Download Szoboszlay, M. Protists vs. purple bacteria

Protists vs. purple bacteria
Márton Szoboszlay
MBL Microbial Diversity course 2014
Abstract
My goal with this project was to learn more about protists. And obtain experience in enriching
and isolating portists. I was able to obtain amoebae that grow anaerobicaly on purple bacterial
lawns. However I could not prove that the amoebae are grazing the purple bacteria. As a sideproject I studied the microbiota of the termite gut with fluorescent in situ hybridization by
cryostat sectioning the intestinal tract of workers and a soldier of Reticulitermes flavipes
(appendix 2)
Introduction
The surface layer of marine coastal sediments in salt marshes can harbor a stratified community
of phototrophic microorganisms. The topmost layer is green due to the abundance of oxygenic
phototrophs. Beneath this a purple, or peach colored layer is found dominated by phototrophic
purple bacteria. The next layer is usually dark green and rich in green sulfur bacteria. Below the
colored layers the sediment is generally black. The sulfate received from the marine water is
utilized by anaerobic sulfate reducing bacteria in the sediment resulting in sulfide production.
The sulfide that moves upwards in the sediment serves as an electron donor for the
photosynthesis of green sulfur bacteria and purple bacteria that occupy layers that are close
enough to the surface to receive sufficient light, but still anoxic. These bacteria are able to form
intra- or extracellular sulfur granules.
There isn’t much information available on the protist community in these sediment. It is known
however that protists, including amoebae are present in marine sediments where they graze on
bacteria (Anderson et al. 2001, First and Hollibaugh 2008). It seems likely that the phototrophic
green and purple bacteria have protist predators, which leads to the question of what happens
with the intracellular sulfur granules of these bacteria once they are engulfed by these protists.
The granules may dissolve in the lysosomes and convert into other sulfur species. If this is not the
case, the predator protist may accumulate the sulfur intracellularly or deposit it via exocytosis. If
the sulfur is not deposited shortly after the consumption of the pray bacteria, considering the
motility of protists and the thin, stratified structure of the surface sediment, these protists could
transport sulfur within layers contributing to the sulfur cycling in the ecosystem.
To investigate this I attempted to isolate protists from salt marsh sediments capable of grazing on
purple sulfur bacteria and observe the fate of sulfur granules within these protists. First I isolated
anoxygenic phototroph bacteria and anaerobic heterotroph bacteria from salt marsh sediments
and microcosms established form salt marsh sediment samples, then used these isolates as pray
organisms to cultivate protists from the same microcosms. Protists forming plaques on lawns of
the pray organism were observed in light and scanning electron microscopy.
Methods
Isolating anoxygenic phototroph bacteria
Marine sulfur-phototroph and marine acetate-degrading nonsulfur-phototroph liquid media were
used to enrich anoxygenic phototroph bacteria from sediment samples from Trunk River and
Little Sippewissett salt marsh. Enrichments were incubated at room T in closed cabinets equipped
with LED lights emitting 631 nm (to enrich green sulfur bacteria) or 850 nm (to enrich purple
bacteria) light. Once turbidity and green / purple color developed, aliquots from the enrichments
were plated to thiosulfate-acetate agar plates under anaerobic conditions. Single colonies were
transferred until pure isolates were obtained, then transferred to 10 ml of anaerobic liquid media
again (figure 1). Agar plates and liquid media tubes were handled in an anaerobic chamber and
incubated in anaerobic jars at room T on the sill of an east-facing window.
Figure 1: Purple phototroph bacterial strains in liquid culture.
Green bacteria did not grow fast on plates, thus pure cultures could not be obtained within the
timeframe of this project. Two strains of purple bacteria were selected for the subsequent
experiments based on their ability to grow fast and form even lawns on thiosulfate-acetate agar
plates under anaerobic conditions. These strains originated from sediment samples from Little
Sippewissett salt marsh.
Salt marsh sediment microcosms
Two microcosms from sediment samples from Little Sippewissett salt marsh were set up by Kurt
Hanselmann. One was from green colored, the other from purple colored sediment (figure 2). The
sediment samples were placed on plastic trays and covered with seawater. The trays were kept at
room T under light, halfway covered with aluminum foil. The water in the trays was not replaced.
