Protocol: Rigorous secondary confirmation of BDM utilization by oligotrophic bacteria and fungi
Overview: BDM degrading fungi and bacteria were originally selected if they grew on minimal medium
containing no carbon source, atop a sterilized square of BDM fabric – but did not grow on minimal
medium that contained no carbon source at all. To repeat and verify these results, we revived
cryopreserved stocks of BDM degrading microbes and grew them in liquid and on solid broth with no
carbon. To our surprise, many of the isolates grew – albeit very slowly. Several factors explain our
observations.
1)
Most of the species we have identified thus far fall into the class of microbes known as oligotrophs
– organisms that grow on very small amounts of nutrients by scavenging from the environment.
Examples are organisms that are able to contaminate bottled water (presumably pure) or grow
upon glass surfaces. Oligotrophs rely on their efficiency at scavenging carbon, nitrogen, and other
nutrients dissolved in the aqueous environment or existing as volatiles in the air. Because
oligotrophs invariably use a wide variety of substrates simultaneously, they are normally nonspecialized regarding substrate utilization. Saprophytes are commonly oligotrophs. It is not
uncommon for such organisms to have a suite of metabolic capabilities available for adaptation to
a range of environments. Thus, it is not surprising that the same microbes we isolated from BDMs
(presumably requiring unusual enzymatic capabilities) seem to have extraordinary oligotrophic
metabolic capabilities. It is also worth considering that these oligotrophs are “keystone species” in
the colonization of recalcitrant nutrient sources like BDMs. Whether they actually degrade them at
all is still in question, though metabolic by products (like organic acids from fermentation) may aid
in weathering the films.
2)
During the course of this research the Brodhagen lab moved. We suspect that the different brand
of agar (Acros) that we used in the new lab may have been contaminated with trace nutrients that
permitted oligotrophs to grow without added C in the solid medium though they had not,
previously. Examples of other possible sources of carbon reported to be available for oligotrophic
microbes include:
i. Traces of detergent on glassware
ii. Trace volatiles in the air
iii. Dust in air, on spatulas, and in reagents
iv. Finger grease
A concern surrounding the isolation of oligotrophs is that we are not sure that organisms growing on the
BDMs are truly utilizing the BDMs for carbon. They could be utilizing dust, volatiles, or some other scant
C source. Thus, we have revamped our procedures to ensure that we were not inadvertently providing
any unknown carbon source to microbes in control treatments (minimal medium with no carbon source
added). We now use stringent protocols and special glassware and reagents. Experimental samples are
incubated in a specially built air-tight box with activated carbon filters at the inflow.
Precautions for glassware: We noted that many of our BDM degraders colonize polypropylene used for
disposable culture tubes. Therefore, we use only glass culture tubes. To avoid any residues from soap,
and to avoid scratches in glass which facilitate colonization, we culture microbes in brand-new glass
culture tubes and do not reuse them for this purpose. Caps are also brand new. They are rinsed well in
distilled water and autoclaved prior to use. Glassware for media-making is critically cleaned: it is
soaked overnight in 1-10% HCl and rinsed 14X in distilled water prior to use.
Precautions for media: We have purchased a set of new reagents (mineral salts and agar) that are
dedicated solely to this project. No spatulas are used in the weighing of reagents – rather, powders are
tapped gently from the container, in order to avoid cross-contamination via spatulas. To avoid
introduction of contaminants from high-pressure steam forced into containers during autoclaving, we
filter-sterilize all media into virgin plastic receptacles.
Precautions for incubation: In order to avoid the introduction of organic volatiles from the room air in
our Chemistry building, we constructed an airtight chamber in which inflow must pass through an
activated carbon (respirator-type) filter. After samples are added to the chamber, air is purged for two
hours by pulling a vacuum on the outflow to force old air out and new air in (over the filter). Because
microbes themselves emit volatiles, air is purged for one our daily throughout the incubation period.*
*In a side-by-side experiment using all new glassware and chemicals, purging the air did not cause a
difference in growth on minimal medium with no C – all cultures, regardless of air “cleanliness”, failed to
grow without an added C source. However, this experiment was conducted over Christmas break when
little work was taking place in the building and presumably fewer volatiles were being released into the
air.
Precautions for BDM preparation: BDMS are handled only with
clean gloves or dedicated (previously unused) forceps not used for
any other purpose, and cut with an unused razor blade on a
polypropylene cutting board. BDM squares are sterilized under a
germicidal UV lamp (253.7 nm) in a biosafety cabinet that has
been thoroughly wiped down with 70% EtOH and then presterilized by running the UV light for 2 hours before use.
A
Cutting BDMs
Materials Required:
1
4
1
4
4
1
Box of nitrile gloves
500 mLs Beakers (critically clean)
polypropylene cutting board, cleaned and surface sterilized
with 70% EtOH
Unused razors
Forceps (dedicated to BDM project)
Metal ruler, cleaned
B
Sizes for BDM pieces to equal 0.0175 grams each:
Cellulose Control:
SB PLA-2010:
BioBag:
BioTelo:
1.
2.75cm x 0.6cm
2.75cm x 0.6cm
5.5cm x 1.7cm
5.5cm x 1.5cm
Wearing gloves, place BDM on polypropylene tray, aka
“cutting board” (Fig. A)
C
D
2.
