Planctopirus ephydatiae, a novel Planctomycete isolated from a freshwater sponge

Journal Pre-proof Planctopirus ephydatiae, a novel Planctomycete isolated from a freshwater sponge T. Kohn, S. Wiegand, C. Boedeker, P. Rast, A. Heuer, M.S.M. Jetten, M. Schuler, ¨ S. Becker, C. Rohde, R.-W. Muller, ¨ F. Brummer, ¨ M. Rohde, H. Engelhardt, M. Jogler, C. Jogler

PII:

S0723-2020(19)30317-0

DOI:

https://doi.org/10.1016/j.syapm.2019.126022

Reference:

SYAPM 126022

To appear in:

Systematic and Applied Microbiology

Received Date:

29 June 2019

Revised Date:

30 September 2019

Accepted Date:

2 October 2019

Please cite this article as: Kohn T, Wiegand S, Boedeker C, Rast P, Heuer A, Jetten MSM, Schuler ¨ M, Becker S, Rohde C, Muller ¨ R-W, Brummer ¨ F, Rohde M, Engelhardt H, Jogler M, Jogler C, Planctopirus ephydatiae, a novel Planctomycete isolated from a freshwater sponge, Systematic and Applied Microbiology (2019), doi: https://doi.org/10.1016/j.syapm.2019.126022

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Planctopirus ephydatiae, a novel Planctomycete isolated from a freshwater sponge Kohn T.1, Wiegand S.1, Boedeker C.2, Rast P.2, Heuer A.2, Jetten M. S. M. 1, Schüler M.5,Becker S.6, Rohde C.2, Müller R.-W.4, Brümmer F.4, Rohde M.3, Engelhardt H.5, Jogler M.2 and Jogler C.1,7* Department of Microbiology, Radboud University, Nijmegen, Netherlands

2

Leibniz-Institut Deutsche Sammlung von Mikroorganismen und Zellkulturen, Braunschweig, Germany

3

Central Facility for Microscopy, Helmholtz-Centre for Infection Research (HZI), Braunschweig,

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1

Germany 4

Institute of Biomaterials and Biomolecular Systems, University of Stuttgart, Germany

5

Department of Molecular Structural Biology, Max Planck Institute of Biochemistry, Martinsried,

Germany University of Veterinary Medicine Hannover, Germany

7

Department of Microbial Interactions, Friedrich Schiller Universität Jena, Germany

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* Correspondence: [email protected]

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Abstract

The microbiome of freshwater sponges is rarely studied, and not a single novel bacterial species has been isolated and subsequently characterized from a

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freshwater sponge to date. A previous one study showed that 14.4% of the microbiome from Ephydatia fluviatilis belongs to the phylum Planctomycetes. Therefore, we sampled an Ephydatia sponge from a freshwater lake and employed enrichment techniques targeting bacteria from the phylum Planctomycetes. The obtained strain spb1T was subject to genomic and phenomic characterization and

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found to represent a novel planctomycetal species proposed as Planctopirus ephydatiae sp. nov. (DSM 106606 = CECT 9866). In the process of differentiating spb1T from its next relative Planctopirus limnophila DSM 3776T, we identified and characterized the first phage - Planctopirus phage vB_PlimS_J1- infecting planctomycetes that was only mentioned anecdotally before. Interestingly, classical chemotaxonomic methods would have failed to distinguish Planctopirus ephydatiae strain spb1T from Planctopirus limnophila DSM 3776T. Our findings demonstrate and

underpin the need for whole genome-based taxonomy to detect and differentiate planctomycetal species. Keywords Planctomycetes; Bacteriophage;

Planctopirus Lake

limnophila;

Constance;

Sponge;

Ephydatia

Freshwater;

fluviatilis,

Taxonomy;

Planctopirus

phage

vB_PlimS_J1

Introduction Members of the phylum Planctomycetes are unique among bacteria in terms of their

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unusual cell biology [1-4]. The cells comprise an enlarged periplasmic space that is thought to be involved in the breakdown of otherwise recalcitrant carbon compounds [2]. Furthermore, most described planctomycetes divide via polar budding, employing a yet unknown molecular mechanism. This mechanism does not rely on otherwise essential bacterial divisome proteins such as FtsZ [5]. Many species of the order

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Planctomycetales show a life cycle, switching between a planktonic flagellated swimmer and a sessile reproducing stage [6-8]. Planctomycetes are ubiquitous

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bacteria that dwell in various aquatic and terrestrial habitats [1]. They are predominantly found associated with all kinds of surfaces to which they can attach

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utilizing holdfast structures [1]. In particular, planctomycetes are abundant on marine surfaces where they can dominate bacterial biofilms, for example on clusters of polysaccharide-rich debris particles, so-called marine snow [9], cyanobacterial

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aggregates [10] or macroalgae [11, 12]. To explain the high abundance of rather slow-growing planctomycetes in these competitive nutrient-rich habitats, the production of small (antibiotic) molecules was suggested [13-15]. Besides aquatic habitats, planctomycetes also represent a significant proportion of the bacterial community in the gut microbiome of mammals [16, 17], fish [18], shrimp [19] and soil-

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feeding termites [20]. .In animals without a digestive system, like tubeworms [21], corals [22] or marine and freshwater sponges [23, 24], Planctomycetes can represent a significant fraction of the microbiome as well. For example, 14.4% of the microbiota associated with the globally occurring freshwater-brackish sponge Ephydatia fluviatilis [25, 26] was found to belong to the phylum Planctomycetes [27]. Also, planctomycetes were isolated and characterized from marine sponges [28-30], but have not been validly described.

Sponges from the phylum Porifera are ancient multicellular animals (Metazoa) that lack a digestive tract as well as organs and a nervous system. They are, with few exceptions, sessile filter feeders [31, 32] with a body structure build around a network of water channels and chambers. The body structure consists of three different tissue-like layers, the pinacoderm (outside), the mesohyl (in-between), and the choanoderm (inside) [33]. Surrounding water, containing food particles like algae, bacteria, viruses and protists, enter the animals' water channels through small pores (ostia) situated within the otherwise dense monolayer of cells forming the pinacoderm. Specialized collar-flagellated cells (choanocytes) actively drive the water

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influx. Besides establishing a steady water current, choanocytes do also capture the prey in reach. The prey particles are internalized by endocytosis and then digested or transferred to the mesohyl, where they are engulfed by phagocytic amoebocytes [34]. Subsequently, near-sterile water, with up to 96% of microbial cells removed, [35] is released into the surrounding water through the osculum.