Samples were taken regularly from the sediment and the overlying water in these microcosms
and examined with phase contrast microscopy.
Figure 2: Salt marsh sediment microcosms after 37 days of incubation. Left: from green sediment; right: from purple
sediment.
I chose to use these microcosms in subsequent experiments instead of fresh sediment samples
because they become enriched in eukaryotic microorganisms over time.
Isolating anaerobic heterotrophic bacteria
Protists that are able to graze on purple bacteria may grow better on other pray organisms, thus
their isolation could be easier using other anaerobic bacteria. To obtain anaerobic bacterial
isolates ~ 1 ml sediment samples were taken with pipet tips to 1.5 ml eppendorf tubes from the
salt marsh sediment microcosms 21 days after they were established. The headspace in the tubes
was completely filled with water from the microcosms. The samples were brought into an
anaerobic chamber, shaken and diluted 10x and 100x with anoxic distilled water. 100 µl aliquots
from the dilutions and the undiluted samples were plated on anoxic SWC and LB plates. After
incubation single colonies were transferred to 10 ml anaerobic liquid SWC or LB media. Once
the cultures become turbid aliquots were plated on SWC or LB plates and single colonies were
reisolated in 10 ml anaerobic liquid SWC or LB media again. All cultures were incubated at 30
o
C and handled in the anaerobic chamber. Plates were incubated in anaerobic jars.
Two isolates, one growing on LB and one on SWC were chosen for the subsequent experiments
based on their ability to grow fast. These were obtained from the microcosm from purple colored
salt marsh sediment.
Sequencing the 16S gene of the selected isolates
16S sequencing was used to identify the selected two purple bacteria and two anaerobic
heterotroph strains. DNA was isolated from 100 µl aliquots of liquid cultures by incubation at 98
o
C for 5 minutes in a thermocycler. For the PCR 2 µl of the DNA extract was mixed with 12.5 µl
Promega Go-Taq Green 2X mix, 2 µl (15 pmol) 27f primer, 2 µl (15 pmol) 1492r primer and 6.5
µl nuclease-free water. The temperature profile of the reaction was 95 oC for 2 minutes followed
by 30 cycles of 95 oC for 30 seconds, 55 oC for 30 seconds and 72 oC for 90 seconds. The
reaction was finished with 7 minutes final extension at 72 oC. The PCR products were stored at 4
o
C. To check for successful amplification of the 16S gene, 5 µl aliquots were run in a 1% agarose
gel and visualized by Sybr Safe staining. PCR products were cleaned with the Promega Wizard
PCR Preps DNA purification system and submitted for sequencing.
Isolating predator protists
The salt marsh sediment microcosms were sampled for protist isolation 30 and 32 days after they
were established. About 1 ml sediment samples were collected with pipet tips in 1.5 ml eppendorf
tubes. The headspace in the tubes was completely filled with water from the microcosms. The
samples were brought into an anaerobic chamber, shaken and 100 µl aliquots were mixed with
100 µl of the liquid culture of the pray bacterium. These mixed cultures were diluted 10x and
100x in the liquid culture of the pray bacterium. 100 µl aliquots of the mixed cultures and their
dilutions were plated on anoxic thiosulfate-acetate agar plates when purple bacteria were used as
pray, or SWC and LB when the anaerobic heterotrophs were used. The plates were incubated at
room T on the sill of an east-facing window or at 30 oC in the dark respective to the pray
organism in anaerobic jars. Wet-mounts for phase contrast microscopy were prepared from
plaques daily as they appeared by gently picking material from the edge of the plaques with a
pipet tip and suspending the material in sterile anoxic distilled water.
