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  1. Daily Culture Maintenance

  2. Clonal Isolation

  3. Cryopreservation


Daily Culture Maintenance

Sampling for Flow Cytometry Counts

  1. Turn on computer and flow cytometer.
  2. Under owncloud/MorrisLab, “LTPE_Fluorescence_FCM.xlsm” spreadsheet. Make a list in your lab notebook of all the cultures that say “YES” under “Transfer?”
  3. Remove indicated tubes in order. DON’T TURN OFF THE HOT DOG ROLLERS! They might not come back on again.*
  4. Take a 96 well plate with a lid. You will fill the plate from 1A to 12H, by columns.
  5. Mark the first empty well with a sharpie dot on the lid.
  6. Wipe down the worksurface in the hood with ethanol. Put the tubes and the 96-well plate in the hood. Keep the tubes in the order you wrote the names down in your lab notebook.
  7. Wipe down the P100 pipetor with ethanol.
  8. Put 90 uL of ASW into one well for each sample to be checked.
  9. Unscrew all the tube caps. Put 10 uL from the first tube in the first well; pipet up and down to mix. Repeat with the second tube, moving down the column.
  10. When you’re done, put the lid on the plate and screw all the tubes tightly closed. Return tubes to incubator.

Flow Cytometry Counts

  1. Check waste and wash vials on Guava. If the waste is full, empty it; if the wash is empty, fill it with milli-Q water.
  2. On the FCM computer, open the guavaSoft application and click “InCyte”.
  3. Click the “Edit Worklist” icon — a scroll icon at the top left of the InCyte screen.
  4. In the 96-well plate schematic, select the wells where you put your samples. Hold down “ctrl” to select multiple wells.
  5. Click the check box beside “Acquire this sample”.
  6. Enter “1000” by “Events to Acquire”
  7. Enter “150” by “Time to Acquire”
  8. At the top left, click the check box beside “By Column”
  9. Click the check box beside “Park Capillary in” and then in the drop-down menu select “T08”. THIS IS VERY IMPORTANT AND IF YOU DON’T DO IT YOU WILL CAUSE $100’s IN DAMAGE.
  10. Click “Run Worklist”
  11. Under “Analysis Method”, click “Choose”. If your work run only includes Pro and Syn samples, select “ProSyn.gsy”. If your work run only includes Syn and Peuk samples, select “SynPeuk.gsy”. Note that you can’t run Pro and Peuk samples in the same run; if you have both of these, you’ll have to run one of them separately.
  12. Under “Settings” click Choose and select ProSyn.gst or SynPeuk.gst based on your above choice for Analysis Method.
  13. Click “Acquire”. The plate ejects.
  14. Tubes W2, W4, and W5 should be filled with milli-Q water; top them off if necessary.
  15. Tube W1 is for backflushed waste. If it’s more than half full, empty it into the biohazard bag or a dirty flask bound for the autoclave.
  16. Tubes W3 and W4 should be empty.
  17. Tube 10 should be filled with milli-Q water.
  18. Tube 8 should be filled with milli-Q water. This one is VERY IMPORTANT AND IF YOU DON’T DO IT YOU WILL CAUSE $100s IN DAMAGE.
  19. Click “OK”. Your program will now run.

Analyzing FCM Data

  1. Go to “Analysis” in InCyte and click on your data set (bottom left box). You should see each well that was analyzed highlighted in blue.
  2. Click on a well. Drag the “Count” box over the population of cells you want to count. It’s usually pretty obvious. In some cases, particularly with Pro, you’ll have to be careful to avoid counting “Noise”.
  3. Write down the concentration (in cells/mL) of the sample in your lab notebook. Continue with remaining samples.
  4. Go back to the LTPE_Fluorescence_FCM spreadsheet. Type in your measured concentrations under “Today’s reading”. If the “Extra Days” column says “Transfer!” then transfer:
    • Click the “Transfer” button. Type in cell count and current media batch # at the prompts”
    • Put a check by the transfer in your lab notebook so you know to transfer it.
  5. If it doesn’t say “Transfer!” but instead has a number, add that number to the number in the “deltaT” column.
  6. Save and close out the worksheet (Ctrl_S). Proceed to transfers.