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In contrast to the pinaco- and choanoderm, the mesohyl is not composed of a single layer of cells. In most species, it is a matrix consisting of collagen and that

harbors

a

bacterial

community

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polysaccharides

comprising

105-1010

microorganisms per gram sponge tissue [36, 37]. However, the ecological role of the

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mostly uncultivated sponge microbiome remains enigmatic [38]. There is growing evidence that the production of (toxic) small molecules, contributing to host defense, is a crucial function of this microbiome [39]. For instance, the uncultivated candidate

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genus Entotheonella of the candidate phylum Tectomicrobia dwells in marine sponges and produces novel small molecules with interesting chemistry and numerous unusual biological activities [38, 40]. Motivated by prospects of drug discovery, significant efforts have been made to obtain axenic cultures of small molecule-producing sponge symbionts, unfortunately, with limited success [34, 38].

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Most of the 8897 different sponge species known thus far appear in marine habitats, while only about 250 freshwater species are described [41-44]. Those are either endemic or ubiquitous [45] and belong to the single monophyletic order Spongilina [46]. Until now, only a few studies focused on bacteria related to these freshwater sponges and their potential in small molecule production [27, 47-53]. Even though different bacterial strains belonging to the phyla Actinobacteria and Proteobacteria were isolated from freshwater sponges before, no strain has been characterized and

validly described [52, 54-56]. Regarding planctomycetes, we only know about their high abundance in the freshwater sponge E. fluviatilis [27]. In this study, we describe the novel species Planctopirus ephydatiae isolated from the freshwater sponge Ephydatia sp. and compare it to the close relative Planctopirus limnophila DSM 3776T by genomic and phenomic analyses. This comparative approach led to the identification and characterization of Planctopirus phage vB_PlimS_J1 from P. limnophila DSM 3776T. This study presents the first detailed characterization of a novel bacterial species from a freshwater sponge. Furthermore,

this

is

the

first

comprehensive

report

of

a

planctomycetal

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bacteriophage.

Materials and Methods Sampling

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Ephydatia sp. sponge samples were harvested at Lake Constance (47°46'12.6" N, 9°08'18.1" E) on August 25th, 2015 via scientific scuba diving according to CMAS

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standards in 15.6 m water depth (Fig. 1 a). The animal (Fig. 1 b) was washed three times in sterile-filtered lake water (Fig. 1 c) and transferred to the laboratory in sterile-

Isolation and Cultivation

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filtered lake water at 4 °C.

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A piece of the sponge was streaked on solid MM1 medium supplemented with Nacetyl-D-glucosamine (NAG) at pH 7.0. Medium MM1 was prepared as previously described [14]. In brief the medium consists of artificial freshwater (0.00053 g/L NH4Cl, 0.0014 g/L KH2PO4, 0.010 g/L KNO3, 0.049 g/L MgSO4 CaCl2

.

.

7 H2O, 0.015 g/L

2 H2O, 0.025 g/L CaCO3, and 0.025 g/L NaHCO3) complemented with 20

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mL/L mineral salt solution and 5 mL/L vitamin solution (double concentrated) and was buffered with 10 mM HEPES. Mineral salt solution was composed of 10 g/L nitrilotriacetic acid (NTA), 29.70 g/L MgSO4 . 7 H2O, 3.34 g/L CaCl2 . 2 H2O, 12.67 mg/L Na2MoO4

.

2 H2O, 99 mg/L FeSO4

.

7 H2O and 50 mL/L metal salt sol. ‘44’.

Metal salt sol. ‘44’ was composed of 250 mg/L Na-EDTA, 1.095 g/L ZnSO4 . 7 H2O, 0.5 g/L FeSO4 . 7 H2O, 154 mg/L MnSO4 . H2O, 39.20 mg/L CuSO4 . 5 H2O, 24.80 mg/L Co(NO3)2

.

6 H2O and 17.70 mg/L Na2B4O7

.

10 H2O. The vitamin solution

contained of 4 mg/L biotin, 4 mg/L folic acid, 20 mg/L pyridoxine-HCl, 10 mg/L

riboflavin, 10 mg/L thiamine-HCl

.

2 H2O, 10 mg/L nicotinamide, 10 mg/L D-Ca-

pentothenate, 0.2 mg/L vitamin B12 and 10 mg/L p-aminobenzoic acid. Solid medium was prepared by adding 15 g/L of triple prewashed Bacto-agar (BD) to the medium. 5 mg/L of N-acetyl-D-glucosamine (NAG) and vitamin solution were supplemented to the medium after autoclaving. Inoculated MM1 plates were incubated at 20 °C and periodically checked for colony formation. Samples from colonies were inspected by light microscopy for typical planctomycetal cell shapes and cell division via budding. Promising isolates were verified as planctomycetes, employing a 16S rRNA gene targeting PCR, using the primer set 8f (5’–AGA GTT TGA TCM TGG CTC AG–3’)

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and 1492r (5’–GGY TAC CTT GTT ACG ACT T–3’) modified from [57]. PCR amplifications were performed employing a Veriti™ 96-Well Thermal Cycler (Applied Biosystems) using the following conditions: initial denaturation at 94 °C for 5 min, followed by 10 cycles of denaturation at 94 °C for 30 s, annealing at 59 °C for 30 s and elongation at 72 °C for 60 s. Following these first 10 cycles, 20 cycles of

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denaturation at 94 °C for 30 s, annealing at 54 °C for 30 s, and elongation at 72 °C for 60 s and a final elongation at 72 °C for 5 min were performed. Amplification

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products were subject to 16S rRNA gene sequencing and NCBI database comparison to identify novel strains. Based on this analysis, strain spb1 was

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confirmed to represent a Planctomycete. For further analyses, strain spb1T was cultivated employing V2 culture broth at pH 7.0. V2 was composed of medium MM1 supplemented with 0.25 g/L yeast extract, 0.25 g/L peptone, 0.025 g/L glucose, and 1

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mL/L trace elements solution. Trace elements solution was composed of 1.5 g/L N(CH2COONa)3 . H2O, 500 mg/L MnSO4 . H2O, 100 mg/L FeSO4 . 7 H2O, 100 mg/L Co(NO3)2 . 6 H2O, 100 mg/L ZnCl2, 50 mg/L NiCl2 . 6 H2O, 50 mg/L H2SeO3, 10 mg/L CuSO4 . 5 H2O, 10 mg/L AlK(SO4)2 . 12 H2O, 10 mg/L H3Bo3, 10 mg/L NaMoO4 . 2 H2O and 10 mg/L Na2WO4 . 2 H2O.

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Genome Analysis

Wiegand et al. [58] published the analyzed genome of strain spb1T, which is available from NCBI under GenBank accession number CP036299. Genomic islands were predicted and visualized employing IslandViewer 4 [59]. The average nucleotide identities (ANI) were determined using the software OrthoANI [60]. Genomes were further compared using the Genome-to-Genome distance calculator GGDC [61]. The genome of strain spb1T was analyzed towards its potential to produce secondary metabolites employing antiSMASH 4.0 [62]. The analysis of orthologous groups to

determine proteins present in all compared genomes was done with ProteinOrtho [63].

