TITLE: Quantify photophysiology of endosymbiotic dinoflagellates (Symbiodiniaceae) using the Guava Flow Cytometer
AUTHORS: Colin J Anthony, Colin Lock, Bastian Bentlage

[DESCRIPTION]
This protocol quantifies light-harvesting complex (LHC) and antioxidant pigments, while simultaneously determining cell density of endosymbiotic dinoflagellates using the Guava Flow Cytometer. Red fluorescence excited by a blue laser represents peridinin, while the red fluorescence excited by the red laser represents both chlorophyll a and chlorophyll c2 , providing insight into Symbiodiniaceae LHC prioritization. Additionally, green fluorescence off of the blue excitation laser target antioxidant-associated pigments (diadinoxanthin, diatoxanthin, beta-carotene, and flavin-based fluorescent proteins). As flow cytometry continues to gain traction as a robust cell counting methodology, the integration and expansion of this protocol into coral or micro-algal workflows will serve as a valuable tool to quantify the physiology of single photosymbiotic cells in-hospite.


[BEFORE_START]
Ensure filtered seawater (FSW) is fresh

Heat SDS solution with benchtop shaking incubator to resuspend salt   
50 mL SDS solution = 7 mL FSW + 43 mL DI + 0.04 g SDS 

Clean the flow cytometer before and after each use 
GuavaSoft 4.0 uses Guava Clean 3.4, which walks you through cleaning process
Two pre-cleans is helpful and if the machine sat over the weekend, cleaning the capillary is also necessary
Throw out waste bottle and make sure other bottle has at least ⅔ ICS

easyCheck may also be necessary if machine has sat for more than a week
reagents and instructions come with easyCheck kit

Clean air brush and make sure needle is still present*

Chill centrifuge 5000 rcf, 0 °C 

Fill cooler with ice

Locate samples and label all tubes (50 mL falcon tubes* and 1.5 mL screw top tubes)

[GUIDELINES]
We focus on the characterization of Symbiodiniaceae associated with reef-building corals (Scleractinia); however, we have has success using this for upside-down jellyfish (Cassiopea). Therefore, with slight modifications this protocol may be used for flash-frozen and live cells for both calcifying and gelatinous cnidarians.

Each instrument, environment, and organism is different, so it is important to optimize this protocol locally.

Samples are prone to degradation, so it is important to work efficiently and consistently.

It is vital the same equipment is used in the same environment each time to avoid batch effect.

If using in a large multi-factorial project, mixed batches are recommended to avoid batch effect.

This protocol pairs well with cell counting methods, ITS2 metabarcoding, photopigment spectrophotometry, and coral morphometric methods, so we would suggest integrating this into more complex data structures as necessary.

Krediet et al. (2015) was pivotal in providing a framework for developing this protocol.
 


[STEPS]
SECTION: Sample Preservation
1. Sample tissue from cnidarian
If calcifying coral, we suggest 3 pieces at around 2 cm3 sampled with wire cutters or hammer and chisel 
If gelatinous cnidarian, 0.05 - 0.1 g of tissue sampled with sterile scissors works well


 


SECTION: Sample Preservation
2. Store tissue in bag or tube
For calcifiers, Whirl-Pak sample bags work well
For gelatinous cnidarians, 1.5 mL screw top tubes are more effective (this allows for immediate processing when ready)
SECTION: Sample Preparation
4. Before Start:
Ensure filtered seawater (FSW) is fresh (~1 L of filtered seawater is sufficient)

Heat SDS solution with benchtop shaking incubator to resuspend salt   
50 mL SDS solution = 7 mL FSW + 43 mL DI + 0.04 g SDS 
Make sure solution cools to ambient temperature before use

Clean the flow cytometer before and after each use 
GuavaSoft 4.0 uses Guava Clean 3.4, which walks you through cleaning process
Two pre-cleans is helpful and if the machine sat over the weekend, cleaning the capillary is also necessary
Throw out waste in waste bottle and make sure other bottle has at least ⅔ ICS

easyCheck may also be necessary if machine has sat for more than a week
reagents and instructions come with easyCheck kit

Clean air brush and make sure needle is still present*

Chill centrifuge 5000 rcf, 0 °C 

Fill cooler with ice

Locate samples and label all tubes (50 mL falcon tubes* and 1.5 mL screw top tubes)
For 12 samples, you will need 12 falcon tubes, 12 1.5 mL screw top tubes, and 1 microwell plate
Label one falcon tube and one 1.5 mL tube (If desired, add more 1.5 mL tubes as added technical replicates) for each sample being processed
SECTION: Sample Preparation
5. Prepare tissue slurry (calcifying cnidarians only)
Before start:
Make sure spray gun compressor is turned on
Have somewhere to store skeletal fragments after air-brushing (e.g. small weigh boats)
Be prepared to work efficiently as samples degrade quickly once removed from the freezer

