solution; incubate for at least 1 hour 8. use forceps or a large bore transfer pipette to carefully transfer each tissue sample to a new 1.5mL tube 9.
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