Miscellaneous Experiments with
Phototrophic Communities:
Photomethanotrophs
Phototroph enrichments and pigment modification
under varied light environments
Spatial makeup of the Purple Berries
Carie Marie Frantz
Ph.D. Student, Geobiology
University of Southern California
3651 Trousdale Parkway, Los Angeles, CA 90031
Summer 2009
Microbial Diversity Course
Marine Biological Laboratory, Woods Hole, Massachusetts
Abstract
For my mini-project I decided to attempt three and a half more or less unrelated (except under the broad
umbrella of phototrophy) mini- mini-projects, and carried them out with varying degrees of success. This
report’s purpose is to serve as a guide for future, more focused investigations of the same or similar topics
and it is broken down by mini-project.
I.
Phototrophic Methane Oxidation
Phototrophic methane oxidation is a tantalizing example of a thermodynamically favorable process that so
far no organism has been found to exploit. One environment where this process might be expected to
occur is the Trunk River near Woods Hole, Massachusetts where shallow sandy mats of purple sulfur
bacteria are bubbled with methane produced by underlying organisms. Inocula of purple sulfur bacteriarich sediments from the Trunk River mats were incubated in an anoxic, carbonate-buffered minimal
medium with methane gas as the sole electron donor. Methane concentrations in the headspace were
monitored over the course of 5-10 days using gas chromatography in order to look for methane
consumption. Methane consumption was observed both in samples incubated in the light as well as
samples incubated in the dark, suggesting that non-phototrophic methanotrophy occurred in these
enrichments (perhaps aerobic methanotrophy due to the presence of trace amounts of oxygen, nonphototrophic anaerobic methane oxidizing consortia, or an as-yet unknown form of anaerobic methane
oxidation).
II.
Phototroph Enrichments & Pigment modification under varied light conditions
Light quality can be an important selective factor for photosynthetic organisms. In these experiments I
used light-emitting diodes of various wavelengths to investigate the response of phototrophs to different
wavelengths of light.
(a) Phototroph enrichments under suboptimal light wavelengths
Inocula of purple phototrophic consortia-containing sediments from the Little Sippewissett salt marsh
were incubated in acetate- or sulfide-containing medium with 470nm (blue), 525nm (green), and 595nm
(yellow) peak wavelength LEDs as their sole light source. A green sulfur bacterium grew to moderate
densities in the 470nm-incubated sulfide medium. Purple bacteria formed dense biofilms in the 470 and
595nm acetate enrichments. These purple bacteria grew in interesting circular and vertical striped
patterns on the bottle front (directly next to the LED board) and back walls; this was determined to be due
to the strong directionality of the LEDs. Surprisingly, no turbidity or pigmented growth was visible in the
595nm enrichments. However, white circles formed on the front walls of the bottle (much like the purple
growth in the 470 and 525nm enrichments) that appeared to be elemental sulfur formed perhaps by metalmediated photooxidation of sulfide or perhaps due to the influence of an unseen (either unpigmented or
extremely dilute) sulfur-oxidizing organism.
(b) Response of a mixed culture of phototrophs to a slightly shifted light environment
The selective nature of changing wavelengths of light was tested in this experiment by incubating
subcultures of purple phototroph enrichments grown for several weeks at 880nm under somewhat shifted
light environments using 870nm and 900nm peak wavelength LEDs. Shift in absorbance was monitored
by looking at in vivo and in extracto pigment absorbances. Community shifts were also observed
2
qualitatively using microscopy and terminal restriction fragment length polymorphism (TRFLP).
Significant changes were observed in extracted pigment absorbances, some of which was not explainable
by the community shifts alone, suggesting that one (or several) type of organism actively increased its
proportion of carotenoids to Bchl a pigments as a response to the shifted light environment.
III.
Spatial makeup of the Purple Berries
The purple aggregates found in shallow ponds of the Little Sippewissett Marsh have been extensively
investigated by previous students in this course, yet their community structure and origin remain
something of a mystery. I attempted to use several methods to examine the physical distribution of
different key players within the berries. Berries were frozen and cross-sectioned and used for DNA
extraction and catalyzed reporter deposition-fluorescence in situ hybridization (CARD-FISH) procedures.
I was unable to amplify DNA extracted from the interior and exterior of berries and CARD-FISH also
proved problematic, but the minimal results obtained suggest that these methods may yet yield at least a
partial solution to the berry problem.
3
Phototrophic Methane Oxidation
Background
Phototrophs are known that oxidize water, sulfide, molecular hydrogen, iron(II), and many other
substances, but, despite favorable thermodynamics, so far no phototrophs have been conclusively shown
to oxidize methane. The CH4/CO2 redox couple in biologically-relevant systems of +0.24V is only
slightly less than that for H2S/S0 (+0.27V) and phototrophic sulfide oxidation is a widespread process in
the anaerobic photic zone. Photosynthesis an anoxygenic phototrophs is coupled to far less favorable
redox pairs than methane and CO2 including the oxidation of nitrate to nitrite (-0.43V) [1].
The favorable thermodynamics of phototrophic methane oxidation was championed by the astrobiologist
Wolf Vishniac [2]. Vishniac’s attempts to demonstrate the process led to the isolation of a methaneutilizing purple nonsulfur bacterium Rhodopseudomonas gelatinosa which appeared to photoassimilate
methane [3], however this result was never replicated and growth using methane as the sole electron
donor was never demonstrated. A previous course participant, Stefan Ratering (1996), attempted to
isolate a photosynthetic methanotroph but saw no growth in his inoculated media bottles after two weeks
[4]. Light-regulated methane oxidation has been observed in natural environments, but this could be
explained by the dependence of aerobic methanotrophs on oxygen-producing cyanobacteria [5].
This experiment represents another attempt to capture a theoretical organism that has thus far remained
elusive.
Methods
Sampling
Shallow, purple-sulfur bacteria-containing sediment samples from Trunk River, Massachusetts were
collected (41⁰32'06.7"N, 70⁰38'28.5"W; see Figure 1) in anaerobic bottles on 7/13/09 (Set 1) and in 50mL
Falcon tubes on a second sampling trip four days later (Set 2). Care was taken to get rid of oxygen
bubbles to minimize oxygenation of the sediments and porewater between sampling and inoculation.