  1. Label each tube’s cap with a dot label. Make label exactly the same as the old tube is labeled, but add today’s date.
  2. In the hood with the Bunsen burner, flame and aseptically decant the sterile milli-Q water in the tube. Flick the tube vigorously 5 times to remove as much of the water as possible.
  3. Add 12.3 mL of the appropriate medium to each tube.
  4. If the tube is C-, add the indicated amounts of sterile milli-Q water and NaOH. If it is C+, add the indicated amount of H2O, HCO3-, and HCl for the given media batch. Each aliquot of HCl needs to be added with a separate pipet tip and pipet up and down to mix when you add it. Wipe down pipettor with ethanol before use. Note that you don’t need filter tips for these additions.
  5. Loosen the caps of the old cultures. Don’t flame! The media goes up to the lip and you will kill all the cells if you try to flame it.
  6. Transfer 500 mL of each culture using P1000 filter tips. Wipe down pipettor between each culture. Using the same tip, transfer 500 mL culture to a YTSS purity tube w. same label. Note that the same purity tube may be used for several subsequent transfers.
  7. Seal tubes VERY TIGHTLY. Wipe off any moisture that got on the outside of the tube.
  8. Place new tubes in the incubator in the same position as the old tubes.
  9. Place old tubes on the “dark shelf” by the purity tubes. Keep 2 generations of old tubes at any given time just in case.


* If you accidentally turn off the Hot Dog Roller and it won’t start again, turn on the rollers and gently rotate the second rollers from the outside counterclockwise until they start moving again.


  • All cultures should be maintained in at least 2 different incubators at all times, preferably either on backup electricity or on separate power grids
  • Cultures are grown in disposable 14 mL polyethylene culture tubes (VWR 60818-667) in 10 mL volume and at low (~10 mEin) light, 22 C
  • Each month, cultures must be transferred:
    1. Wear lab coat and gloves throughout transfer procedure. Ethanol gloves if you touch anything outside of the hood. BE VERY PARANOID about contamination at all steps.
    2. Take 1 fresh tube for each culture. Print out labels on return address labels and stick on before adding media
    3. Add 10 mL of appropriate medium to each tube. After all tubes have been filled for a particular medium, add ~0.5 mL of medium to a YTSS purity tube.
    4. Transfer 200 mL previous culture into fresh medium using a P200 filter tip. Wipe down pipettor barrel between tubes.
    5. For axenic cultures ONLY: With the same tip add 200 mL culture to a YTSS purity tube.
    6. Incubate purity tubes at room temperature.
  • In case of contamination:
    1. If medium was contaminated, ASAP re-transfer culture into a new batch of media.
    2. If culture was contaminated in one set but not in the other set of tubes, ASAP transfer the non-contaminated copy into the second incubator.
    3. If both cultures are contaminated:
      • Use antibiotics if possible to remove contaminating organisms; dilute culture 50X into antibiotic-containing medium and check for purity as soon as growth is visible.
      • If antibiotics aren’t an option, go back to freezer stocks (see section on cryopreservation)
      • If no stocks, plate culture and select the first dilution that has no heterotrophic contaminants.



Prior to starting evolution experiments, or at 100-generation intervals, clones must be isolated from algal populations. Two methods are used: colony isolation on agarose plates and dilution-to-extinction in liquid media using 96-well plates.

Liquid protocol:

In order to be reasonably certain of clonality, dilute until ~90% of wells have no growth. Use Guava flow cytometry to determine cell counts.

  1. For Pro: need overnight EZ55 (JJM310) culture. On day before plating, wash EZ55 twice in sterile ASW and dilute 250 mL into 24.75 mL Pro99 for each organism to be plated.
  2. Dilute culture to ~ 50 cells/mL, then do 2 more 10x dilutions. For Pro, do all dilutions in EZ55-treated Pro99. Last 3 dilutions are in 25 mL volume.
  3. For final 3 dilutions, fill a plate with 200 mL/well using sterile reservoirs and multichannel pipet
  4. Put in Ziploc bag and incubate at whatever light level original culture was growing at.

Plating procedure:

  1. Day before: grow EZ55 (JJM310) if plating Pro
  2. Make soft agar using Invitrogen LMP agarose in ASW: .35%, need 5 mL per 60 mm plate. Make just enough for one set of experiments.
  3. Sterilize agar by autoclaving. Cool for a few minutes and add appropriate nutrients plus sulfite: (0.6302 g/5mL water, use 100 mL/L agar final concentration)
  4. Allow agar to cool in 34C water bath for an hour or two.
  5. Wash EZ55 twice in sterile ASW (if growing Pro).
  6. Dilute phyto cells to ~500 cells/mL in ASW, then 2 more 10x dilutions
  7. Put agarose on bench and let cool to ~29 C. Can use contact or infrared thermometer.
  8. Put 50 mL EZ55 (for Pro) in every plate
  9. Put 1.2 mL last 3 phyto dilutions in ever plate
  10. Add 4.8 mL agarose using 5 mL disposable pipete and mix by swirling.
  11. Let dry in hood for ~1 h
  12. Put in Ziploc bags at low light. Don’t invert!



