dH2O vs. saline: rudimentary fungi culture storage
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README.md

Introduction

Castellani's method for storing fungi in distilled water1234 leverages their ability to enter into stasis absent air, light, and nutrients. This paper describes my experience working with water cultures for one year as part of an emerging strategy for genomic data integrity.

Fungi are resilient and can adapt to extreme environments, including those so hostile as to halt their metabolism. Successful long-term storage depends on the cultivator's ability to consistently and aseptically induce cryptobiosis with the fewest cell divisions.

Unfortunately for many independent researchers and smaller labs, the current state of the art is prohibitively expensive. Saline vials designed after Castellani are part of a larger strategy that cycles cultures every 1, 5, and 10 years: 4 °C agar slants, 4 °C saline, and –80 °C cryovials.

Cavalcanti et al.'s large sample sizes and Bezerra et al.'s PCR analysis show that water cultures can be viable after 20 years.56

Materials and methods

My goal was to store gourmet and medicinal fungi for one year at 25 °C with the least expensive materials. The 3rd generation used distilled water at 25 °C in the homemade vials I describe below. The 4th generation uses 0.6% saline with basal salts at 4 °C in 10 mL glass BD Vacutainer vials (red top).

Making the dram vials

A case of 144 vials stores 18 isolates kept in quadruplicate at 2 locations, on Stamets' recommendation.7

The vials come with foam pads on the underside of the lids: contamination vectors. I replaced the pads with 1/8" of silicone and drilled 1/8" injection ports. Incidentally, drilling the ports made the pads easier to remove.

I wrapped the vials in Parafilm to protect the glass-silicone interface and used parchment paper to keep the Parafilm from sticking together. I keep labelled 4-packs in small plastic bags. The material stack that sheathes the water column is, from inside out: glass, silicone, Parafilm, parchment paper, and plastic, in a glass container.

Drilling the 3rd generation vial caps

Suspending the isolates

I tried to consistently suspend the mycelium at 90% colonization up to Stamets P-3 Value,8 but it varies because I took samples on a rolling basis. The culture library contains roughly a 70:30 mix of Aloha Medicinals strains and wild specimens, and includes a mycorrhiza and a sclerotium.

Then I autoclaved for 20 min at 15 psi the dram vials with loose lids and foil wraps, Poland Spring water at pH 6.4 and 37 TDS,9 and one 60 mL syringe with 16G needle per strain. I squirted water into a ported 4 oz agar plate, took up a syringe of mycelium, and filled 8 drams with liquid inoculant, sealing each vial at once.

I flame sterilized the needle between each movement, kept the drained plate to verify my technique, and plated out the suspension to verify the medium.

Results and discussion

todo: Briefly describe both revival efforts.

3rd generation results

I revived 1 mL dH2O solution for 1 attempt in December 2017 and incubated each culture at 25 °C for 30 days. I visually determined that no plates were viable except L. tigrinus, which contaminated with red and yellow bacteria.

G. lucidum, P. erygnii, and P. nameko are presumed dead, to revive later or acquire new strains.

Species P Value Date Viability
A. blazei P-2 2017-11
G. lucidum P-2 2017-04 nil
G. sessile P-2 2017-04
G. frondosa HI P-2 2017-05
G. frondosa NH P-2 2017-11
H. tessellatus P-2 2017-04
H. americanum P-2 2017-11
I. resinosum P-2 2017-11
L. tigrinus P-2 2017-05
P. columbinus P-3 2017-11
P. erygnii P-1 2017-04 nil
P. ostreatus P-1 2017-04
P. nameko P-2 2017-04 nil
S. rugoso-annulata P-2 2017-04

4th generation results

I started using 0.6% saline and BD Vacutainer vials in January 2018 and reconstructed my culture library from new isolates and backup plates.

todo: Revive the vials for growing out and resuspension into saline and glycerol.

Species P Value Date Viability
A. aegerita P-5 2018-02-13 nil
C. gigantea P-1 2018-02-13 nil
G. lucidum P-1 2018-02-14 nil
G. sessile P-1 2018-01-30 nil
G. frondosa HI P-3 2018-02-13 nil
G. frondosa NH P-1 2018-02-13 nil
H. abietis P-4 2018-02-14 nil
H. americanum P-2 2018-02-14 nil
I. obliquus P-1 2018-02-13 nil
I. resinosum P-1 2018-02-13 nil
L. edodes P-4 2018-02-13 nil
P. roqueforti P-2 2018-02-13 nil
P. nameko P-1 2018-01-30 nil
P. tuber-regium P-2 2018-02-13 nil

Which water to use?