Scanning electron microscopy (SEM)
Scanning electron microscopy coupled with elemental analysis by energy dispersive x-ray
spectroscopy (EDS) was used to detect sulfur granules inside protist. Agar cylinders with
approximately 2 mm diameter containing a plaque were cut out of the protist isolation plates and
fixed in 100 µl drops of 2.5 % glutaraldehyde in phosphate buffer solution at room T for 80
minutes. The top parts of the agar cylinders that contained the plaque surface were cut off with a
razor blade under a dissecting microscope and the rest of the agar cylinders were discarded. The
obtained agar discs were either glued directly to an aluminum sample holder or first stained in a
drop of 2 % silver nitrate for 10 minutes followed by two washes with distilled water. The
samples were examined in a Hitachi TM3030 bench top scanning electron microscope capable of
energy dispersive x-ray spectroscopy.
Results and discussion
16S sequencing
One of the purple bacterium strains used in the protist isolation experiment was identified to be
Rhodovulum sp. while 16S sequencing was not successful from the other. The anaerobic
heterotroph growing on LB medium was classified as Lucibacterium sp. 16S sequencing from the
one growing on SWC gave inconclusive results.
Protist isolation from the salt marsh sediment microcosms
The anaerobic heterotroph bacteria didn’t produce even lawns on the anoxic SWC and LB plates
and yielded no plaques. This could be due to the quality of the lawn, which could make the
detection of the plaques difficult, or the lack of time as the timeframe of this project only allowed
4 days of incubation. An other possibility is that the SWC and LB plates don’t provide a
favorable environment for the protists present in the samples. Considering that the pray bacteria
were isolated from the same salt marsh sediment microcosms it seems unlikely that there were no
portist in the samples that could feed on them.
The lawns of purple bacteria first developed visible plaques after 3 days of incubation and new
plaques emerged continuously until the 6th day when the experiment had to be terminated (figure
3). Plates from the undiluted samples yielded numerous plaques while the ones from the 10x and
100x dilutions developed zero to five.
Figure 3: Plaques on purple bacterial lawns after 6 days of incubation.
This method of isolating protists has its caveats: Beside protist grazing, plaque formation can be a
result of the presence of bacteriophages, which are too small to be detected with the conventional
light microscope. Antagonism with various microorganisms present in the sample could inhibit
the growth of the purple bacteria resulting in a plaque too. Also, if the grazing and growth of the
predator protist is significantly slower than the growth of the pray bacterium, it will not be able to
develop a visible plaque. Using agar plates selects for protists capable of grazing on a solid
surface, which are primarily amoebae. I assume however that the majority of predator protists in
salt marsh sediments are grazing on a surface considering the large surface area of the sand
particles that make up the sediment’s matrix and the small diameter of the pores in between them.
Phase contrast microscopy of plaques
Beside the purple bacteria, I was able to find various bacterial morphotypes. Long rods were
abundant in all examined plaques and usually formed a coating layer around the purple bacterial
aggregates in the wet mounts (figure 4).
Figure 4: Phase contrast micrograph of various bacterial morphotypes observed in the plaques on purple bacterial
lawns. Purple bacteria tended to form spherical aggregates often coated by long rods. Arrows: sulfur granule in a
purple bacterium.
Amoebae were found in several, but not all plaques (figure 5). They were five to ten µm in size.
Other types of eukaryotic microorganisms were not found. None of the amoebae showed any
activity, thus I couldn’t observe their grazing. The wet mounts were prepared in the anaerobic
chamber, but the slides had to be removed from the chamber for microscopy, which exposed the
cells to oxygen. Carefully sealing the wet mount with nail polish could have preserved the anoxic
conditions in the sample and enabled the observation of the activity of the amoebae.
Figure 5: Phase contrast micrograph of amoebae found in a plaque on a purple bacterial lawn.
Due to the lack of activity of the amoebae and the other bacterial morphotypes present in every
examined plaque beside the purple bacterium I can’t conclude that the amoebae grazed on the
purple phototroph. Several transfers of the amoebae with the culture of the purple bacterium
would be necessary until the other bacteria are diluted out. Some amoebae however contained
granules showing up as bright spots in the phase contrast microscope that could be sulfur
granules from the purple bacteria (figure 6).
Figure 6: Phase contrast micrograph of amoebae found in plaques on a purple bacterial lawn. Arrows indicate bright
spots that could be sulfur granules.