Figure 1 - Location of the Trunk River sampling site.
4
Culturing
Set 1
The medium for the first set of inocula (Set 1) consisted of a standard lab freshwater salts base (17 mM
NaCl, 2 mM MgCl2, 0.15 mM CaCl2, 6.7 mM KCl) supplemented with 10mM NH4Cl as a nitrogen
source, 1.5mM KH2PO4 as a phosphate source, 1mM Na2SO4 as a sulfur source, trace elements (from a
stock containing μM levels of Fe, BO3, Mn, Co, Ni, Cu, Zn, MoO4, VO3, WO4, SeO3), and trace amounts
of vitamins (1 μg/mL each of riboflavin, thiamine, L-ascorbic acid, d-Ca-panthothenate, folic acid,
nicotinic acid, 4-aminobenzoic acid, pyrodixine, lipoic acid, NAD, and thiamine pyrophosphate; 0.3
μg/mL biotin; and 10 μg/mL Vitamin B12). The medium was buffered to pH ~7 using 31.5mM NaHCO3
and a 0.2 atm CO2 headspace. The basal salts and trace elements were autoclaved together, the medium
cooled, and the vitamins and carbonate added after the medium had cooled to room temperature.
The medium was dispensed into Balch tubes and bubbled for 20-30 minutes under a stream of N2/CO2
(20% CO2) gas. Tubes were then inoculated with approximately 1 mL (5 tubes) or 5 mL (5 tubes) of
Trunk River sediment, bubbled an additional 10 minutes with N2/CO2, and then stoppered and crimpcapped. Two additional tubes were left uninoculated as controls. 1mL of CH4 gas (to make up a 20%
methane headspace) was added to each stoppered tube by filling a syringe with methane and injecting
1mL of methane into each tube (all tubes had 5mL of headspace).
The gas chromatograph (GC) was not working on the day that the tubes were inoculated, so the tubes
were placed in the dark until the GC was available again. Once initial measurements were taken, the
tubes were moved to a sunlit location in a SW-facing window.
Set 2
The medium for the second set of inocula (Set 2) was the same as for the first except for the addition of
50uM 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) to inhibit the growth of cyanobacteria (which
would introduce oxygen and favor aerobic methanotrophy). Also, this medium was cooled under a
stream of N2/CO2 (20% CO2) gas immediately after autoclaving and the addition of vitamins, carbonate,
and inoculums were all performed under anoxic gas in order to minimize the oxygen content of the
medium.
Twelve tubes were inoculated with 4mL each of freshly-sampled Trunk River sediments and six were left
uninoculated as controls. The tubes were then stoppered and crimped, and 1mL CH4 gas was added (in
the same manner as for Set 1) to eight of the inoculated tubes and all six of the uinoculated tubes. Four
inoculated tubes were left CH4 -free as controls. Initial methane measurements were made within two
hours of CH4 addition and the tubes were then placed in the SW-facing window with the Set 1 samples.
Table 1 gives a summary of the different enrichment conditions.
5
Table 1 - Summary of Enrichment Tubes
Tubes
TR1-3
TR4-5
TR6-8
TR9-10
TR11
TR21-24
TR25-28
TR29-32
TR33-38
Set
1
1
1
1
1
2
2
2
2
Sample &
Inoculation
Date
7/13/09
7/13/09
7/13/09
7/13/09
7/13/09
7/17/09
7/17/09
7/17/09
7/17/09
Inoculum
Volume
(mL)
1
5
1
5
0
4
4
4
0
Methane
Headspace
(atm)
0.1
0.1
0.1
0.1
0.1
0.1
0.1
0
0.1
Light
Light
Environment Environment
Before 7/17
After 7/17
Dark
Light
Dark
Light
Dark
Dark
Dark
Dark
Dark
Light
n/a
Light
n/a
Dark
n/a
Light
n/a
Light
Methane Measurements
Methane concentrations in the enrichment tubes were monitored periodically using a GC (GC 2014
Shimazu Gas Chromatograph, Shimazu Scientific Instruments, Inc., Nowell, MA) by injecting 100μL of
the gaseous headspace onto the column. The syringe used was flushed thoroughly with nitrogen before
each sampling from the tubes in order to minimize oxygen contamination. Retention times for methane
on the column were consistent (0.31-0.34 seconds) across all days that measurements were made,
however, significant variation was observed when measurements were done for the same sample multiple
times in one sampling session. This was true for standards as well as for samples. This is possibly due to
problems with syringe clogging or leaking, despite using only syringes that appeared to be leak- and clogfree. An alternate possibility proposed by other students in the course is that the GC was possessed by an
evil force and was due for an exorcism.
Figure 2 - Daily variation in methane standard measurements. Don’t let the apparent lack of variation in the
earlier measurements fool you. Multiple measurements were only made for the standards on July 20 (10mM
standard), July 22 (10 and 1 mM standards) and July 23 (all standards).
6
Results
Results of the methane measurements are shown in Figure 3 below.
Methane consumption was seen in all samples (for both sets) between the first and second measurements
(the sole exception being TR11 for which no measurement was made during the period where the
decrease was seen for the other samples). This is followed by a dramatic methane increase for most
samples on 7/22 (214 hours for Sample 1, 118 hours for Sample 2); an increase is also visible on the same
date in the standard measurements (Figure 2) suggesting that this may be due to instrumental variation.
Discussion
While the samples appeared to be subject to significant variation in instrument (and syringe) function,
some significant trends were apparent. First, no significant differences were observed between samples
incubated under dark and light conditions, strongly suggesting that the methane consumption observed
was not influenced by light. Second, methane consumption in the uninoculated samples was significantly
slower than in the inoculated samples. This suggests that the methane consumption visible in nearly all
samples was at least in part biotic in origin.