Every strain in the culture collection should be cryopreserved. Also, mixed populations from each 25-transfer mark in the evolution experiment should be given LTPE #’s and cryopreserved both in the evolution media (i.e. straight out of the evolution tube) and in rich media grow-out tubes. There are two methods, one for cyanobacteria and one for eukaryotes.

Cyanobacterial protocol

  1. Open the “LTPE_Strain_Collection.xlsx” spreadsheet in the OwnCloud/LTPE_Stuff folder. We freeze all the lines from a given generation under a contiguous set of LTPE numbers, so the first thing to do is to look to see if any other lines from this generation have been frozen. If so, figure out which reserved LTPE number applies to the line you are about to freeze. If not, input a new set of numbers corresponding to all of the + and – lines for the line/generation you are about to freeze. NOTE: W and M lines each need 24 reserved numbers per generation, as they will be frozen both directly from the evolution tubes AND from the rich media grow-out tubes.
  2. For each line to be frozen, label three 2 mL screwcap cryovials. Label the cap with the strain number and the side with strain number, date, and line. For instance for LTPE151, put a “151” on the cap and on the side write “LTPE151 5/15/17 M+1”.
  3. In the hood using filter tips, place 150 mL of sterile filtered cell-culture grade DMSO into each tube. It’s easiest to do this if you put the cryovials in a cryobox and mostly (but not completely) unscrew the caps. NOTE: if you need to make more DMSO, it must be syringe filtered with a NYLON filter.
  4. If you are freezing a W or M line, also get a 15 mL disposable plastic culture tube and fill it with 9 mL of SN or Pro99 media, respectively. Label this tube the same way you labeled the side of the cryovial but also include the LTPE number reserved for the rich media grow-out line.
  5. Using a 5 mL disposable serological pipet, withdraw 7 mL of the evolution culture. Place 1 mL into the grow-out tube and 2 mL into each cryovial.
  6. Place the grow-out tube into the incubator on the low-light shelf.
  7. Seal the cryovials. Put them in a cryobox, and with the lid on, invert them to mix. Incubate in the dark for 5-15 minutes.
  8. Open the freezer and withdraw one of the racks (make sure to wear cryogloves). Carefully pour off the liquid nitrogen from the lower parts of the rack that are submerged. Put the sealed cryobox in the bottom slot of the rack, replace the restraining rod, and immerse it in the liquid nitrogen for 5 minutes.
  9. Remove the cryobox. One by one, remove the main collection box (Rack 1), the backup collection box (Rack 10), and the “O.S.” collection box (Rack 6). Place one copy of each tube into each box and return to the freezer.
  10. If you have to start a new storage cryobox, label it appropriately on all four sides of the lid as well as on the body of the box.
  11. Record the date of freezing and the cryobox number in the spreadsheet and save it.
  12. Repeat for the grow-out tubes when they show robust color.

Eukaryote protocol

  1. Prepare cryovials and spreadsheet as described above for cyanobacteria.
  2. Add 2 mL of acclimated CCMP371 culture to each cryovial. Seal and invert. Let stand in the dark for at least 5 minutes.
  3. Open Mr. Frosty and throw out the foam insert if it’s still in there. Fill “Mr. Frosty” to the line with isopropanol. Place the tube-holding rack in the alcohol and add the tubes to be frozen. Screw Mr. Frosty shut.
  4. Place Mr. Frosty at the back of our -80, or near the bottom of a chest -80. Let sit for 4-6 hours.
  5. Place the tubes into cryoboxes in the liquid nitrogen freezer as described above for flash-frozen cyanobacteria.

Resurrection protocol

  1. As quickly as possible, bring cryovial to room temperature. Immersion in water is a good way to achieve this. Do this in the dark.
  2. Place 100 mL of frozen culture into 9.9 mL of fresh medium (whatever it was frozen in).
  3. Incubate at VERY low light for 1 day (~ 5 uE)
  4. Move to low light (<= 30 uE) until ready to transfer. When transitioning to high light, maintain original grow-out culture at low light (just in case).