Protists Vs. Purple Bacteria - MBL
Protists vs. purple bacteriaMárton SzoboszlayMBL Microbial Diversity course 2014AbstractMy goal with this project was to learn more about protists. And obtain experience in enrichingand isolating portists. I was able to obtain amoebae that grow anaerobicaly on purple bacteriallawns. 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 bycryostat sectioning the intestinal tract of workers and a soldier of Reticulitermes flavipes(appendix 2)IntroductionThe surface layer of marine coastal sediments in salt marshes can harbor a stratified communityof phototrophic microorganisms. The topmost layer is green due to the abundance of oxygenicphototrophs. Beneath this a purple, or peach colored layer is found dominated by phototrophicpurple bacteria. The next layer is usually dark green and rich in green sulfur bacteria. Below thecolored layers the sediment is generally black. The sulfate received from the marine water isutilized 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 thephotosynthesis of green sulfur bacteria and purple bacteria that occupy layers that are closeenough to the surface to receive sufficient light, but still anoxic. These bacteria are able to formintra- or extracellular sulfur granules.There isn’t much information available on the protist community in these sediment. It is knownhowever that protists, including amoebae are present in marine sediments where they graze onbacteria (Anderson et al. 2001, First and Hollibaugh 2008). It seems likely that the phototrophicgreen and purple bacteria have protist predators, which leads to the question of what happenswith 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 thecase, the predator protist may accumulate the sulfur intracellularly or deposit it via exocytosis. Ifthe sulfur is not deposited shortly after the consumption of the pray bacteria, considering themotility of protists and the thin, stratified structure of the surface sediment, these protists couldtransport 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 onpurple sulfur bacteria and observe the fate of sulfur granules within these protists. First I isolatedanoxygenic phototroph bacteria and anaerobic heterotroph bacteria from salt marsh sedimentsand microcosms established form salt marsh sediment samples, then used these isolates as prayorganisms to cultivate protists from the same microcosms. Protists forming plaques on lawns ofthe pray organism were observed in light and scanning electron microscopy.MethodsIsolating anoxygenic phototroph bacteriaMarine sulfur-phototroph and marine acetate-degrading nonsulfur-phototroph liquid media wereused to enrich anoxygenic phototroph bacteria from sediment samples from Trunk River andLittle Sippewissett salt marsh. Enrichments were incubated at room T in closed cabinets equippedwith LED lights emitting 631 nm (to enrich green sulfur bacteria) or 850 nm (to enrich purplebacteria) light. Once turbidity and green / purple color developed, aliquots from the enrichmentswere plated to thiosulfate-acetate agar plates under anaerobic conditions. Single colonies weretransferred until pure isolates were obtained, then transferred to 10 ml of anaerobic liquid mediaagain (figure 1). Agar plates and liquid media tubes were handled in an anaerobic chamber andincubated 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 thetimeframe of this project. Two strains of purple bacteria were selected for the subsequentexperiments based on their ability to grow fast and form even lawns on thiosulfate-acetate agarplates under anaerobic conditions. These strains originated from sediment samples from LittleSippewissett salt marsh.
Salt marsh sediment microcosmsTwo microcosms from sediment samples from Little Sippewissett salt marsh were set up by KurtHanselmann. One was from green colored, the other from purple colored sediment (figure 2). Thesediment samples were placed on plastic trays and covered with seawater. The trays were kept atroom 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 microcosmsand examined with phase contrast microscopy.Figure 2: Salt marsh sediment microcosms after 37 days of incubation. Left: from green sediment; right: from purplesediment.I chose to use these microcosms in subsequent experiments instead of fresh sediment samplesbecause they become enriched in eukaryotic microorganisms over time.Isolating anaerobic heterotrophic bacteriaProtists that are able to graze on purple bacteria may grow better on other pray organisms, thustheir isolation could be easier using other anaerobic bacteria. To obtain anaerobic bacterialisolates 1 ml sediment samples were taken with pipet tips to 1.5 ml eppendorf tubes from thesalt marsh sediment microcosms 21 days after they were established. The headspace in the tubeswas completely filled with water from the microcosms. The samples were brought into ananaerobic chamber, shaken and diluted 10x and 100x with anoxic distilled water. 