Most DIY biologists and community labs lack either the space, funds, or electrical infrastructure to support −80°C storage of their cell cultures. (BosLab's issue is mostly lack of space followed by lack of funds to pay the increased electric bill.) While it seems that storing E. coli, the most common molecular biology workhorse organism, for indefinite time may be possible in consumer-grade −20°C chest freezers, here we focus our attention on dry storage at ambient temperatures to gain these additional benefits:
Ability to ship cells by mail, even by international mail that may take many months to deliver
Usability in places without electricity, either completely “off-grid” labs or labs without reliable electricity
A survey of the literature indicates that intracellular trehalose accumulation protects living cells against dessication damage. In particular, Garcia de Castro and Tunnacliffe (Ref. 1) noted greatly increased viability of E. coli cells dried, stored for 7 days, then rehydrated, if they were first cultured in an osmotically stressed condition then vacuum dried from a 1 M trehalose solution. Culturing under osmotic stress, such as by culturing in 0.5 M NaCl, induces some bacteria to produce and accumulate trehalose within themselves. Pelleting these cultures and resuspending in 1 M trehalose before drying appears to ensure sufficient trehalose both inside and outside the cell membrane to enable the bacteria to survive the drying process.
23 Nov 2021 Tuesday: Frank Lee and Chris Watt did the initial trial of the draft protocol at BosLab during weekly Novice Night. Cells didn't grow in any way as to be detectable as cloudiness in the culture media either by eye or by OD600 measurements. Snapshot of the protocol used along with results is linked here.
Summary: Looks like the DH5α grew but the GFP strain didn't.
Plan: Consider starting the cultures next time in LB until visible growth, then subculturing in BLOSMM.
27 Nov 2021 Saturday: Trying the approach mentioned in the 23 Nov 2021 status update, which is to first start the cells growing in LB, then add BLOSMM v0.1 to the culture to stress them osmotically. Today's focus is on seeing if transformed E. coli cells like our GFP Agar Art strain can be made to grow on BLOSMM v0.1. Snapshot of the protocol used along with results is here.
Summary: Contamination in the experiment was detected by the controls. Results inconclusive.
5 Apr 2022 Tuesday Novice Night: Planning to restart this project. We'll try simplifying the protocol to use LB instead of M9 because from reading refs. 1 and 4 it seems the exact growth media isn't as important as the need to stress the bacteria with NaCl so they produce and accumulate trehalose. The protocol needs some rework to reflect this new direction.
22 Apr 2022 Friday: Kate, Chris W., Ross and I inoculated cultures per steps 1-9 of this protocol snapshot.
26 Apr 2022 Tuesday Novice Night: Kate, Chris W., and Frank completed steps 9-15 of the procedure for drying E. coli in this protocol snapshot. Fingers crossed that some bacteria will survive the process as intended.
3 May 2022 Tuesday Novice Night: Chris and Frank took the dried paper cultures and rehydrated them in both LB broth liquid cultures and on LB agar plates.
10 May 2022 Tuesday Novice Night: Second attempt at bacterial drying by growing E.coli in LB broth instead of BLOSMM. We performed steps 1-9 of the drying protocol. Protocol snapshot here.
11 May 2022 Wednesday: Frank completed steps 10-12 of the drying protocol but had to leave, so froze the trehalose cultures at -20°C.
17 May 2022 Tuesday Novice Night: Deposited agar art bacteria on a second laser-printed paper template to dry over the week in the 37°C plate incubator. Frank was inattentive and accidentally used glycerol cultures instead of the trehalose cultures from 11 May 2022. (Sigh.)
3 Jun 2022 Friday: Began edits for use of 3M NaCl. Intend to dry the Blue agar art strain which we haven't successfully cultured for some time.
Work in a sterile airspace such as the BosLab PCR workstation (a.k.a. “PCR hood”).
Labeled 6 culture tubes with your initials, the date, and as follows:
LB+chloro Pink
LB+chloro GFP
LB+chloro Teal
LB+chloro Blue
LB+chloro Purple
LB only, DH5α control
With a 5 mL serological pipette, sterile transfer 1 mL of 3M (3 molar) NaCl to each tube.
It is OK to reuse the pipette from the previous step for this one. Sterile transfer 3.8 mL LB media to each tube.
It is OK to reuse the 5 mL pipette because the 3M NaCl is sterile and any residue it leaves in the pipette would be something that is in LB media anyway, i.e. NaCl and water. This is one tactic for designing protocols that reduce plastic waste.
