MBL Embryology -- 2010 , , and origins Nicole King and Stephen Fairclough

1. Compare the cell morphology of choanoflagellates and 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 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 • 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. wash each sample 4 x 5 minutes in 1mL block solution 10. ** use forceps or a large bore transfer pipette to carefully return each sample to a 0.5mL tube containing 200µl of secondary antibody solution; incubate for at least 1 hour (in the dark; e.g., in foil) 11. use forceps or a large bore transfer pipette to carefully transfer each sample to a new 1.5mL tube 12. wash the tissue samples 4 x 5 minutes in 1mL 1xPBS 13. use forceps to carefully transfer tissue samples to new 0.5mL tubes each containing 200µl phalloidin solution 14. incubate for 30 minutes (wrapped in foil) 15. use forceps or a large bore transfer pipette to transfer each tissue sample to a new 1.5mL tube 16. wash each sample 2 x 5 minutes in 1mL 1xPBS (keep wrapped in foil) 17. replace 1xPBS with 75% glycerol. incubate 5 minutes 18. transfer each sample to 1mL 75% glycerol + DAPI (provided) 19. when ready to image, transfer into a depression slide containing 75% glycerol + DAPI solution OR onto a regular slide with lab tape, modeling tape, or coverslip shards as a spacer. The idea is to have the surface of the sponge in direct contact with the coverslip, but not so compressed that the tissue/embryo morphology is disrupted.

**You can stop for a while at these points; for antibodies, can leave at 4˚ overnight.

Protocol 3: Live staining with the lectin WGA to detect glycosyl groups

Overview 1. Label live cells with Lectin 2. Fix Cells 3. Image at microscope

Reagents/Materials − P20 and P200 pipettes and pipette tips − 5 ug/mL Alexafluor488-WGA (kept on ice in the dark) − 5% glut + 100mM HEPES pH 8.0 − Coverslip − Slide

Notes Lectins are highly-specific sugar binding proteins, usually derived from plants and bacteria. Wheat germ agglutinin (WGA) binds specifically to sialic acid and N-acetylglucosaminyl residues.

Cells that are labeled live should be imaged within 5 - 10 min. You should ensure that a microscope is available before staining the cells.

Live cell labeling Labeling live cells is the best way to ensure that you can distinguish cell types 1. Pipette 50µL of 5 ug/mL Alexafluor488-WGA into the bottom of a 1.5ml eppendorf tube, then add 1ml of cell culture to the tube using a wide-bore pipette and gently triturate until well mixed. (final WGA concentration is 0.25µg/ml (5*50/1000)) 2. Gently pipette labeled cells onto a slide and cover with a coverslip and view in GFP channel. You have ~5-10 minutes to view the cells before they start to accumulate significant amounts of WGA in the vacuole.

Fixed cell labeling You can also fix the labeled cells. Fix by mixing 1:1 with 5% glut + 100mM HEPES pH 8.0 in ASW, which is the fix for the EM and seems to preserve the morphology well.

Thought experiment: You will observe that a subset of slow swimming cells stains with the WGA. Can you design an experiment to test whether the slow swimming cells that stain are the cells from which colonies will develop? Protocol 4: Monitoring prey capture by live choanoflagellates

Overview 1. Settle choanoflagellates in glass-bottom dish with galena crystals (we have done this for you) 2. Mount dish on microscope for imaging 2. Add small amount of fluorescent bacteria and image instantly

Reagents/Materials − P20 and P200 pipettes and pipette tips − galena crystals − glass-bottomed dishes − GFP-expressing Enterobacter aerogenes

0. We will culture choanoflagellates in the presence of galena crystals to encourage them to orient and attach parallel to the plane of the microscope stage.

1. Place culture dish on microscope stage and focus on cells that have attached to the vertical surfaces of the galena crystals (so that you are imaging them from the side)

2. Pick a colony of GFP-expressing Enterobacter aerogenes into 100 ul of choanoflagellate cereal grass media and mix.

3. Add a small volume (1-2 ul) of bacteria to choanoflagellate culture on microscope. Watch them feast! It happens fast, so don't blink.