Phylogenetic analysis and tree reconstruction 16S rRNA gene phylogeny was computed for strain spb1T (GenBank acc. no. MK559993), the type strains of all described planctomycetal species (May 2019) and all isolates recently published by Wiegand et al. [58]. The 16S rRNA gene sequences were aligned with SINA [64]. The phylogenetic analysis was done employing a maximum likelihood (ML) approach with 1,000 bootstraps, the nucleotide substitution

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model GTR, gamma distribution and estimation of proportion of invariable sites (GTRGAMMAI option) [65]. Three 16S rRNA genes of bacterial strains from the PVC superphylum served as outgroup. The rpoB nucleotide sequences were taken from the above-mentioned as well as other publicly available genome annotations, and the

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sequence identities were determined as described by Bondoso et al. [66]. Upon extracting only those parts of the sequence that would have been sequenced with the

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described primer set the alignment and matrix calculation was done with Clustal

Light Microscopy

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Omega [67]

Phase contrast (Phaco) analysis was performed using a Nikon Eclipse Ti inverted microscope with a Nikon N Plan Apochromat λ 100x/1.45 Oil objective and a Nikon

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DS-Ri2 camera. Cells were immobilized, employing a 1% agarose cushion in MatTek glass-bottom dishes (35 mm, No. 1.5). Images were analyzed using the Nikon NISElements software (Version V4.3).

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Electron Microscopy

For field emission scanning electron microscopy (FESEM) bacteria were fixed with 1% formaldehyde in HEPES buffer (3 mM HEPES, 0.3 mM CaCl2, 0.3 mM MgCl2, 2.7 mM sucrose, pH 6.9) for 1 h on ice and washed one time with the same buffer. Coverslips with a diameter of 12 mm were coated with a poly–L–lysine solution (Sigma–Aldrich) for 10 min, washed in distilled water and air-dried. Fifty microliters of the fixed bacteria solution were placed on a coverslip and allowed to settle for 10 min. Coverslips were then fixed in 1% glutaraldehyde in TE-buffer (20 mM TRIS, 1

mM EDTA, pH 6.9) for 5 min at room temperature and subsequently washed twice with TE–buffer before dehydrating in a graded series of acetone (10, 30, 50, 70, 90, and 100%) on ice for 10 min at each concentration. Samples from the 100% acetone step were brought to room temperature before placing them in fresh 100% acetone. Samples were then subjected to critical–point drying with liquid CO2 (CPD 300, Leica). Dried specimens were covered with a gold/palladium (80/20) film by sputter coating (SCD 500, Bal–Tec). Examination in a field emission scanning electron microscope (Zeiss Merlin) was done using the Everhart Thornley HESE2–detector and the inlens SE–detector in a 25:75 ratio at an acceleration voltage of 5 kV. TEM

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micrographs were taken after negative staining of the cells with aqueous 0.1–2% uranyl acetate, using a Zeiss transmission electron microscope EM 910 at an acceleration voltage of 80 kV at calibrated magnification as previously described [68]. Cryo-electron tomography (CET) of P. limnophila cells was performed as previously

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described [2]. In brief, a late-exponential-phase culture was gently filtered (10 µm membrane filter, Whatman Nuclepore) to exclude aggregated cells. Aliquots (3 µl) of the filtered cell suspension where mixed with the same volume of BSA-stabilized 15

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nm colloidal gold solution (Aurion) and placed on holey carbon-coated 200 mesh copper grids (R2/1, Quantifoil, Jena, Germany) directly before thin-film vitrification by

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plunge-freezing in liquid propane (63%)/ethane (37%)[69]. Typically, grids with frozen-hydrated samples were mounted in Autogrids [70] and ~200 nm thin lamellae of vitrified material were milled with 30 keV gallium ions after application of a

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protective platinum layer in a dual-beam (FIB/SEM) instrument (Quanta 3D FEG, FEI, Hillsboro, OR, USA) equipped with a Quorum cryo-stage maintained at -185°C (PP2000T, Quorum, East Sussex, UK). Milling was carried out at nominal incident ion beam angles of 16° to 20° (9° to 13° effectively) using gallium beam currents of 300 pA, 100 pA and 30 pA in sequential milling steps [71]. Subsequently tomographic tilt

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series were recorded under low dose conditions (typically 150 e/Å2 total dose) on a Tecnai G2 Polara (FEI, Eindhoven, The Netherlands) equipped with a post-column energy filter and a 2k CCD camera (MultiScan) or a K2 summit direct electron detector (Gatan, Pleasanton, CA, USA). Tilt series recorded with the direct detection device, dose fractionation mode was employed, and subframes of each projection were sampled, which were then aligned to compensate for beam-induced object drift, using an in-house implementation of the algorithm described by Li, et al.[72]. Typically, tilt series were recorded at a nominal defocus of -5 or -6 µm and a primary

magnification of 27,500 x (corresponding to pixel sizes on the object level of 0.427 nm [K2] and 0.805 nm [2k CCD]) and covered an angular range of  60° in increments of 1.5° or 2°, respectively. IMOD v4.7.8 [73] was used for 3D reconstruction, and MatLab8 (MathWorks) incorporating the TOM toolbox[74] for all image processing. Segmentation of three times binned volumes was performed in Amira v6.0.1 (FEI, Eindhoven, The Netherlands) with specific automatic membrane segmentation [75].

Physiological Tests

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Physiological tests such as pH and temperature tolerance were performed in liquid medium V2 as previously described [76]. Growth was detected by monitoring the optical density at 600 nm using a Photometer Ultrospec II (LKB Biochrom). Enzymatic activities were tested using the API® ZYM method according to the

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manufacturer instructions (bioMérieux).

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Fatty Acid Analysis

For fatty acid analysis, biomass of both strains (Planctopirus limnophila DSM 3776T

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and spb1T) was obtained from agar plate cultures, grown on V2 medium, at 28°C. The obtained biomasses were processed according to the standards of the Identification Service of the German Collection of Microorganisms and Cell Cultures

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(DSMZ) [77-79].