With a P20 micropipette, transfer 10 µL of 1000X chloramphenicol stock solution to each of the “LB+chloro” tubes, being sure to skip the “LB only, DH5α control” tube.
Vortex each tube for 10 seconds so contents are well mixed.
Using a P200 micropipette to transfer 190 µL GFP Agar Art E. coli liquid culture to the LB+chloro GFP tube.
Same with the other colors into the corresponding LB+chloro color tube, except that for the “LB only, DH5α control” tube, add 200 µL DH5α liquid culture to it to equalize volume with the other tubes which received 10 µL chloramphenicol.
Incubate tubes in the orbital shaker at 37°C and 250 RPM for 36 hours. See Ref. 5 below for justification for the 36 hours.
While incubating, label six sterile microfuge tubes with your initials, the date, and:
LB+chloro Pink
LB+chloro GFP
LB+chloro Teal
LB+chloro Blue
LB+chloro Purple
LB only DH5α
If you're dry-storing the bacteria onto paper in petri dishes, then:
Label 6 sterile Petri dishes the same way as the tubes.
Sterile transfer a piece of autoclaved paper into each Petri dish.
If you're dry-storing to a laser-printed paper template, then be sure you've autoclaved it and dried it in a clean container.
I mounted my paper template using autoclave tape onto the inner compartment of a an all-metal clipboard, the kind that construction contractors use, and autoclaved it in that. Stainless-steel clipboards don't seem to corrode much when autoclaved. I then removed the autoclaved clipboard and moved it directly into a 37°C incubator to dry overnight without opening, thereby attempting to keep the template sterile.
Pipette 1500 µL from each incubated culture tube to the correspondingly labeled microfuge tube.
We used a P1000 and pipetted 750 µL twice.
Centrifuge the microfuge tubes to pellet cells. Discard supernatant into liquid biohazard waste.
Resuspend cells in 200 µL of 1 M trehalose by vortexing for 15-30 seconds, or until the pellet is gone.
Pipette 40 µL per piece of paper. This covers an area of approximately 1 cm x 1 cm.
Put paper in sterile petri dishes with lids closed but NOT parafilmed or sealed. Use two small pieces of tape to keep the lids from opening if the dishes are dropped.
Or put paper in new unused zip lock bags (leave bags open). This might work also but we haven't tried it yet.
Place petri dishes or bags in plate incubator to dry. May take 1-3 days depending on ambient humidity.
Due to scheduling, we incubated for a full 7-day week.
Remove petri dishes and/or bags and store in a cool dry place.
Let's say we want to rehydrate and revive N cultures. For today, N=5.
Autoclave a pair of scissors and two pairs of tweezers.
Perform all steps below in a sterile atmosphere.
Label N 14 mL culture tubes with your initials, date, “LB+chloro”, and the name of the bacterial strain you're reviving.
For today, there are five colors: (N=5)
“Pink Agar Art E.coli”
“GFP Agar Art E.coli”
“Teal Agar Art E.coli”
“Blue Agar Art E.coli”
“Purple Agar Art E.coli”
and one plain old “DH5α E.coli”
Sterile transfer (N+1) × 2 mL of LB to a sterile container. This could be another 14 mL culture tube if the quantity fits, else use sterile glassware such as an autoclaved 100 mL bottle.
For today, N=2, so use a sterile 10 mL serological pipette to transfer 6 mL LB to a sterile culture tube that we'll label “LB+chloro”.
Antibiotic stock solutions at BosLab are made at 1000X concentration and stored in the “Antibiotics” box inside the −20°C chest freezer. Sterile transfer (N+1) × 2 µL of 1000X chloramphenicol stock solution to the LB. Mix the LB and chloramphenicol thoroughly.
For today, use a P20 micropipette to transfer 6 µL of chloro stock to the LB. Use the vortex to help mix the LB and chloro.
Using the sterile tweezers and scissors, cut small squares of dried cultured paper and insert into the correspondingly-labeled culture tube. Close and tap each tube as necessary to ensure the paper goes to the bottom of the tube.
Sterile transfer 2 mL of the LB+chloro into each culture tube to wet the dried paper within.
Cap each tube but be sure the cap is in the “loose” position so air can get in.
Incubate tubes in the shaking incubator at 250 rpm and 37°C for 1-2 days or until desired growth stage is reached.