Scanning electron microscopy
To verify the presence of sulfur granules in the amoebae I attempted to use SEM with EDS, but
without success. The vacuum in the SEM dehydrated the sample leading to the precipitation of
sodium and calcium salts and disruption of the cells. No amoeba or bacterial cell was found in the
samples. The silver nitrate treatment didn’t stain the sample, but led to the precipitation of silver
containing white crystals over the entire surface of the samples. Unfortunately we didn’t have
access to better sample preparation methods for SEM like carbon or gold coating or osmium
tetroxide staining.
Acknowledgements
I owe thanks to the microbial diversity students and teaching team for their invaluable help and
advice especially Kurt Hanselmann, Arpita Bose, Scott Dawson and Emil Ruff who guided me
through this project.
References
Anderson, O. R., Gorrell, T., Bergen, A., Kruzansky, R., & Levandowsky, M. (2001). Naked
amoebas and bacteria in an oil-impacted salt marsh community. Microbial Ecology, 42(3), 474481.
First, M. R., & Hollibaugh, J. T. (2008). Protistan bacterivory and benthic microbial biomass in
an intertidal creek mudflat. MARINE ECOLOGY-PROGRESS SERIES-, 361, 59.
Appendix 1 – media recipes
Marine sulfur-phototroph liquid media
360 ml sea water base solution
342.2 mM NaCl
14.8 mM MgCl2
1.0 mM CaCl2
6.71 mM KCl
2 ml 1 M NH4Cl
4 ml 100 mM K phosphate buffer pH 7.2
4.0 g/l KH2PO4
12.7 g/l K2HPO4
2 ml 1 M MOPS buffer pH 7.2
0.4 ml trace element solution
20 mM HCl
7.5 mM FeSO4
0.48 mM H3BO3
0.5 mM MnCl2
6.8 mM CoCl2
1.0 mM NiCl2
12 µM CuCl2
0.5 mM ZnSO4
0.15 mM Na2MoO4
25 µM NaVO3
9 µM Na2WO4
23 µM Na2SeO3
flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclaving
the following ingredients were added during flushing with N2 / CO2 gas
4 ml sterile multivitamin solution
10 mM MOPS buffer pH 7.2
0.1 g/l riboflavin
0.03 g/l biotin
0.1 g/l thiamine HCl
0.1 g/l L-ascorbic acid
0.1 g/l d-Ca-pnathotenate
0.1 g/l folic acid
0.1 g/l nicotinic acid
0.1 g/l 4-aminobenzoic acid
0.1 g/l pyridoxine HCl
0.1 g/l lipoic acid
0.1 g/l NAD
0.1 g/l thiamine pyrophosphate
0.01 g/l cyanocobalamin
28 ml sterile 1 M NaHCO3
4 ml sterile 1 M Na2S2O3
0.4 ml sterile 0.2 M Na2S added in the anaerobic chamber
Marine acetate-degrading nonsulfur-phototroph liquid media
390 ml sea water base solution
0.68 g sodium acetate
2 ml 1 M NH4Cl
0.1 ml 1 M Na2SO4
4 ml 100 mM K phosphate pH 7.2
0.5 ml 1 M MOPS buffer pH 7.2
0.4 ml trace element solution
flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclaving
the following ingredients were added during flushing with N2 / CO2 gas
4 ml sterile multivitamin solution
1 ml sterile 1 M NaHCO3
Thiosulfate-acetate agar
1 l sea water base solution
5 ml 1 M NH4Cl
10 ml 100 mM K phosphate pH 7.2
5 ml 1 M MOPS buffer pH 7.2
1 ml trace element solution
0.25 ml 1 M Na2SO4
1.75 g sodium acetate
17 g agar washed in distilled water
flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclaving
the following ingredients were added during flushing with N2 / CO2 gas
10 ml sterile multivitamin solution
70 ml sterile 1 M NaHCO3
10 ml sterile 1 M Na2S2O3
Anoxic LB
1 l distilled water
10 g tryptone
5 g yeast extract
10 g NaCl
15 g agar
set pH to 7.0
flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclaving
Anoxic SWC
1 l sea water base solution
5 g tryptone
1 g yeast extract
3 ml glycerol
15 g agar
set pH to 7.0
flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclaving
Appendix 2 – Detection of Bacteria and Archaea by fluorescent in situ hybridization (FISH)
in the termite hindgut
The hindgut of wood-eating termites is rich in protists, Bacteria and Archaea. Bacteria are present
not only as free living members of the termite hindgut microbiota but also as exo- and
endosymbionts of protists. The diet of wood-eating termites is low in nitrogen, and N2 fixing
bacteria living as endosymbionts in protists are abundant in the hindgut (Breznak and Pankratz
1977, Inoue et al. 2008, Noda et al. 2005). The goal of this project was to investigate the
localization of nitrogenase activity in the hindgut of Reticulitermes flavipes.
