TITLE: Isolating non-axenic monoclonal Symbiodinium cultures from Aiptasia pallida
AUTHORS: Emily Aguirre

[DESCRIPTION]
To isolate non-axenic, monoclonal Symbiodinium from the anemone Aiptasia pallida (CC7). Although there is no guarantee that all bacterial associates will be conserved using this method, 16S rDNA sequencing (unpublished data) suggested that previously documented bacteria1 in the Symbiodiniaceae phycosphere (in a study not affiliated with our group, done on 5 clades of Symbiodiniaceae) were also prevalent in our monocultures even after three years of consistent subculturing with L1 (no Si) media.


1  Lawson, C. A., Raina, J. B., Kahlke, T., Seymour, J. R., & Suggett, D. J. (2018). Defining the core microbiome of the symbiotic dinoflagellate, Symbiodinium.Environmental Microbiology Reports,10 (1), 7-11.

NOTE: This protocol requires an inverted microscope. Please assure you have access to one for the entirety of the process.

[BEFORE_START]
This protocol will take a minimum of 12 weeks. Symbiodinium growth takes time but it is mostly a background process, and patience is key. This project requires minimal expenditure on the human side after the initial isolations. 

[GUIDELINES]
This protocol requires an inverted microscope. Please assure you have access to one for the entirety of this protocol.

[STEPS]
SECTION: INITIAL ISOLATION FROM ANEMONE
1. Rinse an anemone (preferably ~ 1cm + ) with 0.2 µm FSW at least 3x, prior to disruption with a homogenizer. Place the anemone in a 15 mL Eppendorf tube with ~ 5mL of 0.2 µm FSW and homogenize at "medium" speed for >30 seconds or until all visible anemone tissue has been completely disrupted. 
SECTION: INITIAL ISOLATION FROM ANEMONE
2. Remove any chunky, clear floating tissue. Transfer the slurry to three, 2 mL microcentrifuge tubes. Aliquot evenly and centrifuge at 8000 rpm for 2 minutes.  This will bring the algal cells to the bottom. 
SECTION: INITIAL ISOLATION FROM ANEMONE
3. Remove the supernatant and add 1 mL of 0.2 µm FSW to each tube. Centrifuge at 8000 rpm for 1 minute. Repeat 3x. 
SECTION: INITIAL ISOLATION FROM ANEMONE
4. Prepare a 6-well plate with 3mL of 1:1 L1 media (no Si added) and 0.2 µm FSW.  
SECTION: INITIAL ISOLATION FROM ANEMONE
5. Transfer the pellets of one tube to a well in the 6-well plate. Allow to incubate for up to 5 days (14:10, light: dark cycle, at 25-27ºC)  or use an inverted microscope to check whether some Symbiodinium have expressed flagella and returned to a motile state. 

*these Symbiodinium cells can range in size from 6-10 µm. The magnification you use will be limited to your microscope but please keep in mind their size so you can adjust your own settings.

  
 

SECTION: INITIAL ISOLATION FROM ANEMONE
6. While waiting for Step 5, prepare 1% agar according to the manufacturer's instructions, but modify with 0.2 µm FSW, instead of DI water. After the agar has cooled down and it is at a temperature of ~60-65ºC, add the components for L1 media, including the F/2 vitamins, no Si. Deposit plates in incubator at 27ºC for 24 hours. Store at 4ºC in a plastic sleeve. 
SECTION: INITIAL ISOLATION FROM ANEMONE
7. After ~ 5 days, check for motile Symbiodinium in the 6-well plates using an inverted microscope, and if any are spotted swimming around, add a couple drops (or as needed) of Protoslo (or methyl cellulose) a protozoa slowing agent, to "catch them". 
 
Symbiodinium, freed from the cnidarian symbiosome, ~ five days after separation from the host and after incubation (step 6). An accumulation of debris from anemone tissue degradation has occurred, which is why the many rinsing steps are crucial in this protocol. Video by Emily Aguirre.
SECTION: INITIAL ISOLATION FROM ANEMONE
8. Pipet motile Symbiodinium and coccoid Symbiodinium into a sterile, small petri dish containing 1 mL of 0.2 µm FSW. 
SECTION: FIRST TRANSFER/INCUBATION STEP (AGAR)
10. Using an inverted microscope, transfer 1 or up to 10 individual cells onto a microscope slide with concave wells, each containing 20-50 µL of 0.2 µm FSW. Transfer the cells, GENTLY, through 3 wells to further rinse them. When the last well is reached, pipet the liquid (including the cell/s) and aliquot onto a warmed L1 agar plate (from step 9). 