Phage detection and classification For phage-isolation experiments, P. limnophila DSM 3776T was cultivated at 28 °C in

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medium M3. M3 was composed of 1 g/L peptone, 1 g/L yeast extract, 1 g/L glucose, 5 mL/L vitamin solution (double concentrated) and 20 mL/L mineral salt solution (for composition see section Isolation and cultivation). Furthermore, the medium was buffered with 10 mM HEPES, and the pH adjusted to 7.5. To isolate lytic phages, the supernatant of a P. limnophila DSM 3776T culture, showing classic signs of bacterial lysis indicated by loss of turbidity and color, was filtered through a 0.45 µm filter. As a negative control, the filtered supernatant of a P. limnophila culture, containing lysogenic phages, was used. To detect the P. limnophila phage PI-89 in the

supernatant a phage-specific PCR using the primer set CJ334 (5’–CAGT CGG GCT TTG ATA CGA T–3’) and CJ335 (5’–AGA TGA GTC ACG GCT TGC TT–3’) was employed. PCR amplifications were performed using a Veriti™ 96-Well Thermal Cycler (Applied Biosystems) applying the following conditions: initial denaturation at 95°C for 5 min, followed by 40 cycles of denaturation at 95°C for 30 s, annealing at 50°C for 30 s and elongation at 72°C for 2 min. A final elongation step was performed at 72°C for 5 min. Amplicons were visualized on a 1% agarose gel by staining with ethidium bromide and exposure to UV light. Proteins of phage PI-89 (Accession number: CP001745) were compared against the

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NCBI database using BLASTp [80] with viruses, Siphoviridae, Myoviridae, or Podoviridae as taxon filters. Homologs were identified using initial thresholds of identity ≥ 30%; coverage ≥ 60% and e-value of ≤ 10-5 for at least one of the filtered taxa, and results were confirmed by reciprocal blast analysis. PASC (PAirwise

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Sequence Comparison) was used for analysis of pairwise identity distribution within viral families [81].

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Results and Discussion Isolation of strain spb1T

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Sampling took place on Lake Constance (47°46'12.6" N, 9°08'18.1" E) on August 25th, 2015. A freshwater sponge of the genus Ephydatia was chosen as a specimen (Fig. 1), based on previous reports of high planctomycetal abundance in these

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metazoa [27]. Employing an enrichment based on 5 mg/L of N-acetyl-D-glucosamine (NAG) as sole carbon source, we obtained a planctomycetal strain from Ephydatia sp. that formed pinkish colonies on solid agar plates. 16S rRNA gene sequencing and BLAST analysis revealed that strain spb1T shared 99.99% 16S rRNA gene sequence identity with Planctopirus limnophila DSM 3776T showing only two distinct

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mismatches. Despite the close relation of spb1T with P. limnophila DSM 3776T (Fig. 2), we decided to characterize this strain for the following reasons further: i) P. limnophila DSM 3776T is the most important model Planctomycete with genetic tools being readily available [82]. ii) Thus far, nothing is known about the natural ecology of P. limnophila DSM 3776T, which was initially isolated from surface water [83, 84], but escaped detection via amplicon sequencing in comparable habitats (unpublished observation). iii) Strain spb1T would represent the first validly described bacterium

isolated from a freshwater sponge, and its characterization might help to understand the interplay between freshwater sponges and their associated bacteria. Phenotypic characterization of strain spb1T In wide-field light microscopic phase-contrast micrographs, cells of strain spb1T look like the closely related type strain DSM 3776T. Mother cells are ovoid to spherical and 1.1-1.5 x 1.5-2.5 µm in size; daughter cells are flagellated and motile. Cells form rosettes and aggregates in liquid culture (Fig. 3). However, if scanning electron microscopy (SEM) is applied, differences become visible (Fig. 4): while both strains

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form aggregates, those of strain spb1T appear more densely packed (Fig. 4 a/d). Fibers become visible at higher magnification. To ensure that these observations are not methodical artifacts of critical point drying necessary for SEM specimen preparation, we also performed transmission electron microscopy (TEM) where such invasive methods are not needed (Fig. 5). Both microscopic methods show a similar

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picture: Fibers can be observed considerably more frequent on spb1T cells than on P. limnophila DSM 3776T cells (Fig. 4 b/e). Both strains possess characteristic

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crateriform structures that are the origin of the fibers (Fig. 4 c/f, white arrowheads). Crateriform structures are a unique trait among bacteria and are exclusively found in

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the phylum Planctomycetes [1, 2]. We recently proposed that they are involved in the uptake of high molecular weight carbon substrates [2]. It was shown for P. limnophila DSM 3776T, that fibers arising from crateriform structures can bind dextran that is

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then incorporated - by a still unexplored process - into the enlarged periplasmic space where it seems to be degraded [2]. The environmental significance of uptake mechanisms for polysaccharides for marine habitats was recently demonstrated [85]. Given that cells of strain spb1T produce even more fibers than P. limnophila DSM 3776T, it is tempting to speculate that this organism ‘filters’ complex organic

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compounds out of the lake water, analogous to its animal host. In addition to variations in cell biology, strain spb1T also showed slightly different physiological characteristics compared to P. limnophila DSM 3776T. Strain spb1T grows between pH 7.0 and 9.0, with an optimal growth rate at pH 9.0 (Fig. S1 a). In contrast, P. limnophila DSM 3776T showed growth between pH 6.0 and 10.0, with an optimal growth rate at pH 6.0. Thus, P. limnophila DSM 3776T seems to be adapted to typical freshwater environments, while strain spb1T prefers higher pH values. A typical freshwater lake has a pH of 7.0-8.0, and even typical marine habitats do

barely exceed pH 8.0 [86, 87]. Thus, future enrichment strategies targeting planctomycetes from sponges might use pH 9.0 buffered medium as an additional selection factor. While strain spb1T grows at temperatures between 15 and 33 °C (optimum 30 °C, Fig. S1 b), P. limnophila DSM 3776T grows in a range from 10 to 36 °C (optimum 27 °C). Despite differences in optimal growth conditions, the tested enzymatic repertoires of both strains were identical: they comprise alkaline phosphatase, esterase, esterase lipase, naphthol-AS-BI-phosphohydrolase, and α-galactosidase activities. The same is true for the lipid composition. Both strains share the major fatty acids C16:0 (40.71

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and 37.17%), C16:1 6c/16:1 7c (21.46 and 22.8%) and C18:1 9c (18.22 and 19.37%). The complete fatty acid profiles are listed in Supplementary Table S1. Genome analysis and phylogeny of strain spb1T

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The chromosome of strain spb1T comprises 5,214,061 bp with a GC content of 53.66% (Table S2). Strain spb1T does not possess any plasmid. We could annotate

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4,127 genes, and 4,038 of that were identified as protein-coding. The analysis of the genomes of P. limnophila DSM 3776T and strain spb1T using IslandViewer 4 [59]

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revealed several genomic islands that represent regions of horizontal gene transfer (Fig. S2). The secondary metabolite potential of the two analyzed strains was predicted using antiSMASH 4.0 [62]. P. limnophila DSM 3776T comprised seven

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putative secondary metabolite-related gene clusters, three terpenes, two type1-PKS, one type3-PKS and one bacteriocin. Strain spb1T shared most of the clusters predicted in the strain DSM 3776T, with one exception. The bacteriocin cluster is missing in strain spb1T, but a type1-PKS-resorcinol-arylpolyene cluster was predicted

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instead (Fig. S3).