The intestinal tract of workers and one soldier of Reticulitermes flavipes was dissected out and
directly, or after 50 minutes of fixation at room T in a drop of 3.2 % formamide, embedded in
Tissue-Tek resin, frozen in liquid nitrogen and stored at -20 oC. The frozen blocks were cut to 20
µm thick slices with a microtome in a cryostat. The slices were collected on sterile 0.2 µm GTTP
filter membranes. The membranes were sprayed with 0.2 % low melting point agarose, dried at
room T and stored at -20 oC. The filter membranes were not transparent which prevented the
examination of the samples with transmission light microscopy, but it was much easier to handle
them than glass slides during the FISH procedure, also the majority of slices collected on regular
or poly-L-lysine coated glass slides were lost during the washing in the FISH protocol.
First I stained sections of the membranes with DAPI, which was successful. I attempted CARDFISH with the general bacterial and archaeal probes, but almost all the termite gut sections were
lost from the membranes. Most likely the permeabilization and H2O2 treatments damaged the
insect tissue or the samples were lost during the many washing steps in the protocol. I tried
mono-FISH using the general bacterial and archaeal probes again, which worked, but the
fluorescent signal was often hard to see. Parts of the gut content bound DAPI and the fluorescent
probes resulting in a foggy background. This could likely be circumvented by slicing the samples
to thinner sections. I also observed chitin autofluorescence. As the next step I was planning to do
hybridization chain reaction with NifH specific probes on the membranes to localize NifH
expression in the termite gut, but eventually didn’t have the time.
Figure 7: Section of the gut of a termite worker stained with DAPI. This section is most likely from the midgut
because the gut wall is thin and the lumen is empty.
Figure 8: Section of the hindgut of a termite worker stained with DAPI. The lumen is full with “foggy” content and
also bacteria and nuclei of protists.
Figure 9: Higher magnification of the gut lumen (same gut section as on the previous figure). Spirochetes and other
bacteria are present in high numbers.
Figure 10: Section of the gut of a termite soldier stained with DAPI. This is likely the fore-gut because the lumen is
empty and the gut wall is thick.
Figure 11: Mono-FISH general bacterial probe signal from an area close to the gut wall in the hindgut of a worker
termite.
Figure 12: Mono-FISH general bacterial probe signal from the gut wall in the hindgut of a worker termite. Many
bacteria appear to be within the insect tissue. This was only observed in some regions and I didn’t find any indication
in the literature that bacteria would be penetrating the insect tissue, therefore it is likely an artifact due to the
sectioning of the sample.
References:
Breznak, J. A., & Pankratz, H. S. (1977). In situ morphology of the gut microbiota of
wood-eating termites [Reticulitermes flavipes (Kollar) and Coptotermes formosanus
Shiraki]. Applied and environmental microbiology, 33(2), 406.
Inoue, J. I., Noda, S., Hongoh, Y., Ui, S., & Ohkuma, M. (2008). Identification of
endosymbiotic methanogen and ectosymbiotic spirochetes of gut protists of the termite
Coptotermes formosanus. Microbes and Environments, 23(1), 94-97.
Noda, S., Iida, T., Kitade, O., Nakajima, H., Kudo, T., & Ohkuma, M. (2005).
Endosymbiotic Bacteroidales bacteria of the flagellated protist Pseudotrichonympha
grassii in the gut of the termite Coptotermes formosanus. Applied and environmental
microbiology, 71(12), 8811-8817.