One explanation for these trends would be the aerobic oxidation of methane using small amounts of
oxygen present in the sediment inoculum’s porewater, the medium, or introduced during headspace
sampling. However, I believe it is important to note that Sample Set 1, the sample set that probably had a
higher oxygen content (due to less carefully-controlled anoxic media preparation, more time for gas
leakage, and the growth of cyanobacteria in the tubes after several days of light incubation), exhibited a
significantly slower rate of methane consumption than the probably more anoxic cultures of Sample Set 2.
An alternative explanation for these results is the presence of non-phototrophic anaerobic oxidation of
methane (AOM). This process is widespread and well-documented in anoxic sediments and is thought to
be coupled with the reduction of sulfate [6] and nitrate [7], and was very recently (less than two weeks
prior to the writing of this paper) reported to be coupled to the reduction of manganese and iron [8]. The
emission of sulfide from Trunk River suggests a high level of sulfate reduction in the anoxic sediments,
and if sulfate reducers were present in the initial inoculula for this experiment, they would have had 1mM
sulfate (15 μmol total in 15mL medium) available to them in the culturing medium. Theoretically, one
mole of sulfate oxidation could be coupled to one mole of methane oxidation by the following AOM
reaction:
Equation 1
CH4 + SO42- → HCO3- + HS- + H2O
If this were the case in my enrichments, one would expect a maximal oxidation of 15μM of CH 4, far less
(by several orders of magnitude if the GC calibration and my late-night back-of-the-envelope calculations
are to be believed) than the level of CH4 oxidation observed. No oxidized manganese, nitrate, or iron was
supplied to the cultures. This, however, assumes a negligible amount of sulfate or other potential electron
acceptors for AOM processes in the actual sediment inoculums.
In short, I’m not sure how to interpret these results and can’t come up with a good explanation right now,
on the last night, after several weeks of far too little sleep and time to think. If someone figures this out or
has any ideas, tell me; I’d be really interested.
7
Figure 3 - Methane consumption in photomethanotroph enrichment cultures. Top: Sample Set 1, Bottom:
Sample Set 2. Yellow and orange lines represent light-incubated inocula, blue lines represent dark-incubated
inocula, green lines represent uninoculated controls, and red lines represent inoculated samples incubated with a
20% CO2 (methane-free) headspace. The tube containing Sample TR24 was cracked, and lost all methane within
three days.
8
Suggestions for future work
Photomethanotrophs are still waiting to be conclusively demonstrated to exist. This is not an easy thing
to do in a few short weeks, but should any future course participants wish to attempt this project again,
here are a few suggestions:
1. Leave yourself time to do this project right.
2. Know what your electron donors and acceptors are in your medium and calculate how much methane
non-phototrophic AOM could possibly consume. Test the concentrations of relevant ions and oxygen
in the sediment inoculum before inoculating.
3. Inoculate your samples as early as possible in order to give any phototrophic methanotrophs a good
amount of time to grow and start –trophing the methane.
4. Try inoculating with and without an oxygen scavenger + oxygen indicator. Oxygen scavengers can
also act as electron donors (and could theoretically supplant methane), but it would be good to know
whether oxygenation of your samples over time is a problem.
5. Incubate samples in tubes on their sides so that (a) gas exchange is maximized (this is very important
as methane is very insoluble in water and gas exchange may be a key limiting factor in the existence
of photomethanotrophs and (b) fluid covers the stoppers and helps minimize gas leakage. Another
option would be to incubate the tubes upside-down or tilted on a shaker.
6. Incubate under wavelengths that select for purple bacteria (e.g. Bchl a at 860-880nm).
7. Leave lots of time for messing with the GC, i.e. 2-3 hours/sampling day. Take multiple
measurements for each sample, make good standard curves, and check your syringe (and flush it
thoroughly with something inert) before each injection.
8. Take lots of timepoints.
9. Calculate peak areas, not just peak heights. This helps account for any variations in peak shape that
can occur on different days.
10. If time permits, look for AOM organisms in your inocula using clone libraries, FISH, and/or
microscopy.
9
Phototroph enrichments & pigment modification under varied light
environments
Background
One key limiting factor for different types of photosynthetic life is the amount and quality of light
available in a given environment. The ability to use light of a particular quality can be very important in
some ecosystems, such as deep in the water column or within soils, rocks, or microbial mats where a
significant amount of the solar spectrum has been scattered, absorbed, or filtered out by overlying
phototrophic organisms and only small color bands of light are available; organisms that are able to
efficiently use light at these remaining wavelengths have a competitive advantage over other organisms.
Photosynthetic pigments absorb fairly discrete bands of light and only these energy transitions can be
used to convert light energy to chemical redox potential. Organisms get around this by having antenna
complexes that use different pigments that absorb light at shorter wavelengths and then funnel that energy
via resonant energy transfer (with some loss) to the pigments involved in the chemical reactions. The
absorbance of these pigments can be shifted somewhat by the presence of different bound organic chains
and by the protein environment, and these modifications can be very important for organisms under light
stress (e.g. [9]). Organisms can also modify their ability to absorb in certain parts of the spectrum by
beefing up the number of certain pigments in their membranes [10]. Despite this degree of absorption
flexibility, light color appears to be a limiting factor in some environments and can dramatically influence
the makeup of phototrophic communities (e.g. [11, 12]).
In the following two mini-mini-projects, I sought to demonstrate the concept of light color as a limiting
factor in photosynthesis and look at color as (1) a selecting factor for organisms with particular pigments
and (2) a selective pressure that causes phototrophs to modify the relative abundance or properties of their
pigments.
Part 1: Phototroph enrichments under suboptimal light wavelengths
Methods
Experimental Setup
Light-emitting diodes (LEDs) were obtained at three “suboptimal” (i.e. not corresponding with the peak
absorbances of major pigments) peak wavelengths: 470nm (5mm, 30⁰ spread, 2400mcd at 20mA; RL5B2430), 525nm (5mm, 32⁰ spread, 7500mcd at 20mA; RL5-G7532), and 595nm (5mm, 30⁰ spread,
5000mcd at 20mA; RL5-Y5030) all ordered from Super Bright LEDs, Inc. (St. Louis, MO). The spectral
output of the LEDs is a normal distribution centered about the listed peak wavelength with an
approximately 50nm full width at half maximum value. Light boxes (Figure 4, Figure 5) were built using
cardboard boxes equipped with LEDs on a large solderless breadboard (Figure 6). LEDs were spaced
such that two Pfennig bottles could be illuminated. All LEDs were wired in parallel with a variable
current/variable voltage power supply. Voltages were set to values slightly below their continuous
forward voltage ratings and the current from the power supply was increased until the LEDs turned on.