Phylogenetic characterization As shown in Fig. 2 strain spb1T clusters monophyletically with the strains Planctopirus limnophila DSM 3776T and Planctopirus hydrillae JC280T. Initially, 16S rRNA gene sequencing revealed that strain spb1T shared 99.99% 16S rRNA gene sequence identity with P. limnophila DSM 3776T, including two distinct mismatches. The genome of strain spb1T harbors two distinctive 16S rRNA genes that differ in one transition of A to G, but shares the two mismatches compared to P. limnophila DSM

3776T. The 16S rRNA sequence identity between strain spb1T and P. hydrillae JC280T is 99.7% (Fig. 6). If based solely on 16S rRNA similarity thresholds, all three strains would belong to the same species as the species threshold was proposed to be 98.7% [88]. However, the 16Ss rRNA gene is known to be of limited benefit when planctomycetal species need to be differentiated [66]. Otherwise, P. limnophila DSM 3776T and P. hydrillae JC280T would also belong to the same species. Therefore the rpoB gene (encoding the β-subunit of the bacterial RNA polymerase) was proposed as an alternative marker [66]. P. limnophila DSM 3776T and strain spb1T share 93.3% rpoB gene identity and P. hydrillae JC280T and spb1T share 93.6%, both

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values being below the 96.3% cutoff proposed to distinguish between different planctomycetal species [66] (Fig. 6). To clarify these conflicting results, we employed further well-established genomic analyses. The reciprocal average nucleotide identities (ANI) of P. limnophila DSM 3776T and strain spb1T, and P. hydrillae JC280T and spb1T were determined to be 93.1% and 91.9%, respectively, and are also

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below the 95-96% identity species border defined for this method (Fig. 6) [89]. Based on digital DNA-DNA hybridization employing the Genome-to-Genome Distance

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Calculator GGDC [61], the DDH estimate for P. limnophila DSM 3776T and strain spb1T yields 49.8%, far below the 70% cutoff for identical species [90, 91]. Finally,

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the analysis of conserved proteins between the three strains also points towards a similar relatedness amongst them (Fig. 6).

Except for the 16S rRNA gene identity, which can fall short to distinguish

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planctomycetal species [66]. Thus, all genome-based analyses suggest that strain spb1T is a novel species within the genus Planctopirus.

Planctopirus phage vB_PlimS_J1 Horizontal gene transfer (HGT) can be mediated by plasmids, transposons, and -

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very commonly - by bacteriophages [92]. However, only a few planctomycetes possess plasmids [93], and we could only annotate 3 and 5 transposases in the genomes of P. limnophila DSM 3776T and strain spb1T, respectively. Thus, transposition might contributes to HGT in the genus Planctopirus. However, species with intensively studied transposon-mediated HGT, such as magnetotactic bacteria, contain far more transposase genes [94-96]. In contrast to the existence of plasmids and transposons, even less is known about bacteriophages infecting planctomycetes. Thus far, only two planctomycetal bacteriophages were anecdotally reported [97],

phage PI-89 from P. limnophila DSM 3776T and phage Pi-57 found in Pirellula sp. IFAM 1358. Both phages were preliminarily classified as members of the family Siphoviridae morphogroup B1 and B2, respectively, but not further analyzed [97]. When the genome sequence of P. limnophila DSM 3776T became available, a 37 kb extrachromosomal element was identified [98]. Since this extrachromosomal element is absent in strain spb1T, we examined it further. Using BLASTp analysis, we identified the extrachromosomal element as a prophage (Supplementary Table S3) with closest relationship to the family Siphoviridae. This classification was also supported by analysis of pairwise identity distribution within viral families with the

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PASC webtool. The sequence shows 8.97% identity to the closest relative Burkholderia phage KS9 belonging to the family Siphoviridae. The prophage displayed less than 50% sequence identity to any other known phage and consequently represented a new genus [99].

We did not detect phage DNA in the cell-free supernatant of exponentially growing P.

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limnophila DSM 3776T cultures employing PCR with primers deduced from the prophage sequence. Therefore, the phage seems to be in the lysogenic stage under

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standard laboratory conditions. In contrast, when stimuli such as nutrient limitation or high temperatures are stressing the cells, the phage enters the lytic cycle. The deep-

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pinkish liquid culture of P. limnophila DSM 3776T lost color and turbidity within a single day when the phage entered the lytic cycle, and phage DNA was detected in the cell-free supernatant. Furthermore, the cell-free supernatant from those cultures

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induced lysis in actively growing pinkish P. limnophila DSM 3776T cultures within a day. Thus, the phage can switch between a lysogenic and lytic cycle. Cultures with lytic phages were used for morphological characterization. Transmission electron microscopy showed an icosahedral capsid of Caudovirales with a diameter of 54 nm and a tail of 100 nm length (Fig. 7 a+b). Many of the observed phages were attached

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to cells with tails of the identical length (Fig. 7 b). The phage tails seem therefore non-contractile and also smaller in diameter than in the family Myoviridae (Fig. 7 b). Furthermore, we visualized the tailed phage employing cryo-electron tomography and verified the icosahedral capsid structure after three-dimensional reconstruction (Fig. 7 c and supplementary movie). Based on our data, the tailed dsDNA phage belongs to the family Siphoviridae and represents a new undescribed genus. While the isolated phage is likely identical to

the earlier anecdotally reported phage PI-89, we name it Planctopirus phage vB_PlimS_J1 to meet ITCV recommendations [99]. As phages are amongst the main drivers of horizontal gene transfer, the Planctopirus phage could be responsible for some of the differences in the two genomes of P. limnophila DSM 3776T and strain spb1T. The HGT related unique areas that we found in both genomes can explain the low DDH estimate of 49.8%. However, HGT cannot explain the low ANI values. While strain spb1T represents a novel species based on our genomic analysis, it remains enigmatic whether HGT is a general driving force in planctomycetal speciation. Furthermore, this new Phage might be interesting in light

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of the growing field of phage therapy and gut microbiome modeling [100]. As stated above Planctomycestes can colonize the gut of several species including humans[1]. Thus phages specific to planctomycetes might become an interesting additional tool

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for microbiome modeling.