100 µl aliquotsfrom the dilutions and the undiluted samples were plated on anoxic SWC and LB plates. Afterincubation single colonies were transferred to 10 ml anaerobic liquid SWC or LB media. Oncethe cultures become turbid aliquots were plated on SWC or LB plates and single colonies werereisolated in 10 ml anaerobic liquid SWC or LB media again. All cultures were incubated at 30oC 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 experimentsbased on their ability to grow fast. These were obtained from the microcosm from purple coloredsalt marsh sediment.Sequencing the 16S gene of the selected isolates16S sequencing was used to identify the selected two purple bacteria and two anaerobicheterotroph strains. DNA was isolated from 100 µl aliquots of liquid cultures by incubation at 98oC for 5 minutes in a thermocycler. For the PCR 2 µl of the DNA extract was mixed with 12.5 µlPromega Go-Taq Green 2X mix, 2 µl (15 pmol) 27f primer, 2 µl (15 pmol) 1492r primer and 6.5µl nuclease-free water. The temperature profile of the reaction was 95 oC for 2 minutes followedby 30 cycles of 95 oC for 30 seconds, 55 oC for 30 seconds and 72 oC for 90 seconds. Thereaction was finished with 7 minutes final extension at 72 oC. The PCR products were stored at 4oC. To check for successful amplification of the 16S gene, 5 µl aliquots were run in a 1% agarosegel and visualized by Sybr Safe staining. PCR products were cleaned with the Promega WizardPCR Preps DNA purification system and submitted for sequencing.Isolating predator protistsThe salt marsh sediment microcosms were sampled for protist isolation 30 and 32 days after theywere established. About 1 ml sediment samples were collected with pipet tips in 1.5 ml eppendorftubes. The headspace in the tubes was completely filled with water from the microcosms. Thesamples were brought into an anaerobic chamber, shaken and 100 µl aliquots were mixed with100 µl of the liquid culture of the pray bacterium. These mixed cultures were diluted 10x and100x in the liquid culture of the pray bacterium. 100 µl aliquots of the mixed cultures and theirdilutions were plated on anoxic thiosulfate-acetate agar plates when purple bacteria were used aspray, or SWC and LB when the anaerobic heterotrophs were used. The plates were incubated atroom T on the sill of an east-facing window or at 30 oC in the dark respective to the prayorganism in anaerobic jars. Wet-mounts for phase contrast microscopy were prepared fromplaques daily as they appeared by gently picking material from the edge of the plaques with apipet 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-rayspectroscopy (EDS) was used to detect sulfur granules inside protist. Agar cylinders withapproximately 2 mm diameter containing a plaque were cut out of the protist isolation plates andfixed in 100 µl drops of 2.5 % glutaraldehyde in phosphate buffer solution at room T for 80minutes. 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. Theobtained agar discs were either glued directly to an aluminum sample holder or first stained in adrop of 2 % silver nitrate for 10 minutes followed by two washes with distilled water. Thesamples were examined in a Hitachi TM3030 bench top scanning electron microscope capable ofenergy dispersive x-ray spectroscopy.Results and discussion16S sequencingOne of the purple bacterium strains used in the protist isolation experiment was identified to beRhodovulum sp. while 16S sequencing was not successful from the other. The anaerobicheterotroph growing on LB medium was classified as Lucibacterium sp. 16S sequencing from theone growing on SWC gave inconclusive results.Protist isolation from the salt marsh sediment microcosmsThe anaerobic heterotroph bacteria didn’t produce even lawns on the anoxic SWC and LB platesand yielded no plaques. This could be due to the quality of the lawn, which could make thedetection of the plaques difficult, or the lack of time as the timeframe of this project only allowed4 days of incubation. An other possibility is that the SWC and LB plates don’t provide afavorable environment for the protists present in the samples. Considering that the pray bacteriawere isolated from the same salt marsh sediment microcosms it seems unlikely that there were noportist in the samples that could feed on them.The lawns of purple bacteria first developed visible plaques after 3 days of incubation and newplaques emerged continuously until the 6th day when the experiment had to be terminated (figure3). Plates from the undiluted samples yielded numerous plaques while the ones from the 10x and100x 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 aresult of the presence of bacteriophages, which are too small to be detected with the conventionallight microscope. Antagonism with various microorganisms present in the sample could inhibitthe growth of the purple bacteria resulting in a plaque too. Also, if the grazing and growth of thepredator protist is significantly slower than the growth of the pray bacterium, it will not be able todevelop a visible plaque. Using agar plates selects for protists capable of grazing on a solidsurface, which are primarily amoebae. I assume however that the majority of predator protists insalt marsh sediments are grazing on a surface considering the large surface area of the sandparticles that make up the sediment’s matrix and the small diameter of the pores in between them.Phase contrast microscopy of plaquesBeside the purple bacteria, I was able to find various bacterial morphotypes. Long rods wereabundant in all examined plaques and usually formed a coating layer around the purple bacterialaggregates in the wet mounts (figure 4).
