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 side-project 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 oC 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 oC 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 oC. 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), 474-481. 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 CARD-FISH 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.