Output voltages remained approximately constant throughout the duration of the incubation experiments.
10
Figure 4 - Experimental setup with the light
boxes and power sources shown.
Figure 5 - Light boxes (shown opened) for the
phototroph enrichments. Boxes were closed during
the experiments.
Figure 6 - Breadboard circuit with LEDs used for the phototroph enrichments. The colored wires in the upper
right are not connected/not part of the circuit. Red wires are positive leads that are connected to the positive
terminal of the power supply and black wires are negative leads that are connected to the negative terminal of the
power supply.
11
Enrichments
Two media were used for the enrichments in study, both based on a minimal seawater medium (342mM
NaCl, 15mM MgCl2, 1mM CaCl2, 6.7 mM KCl, 10M NH4Cl, 1.5mM K2HPO4, and the trace elements and
vitamins described in the photomethanotroph experiments) buffered and supplemented with an electron
donor (acetate or sulfide).
Acetate Medium: In seawater medium, 0.25mM Na2SO4, 2.5mM MES Buffer (pH 5.5), 20mM Sodium
Acetate, and 50μM DCMU (to inhibit cyanobacteria).
Sulfide Medium: In seawater medium, 1.5mM MOPS Buffer (pH 7.2), 70mM Bicarbonate, 10mM
Sodium Thiosulfate, 1mM Sulfide, and 50μM DCMU (to inhibit cyanobacteria).
In both cases, the minimal seawater medium was prepared with buffer (and sulfate for the acetate
medium) and autoclaved, cooled under a stream of gas (N2 for acetate, N2/CO2 for sulfide), and then the
additional components added after the medium had cooled. The prepared medium was dispensed into
Pfennig bottles which were then gassed to remove oxygen and capped.
Prepared bottles (one of each medium for each of the three light environments) were then inoculated with
~1g of purple berry-containing sediment from the Little Sippewissett Salt Marsh (Figure 7). Bottles were
then placed in their respective light environments (with bottles placed directly adjacent to the LEDs),
although a bottle mixup resulted in two acetate bottles being placed under 525nm and two sulfide bottles
being placed under 595nm. After nine days of incubation (with periodic checking and movement of the
bottles), the bottles were oriented with their labeled sides facing away from the lights and kept in this
position for the remaining four days of the incubation period.
Figure 7 - Pfennig bottles with prepared media inoculated with Little Sippewissett Purple Berry sediment.
Photo taken immediately after inoculation, before light box incubation.
Sample bottles were photographed and wet-mounted slides were prepared of the medium (for planktonic
growth) and wall scrapings (for attached growth) and observed.
Results & Discussion
Significant growth was seen in all of the 470nm and 525nm cultures, but no visible turbidity was present
in the 595nm cultures (Figure 9).
Green sulfur bacteria (Figure 10A) grew to moderate density in the 470nm sulfide enrichment bottle,
giving it a turbid green color. 470nm is outside the range of typical absorbance peaks for green sulfur
bacteria, however the full range of spectral output from the LED (giving appreciable light at 20-30nm
12
below and above the peak wavelength) overlaps with the in vivo absorbance peak ranges for Bchl c, Bchl
d and Bchl e, all pigments found in the green phototrophic bacteria.
Purple sulfur bacteria (with beautiful internal sulfur granules) and other bacteria grew in all of the acetate
enrichments; the apparent density and diversity of the 525nm enrichments (as determined by light
microscopy), however, was higher than for the 470nm enrichments (Figure 10B-D). All purple sulfur
bacteria formed circular growth patterns on the front of their enrichment bottles which corresponded with
the range of light spread from the LEDs. Additional striped growth patterns appeared on the back which
probably resulted from the pattern of light scattering through the glass bottle and medium. No significant
planktonic growth was observed. The output spectrum of the 470nm and 525nm LEDs would not be
expected to overlap with the bacteriochlorophyll pigments of purple sulfur bacteria. However, 470nm
falls well within the range of carotenoid absorbances and this is likely what the purple bacteria were using
to harvest light in the 470nm enrichment. 525nm would have provided light only at the edge of the
carotenoid absorbance capability, so it is somewhat surprising that so much growth (and apparent
diversity) was present in the 525nm enrichment cultures.
No visible turbidity was present in the 525nm enrichments, however, white spots formed on the front wall
of both sulfide bottles that mimicked the purple spots of the shorter-wavelength acetone enrichments in
both size and shape (Figure 8). Phototrophic growth would not be expected to be white or invisible when
growing at these wavelengths because that would imply that they are not absorbing an appreciable
amount of visible light (including light at and around 525nm). Microscopy revealed no significant cell
density in the medium and only very few recognizable cells (including some with what appeared to be
internal sulfur granules) were present in the wall scrapings (Figure 10E; most of the dark spots in the
image are mineral precipitates but the three bright spots are cells). Aggregates of bright white spheres of
varying diameter were abundant in the wall scrapings (Figure 10F), and may be elemental sulfur deposits.
Who or what is responsible for making these deposits, however, remains unclear.
Figure 8 - White spots on the front wall of a 595nm H2S phototroph enrichment bottle.
Future Work
Future studies could attempt to investigate the composition and origin of the white spots. Techniques
such as the construction of clone libraries, terminal restriction fragment length polymorphism (TRFLP),
and fluorescence in situ hybridization (FISH) could be used to identify the major (and minor) players in
the various cultures.
13
Figure 9 - Phototroph enrichments after 13 days incubation in restricted light conditions.