Figure 4: Phase contrast micrograph of various bacterial morphotypes observed in the plaques on purple bacteriallawns. Purple bacteria tended to form spherical aggregates often coated by long rods. Arrows: sulfur granule in apurple 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 anyactivity, thus I couldn’t observe their grazing. The wet mounts were prepared in the anaerobicchamber, but the slides had to be removed from the chamber for microscopy, which exposed thecells to oxygen. Carefully sealing the wet mount with nail polish could have preserved the anoxicconditions 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 everyexamined plaque beside the purple bacterium I can’t conclude that the amoebae grazed on thepurple phototroph. Several transfers of the amoebae with the culture of the purple bacteriumwould be necessary until the other bacteria are diluted out. Some amoebae however containedgranules showing up as bright spots in the phase contrast microscope that could be sulfurgranules from the purple bacteria (figure 6).Figure 6: Phase contrast micrograph of amoebae found in plaques on a purple bacterial lawn. Arrows indicate brightspots that could be sulfur granules.Scanning electron microscopyTo verify the presence of sulfur granules in the amoebae I attempted to use SEM with EDS, butwithout success. The vacuum in the SEM dehydrated the sample leading to the precipitation ofsodium and calcium salts and disruption of the cells. No amoeba or bacterial cell was found in thesamples. The silver nitrate treatment didn’t stain the sample, but led to the precipitation of silvercontaining 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 osmiumtetroxide staining.AcknowledgementsI owe thanks to the microbial diversity students and teaching team for their invaluable help andadvice especially Kurt Hanselmann, Arpita Bose, Scott Dawson and Emil Ruff who guided methrough this project.ReferencesAnderson, O. R., Gorrell, T., Bergen, A., Kruzansky, R., & Levandowsky, M. (2001). Nakedamoebas 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 inan intertidal creek mudflat. MARINE ECOLOGY-PROGRESS SERIES-, 361, 59.Appendix 1 – media recipesMarine sulfur-phototroph liquid media360 ml sea water base solution342.2 mM NaCl14.8 mM MgCl21.0 mM CaCl26.71 mM KCl2 ml 1 M NH4Cl4 ml 100 mM K phosphate buffer pH 7.24.0 g/l KH2PO412.7 g/l K2HPO42 ml 1 M MOPS buffer pH 7.20.4 ml trace element solution20 mM HCl7.5 mM FeSO40.48 mM H3BO30.5 mM MnCl26.8 mM CoCl21.0 mM NiCl212 µM CuCl2
0.5 mM ZnSO40.15 mM Na2MoO425 µM NaVO39 µM Na2WO423 µM Na2SeO3flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclavingthe following ingredients were added during flushing with N2 / CO2 gas4 ml sterile multivitamin solution10 mM MOPS buffer pH 7.20.1 g/l riboflavin0.03 g/l biotin0.1 g/l thiamine HCl0.1 g/l L-ascorbic acid0.1 g/l d-Ca-pnathotenate0.1 g/l folic acid0.1 g/l nicotinic acid0.1 g/l 4-aminobenzoic acid0.1 g/l pyridoxine HCl0.1 g/l lipoic acid0.1 g/l NAD0.1 g/l thiamine pyrophosphate0.01 g/l cyanocobalamin28 ml sterile 1 M NaHCO34 ml sterile 1 M Na2S2O30.4 ml sterile 0.2 M Na2S added in the anaerobic chamberMarine acetate-degrading nonsulfur-phototroph liquid media390 ml sea water base solution0.68 g sodium acetate2 ml 1 M NH4Cl0.1 ml 1 M Na2SO44 ml 100 mM K phosphate pH 7.20.5 ml 1 M MOPS buffer pH 7.20.4 ml trace element solutionflushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclavingthe following ingredients were added during flushing with N2 / CO2 gas
4 ml sterile multivitamin solution1 ml sterile 1 M NaHCO3Thiosulfate-acetate agar1 l sea water base solution5 ml 1 M NH4Cl10 ml 100 mM K phosphate pH 7.25 ml 1 M MOPS buffer pH 7.21 ml trace element solution0.25 ml 1 M Na2SO41.75 g sodium acetate17 g agar washed in distilled waterflushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclavingthe following ingredients were added during flushing with N2 / CO2 gas10 ml sterile multivitamin solution70 ml sterile 1 M NaHCO310 ml sterile 1 M Na2S2O3Anoxic LB1 l distilled water10 g tryptone5 g yeast extract10 g NaCl15 g agarset pH to 7.0flushed with sterile N2 / CO2 (80% / 20%) gas for 2 hours after autoclavingAnoxic SWC1 l sea water base solution5 g tryptone1 g yeast extract3 ml glycerol15 g agarset pH to 7.0flushed 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 hindgutThe hindgut of wood-eating termites is rich in protists, Bacteria and Archaea. Bacteria are presentnot only as free living members of the termite hindgut microbiota but also as exo- andendosymbionts of protists. The diet of wood-eating termites is low in nitrogen, and N2 fixingbacteria living as endosymbionts in protists are abundant in the hindgut (Breznak and Pankratz1977, Inoue et al. 2008, Noda et al. 2005). The goal of this project was to investigate thelocalization of nitrogenase activity in the hindgut of Reticulitermes flavipes.The intestinal tract of workers and one soldier of Reticulitermes flavipes was dissected out anddirectly, or after 50 minutes of fixation at room T in a drop of 3.2 % formamide, embedded inTissue-Tek resin, frozen in