PHYTOPLANKTON EVOLUTION PROTOCOLS
J. Jeffrey Morris, last updated 6/4/15
Daily Culture Maintenance
I.
Fluorometer Readings
a. Turn on computer and open “Trilogy” program. This should bring up an Excel spreadsheet.
b. Open fluorometer door and use metal spatula to defeat the door-latch mechanism.
c. Start w. hot dog roller cultures. Remove all tubes in order (S1 through S5, C- first, then
C+). DON’T TURN OFF THE ROLLERS! They might not come back on again.*
d. Try to keep the tubes in order, since the order they’re on the table is the order of the labels
in the spreadsheet.
e. Read each tube. Drop tube into 3D-printed adaptor and touch the “Measure Fluorescence
Raw” button on fluorometer and wait. Reading should appear on the next free line in the
spreadsheet.
f. Put the tubes back in the incubator.
g. Open the "LTPE_Fluorescence.xlsm" worksheet (on the Desktop).
h. Copy and paste readings into the last empty column. Type in the time transfers were done
beneath the last reading.
i. Any cultures that are past the fluorescence cutoff will now say "YES" under the "Transfer?"
heading. Retrieve these cultures from the incubator and pair each one with a fresh test tube.
j. Return to the original spreadsheet. Measure the fluorescence of each new tube.
k. Now go back to "LTPE_Fluorescence". Click the "Transfer" button one by one for each
tube that's ready to transfer. When prompted, input the new tube's blank value. Also input
the current media batch #.
l. Save the worksheet (Ctrl_S) and close out all programs. Proceed to transfers.
II.
Transfers
a. 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.
b. At the bench 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.
c. Add 12.3 mL of the appropriate medium to each tube. Media bottles are stored in the
cabinet to the right of the Guava flow cytometer.
d. If the tube is C-, add 200 L sterile milli-Q water. If it is C+, add the indicated amount of
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. The HCO3 and
HCl stocks are stored on the shelf over Jeff's bench.
e. 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.
f. Transfer 500 L of each culture using P1000 filter tips. Wipe down pipettor between each
culture. Using the same tip, transfer 500 L culture to a YTSS purity tube w. same label.
Note that the same purity tube may be used for several subsequent transfers.
g. Seal tubes VERY TIGHTLY. Wipe off any moisture that got on the outside of the tube.
h. Place new tubes in the incubator. Don’t worry about reading them after transferring.
* 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.
CULTURE COLLECTION MAINTENANCE
1. All cultures should be maintained in at least 2 different incubators at all times, preferably
either on backup electricity or on separate power grids
2. Cultures are grown in disposable 14 mL polyethylene culture tubes (VWR 60818-667) in
10 mL volume and at low (~10 Ein) light, 22 C
3. Each month, cultures must be transferred:
a. Take 1 fresh tube for each culture. Print out labels on return address labels and
stick on before adding media
b. 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.
c. Transfer 200 L previous culture into fresh medium using a P200 filter tip. Wipe
down pipettor barrel between tubes.
d. For axenic cultures ONLY: With the same tip add 200 L culture to a YTSS
purity tube.
e. Incubate purity tubes at room temperature.
4. In case of contamination:
a. If medium was contaminated, ASAP re-transfer culture into a new batch of media.
b. 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.
c. If both cultures are contaminated:
i. 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.
ii. If antibiotics aren’t an option, go back to freezer stocks (see section on
cryopreservation)
iii. If no stocks, plate culture and select the first dilution that has no
heterotrophic contaminants.
CLONAL ISOLATION PROCEDURE
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 dilutionto-extinction in liquid media using 96-well plates. Both methods should be used simultaneously
to maximize the probability of success. Colonies are preferable to positive wells in liquid
because of the greater certainty of clonality.
Liquid protocol:
In order to be reasonably certain of clonality, dilute until ~90% of wells have no growth. A
green culture of Pro/Syn has ~ 108 cells/mL and a stationary phase F/50 culture has ~106. In
contrast, Evolution Media at transfer point have about 106 cells for Pro and Syn and 105 cells/mL
for Euks. If available, use Flow Cytometry to determine cell counts.