14
(A)
470nm H2S Medium
(B)
470nm Acetate Medium
(C)
525nm Acetate Medium
(D)
525nm Acetate Medium
(E)
595nm H2S Medium
(F)
595nm H2S Medium
Figure 10 - Representative microscopy images of the phototroph enrichment cultures after 13 days incubation
in restricted light environments. All images were taken at 100x magnification under phase contrast except (F),
which was taken at 100x under differential interference contrast (DIC).
15
Part 2: Response of a mixed culture of phototrophs to a slightly shifted light
environment
Method
Experimental Setup
Light-emitting diodes (LEDs) with peak emission wavelengths at values slightly spaced from 880nm
(Figure 11; peak emission wavelength of the lamp used for class purple sulfur bacterial enrichments) were
purchased from Roithner LaserTechnik GmbH (Vienna, Austria): 870nm (5mm, 20⁰ spread, 24mW at
50mA; LED870-01UP) and 900nm (5mm, 8⁰ spread, 16mW 100mA; B5B-900-8). The spectral output of
the LEDs is a normal distribution centered about the listed peak wavelength with an approximately 70nm
full width at half maximum value. Light boxes (See Figure 4, Figure 5) were built using cardboard boxes
equipped with LEDs on small, adhesive-backed solderless breadboards (Figure 12; breadboards
purchased from FunGizmos.com). For each breadboard, five LEDs were wired in parallel with a variable
current/variable voltage power supply. Data sheets for the LEDs purchased were not available, so
appropriate forward voltages were estimated from similar LEDs and power supply voltages were set to
values at least 0.5V below these estimated values. The current from the power supply was then increased
until the LEDs turned on (light was outside the visible range but could be monitored using the CMOS
display of a camera phone). Output voltages remained approximately constant throughout the duration of
the incubation experiments.
Figure 11 - LED output spectra compared to the in vivo and in vitro pigment
absorption spectra of Bchl a-containing purple sulfur bacteria. Pigment spectra
are from an 880nm incubated culture after two days of incubation with pigments in
30% sucrose (in vivo) and extracted in 7:2 acetone/methanol (in vitro). Normalized
LED emission spectra estimated using a normal distribution and the specificication
data provided by the vendor (Roithner LaserTechnik GmbH).
16
Figure 12 - Breadboard
circuit with LEDs used
for
pigment
shift
experiments.
Cultures
The acetate medium described for the previous mini-project (phototroph enrichments) was also used for
this experiment. A relatively fast-growing enrichment culture that formed densely pigmented planktonic
growth in acetate medium under an 880nm peak wavelength lamp was selected for this experiment. The
25-day-old enrichment culture was from the class anoxygenic phototroph enrichment exercise and the
original inoculums was a slice through the upper ~3cm (including all of the pigmented bands) of a sandy
microbial mat from the Little Sippewissett (Figure 13). The original culture was subcultured into fresh
media in a 225mL Meplats bottle and had formed a dense planktonic culture within two days (Figure 14).
Figure 13 - Little Sippewissett Mat, the likes of
which were sliced up and used to inoculate the 880nm
acetate medium enrichment culture that was
subcultred for use in this light environment shift
experiment.
Figure 14 - Subculture of the Sippewisset Mat
enrichment culture after two days of growth. This
culture was used to inoculate the light environment
shift experiment bottles.
Once the LEDs had arrived from Austria and the light boxes set up (eleven days later), three Pfennig
bottles containing 35mL of acetate medium were inoculated with 20mL of the dense subculture, bubbled
for 20 mins with N2/CO2, capped, and put in a dark box for 4 hours before putting them into the light
boxes (oriented with the wall with label tape facing away from the LEDs).
17
DNA Extraction and TRFLP
On the same day that the original subculture was subcultured for incubation in the light boxes, a DNA
extraction kit (UltraClean™ Microbial DNA Extraction Kit, Mo Bio Laboratories, Inc., Carlsbad, CA)
was used to extract DNA from three identical 2mL samples of the original subculture. The following
modifications (mistakes) were made to the kit’s procedure: only 30 μL (rather than 50 μL) of solution
MD1 was added prior to bead-beating, and bead tubes were centrifuged twice for one minute at 8,000g
(rather than once at 10,000g) following the bead-beating step. The extracted DNA was successfully
amplified using a PCR reaction with fluorescent 16S-8F and non-fluorescent 16S-1492R primers. The
resulting amplified DNA was combined and cleaned using a cleanup kit (QuickStep™ 2 PCR Purification
System, Edge Bio Systems, Gaithersburg, MD). DNA concentrations were measured using a NanoDrop
and deemed sufficient for TRFLP. A digestion reaction was run using RSA1 and Sau96 restriction
enzymes, after which Hinsby worked his magic and TRFLP results materialized on the interwebs.
The same extraction procedure was repeated four days later at the end of the experiment period with a few
important difference: no more UltraClean™ kit bead vials were left, so I instead used the PowerSoil™
DNA Isolation Kit (Mo Bio Laboratories, Inc., Carlsbad, CA) bead tubes with 500μL of the fluid in the
tubes removed before adding the culture material for bead-beating. After bead-beating, the tubes were
centrifuged 30s at 10,000g, and the rest of the steps were followed as in the previous extraction. No more
Sau96 was available on the final day, so only RSA1 was used for the restriction enzyme digest.
Pigment Analysis
In addition to TRFLP, material was regularly removed from the incubated bottles for pigment analysis.
2mL of culture was centrifuged at 13,500g for 2 minutes to pellet out the cells and the supernatant
removed. For in vivo measurements, cell pellets were re-suspended (by vortexing) in a 30% w/v sucrose
in mqH2O solution to minimize the effects of cell scattering. I also attempted to add glycerol to fresh
media suspensions of cell pellets until cell scattering was reduced, but found the method to be messy,
tedious, and inaccurate and abandoned it. For in vitro measurements, pellets were re-suspended (by
vortexing) in a 7:2 acetone/methanol solvent, soaked in the dark for 5 minutes, centrifuged at 13,500g for
2 minutes to pellet the cells, and the pigment extract was then immediately scanned in quartz cuvettes
using the spectrophotometer. All scans were done from 350-1100nm using a baseline-zero correction.