Use metal ruler to measure length, 2.75cm or 5.5cm, and make
tick marks on each end with fresh razor. With metal ruler cut a
clean straight line between the two tick marks. (Fig. B)
3.
Measure out width with ruler making tick marks on either side.
With the ruler cut a clean straight line between the two tick
marks. You will end up with a rectangular piece of film that
eventually will be placed longitudinally in a culture tube,
supported with a sterile paper clip. (Fig. C)
4.
With forceps, transfer cut BDM into beaker.
5.
Repeat steps 1-4 until all BDMs are cut.
E
Sterilizing BDMs
Materials required:
70% EtOH
Box of nitrile gloves
Four 500 mL beakers (critically clean & sterile)
Forceps, sterile (dedicated to BDM project)
1.
Turn blower on and thoroughly wipe down all surfaces (base,
sides, window) of biosafety cabinet using 70% EtOH. Let fan
blow for 15 minutes to ensure 100% turnover of stale air with
freshly filtered, sterile air.
2.
Close sash and turn UV germicide lamp on. Let UV sterilize for
two hours.
3.
Turn UV off and blower on.
4.
Use sterilized forceps to lay out 0.0175g BDM pieces. Prevent
excessive contamination from having your hands/arms pass
over the BDMs by starting in the back left corner and laying
pieces out in a “Z” pattern (Fig. D & Fig. E).
5.
Close hood and turn UV germicide lamp on (Fig. F). If sash
closes fully with blower on the BDMS fly everywhere! To
prevent this, turn blower off when the sash is 1 in from closing.
6.
Let UV sterilize BDM for 2 hours.
7.
Turn UV off and blower on.
8.
Use sterile forceps to flip over BDMs. Now the side that will
become the bottom side is sterile - prevent contamination of
F
G
H
I
these prior to flipping them (from passing hands/arms above them) by starting in the front right
corner and flipping pieces over in a reverse “Z” formation (Fig. G & Fig. H).
9.
Close sash and turn UV germicide lamp on. Let UV sterilize for two hours. Use sterilized forceps to
collect the sterilized BDMs and place them into a sterile beaker. Prevent contamination by starting
in the front right corner and collecting in a reverse z formation (Fig. G and Fig. H).
10. Cover beaker with its sterile aluminum foil cover while still in sterile hood.
Inoculating:
Materials required:
--Spores harvested from each fungal species, suspended in 0.01% Triton X-100* and enumerated via
hemacytometer
--New, rinsed, sterile glass culture tubes filled with 5 mL FMM, GMM, M9, or M9 + 0.2% Glucose (see
accompanying protocol for media preparation for mesh bag study, for recipes)
--Sterile paper clips (stainless steel, rinsed with ddH2O, autoclaved)
--Beakers containing all four BDMS, cut into 0.0175 g pieces and UV-sterilized
--Biosafety cabinet
--P10, P20, P200, P1000 pipettors and filter tips, cut 1mm from ends to permit spore passage
*This potential C source (traces of Triton X-100) has not seemed to permit fungal growth in past
experiments.
Experimental setup: Glass culture tubes are filled with 5 mL fungal minimal medium or M9 (bacterial)
minimal medium and carbon sources are supplied according to the table below.
Carbon source treatments
No carbon (negative control)
Glucose (0.2% w/v) (positive control)
BDM (0.0175 g of film)  cellulose control, SB PLA-2010, BioTelo, and BioBag
Three replicates of each treatment are performed for each fungal or bacterial isolate. To ensure that no
pre-existing contaminant serves as an inoculant for each film type, five replicates of medium plus BDM
film are left uninoculated and incubated along with all other samples. Five tubes of uninoculated
minimal medium and minimal medium + 0.2% glucose are also inbuated as controls (negative).
BDMs are supported by paper clips so that they are partially in, and partially out of the growth medium.
This is because some of the BDM degrading microbes prefer to grow on the BDM while it is submerged
(anaerobic) and others prefer to grow on it when it is exposed to the air (aerobic).
1.
Films must be attached to paper clips using sterile forceps, in a decontaminated biosafety
cabinet. Plan ahead – to do ~100 tubes will require about 8 hours of painstaking work (the
BDMS flutter in the hood air and frequently come into contact with the outside of the culture
tubes, rendering them unsterile). It is easiest to do this task with two people: one to handle the
BDM pieces and one to open and close the culture tubes.
2. 5 mLs of the appropriate medium is added to each tube
3. Culture tubes are inoculated with 5 x 103 spores/mL of growth medium or with a known starting
OD of yeast or bacterial culture (to be determined).
4. Cultures are incubated at 20oC for 5 days in an airtight chamber permitting filtration through
activated carbon to eliminate most organic volatiles from the air.
5. Air in the culture box is purged for 2h at the outset and for 1h every day afterwards.
6. Cultures are observed visually for growth after 5d (longer or shorter incubation times may be
performed with certain isolates).
7. Optical density readings can be made of the positive and negative controls, but are meaningless
for the BDM samples as the film blocks the passage of light*.
*Future protocols may include a method for safranin staining followed by safranin recovery as a crude
measurement of overall biomass, but because species are expected to take up stain differently, it is not a
good method for across-species comparisons of BDM colonization.