Benedek used normal saline (0.9% NaCl) in 1962 with apparent success,10 and Tresner and Hayes found in 1971 that over half their basidiomycetes died in 2% salt.11 Osmobiosis is a relatively unknown phenomenon as of 201112 and evaporation is a long-term worry. The normal saline ratio is arbitrary or derived from an osmotic coefficient calculation specific to human blood in vivo.13

For these reasons I use a 0.6% sea salt–dH2O solution that mimics the composition of human blood. This "blood saline" fits an emerging concept to use natural entropy, as measurements become asymptotically precise, to correct for intrinsic flaws in human perception.

An increasingly accurate salt ratio would be the mean blood salinity of all tested mammals or animals. The best fit may be mineral water obtained close to the strain's origin. dH2O doesn't suffice because mushrooms are vulnerable to hypertension even though they employ high turgor pressures to colonize lignins.14

Sealing the 3rd generation vials with silicon

Assessing the vacuum's quality

The silicone–glass interface between vial and cap is the main entry point for contaminants. An airtight seal makes a weak vacuum to ensure that with proper care, only sterile air and water can enter through the inoculation port.

Good materials and technique enforce the vacuum's guarantee of a single entry point. My vessels are handmade and the water cheap, but my method is designed to succeed even in open air.

BD Vacutainer vials are purpose-made to hold serum-like substances such as water cultures. Breaking the vacuum automatically and efficiently fills 10 mL with an adequate air reservoir.

Future work

We need effective solutions to store rare fungi cultures as soon as they're isolated. Wild polypores such as Laricifomes officinalis that inhabit old-growth forests may have medically and industrially significant compounds. Live cell cultures can also be transformed to produce related variants of the compounds of interest.

There's a a security aspect to safeguarding old-growth forests that may likely contain the next generation of drugs. Possible attacks against a strategically fruitful biome might include introducing a predator or altering a niche. Harmful stewardship doesn't require malicious intent, however, and we can attribute much biodiversity loss to industrial progress.

Appendix 2020: Storage methods reviewed

The text below is a post in a persistent Shroomery junk thread where I discuss the options the paper presents that informed my cryopreservation work. Doing –80 °C storage is technically difficult and fraught with complications such as, what if you can't afford to run a massive 230V appliance 24/7 for years or decades? Mycelium seems more fragile with dry vs. wet methods because it grows as a linear chain of cells rather than budding or dividing.


Since the thread is twice bumped, here's the state of the art: https://sci-hub.tw/10.1016/j.funbio.2013.12.002

Filter paper made the rounds in Fall 2019 as a good way to mail bacteria samples, which it is. This is often true for yeast and other molds so then people look at "fungi storage." We mostly care about mushroom producers (basidiomycetes) that are known to be more fragile than yeast and bacteria, i.e., you can't revive shiitake broth from minus 80 unless you make special preparations first.

The Homolka 2014 paper is divided into every known storage technique and it lists the most recently published storage length. Basidiomycetes are often underrepresented or else they die so grading techniques by "how many lived" is a bad idea. Note that freezing temps reflect the experimental designs described and are not constants. Any non-fluctuating freezer (no auto-defrost) is probably fine.

Subculturing (serial transfer) [agar slants]. Universally used despite their disadvantages. You can mitigate strain degradation with half-strength agar, a mineral or paraffin oil cover, or partially drying it.

Mineral or paraffin oil. Agar under mineral oil at minus 20 is good for our fungi. The linear growth of mycelium vs. budding of yeast and bacteria seems to require moisture for reliable storage. "Piaggio (1996) reported quite high viability of 127 fungal strains (mainly wood-rotting basidiomycetes) stored under oil and retrieved after 10y and 20y. A general survival rate was 80% after 10y and 93% of the surviving after 20y."

Storage in sterile water. It generally seems to work well for basidiomycetes provided the solution is isotonic (saline) and preferably not all aqueous. This is one of the first "solid teks" from 1939. My pet method, details at the end of the post.

Drying of fungal cultures. Drying them as on filter paper is good for spores, i.e., spore prints can last up to 5 years or more. For mycelium, you'd need silica gel to achieve worse results than water.

Freeze-drying (lyophilization). Large and often mold/bacteria libraries like ATCC freeze-dry their stuff in milk with a cryoprotectant. Not good for our goals: to reliably store a diverse library of hyphae so that we can easily revive a dead agar slant. "The effort to apply lyophilization to fungal hyphae in most cases has met with mixed success. Nevertheless, several results show that freeze-drying of basidiomycetes can be successful in some cases."