Conclusions

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Here we characterized the first new bacterial species from a freshwater sponge. While standard chemotaxonomic measurements, including cellular fatty acid analysis

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fell short in distinguishing strain spb1T from the related species P. limnophila (DSM 3776T). The genomic analysis revealed that strain spb1T represents a hitherto unknown species of Planctomycetes. Consequently, we suggest employing genome-

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based taxonomy for future planctomycetal species descriptions. The closest free-living relative of strain spb1T is P. limnophila DSM 3776T, which was isolated in summer from surface water of Lake Plußsee (Holstein, Germany) [83, 84]. Thus, it might have been related to the typical algal or cyanobacterial blooms that occur summer or winter time in such habitats [10, 101]. Another close relative,

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Planctopirus hydrillae JC280T was isolated from the surface of the plant Hydrilla verticillata collected from an alkaline lake [102]. Attachment to eukaryotes appears to be a general feature of the genus Planctopirus. However, the role and interaction of strain spb1T in the sponge remains elusive. Thus far, only one report indicates that planctomycetes may be related to a sponge disease: Heterotrophic planctomycetes were enriched in abnormal tissue of Carteriospongia foliascens [103]. Whether the interaction of strain spb1T with Ephydatia sp. is mutualistic, parasitic, or represents a more neutral symbiosis requires further investigation.

Since strain spb1T is genetically distant to the closest species P. limnophila, but belongs to the genus Planctopirus and was isolated from the freshwater sponge Ephydatia sp., we propose the name Planctopirus ephydatiae for the new planctomycetal species.

Description of Planctopirus ephydatiae sp. nov E.phy.da’ti.ae. N.L. gen. n. ephydatiae of Ephydatia, a genus of limnic sponge from which the novel species was isolated. Colonies on solid medium are pink colored. Cells are ovoid-shaped, 1.1-1.5 µm in diameter, and 1.5-2.5 µm in length. Cells form

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aggregates in liquid culture and produce large amounts of fibers. Crateriform structures are dominant and spread evenly over the cell surface. Growth occurs between pH 7.0 and 9.0 with optimal growth at pH 9. Temperatures are tolerated in a range from 15 – 33 °C with an optimum at 30 °C. The type strain is spb1T (DSM 106606 = CECT 9866). The genome is available from NCBI under GenBank

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accession number CP036299, the 16S rRNA gene sequence is available under the

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Conflict of Interest Statement

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accession number MK559993.

The authors declare that the research was conducted in the absence of any

interest.

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commercial or financial relationships that could be construed as a potential conflict of

Acknowledgment

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We thank the Deutsche Forschungsgemeinschaft and the Soehngen Institute of Anaerobic Microbiology for funding (DFG grant JO893/4-1, SIAM 024002002).

References

Jo

ur na

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re

-p

ro of

[1] S. Wiegand, M. Jogler, C. Jogler, On the maverick Planctomycetes, FEMS Microbiol Rev, (2018). [2] C. Boedeker, M. Schuler, G. Reintjes, O. Jeske, M.C. van Teeseling, M. Jogler, P. Rast, D. Borchert, D.P. Devos, M. Kucklick, M. Schaffer, R. Kolter, L. van Niftrik, S. Engelmann, R. Amann, M. Rohde, H. Engelhardt, C. Jogler, Determining the bacterial cell biology of Planctomycetes, Nat Commun, 8 (2017) 14853. [3] O. Jeske, M. Schüler, P. Schumann, A. Schneider, C. Boedeker, M. Jogler, D. Bollschweiler, M. Rohde, C. Mayer, H. Engelhardt, S. Spring, C. Jogler, Planctomycetes do possess a peptidoglycan cell wall, Nat Commun, 6 (2015) 7116. [4] T. Kohn, A. Heuer, M. Jogler, J. Vollmers, C. Boedeker, B. Bunk, P. Rast, D. Borchert, I. Glöckner, H.M. Freese, H.P. Klenk, J. Overmann, A.K. Kaster, S. Wiegand, M. Rohde, C. Jogler, Fuerstia marisgermanicae gen. nov., sp. nov., an unusual member of the phylum Planctomycetes from the German Wadden Sea, Front Microbiol, 7 (2016). [5] C. Jogler, J. Waldmann, X. Huang, M. Jogler, F.O. Glöckner, T. Mascher, R. Kolter, Identification of proteins likely to be involved in morphogenesis, cell division, and signal transduction in Planctomycetes by comparative genomics, Journal of bacteriology, 194 (2012) 6419-6430. [6] P.D. Franzmann, V.B. Skerman, Gemmata obscuriglobus, a new genus and species of the budding bacteria, Antonie van Leeuwenhoek, 50 (1984) 261-268. [7] M. Jogler, C. Jogler, Towards the development of genetic tools for Planctomycetes., in: J.A. Fuerst (Ed.) Planctomycetes: Cell Structure, Origins and Biology, Springer, 2013, pp. 141164. [8] D. Gade, T. Stührmann, R. Reinhardt, R. Rabus, Growth phase dependent regulation of protein composition in Rhodopirellula baltica, in, 2005, pp. 1074-1084. [9] E.F. DeLong, D.G. Franks, A.L. Alldredge, Phylogenetic diversity of aggregate-attached vs. free-living marine bacterial assemblages, Limnology and oceanography, 38 (1993) 924-934. [10] H.Y. Cai, Z.S. Yan, A.J. Wang, L.R. Krumholz, H.L. Jiang, Analysis of the attached microbial community on mucilaginous cyanobacterial aggregates in the eutrophic lake taihu reveals the importance of planctomycetes, Microb Ecol, 66 (2013) 73-83. [11] M.M. Bengtsson, L. Øvreås, Planctomycetes dominate biofilms on surfaces of the kelp Laminaria hyperborea, BMC Microbiol, 10 (2010) 261. [12] O.M. Lage, J. Bondoso, Planctomycetes and macroalgae, a striking association, Front Microbiol, 5 (2014) 267. [13] O. Jeske, M. Jogler, J. Petersen, J. Sikorski, C. Jogler, From genome mining to phenotypic microarrays: Planctomycetes as source for novel bioactive molecules, Antonie van Leeuwenhoek, 104 (2013) 551-567. [14] O. Jeske, F. Surup, M. Ketteniss, P. Rast, B. Forster, M. Jogler, J. Wink, C. Jogler, Developing Techniques for the Utilization of Planctomycetes As Producers of Bioactive Molecules, Front Microbiol, 7 (2016) 1242. [15] A.P. Graca, R. Calisto, O.M. Lage, Planctomycetes as Novel Source of Bioactive Molecules, Front Microbiol, 7 (2016) 1241. [16] R.E. Ley, M. Hamady, C. Lozupone, P.J. Turnbaugh, R.R. Ramey, J.S. Bircher, M.L. Schlegel, T.A. Tucker, M.D. Schrenzel, R. Knight, J.I. Gordon, Evolution of Mammals and Their Gut Microbes, Science, 320 (2008) 1647-1651.