SECTION: Sample Preservation
3. Flash-freeze tissue in Cryogenic storage Dewar flask with LN2 -320 °C
Store samples in ultra low freezer -80 °C  for extended preservation


SECTION: Sample Preparation
5.1. Locate samples in ultra-low freezer and remove up to 3 samples at a time
SECTION: Sample Preparation
5.2. Cut a ~1-2 cm piece from coral fragment (this will vary slightly depending on species)
SECTION: Sample Preparation
5.3. Use forceps to hold coral fragment just inside the mouth of a 50 mL falcon tube, then use an airbrush loaded with filtered seawater (FSW) to remove all coral tissue from the skeleton making sure to capture the tissue in the falcon tube.
Depending on the species, this can take 5-20 mL of FSW
Store falcon tube on ice in dim ambient lighting
Place the remaining skeleton on a weigh boat (Keep track of it. You will need it to normalize cell densities at the end)
SECTION: Sample Preparation
5.4. Repeat steps 5.1-5.3  with all samples
Move quickly and take no more than 1 hour for all samples combined
SECTION: Sample Preparation
5.5. Once all samples have been airbrushed, vortex and needle shear each tissue slurry until homogenized
There should be no mucus clumps or visible chunks; however, it is normal for small skeletal fragments to settle at the bottom
SECTION: Sample Preparation
5.6. Once a slurry has been homogenized, transfer 1 mL to a 1.5 mL screw top tube.
SECTION: Sample Preparation
6. Wash tissue slurries to be loaded in flow cytometer
If not using a calcifying cnidarian, add 1 mL of FSW to gelatinous tissue sample that was stored in a 1.5 mL screw-top tube

SECTION: Sample Preparation
6.1. Bead beat 1.5 mL tubes 4 s 

SECTION: Sample Preparation
6.2. Centrifugate samples 5000 rcf, 4 min, 0 °C 
SECTION: Sample Preparation
6.3. Remove 1 mL supernatant using 1000 mL pipette
Do not pour supernatant out. Pellets are often loose and liquid does not empty completely
SECTION: Sample Preparation
6.4. Resuspend pellets via repeated pipetting in 1 mL of FSW

SECTION: Sample Preparation
6.5. Bead beat tubes again 4 s 

SECTION: Sample Preparation
6.6. Centrifugate samples 5000 rcf, 3 min, 0 °C 

SECTION: Sample Preparation
6.7. Remove 1 mL  supernatant using pipette
Do not pour supernatant out. Pellets are often loose and liquid does not empty completely

SECTION: Sample Preparation
6.8. Resuspend pellets via repeated pipetting in 1 mL  of FSW and set samples to the side

SECTION: Load the Cytometer
7. Prepare 96-microwell plate for cytometry
Before start , check for supplies:
50 mL sufficient saltwater-freshwater solution (SFS) 
        25 mL FSW + 25 mL DI H2O = 50 ml SFS
200 uL pipette tips
200 uL pipette
20 uL pipette
Syringe with needle for needle sheering
Vortexer
SECTION: Load the Cytometer
7.1. Load wells of microwell plate with 180 µL SFS.
We describe this protocol with a 10x sample dilution (9 units SFS to 1 unit Sample). This volume of SFS will change as you optimize your dilutions
We also do not recommend loading more than half a plate as fluorescent properties change after an extended period within the machine


 


SECTION: Load the Cytometer
7.2. Bead beat  4 s a washed tissue sample then immediately load 20 µL of tissue homogenate into two wells preloaded with SFS before particulate settles
If there are visible clumps left over from the symbiont pellet, a combination of needle sheering and vortexing properly homogenize the sample; however, we do not recommend bead beating samples again as this risks lysing the algal cells

SECTION: Load the Cytometer
7.3. Repeat step 7.2 until all samples have been loaded. 
We typically process 12-24 samples at a time. We do not recommend processing more than 24 samples in one run.
SECTION: Load the Cytometer
8. Prepare worklist and set cytometry run settings for Guava Flow Cytometer
 

SECTION: Load the Cytometer
8.1. In guavaSoft v4.0, open InCyte v4.0
SECTION: Load the Cytometer
8.2. Click "Edit Worklist"
SECTION: Load the Cytometer
8.3. Select the wells with loaded samples and click "Acquire Samples".
SECTION: Load the Cytometer
8.4. Set each wells setting to acquire for 180s with a maximum gated cell count of 2500.