Cryopreservation. There are many methods that range from minus 80 to liquid nitrogen on a variety of substrates. Homolka's own pet method (a good one except for working with perlite) involves growing a 10% glycerol broth with sterile perlite crystals, then draining the broth and storing the crystals. This section is the bulk of the paper and it's worth reading. I want to focus on techniques you can do with a freezer that doesn't auto-defrost.

Based on all this info, I think there are a few criteria that stand out as a rule of thumb: "basidiomycetes keep best either in cryo conditions or those closest to their natural habitat." Well, where do we usually find the best mushrooms? In places that are wet but not underwater, gritty but not salty, cold but not permafrost, where air and light are either benign or helpful.

A larger strategy would have multiple time scales, including decades if you can afford cryo for that long. The simplest would be two scales: agar slants to make plates from, and frozen glycerol/saline to have reference backups. Trading very low metabolism samples and high quality genetic data would be like downloading a cool file and a crypto signature to verify it.


  1. Aldo Castellani. "The viability of some pathogenic fungi in sterile distilled water." Journal of Tropical Medicine and Hygiene 42. 1939: 225–226. ↩︎

  2. Aldo Castellani. "Further researches on the long viability and growth of many pathogenic fungi and some bacteria in sterile distilled water." Mycopathologia et Mycologia Applicata 20. 1963: 1. ↩︎

  3. Aldo Castellani. "The 'Water Cultivation' of Pathogenic Fungi." Annales de la Société belge de médecine tropicale 44/2. 1964. http://lib.itg.be/open/asbmt/1964/1964asbm0217.pdf ↩︎

  4. Aldo Castellani. "A maintenance and cultivation of the common pathogenic fungi of man in sterile distilled water. Further researches." Journal of Tropical Medicine and Hygiene 70. 1967: 181–184. ↩︎

  5. Bezerra, C. C. F., Lima, R. F. de, Lazera, M. S., Wanke, B., & Borba, C. M. (2006). "Viability and molecular authentication of Coccidioides immitis strains from culture collection of the Instituto Oswaldo Cruz, Rio de Janeiro, Brazil." Revista Da Sociedade Brasileira de Medicina Tropical, 39(3), 241–244. https://doi.org/10.1590/s0037-86822006000300002 ↩︎

  6. Cavalcanti, S. D. B., Vidal, M. S. M., Sousa, M. da G. T. de, & Del Negro, G. M. B. (2013). "Viability and molecular authentication of Coccidioides spp. isolates from the Instituto de Medicina Tropical de São Paulo culture collection, Brazil." Revista Do Instituto de Medicina Tropical de São Paulo, 55(1), 7–11. https://doi.org/10.1590/s0036-46652013000100002 ↩︎

  7. Paul Stamets. Growing Gourmet and Medicinal Mushrooms, chapter 12, pages 110–121. Ten Speed Press, 3rd edition, 2000. ↩︎

  8. Fungi Perfecti. "What is the Stamets P Value System?" Online; accessed 2017-Nov-13. https://www.fungi.com/blog/items/what-is-the-stamets-p-value-system.html ↩︎

  9. Home Reverse Osmosis. "What's in the Water You Buy?" Online; accessed 2017-Nov-27. http://www.h2ro.com/bottled2.htm ↩︎

  10. Benedek, T. (1962). "Fragmenta mycologica." Mycopathologia et Mycologia Applicata, 17(3), 255–260. https://doi.org/10.1007/bf02279298 ↩︎

  11. H. D. Tresner and Jean A. Hayes. "Sodium Chloride Tolerance of Terrestrial Fungi." Applied and Environmental Microbiology 22/2. Aug 1971: 210–213. ↩︎

  12. Møbjerg, N., Halberg, K. A., Jørgensen, A., Persson, D., Bjørn, M., Ramløv, H., & Kristensen, R. M. (2011). "Survival in extreme environments - on the current knowledge of adaptations in tardigrades." Acta Physiologica, 202(3), 409–420. https://doi.org/10.1111/j.1748-1716.2011.02252.x ↩︎

  13. Awad, S., Allison, S. P., & Lobo, D. N. (2008). "The history of 0.9% saline." Clinical Nutrition, 27(2), 179–188. https://doi.org/10.1016/j.clnu.2008.01.008 ↩︎

  14. RJ Howard, MA Ferrari, DH Roach, and NP Money. "Penetration of hard substrates by a fungus employing enormous turgor pressures." Proceedings of the National Academy of Sciences 88/24. Dec 1991: 11281-4. ↩︎