30% sucrose solution was used as the blank for the in vivo measurements, and 7:2 acetone/methanol was
used as the blank for the in vitro measurements. An attempt was made to homogenize the bottles prior to
sampling them.
Microscopy
At the end of the experiment (4 days of incubation), wet-mounted slides were prepared and examined
using 10μL of planktonic sample.
Results
After only four days, visible differences in growth were apparent in the three different enrichments
(Figure 15). The 880 culture was very turbid and a light pinkish color (not apparent in the photos, you
have to trust me). The 870 culture was a dense, deep red/brown with planktonic growth. The 900nm
culture was a lighter red with most of the cell mass at the bottom of the bottle. These visible differences
were further supported by differences seen in microscopy, TRFLP, and pigment absorbance.
18
Figure 15 - Cultures after 4 days of light box incubation. Top images are of the backs (side facing away from the
LEDs), and bottom images are the fronts of the bottles.
Microscopy
Microscope observation of the cultures after 4 days of incubation revealed significantly different
community structures. The culture incubated in an unchanged (880nm) light environment was full of
granule-containing cells (Figure 16A-B). In contrast, the culture incubated at a slightly shorter
wavelength maximum (870nm) had almost no granule-containing organisms (Figure 16C-D) while the
culture grown at a somewhat longer wavelength maximum (900nm) contained a fairly even mix of
organisms with and without internal granules (Figure 16E-F). In addition, the cell density of the 900nm
culture was much less than in the other cultures.
19
(A)
880nm Culture
(B)
880nm Culture
(C)
870nm Culture
(D)
870nm Culture
(E)
900nm Culture
(F)
900nm Culture
Figure 16 - Microscope images of the light environment experiment cultures after 4 days incubation under
slightly different restricted light conditions. All images were taken at 100x under differential interference
contrast (DIC).
20
Pigment Absorbance
Pigment absorbance readings were very successful and peaks matched up to expected locations for Bchl a
and carotenoids. Due to inconsistencies between different dilutions of the same sample, it was not
possible to compare the in vivo spectra in a quantitative fashion (Figure 17). The in vitro (extracted)
pigments, on the other hand, appeared to give semi-quantitative results and differences in in vitro pigment
peaks of the different incubation cultures were significant, with the 870nm culture showing an
exaggerated carotenoids peak and the 900nm culture showing depressed Bchl a peaks compared to the
880nm culture (Figure 18).
Figure 17 - In vivo pigment absorbance spectrum from cultures after 4 days of incubation in slightly different
light environments.
Figure 18 - In vitro pigment absorbance spectrum of pigments extracted in 7:2 Acetone/Methanol from
cultures after 4 days of incubation in slightly different light environments.
21
TRFLP
TRFLP results for the initial 880 culture (not shown) and the culture after four days of incubation were
significantly different, but this could be due in part to the different DNA extraction procedures used. The
TRFLP results for all three cultures after four incubation days are shown in Figures 19-20 below. Most of
the major peaks are present in all samples, but their intensities vary significantly.
Figure 19 - TRFLP Electropherogram showing bands representing different community members.
Figure 20 - TRFLP Fragment abundance peakmap.
22
Discussion
After only four days of incubation in very slightly shifted light environments, significant differences were
seen between cultures in all four of the methods used to differentiate them (macroscopic observation,
microscopy, pigment analysis, and TRFLP).
Cultures moved to a slightly shorter peak wavelength (870nm) appeared to undergo a very significant
community and pigment shift, losing nearly all granule-containing cells (Figure 16C-D) compared to the
culture kept in the light environment (880nm) in which the original culture was enriched (Figure 16A-B).
This may correspond with the significant TRFLP peak drop at ~470 (Figure 20). The pigment results
show a very significantly increased carotenoid absorption peak (Figure 18) which does not appear to
correspond with any relative increase in the presence of a community member (as assessed by the TRFLP
results—no major peak appears much larger in the 870nm culture than in the other two cultures). This
may suggest that one or several community members are not increasing in abundance but are instead
increasing their number of carotenoids pigments. The deep red-brown color visible in the liquid culture
(Figure 15) is also indicative of an increased carotenoid abundance. 870nm is well within the range of the
Bchl a absorbance spectrum and is far outside the absorbance range of carotenoids (Figure 11), so why
this wavelength would select for this behavior is a mystery.
At longer peak wavelengths, a significant difference is also apparent with all of the results (settled cell
material in liquid culture, significantly decreased cell density in microscopy, decreases in all peak heights
in pigment absorbance and TRFLP) indicating a loss of cell density and a corresponding loss in pigment
density. The biomass seen settled to the bottom of the culturing bottle may be due to cell death, however
viable cell counts to test this hypothesis were not performed. Reduced cell activity or survival at 900nm
(compared to 880nm) is not surprising, as a shift to longer wavelengths pushes the light spectrum to the
far shoulder of the Bchl a Qy absorbance peak (Figure 11). Unlike shorter wavelengths, whose energy
can be harvested by accessory pigments in the antenna complex and transferred (with energy loss along
the way) to the reaction center pigments, any light with wavelengths longer than the Q y absorbance band
for an organism cannot be utilized the cell.
Suggestions for future work
One caveat of this work is that I was not able to accurately control light intensity, and it is possible that
the results discussed above are the result of light intensity differences instead of or in addition to the
varied wavelength. LED light intensity can be controlled by varying the current through (by varying the
voltage across) the LED, and current to power output charts are usually available for LEDs. The LEDs
that I purchased, however, did not have available datasheets. Another way of determining light output is
to use a spectroradiometer, and this would be a good thing to do for future experiments.
It was unfortunate that I only had 4 days to run this experiment as it would have been interesting to see if
the trends continued over time and if putting the cultures under even more shifted wavelengths would
amplify the trends seen or give entirely new behavior. Any students who would wish to try this project in
the future should order lights well in advance of when they wish to start their project or, if possible, order
from a more local source (at the time of purchase, Roithner LaserTechnik was the only supplier I could
find with the assortment of LEDs that I needed, but other suppliers may exist).