Jo

ur na

lP

re

-p

ro of

[17] T.D. Leser, J.Z. Amenuvor, T.K. Jensen, R.H. Lindecrona, M. Boye, K. Moller, Cultureindependent analysis of gut bacteria: the pig gastrointestinal tract microbiota revisited, Appl Environ Microbiol, 68 (2002) 673-690. [18] X.M. Li, Y.J. Zhu, Q.Y. Yan, E. Ringo, D.G. Yang, Do the intestinal microbiotas differ between paddlefish (Polyodon spathala) and bighead carp (Aristichthys nobilis) reared in the same pond?, J Appl Microbiol, 117 (2014) 1245-1252. [19] S. Zeng, Z. Huang, D. Hou, J. Liu, S. Weng, J. He, Composition, diversity and function of intestinal microbiota in pacific white shrimp (Litopenaeus vannamei) at different culture stages, PeerJ, 5 (2017) e3986. [20] T. Kohler, U. Stingl, K. Meuser, A. Brune, Novel lineages of Planctomycetes densely colonize the alkaline gut of soil-feeding termites (Cubitermes spp.), Environ Microbiol, 10 (2008) 1260-1270. [21] R. Medina-Silva, R.R. Oliveira, F.J. Trindade, L.G.A. Borges, T.L. Lopes Simão, A.H. Augustin, F.P. Valdez, M.J. Constant, C.L. Simundi, E. Eizirik, C. Groposo, D.J. Miller, P.R. da Silva, A.R. Viana, J.M.M. Ketzer, A. Giongo, Microbiota associated with tubes of Escarpia sp. from cold seeps in the southwestern Atlantic Ocean constitutes a community distinct from that of surrounding marine sediment and water, Antonie van Leeuwenhoek, (2017). [22] J.K. Jarett, M.D. MacManes, K.M. Morrow, M.S. Pankey, M.P. Lesser, Comparative Genomics of Color Morphs In the Coral Montastraea cavernosa, Scientific reports, 7 (2017) 16039. [23] S. Gaikwad, Y.S. Shouche, W.N. Gade, Microbial community structure of two freshwater sponges using Illumina MiSeq sequencing revealed high microbial diversity, AMB Express, 6 (2016) 1-9. [24] R.L. Simister, P. Deines, E.S. Botte, N.S. Webster, M.W. Taylor, Sponge-specific clusters revisited: a comprehensive phylogeny of sponge-associated microorganisms, Environ Microbiol, 14 (2012) 517-524. [25] J. Hooper, R.W.M. van Soest, Systema Porifera: A Guide to the Classification of Sponges, Springer, 2002. [26] M. Unger, L. Asplund, P. Haglund, A. Malmvarn, K. Arnoldsson, O. Gustafsson, Polybrominated and mixed brominated/chlorinated dibenzo-p-dioxins in sponge (Ephydatia fluviatilis) from the Baltic Sea, Environ Sci Technol, 43 (2009) 8245-8250. [27] R. Costa, T. Keller-Costa, N.C. Gomes, U.N. da Rocha, L. van Overbeek, J.D. van Elsas, Evidence for selective bacterial community structuring in the freshwater sponge Ephydatia fluviatilis, Microb Ecol, 65 (2013) 232-244. [28] H. Izumi, E. Sagulenko, R.I. Webb, J.A. Fuerst, Isolation and diversity of planctomycetes from the sponge Niphates sp., seawater, and sediment of Moreton Bay, Australia, Antonie van Leeuwenhoek, 104 (2013) 533-546. [29] D. Sipkema, K. Schippers, W.J. Maalcke, Y. Yang, S. Salim, H.W. Blanch, Multiple approaches to enhance the cultivability of bacteria associated with the marine sponge Haliclona (gellius) sp, Appl Environ Microbiol, 77 (2011) 2130-2140. [30] S. Pimentel-Elardo, M. Wehrl, A.B. Friedrich, P.R. Jensen, U. Hentschel, Isolation of planctomycetes from Aplysina sponges, Aquat Microb Ecol, 33 (2003) 239-245. [31] M. Nickel, Like a 'rolling stone': quantitative analysis of the body movement and skeletal dynamics of the sponge Tethya wilhelma, J Exp Biol, 209 (2006) 2839-2846. [32] J.T. Hestetun, J. Vacelet, N. Boury-Esnault, C. Borchiellini, M. Kelly, P. Rios, J. Cristobo, H.T. Rapp, The systematics of carnivorous sponges, Mol Phylogenet Evol, 94 (2016) 327-345. [33] P.R. Bergquist, Invertebrate Zoology, Oxford University Press, New York, 1998.