I'm adapting Garcia de Castro's and Tunnacliffe's approach as given in Ref. 1 for use in a DIY/community bio lab setting. Paraphrasing/quoting Ref. 1:
Growing E. coli cells under osmotic stress to induce trehalose production and accumulation
Specifically, E. coli MC4100 was grown in M9 medium (Ref. 3) with 1% glucose and trace elements (0.015 mM FeSO4, 0.015 mM ZnSO4, and 0.015 mM MnSO4), with or without osmotic stress (0.6 M NaCl), and harvested in growth and stationary phases. As determined by gas chromatography, stressed E. coli in growth phase contained the highest level of trehalose (230 µg/109 CFU in a typical experiment), while stressed cells from stationary phase contained somewhat less (150 µg/109 CFU). Unstressed cells from either growth phase or stationary phase did not contain detectable amounts of trehalose (<0.5 µg/109 CFU).
Drying the trehalose-loaded E. coli in a trehalose solution
Cells were vacuum dried from a trehalose solution, using a “quick drying” protocol: Samples from both growth and stationary phases were recovered by gentle centrifugation (4,000 relative centrifugal force) and resuspended in 1 M trehalose (~109 bacteria/mL), and 100 µL volumes were divided into 7 mL glass serum vials. Vacuum drying for 2 h at 30 to 100 mTorr with a shelf temperature of 30°C was carried out in a modified freeze dryer (Dura-Stop MP; FTS Systems, Stone Ridge, N.Y.); vials were sealed under vacuum. Samples were either rehydrated and plated immediately after drying or were first stored at 30°C for 7 days (Fig. 1). Bacteria were rehydrated with 1 mL of Luria broth at 20°C, serially diluted, and plated on Luria broth agar. Viability is defined as CFU of rehydrated bacteria/CFU of a nondried control, expressed as a percentage.
Preparing the 1 M trehalose solution for the drying process
The trehalose we have in the lab is food grade and doesn't specify the waters of hydration, but searching “trehalose” on research chemical supplier Sigma-Aldrich's website tells us that the dihydrate is normally what's sold (there's no other option), and they helpfully give the molar mass of trehalose dihydrate as 378.11 g/mol with an empirical formula of C12H22O11 · 2H2O. Note this means the molar mass of pure trehalose is 342 g.
We note that 1 M (i.e. a trehalose concentration of 1 mole per liter of solution) is fairly close to the maximum amount of trehalose that can dissolve in water at ambient temperature, especially since at BosLab we turn down the building heat at night and ambient temperature can drop to 10°C. Ref. 2 gives the solubility of trehalose at 10°C as 42.3 g per 100 g aqueous solution which, if we assume 1 L of saturated aqueous solution weighs 1000 g (check this!), the concentration at saturation is 423 g/L. In terms of molarity, this is 423 g/(342 g/mol)/L or 1.23 mol/L = 1.23 M, meaning a 1 M concentration is around 80% saturation at 10°C.
Wikipedia says that trehalose is heat stable, so we will autoclave sterilize our 1 M trehalose solution.
Drying the E. coli
While we do have a vacuum pump in the basement, for simplicity for our first attempt, we will dry the bacteria as drops of 1 M trehalose solution on filter paper in closed sterile petri dishes in our plate incubator. We've all seen our plates dry out completely if left in the incubator for more than a few days without parafilm wrapping or being enclosed in a zip-lock bag, so I think this will work. We will try drying overnight at 37°C because the incubator is a shared resource and other members may have cells they need to incubate at 37°C, but if the cells fail to revive after rehydrating, we can try lowering the temperature to 30°C to match what was done in the paper.
Mix 1 M trehalose ingredients in another clean 1 L culture bottle.
I found it helpful to first fully dissolve 189.1 g trehalose in 300 mL distilled H2O using a magnetic stirrer and hot plate with either the top-surface temperature set to 80°C, or the solution temperature set to 50°C if you have a probe that can directly measure the solution temperature. Turn off the heat and remove the stir bar once fully dissolved, and allow to cool to room temperature. Pour the solution into a clean graduated cylinder with 500 mL or greater capacity, and add distilled H2O to make 500 mL final volume.
Solubility of trehalose in water: The solubility of trehalose in water at 10, 20, 30, and 40°C was found to be 42.3, 46.6, 52.3 and 59.7% (g trehalose per 100 g solution), respectively.