Also, it would be interesting to work with a pure culture as well as a mix of a few defined organisms to be
better able to distinguish between changes in pigment makeup of individuals and community shifts. Our
class did not produce any pure cultures from our phototroph enrichments, which is why I did not attempt
this.
23
Spatial Makeup of the Purple Berries
Background
The purple “berries” of the Sippewissett are distinctive, purple- or pink-colored macroscopic aggregates
of microorganisms found in the shallow muddy ponds of the salt marshes. The berries have been
extensively studied by students in the MBL Microbial Diversity course, yet their composition and origin
remains somewhat of a mystery. I was interested in investigating the heterogeneity in the composition of
the berry (as suggested by the spatial concentration of Cytophaga and Flavobacteria on the exterior of the
berry from FISH staining of cross-sectioned berries done by Jarrod Scott in 2007; [13]) and investigating
whether or not chemical (particularly sulfur) cycling exists within the berries.
Figure 21 - Purple aggregates from the Little Sippewissett salt marsh. Scales are approximate.
Method
Purple berries were collected from various shallow ponds within the Little Sippewissett salt marsh during
low tide. Once collected, several methods were attempted to preserve, cut, and mount the berries, all of
which met with varying levels of disaster. The key variations of these methods are summarized below:
Method 1: Fix, wash, freeze, mount – Didn’t work
Fixation & Rinse
Whole berries were picked out of the sampled sediment and water and placed into a tube with 1% PFA in
PBS buffer and fixed at 4⁰C overnight. Berries were then picked out of the fixing tube and transferred
sequentially into 1xPBS buffer (10 min), fresh 1xPBS buffer (10 min), 50% EtOH in PBS buffer (10
min), 70% EtOH in PBS buffer (12 min), and 100% EtOH in PBS buffer (25 min). After the EtOH wash
steps, the berries had lost their purple color.
Freezing & Sectioning
The berries were then put into a mold filled with a cryo-embedding medium (TissueTek 4583 O.C.T.
Compound, Sakura Finetek USA, Inc., Torrance, CA) and frozen at -80⁰C for 1 hour. Within two
minutes of being transferred into the embedding medium, the berries turned purple again.
24
An attempt was then made to make cross sections of the berries using a cryotome associated with the
MBL Central Microsopy Facility. However, the berries did not seem to be properly frozen (were gellylike and didn’t cut well), and a technician suggested that this could be due to the ethanol wash step
inhibiting the uptake of the cryo-embedding medium. In addition, the thin sections being cut (10-15μm)
wrinkled immediately, preventing the attachment of flat sections to slides.
Method 2: Freeze, mount, fix, wash – Worked (sort of)
Slide Preparation
Two types of slides were used for mounting cross-sectioned samples: Fischer SuperFrost® Excell precoated slides and normal glass slides hand-coated in a 0.1% gelatin, 0.01% Cr(III)KSO4 in mqH2O
solution heated to 65⁰C (and then allowed to dry for several days). The hand-coated slides appeared to
have fewer problems with material floating off of the slides during washes.
Freezing & Sectioning
For the second attempt, molds to hold individual berries were constructed by sawing off the tapered
bottoms of 2mL microcentrifuge tubes and filling the inverted tubes (caps at the base, sawed open end as
the open top) with cryo-embedding medium (Figure 22). Fresh berries were placed in individual tube
molds and covered in embedding medium. The berries in embedding medium were then incubated at 4⁰C
for 8 hours to allow the solution to penetrate the berries. After this, the berries were frozen at 20⁰C
overnight. This seemed to work well, as the berries now cut nicely with the cryotome. To cut the berries,
the frozen berry was pushed out of the mold by opening up the cap of the mold and using the tapered end
of another cryotube to push the berry out the cap side of the tube. The berry was then cut in sections from
top (cap end) to bottom (open tube end). 20-25 μm sections seemed to be the best thickness for cutting
being thick enough to avoid wrinkling problems while still thin enough to minimize the problem of
material floating off of the slides during rinse steps.
Mounting Sections
The method that I found to work best for attaching sections to the slides was to cut slices with the
cryotome’s glass roll guard down, quickly raise the roll guard and use two points of a forceps to hold the
slice flat, and then allow the edges of the slice to curl around the ends of the forceps, after which the
sample could be picked up and quickly placed onto a warm slide. In this way, I was able to prepare very
nice thin section slides of the berries (Figure 23).
Fixation & Rinse
The following fixation and wash steps were done by gentle pipetting of the medium onto the slides to
prevent samples from floating off of the slides. When I tried dunking the slides into falcon tubes of wash
medium, the samples detached from the slides.
Berries on the slides were then fixed by adding 2mL of 1% PFA in PBS to the tops of the slides and
allowing it to soak for 1 hour at room temperature. Slides were then washed (again by gentle pipetting) in
1xPBS (5 min), fresh 1xPBS (5 min), mqH2O (5 min), and 100% EtOH (5 min). Following the ethanol
rinse, I shook off the excess ethanol and let the slides air-dry.
25
Figure 23 - Thin section slides prepared of the
purple berries.
Figure 22 - Berries in embedding solution in the cut
microcentrifuce tube molds.
Permeabilization & Rinse
Berry sections on the dried slide were circled using a special hydrophobic pen (Liquid Blocker Super Pap
Pen, Daido Sangyo Co., Ltd., Tokyo, Japan) to contain individual reactions on the slides (Figure 24).
Sections were then permeablized in ~20μL/well lysozyme medium (20 mg/mL Lysozyme, 50mM
EDTA, and 100mM TRIS-HCl in 1xPBS) for 1 hour. Lysozyme was shaken off of the slide and slides
were then soaked in mqH2O for 5 min, then 30μL 0.01M HCl for 10 min. HCl was rinsed off by
repeating the post-fixation rinse steps.