liquid nitrogen and stored at -20 oC. The frozen blocks were cut to 20µm thick slices with a microtome in a cryostat. The slices were collected on sterile 0.2 µm GTTPfilter membranes. The membranes were sprayed with 0.2 % low melting point agarose, dried atroom T and stored at -20 oC. The filter membranes were not transparent which prevented theexamination of the samples with transmission light microscopy, but it was much easier to handlethem than glass slides during the FISH procedure, also the majority of slices collected on regularor 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 werelost from the membranes. Most likely the permeabilization and H2O2 treatments damaged theinsect tissue or the samples were lost during the many washing steps in the protocol. I triedmono-FISH using the general bacterial and archaeal probes again, which worked, but thefluorescent signal was often hard to see. Parts of the gut content bound DAPI and the fluorescentprobes resulting in a foggy background. This could likely be circumvented by slicing the samplesto thinner sections. I also observed chitin autofluorescence. As the next step I was planning to dohybridization chain reaction with NifH specific probes on the membranes to localize NifHexpression 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 midgutbecause 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 andalso bacteria and nuclei of protists.
Figure 9: Higher magnification of the gut lumen (same gut section as on the previous figure). Spirochetes and otherbacteria 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 isempty 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 workertermite.Figure 12: Mono-FISH general bacterial probe signal from the gut wall in the hindgut of a worker termite. Manybacteria appear to be within the insect tissue. This was only observed in some regions and I didn’t find any indicationin the literature that bacteria would be penetrating the insect tissue, therefore it is likely an artifact due to thesectioning of the sample.References:
Breznak, J. A., & Pankratz, H. S. (1977). In situ morphology of the gut microbiota ofwood-eating termites [Reticulitermes flavipes (Kollar) and Coptotermes formosanusShiraki]. Applied and environmental microbiology, 33(2), 406.Inoue, J. I., Noda, S., Hongoh, Y., Ui, S., & Ohkuma, M. (2008). Identification ofendosymbiotic methanogen and ectosymbiotic spirochetes of gut protists of the termiteCoptotermes 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 Pseudotrichonymphagrassii in the gut of the termite Coptotermes formosanus. Applied and environmentalmicrobiology, 71(12), 8811-8817.
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
Protists EQ: How do protists live and function? Objectives: 7A1 Identify the major characteristics of protists. 7A2 Classify protists on the basis of major characteristics. 7A3 Summarize how protists move, obtain energy, and reproduce. 7A4 Explain the roles of protists in the environment. 7A5 Evaluate the assumption that protists are lower life .
Animal-like Protists Plantlike Protists Fungus-like Protists Group Example(s) Distinguishing Characteristics II. Protozoans: Animal-like Protists (4A, 4B, 8A, 8C) (pg. 546) a. Main Idea: Protozoans are_, heterotrophic Protists b. Protozoans are
Regarding how they obtain energy, some protists act like plants, some like fungi, and some like animals. Plant-like protists obtain energy from the sun (autotrophs), fungi-like protists obtain energy by absorbing it from other organisms (heterotrophs), and animal-like protists obtain energy by eating other organisms (heterotrohs). Protists also .File Size: 1MB
Catalyst Summer 2015 Volume 10, Number 2 MBL Catalyst is published twice yearly by the Office of Communications at the Marine Biological Laboratory (MBL) in Woods Hole, Massachusetts. The MBL is dedicated to scientific discovery and improving the human condition through research and education in biology, biomedicine, and
protist. This red alga is a multicellular, plantlike protist found on ocean floors. What Is a Protist? Diatoms are only one of the vast varieties of protists. Protists are eukaryotes that cnnot a be classified as animals, plants, o rfungi. Because protists
Fungus-like Protists Water molds and slime molds . Animal-like Protists classified according to how they move: – amoeba-like – ciliated – flagellated . Protists and Human Health and Welfare Many protists
Protists can be divided into three ecological categories: Protozoa—ingestive, animal-like protists Absorptive, fungus-like protists Algae—photosynthetic, plant-like protists. Most protists move with flagella or cilia during some time in their life cycles.
Unit 14: Advanced Management Accounting Unit code Y/508/0537 Unit level 5 Credit value 15 Introduction The overall aim of this unit is to develop students’ understanding of management accounting. The focus of this unit is on critiquing management accounting techniques and using management accounting to evaluate company performance. Students will explore how the decisions taken through the .