*This is a tedious process and will take a couple hours (depending on your pipetting skills) to go through all cells. But please try to be gentle and go as fast as you safely can, since the harsh microscope light (even in dimmer settings) can exacerbate their photosynthetic systems and stress them out, if exposed too long.  
SECTION: INITIAL ISOLATION FROM ANEMONE
9. Meanwhile, place L1 agar plates in a 27ºC incubator for 30 minutes or until they are at room temperature.
SECTION: FIRST TRANSFER/INCUBATION STEP (AGAR)
11. Add as many "cleaned" cells per agar plate (you can mix and match, motile vs coccoid shape) as you'd like. The total aliquot should be no more than 200 µL.  I recommend starting off with 20-60 cells per plate. 

The more cells you rinse and transfer to an agar plate, the faster your colonies will visibly show up on the plate. 

However, transferring too many cells at once may introduce other eukaryotic microbes if not adequately rinsed. These can outcompete the dinoflagellates in the media immediately, leading to no Symbiodinium growth. Rinse plenty. 
SECTION: FIRST TRANSFER/INCUBATION STEP (AGAR)
13. Incubate as before (step 5), but allow cells to grow for 2 weeks to 6 weeks after plating, or until visible, brown clusters appear on the surface of the plates. 
SECTION: SECOND TRANSFER/INCUBATION STEP (AGAR)
14. Pick a colony "cluster" out, using a sterile filter-tip/ pipettor and transfer to a sterile, small petri dish containing 1mL of 0.2 µm FSW. Pipet the cluster up and down, gently, to disperse the cells and make it easier to visualize them. 
SECTION: SECOND TRANSFER/INCUBATION STEP (AGAR)
15. Visibly verify (by ASEPTICALLY taking a 10 µL aliquot from Step 14) it is a Symbiodinium cluster (and not brown diatoms) on an inverted or compound microscope.
SECTION: FOURTH TRANSFER/INCUBATION STEP (LIQUID, L1 MEDIA)
19. Incubate for ~2-6 weeks or until there is sufficient visible, brown biomass. 
SECTION: SECOND TRANSFER/INCUBATION STEP (AGAR)
16. Take 200 µL of the aliquot from step 14 and plate/incubate (as seen in steps 12 and 13) for 2-4 weeks.


SECTION: FIRST TRANSFER/INCUBATION STEP (AGAR)
12. Add 3-4 plating beads, shake until all the liquid has been adsorbed on the surface. Discard beads and wrap in parafilm. 
SECTION: THIRD TRANSFER/INCUBATION STEP (AGAR)
17. The resulting plate should clearly be all brown clusters, evenly distributed throughout the surface. From this plate, pick a colony and streak a new L1 agar plate with this colony. Incubate for 2-4 weeks, as before. 
SECTION: FOURTH TRANSFER/INCUBATION STEP (LIQUID, L1 MEDIA)
18. After incubation, inspect the plate, aseptically pick out the biggest colony and add it to a sterile, 50 mL cell culture flask containing 25 mL of L1 media (no Si). Using a sterile 1000 µL pipet tip, pipet the liquid up and down to disperse the Symbiodinium and associated bacterial cells. Allow oxygen to pass through by loosely capping the cultures. A culture flask with a 0.22 µm filter cap can be tightly capped.
SECTION: FOURTH TRANSFER/INCUBATION STEP (LIQUID, L1 MEDIA)
20. I usually subculture once every 1.5 months when the flasks display a distinguishable brown film.  

To subculture: Disturb the cultures gently with a 1000 µL pipet, and then pipet 500 µL of culture into a new cell culture flask containing 25 mL of L1 media (no Si). Repeat as needed. 
SECTION: VERIFYING IDENTITY OF SYMBIODINIUM
21. To verify and identify Symbiodinium, you can use any DNA extraction protocol for dinoflagellates, but I used DNeasy Plant Mini Kit. Quantify your DNA concentrations (I used Qubit fluorometer).

The ITS2 gene was sequenced and sent in for Sanger sequencing. 
SECTION: VERIFYING IDENTITY OF SYMBIODINIUM
21.1. ITS2 Primers 
ITS2-F: 5'-GTG AAT TGC AGA ACT CCG TG-3'
ITS2-R: 5'-CCT CCG CTT ACT TAT ATG CTT-3'