Hybridization & Wash
Hybridization buffer of the appropriate formamide concentration for each attempted probe was prepared
and warmed to 38⁰C water bath. 1μL of probe was mixed with 99μL hybridization buffer in separate
PCR tubes. 20μL of hybridization buffer + probe mix was added to each well on the slide and the slides
were carefully placed in 50mL Falcon tubes with a half-Kimwipe wetted with the appropriate
concentration of formamide (e.g. 35% formamide in H2O for a 35% formamide hybridization buffer);
tubes were capped and placed in a 46⁰C incubator for 3 hours.
I learned the hard way that attempting to do different probes on a single slide is a bad idea as slides are
easily ruined by probes running together during the hybridization incubation). I attempted to fix this by
soaking the slide in 0.1M HCl for 10 minutes to deactivate the probes . Slides were then rinsed in 1xPBS
(1 min), fresh 1xPBS (8 min), mqH2O (2 min), and 100% EtOH (2 min) and allowed to dry.
Hybridization was repeated from this point.
After successful hybridization, slides were dipped in a wash buffer (5mM EDTA, 20mM TRIS-HCl, and
70mM NaCl in water for a 35% formamide hybridization; recipe is different for different formamide
concentrations) that was preheated to 46⁰C in a water bath and then soaked for 15 minutes in a fresh wash
buffer.
26
Amplification & Wash
The following steps were done in dim conditions to minimize light exposure of the fluorophore. Berries
were incubated with 10μL of amplification buffer with Alexa 594 tyramide Alexa 594 added to each well
at 46⁰C for 40 minutes. Slides were then dipped in 1xPBS then soaked in fresh 1xPBS (5 min) and
mqH2O (2 min), then dipped in100% EtOH and allowed to air dry in a dark box and kept at -20⁰C.
Figure 24 - Individual reactions contained by marked wells around each berry cross section on a slide.
Method 3: Give in and just do the normal filter procedure – Worked (sort of)
While struggling with getting methods to work with the slide-mounted berry cross sections, I decided to
do a “back up” FISH reaction using filters that had been prepared by Alex Petroff (and used in his miniproject) from mashed up purple berries. This was done in order to screen for probes of particular interest
for use on the few slides that remained after all of the problems with Methods 2 & 3. A standard FISH
procedure was followed for filtered samples that is probably described in someone else’s report for their
mini-project this year. I used the following probes: Non338 (35% formamide), EubI-III (35%
formamide), Alf968 (35% formamide), Bet42a (with compliment; 35% formamide), Gam42a (with
compliment; 35% formamide), Delta495 a-c (with compliment; 35% formamide), Dbb660 (with
compliment; 60% formamide), Dss658 (60% formamide), Dsb985 (20% formamide), Dsr651 (35%
formamide), and Dsv689 (with compliment; 40% formamide). During the procedure, the following
mistakes were made that may have impacted the quality of the staining: cells were permeablized for 1.5
hours (rather than 1 hour) and later rinsed with HCl for 20 minutes (rather than 10 min), and the
amplification incubation was run for 27 (rather than 20) minutes.
DNA Extraction
I also attempted to extract DNA from the center and outside of a berry. I did this by slicing off the top
~1/3 of a large berry using the microtome and then using a 10μL pipette tip to stab out a small core of
material from different locations of the berry. Each sample of material was suspended in 20μL NP40 and
boiled using a five-minute boil program on a thermocycler. PCR with 16S 8F and 1492R primers was
performed, but amplification did not appear to be successful and I gave up after several attempts to
amplify DNA extracted from coring of frozen berries. It is possible that the TissueTek cryo-embedding
solution interferes with PCR.
Results & Discussion
CARD-FISH on the berry filters met with moderate success, due in part to insufficient mounting of the
filters to the slides with DAPI. Some slides did turn out well, and it does appear that sulfate reducing
bacteria are present within the berries (Figure 25B,C,F), although autofluorescence (e.g. Figure 25D) may
be confusing the results.
27
(A)
Alf968 (α-Proteos)
(B)
Dsb968 (Desulfovibrio)
(C)
Dsb985 (Desulfobacter?)
(D)
Db985 (Autofluorescence?)
(E)
Dsb660 (Desulfobulbus)
(F)
Dss658 (Desulfosarcina)
Figure 25 - Selected results of CARD-FISH on mashed berry filters.
28
The Desulfobacter (Dsb985) probe was chosen for staining of the few remaining berry cross section slides
that weren’t destroyed from previous, more ambitious (and totally unsuccessful) CARD-FISH attempts.
Comparing the Dsb985 results to the Non338 results suggests that a significant population of
Desulfobacter lives in the berries and that they are concentrated on the exterior of the berry (Figure 26).
This is interesting, because the microelectrode work by Alex Petroff this summer suggests that sulfide
may be being produced in the interior of the berry, so the interior is where one would expect to find the
Desulfobacteria, not the exterior.
So who is in the middle and what are they doing? Hopefully future students will be able use tips
contained in this project report to do a more effective job of investigating the chemical cycling and spatial
makeup of these interesting aggregates.
(A)
Non338 (Nonsense Probe)
(B)
Figure 26 - Selected results of CARD-FISH on berry cross-sections.
29
Dsb985 (Desulfobacter)
Acknowledgements
In addition to generally acknowledging all of the students and staff of the 2009 Microbial Diversity
Course for a tremendous summer and for moral support and help throughout, I would like to acknowledge
the following people specifically for their assistance with this project:

Instructors Steve Zinder, Dan Buckley, Victoria Orphan, and Bill Metcalf for valuable
discussions and advice.

TAs Alexa Price-Whelan (gas chromatography), Heather Fullerton (LED ordering madness),
Cristina Moraru (FISH), and Hinsby Cadillo-Quiroz (PCR & TRFLP).

Student Alex Petroff for help, discussions, and much-needed sanity checks during the often
extremely frustrating work on the berries. Thanks are also due to students Shannon Tronic, Jan
Blees, and Orr Shapiro for always taking time out to show me how to do things when I didn’t
know what I was doing.
I would not have been able to attend this course without the generous financial support and backing of my
advisor, Ken Nealson, as well as the Planetary Biology Internship Scholarship and the Holger &
Friederun Jannasch Scholarship in Microbial Diversity distributed by the Marine Biological Laboratory.
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31