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For Pro: need overnight EZ55 (JJM310) culture. On day before plating, wash EZ55
twice in sterile ASW and dilute 250 L into 24.75 mL Pro99 for each organism to be
plated.
Dilute culture to ~ 5 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.
For final 3 dilutions, fill a plate with 200 L/well using sterile reservoirs and
multichannel pipets
Put in Ziploc bag and incubate at low light.
Plating procedure:
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Day before: grow EZ55 (JJM310) if plating Pro
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.
Sterilize agar by autoclaving. Cool for a few minutes and add appropriate nutrients plus
sulfite:
.6302 g/5mL water, use 100 L/L agar (final concentration!)
Allow agar to cool in 34C water bath for an hour or two.
Wash EZ55 twice in sterile ASW (if growing Pro).
Dilute phyto cells to ~500 cells/mL in ASW, then 2 more 10x dilutions
Put agarose on bench and let cool to ~29 C. Can use contact or infrared thermometer.
Put 50 L EZ55 (for Pro) in every plate
Put 1.2 mL last 3 phyto dilutions in ever plate
Add 4.8 mL agarose using 5 mL disposable pipete and mix by swirling.
Let dry in hood for ~1 h
Put in Ziploc bags at low light. Don’t invert!
CRYOPRESERVATION PROTOCOL
Every strain in the culture collection should be assigned an SEK # and cryopreserved. Also,
mixed populations and clonal isolates from each 100-generation mark in the evolution
experiment should be given LTPE #’s and cryopreserved. There are two methods, one for
cyanobacteria and one for eukaryotes.
Cyanobacterial protocol
1. Transfer 500 L of culture to 5 disposable plastic culture tubes, each containing 9.5 mL
of high-nutrient media (e.g. Pro99) and grow until bright green
2. Pool cultures and centrifuge at 8000 G, room temperature, for 15 minutes
3. Carefully decant spent medium
4. Resuspend pellet in 5 mL of fresh medium
5. Add tissue-culture grade sterile-filtered (make sure to use a nylon filter!) DMSO to final
concentration 7.5%
6. Distribute to 5 sterile 2 mL cryovials
7. In a dark or dimly-lit room, flash-freeze vials in liquid nitrogen in a small dewar.
8. Place vials into cryoboxes and store in liquid nitrogen.
Eukaryote protocol
1. Transfer 500 L of culture to 5 disposable plastic culture tubes, each containing 9.5 mL
of high-nutrient media (e.g. F/2) and grow until significant growth evident
2. The day before freezing, fill Mr. Frosty to the indicated line with isopropyl alcohol and
chill to 4C overnight.
3. Prepare cultures for freezing as in steps 2-6 above
4. Place room-temperature cryovials in Mr. Frosty, seal, and place in -80 freezer (near the
bottom)
5. After 1 hour, remove vials and immerse in liquid nitrogen. Store in cryoboxes in liquid
nitrogen dewars.
Resurrection protocol
1. Work in the dark – at night, using very dim or red light
2. As quickly as possible, bring cryovial to room temperature. Immersion in water is a good
way to achieve this.
3. Place entire content of vial into 9 mL fresh medium (same medium in which sample was
frozen)
4. Immediately make 2 10-fold dilutions
5. Incubate at VERY low light for 2 days (~ 5 E)
6. Move to increasingly higher light afterwards for grow-out
PH/ALKALINITY MEASUREMENT PROTOCOL
pH and alkalinity should be assessed for each batch of culture medium at both CO2 levels.
1. Calibrate probe w. pH 4, 7, and 8.3 standards. Rinse probe in milli-Q and then buffer
before each point.
2. Prime and calibrate pump.
3. >= .05 mEq sample to clean beaker w. stir bar; bring to ~ 50 mL w. freshly autoclaved
milli-Q H2O
4. Titrate. Formula for your sample, where A is machine’s reading and S is volume of
sample you added in L:
𝐴𝑙𝑘𝑎𝑙𝑖𝑛𝑖𝑡𝑦 (
𝑚𝑒𝑞
50𝐴
)=
𝐿
𝑆
To measure pH: if sample for titration < 25 mL, measure undiluted sample separately.
Otherwise can measure prior to dilution. Not necessary to stir but have to swirl; make sure to
rinse probe between samples and get the ceramic junction under the water-line.