 

SECTION: Load the Cytometer
8.5. Also set each well to have 2 technical replicates and 7 seconds of high energy stirring.
SECTION: Load the Cytometer
8.6. Name each well
SECTION: Load the Cytometer
8.7. Click "Run Worklist"
SECTION: Load the Cytometer
9. Load in appropriate method, settings, and compensation files

Starter Files
Method: 
 
Settings & Compensation (Same file for both):
 

 

SECTION: Load the Cytometer
10. Click "Acquire"
(If you would like to verify the integrity of your samples before starting the worklist, click "Adjust Settings")
SECTION: Load the Cytometer
10.1. Follow the plate loading prompt
Load DI H2O, ICF, and Bleach into the appropriate positions
Place plate in the Guava Flow Cytometer in the appropriate orientation, as indicated by the marks in the loading tray
SECTION: Load the Cytometer
10.2. If you clicked "Adjust Settings"
Select the well of interest, 
Verify your cells are in the appropriate region [Image Attached] 
Once verified, click "Next Step"
click "Resume Worklist"





SECTION: Load the Cytometer
11. Acquire samples
A 48-well run should take ~300 min 

 

 

SECTION: Post-run Processing
12. Export single-cell observations to determine dataset-specific binning threshold
SECTION: Post-run Processing
12.1. When a run has finished, click the "Analyse" tab inside of InCyte.
SECTION: Post-run Processing
12.2. In most cases, recent runs will be preloaded in "Analysed Data"; however, if your dataset of interest is missing, you may load your dataset in by clicking the blue folder that says "Open Analysed Group".
Raw files are exported as YEAR-MONTH-DAY_at_HOUR-MINUTE-SECONDpm.fcs (e.g. 2022-10-04_01-38-41pm.fcs)
Make sure the correct Method is applied to the analyzed data (Starter Method File: Method.gsy)
SECTION: Post-run Processing
12.3. Highlight all wells of interest
Click on one well, then click again and drag your mouse across your desired selection
SECTION: Post-run Processing
12.4. Right click your highlighted selection and select "Export List Mode Data"
Sometimes an error pops up saying that the file name has already been written. Don't worry, your files were successfully exported.
SECTION: Post-run Processing
12.5. Locate your exported files of interest
If combining multiple cytometry runs, we recommend placing all .csvs in the same file.
SECTION: Post-run Processing
13. Open RStudio to determine the dataset-specific symbiont binning threshold

Starter R Script is available here:
  
Example (Reduced Wells) List Mode Dataset: 
  
 


SECTION: Post-run Processing
13.1. Install and load in R Packages:
dplyr v1.0.10 (Wickham et al. 2022)
tidyr v1.2.0 (Wickham and Girlich 2022)
readr v2.1.2 (Wickham et al. 2022)
ggplot2 (Wickham 2016)
ggpubr v0.4.0 (Kassambara 2020)
cowplot v1.1.1 (Wilke 2020)
SECTION: Post-run Processing
13.2. Import and combine all list mode data
 


SECTION: Post-run Processing
13.3. Replace "-" with "." and make our fluorescent signature of interest (red fluorescence off of the blue laser) into a numeric

 

SECTION: Post-run Processing
13.4. Plot the density of observations based on their relative red fluorescent intensity off of the blue laser

 


SECTION: Post-run Processing
13.5. Determine your dataset-specific binning threshold to separate cells from other particles by identifying where x equals the minimum number of observations

 

 


SECTION: Post-run Processing
13.6. If desired, remove observations that do not fall within this binning threshold to have dataset, with every fluorescent profile for each fluorescent signature detected
Metadata can be applied to this dataset based on file names
 

 


SECTION: Post-run Processing
13.7. Export filtered data to avoid the need for reprocessing
 

SECTION: Post-run Processing
14. Using the threshold determined in step 13.5, manually adjust the bin titled "Symbiont" on the RED-B Fluorescence Histogram in InCyte:Analyse. 

 


  

SECTION: Post-run Processing
15. Once the appropriate "Symbiont" bin has been applied to a dataset export a Group Stats .csv file


 

SECTION: Post-run Processing
15.1. On the left side of InCyte, click "Show Group Stats"
SECTION: Post-run Processing
15.2. Click "Setup"
SECTION: Post-run Processing
15.3. Remove the checkmarks for each empty field
SECTION: Post-run Processing
15.4. Click "Done"
SECTION: Post-run Processing
15.5. Click "Export to .csv" and save in desired location
SECTION: Concentration Normalization
15.7. 

To determine the cell density, multiply the number exported (Concentration) in step 15 by your dilution and slurry volume, and then normalize your concentration to a surface area for calcifying Cnidaria (e.g. Koch et al. 2021) or protein content for non-calcifying Cnidaria (e.g. Krediet et al. 2015).

  

Starting database for  cell density calculations:

  

Example methods to get you started on cell concentration normalization:

 


 


 



SECTION: Post-run Processing
15.6. The fluorescence readings are now ready to be used! Label your numbers appropriately, combine with other files, and apply any necessary metadata