Jo

ur na

lP

re

-p

ro of

[34] U. Hentschel, J. Piel, S.M. Degnan, M.W. Taylor, Genomic insights into the marine sponge microbiome, Nat Rev Microbiol, 10 (2012). [35] M. Wehrl, M. Steinert, U. Hentschel, Bacterial uptake by the marine sponge Aplysina aerophoba, Microb Ecol, 53 (2007) 355-365. [36] V. Gloeckner, M. Wehrl, L. Moitinho-Silva, C. Gernert, P. Schupp, J.R. Pawlik, N.L. Lindquist, D. Erpenbeck, G. Worheide, U. Hentschel, The HMA-LMA dichotomy revisited: an electron microscopical survey of 56 sponge species, Biol Bull, 227 (2014) 78-88. [37] U. Hentschel, K.M. Usher, M.W. Taylor, Marine sponges as microbial fermenters, FEMS Microbiol Ecol, 55 (2006) 167-177. [38] G. Lackner, E.E. Peters, E.J. Helfrich, J. Piel, Insights into the lifestyle of uncultured bacterial natural product factories associated with marine sponges, Proc Natl Acad Sci U S A, 114 (2017) E347-E356. [39] C.A. Bewley, D.J. Faulkner, Lithistid Sponges: Star Performers or Hosts to the Stars, Angewandte Chemie, 37 (1998) 2162-2178. [40] M.C. Wilson, T. Mori, C. Ruckert, A.R. Uria, M.J. Helf, K. Takada, C. Gernert, U.A. Steffens, N. Heycke, S. Schmitt, C. Rinke, E.J. Helfrich, A.O. Brachmann, C. Gurgui, T. Wakimoto, M. Kracht, M. Crusemann, U. Hentschel, I. Abe, S. Matsunaga, J. Kalinowski, H. Takeyama, J. Piel, An environmental bacterial taxon with a large and distinct metabolic repertoire, Nature, 506 (2014) 58-62. [41] C. Morrow, P. Cardenas, Proposal for a revised classification of the Demospongiae (Porifera), Front Zool, 12 (2015) 7. [42] R.W. Van Soest, N. Boury-Esnault, J. Vacelet, M. Dohrmann, D. Erpenbeck, N.J. De Voogd, N. Santodomingo, B. Vanhoorne, M. Kelly, J.N. Hooper, Global diversity of sponges (Porifera), PloS one, 7 (2012) e35105. [43] R.W.M. Van Soest, N. Boury-Esnault, J.N.A. Hooper, K.d.V. Rützler, N.J.,, B. Alvarez, E. Hajdu, A.B. Pisera, R. Manconi, C. Schönberg, M. Klautau, B. Picton, M. Kelly, J. Vacelet, M. Dohrmann, M.-C. Díaz, Cárdenas, J.L. P.; Carballo, P.D. Ríos, R., World Porifera database, (2018). [44] R.W.M. Van Soest, N. Boury-Esnault, J.N.A. Hooper, K. Rützler, N.J. de Voogd, B. Alvarez, E. Hajdu, A.B. Pisera, R. Manconi, C. Schönberg, M. Klautau, B. Picton, M. Kelly, J. Vacelet, M. Dohrmann, M.-C. Díaz, P. Cárdenas, J.L. Carballo, P. Ríos, R. Downey, World Porifera Database, in, 2019. [45] R. Manconi, R. Pronzato, Global diversity of sponges (Porifera: Spongillina) in freshwater, Hydrobiologia, 595 (2007) 27-33. [46] G. Worheide, M. Dohrmann, D. Erpenbeck, C. Larroux, M. Maldonado, O. Voigt, C. Borchiellini, D.V. Lavrov, Deep phylogeny and evolution of sponges (phylum Porifera), Adv Mar Biol, 61 (2012) 1-78. [47] O.V. Kaliuzhnaia, N.V. Kulakova, B.V. Itskovich, Diversity of polyketide synthase genes (PKS) in metagenomic community of the freshwater sponge, Mol Biol (Mosk), 46 (2012) 887893. [48] O.V. Kaluzhnaya, B.V. Itskovich, G.P. McCormack, Phylogenetic diversity of bacteria associated with the endemic freshwater sponge Lubomirskia baicalensis, World J Microbiol Biotechnol, 27 (2011) 955–1959. [49] O.V. Kaluzhnaya, V.B. Itskovich, Distinctive features of the microbial diversity and the polyketide synthase genes spectrum in the community of the endemic Baikal sponge Swartschewskia papyracea, Russian Journal of Genetics, 52 (2016) 38-48. [50] A. Talevska, B. Pejin, V. Kojic, T. Beric, S. Stankovic, A contribution to pharmaceutical biology of freshwater sponges, Nat Prod Res, 32 (2018) 568-571.

Jo

ur na

lP

re

-p

ro of

[51] A. Talevska, B. Pejin, T. Beric, S. Stankovic, Further insight into the bioactivity of the freshwater sponge Ochridaspongia rotunda, Pharm Biol, 55 (2017) 1313-1316. [52] T. Keller-Costa, A. Jousset, L. Overbeek, J.D. Elsas, R. Costa, The freshwater sponge Ephydatia fluviatilis harbours diverse Pseudomonas species (Gammaproteobacteria, Pseudomonadales) with broad-spectrum antimicrobial activity, PloS one, 9 (2014). [53] B. Pejin, A. Talevska, A. Ciric, J. Glamoclija, M. Nikolic, T. Talevski, M. Sokovic, Antiquorum sensing activity of selected sponge extracts: a case study of Pseudomonas aeruginosa, Nat Prod Res, 28 (2014) 2330-2333. [54] D. Axenov-Gribanov, Y. Rebets, B. Tokovenko, I. Voytsekhovskaya, M. Timofeyev, A. Luzhetskyy, The isolation and characterization of actinobacteria from dominant benthic macroinvertebrates endemic to Lake Baikal, Folia Microbiol (Praha), 61 (2015) 159-168. [55] O.V. Kalyuzhnaya, I.A. Lipko, V.B. Ickovich, O.V. Kalyuzhnaya, V.V. Parfenova, PCRscreening of bacteria isolated from freshwater sponge Lubomirskia baicalensis for detection of genes of secondary metabolites, Water: chemistry and ecology, 7 (2013) 70-74. [56] V.V. Parfenova, I.A. Terkina, T.Y. Kostornova, I.G. Nikulina, V.I. Chernykh, E.A. Maksimova, Microbial community of freshwater sponges in Lake Baikal, Izv Akad Nauk Ser Biol, 4 (2008). [57] E. Stackebrandt, M. Goodfellow, Nucleic acid techniques in bacterial systematics, Wiley, 1991. [58] S. Wiegand, M. Jogler, C. Boedeker, D. Pinto, J. Vollmers, E. Rivas-Marín, T. Kohn, S.H. Peeters, A. Heuer, P. Rast, S. Oberbeckmann, B. Bunk, O. Jeske, A. Meyerdierks, J.E. Storesund, N. Kallscheuer, S. Lücker, O.M. Lage, T. Pohl, B.J. Merkel, P. Hornburger, R.-W. Müller, F. Brümmer, M. Labrenz, A.M. Spormann, H. Op den Camp, J. Overmann, R. Amann, M.S.M. Jetten, T. Mascher, M.H. Medema, D.P. Devos, A.-K. Kaster, L. Øvreås, M. Rohde, M.Y. Galperin, C. Jogler, Deep-cultivation of Planctomycetes uncovers their unique biology, Nature Microbiology (accepted manuscript), manuscript ID # NMICROBIOL-18122682A (2019). [59] C. Bertelli, M.R. Laird, K.P. Williams, G. Simon Fraser University Research Computing, B.Y. Lau, G. Hoad, G.L. Winsor, F.S. Brinkman, IslandViewer 4: expanded prediction of genomic islands for larger-scale datasets, Nucleic Acids Res, (2017). [60] I. Lee, Y. Ouk Kim, S.C. Park, J. Chun, OrthoANI: An improved algorithm and software for calculating average nucleotide identity, Int J Syst Evol Microbiol, 66 (2016) 1100-1103. [61] J.P. Meier-Kolthoff, A.F. Auch, H.P. Klenk, M. Goker, Genome sequence-based species delimitation with confidence intervals and improved distance functions, BMC bioinformatics, 14 (2013) 60. [62] K. Blin, T. Wolf, M.G. Chevrette, X. Lu, C.J. Schwalen, S.A. Kautsar, H.G. Suarez Duran, E.L.C. de Los Santos, H.U. Kim, M. Nave, J.S. Dickschat, D.A. Mitchell, E. Shelest, R. Breitling, E. Takano, S.Y. Lee, T. Weber, M.H. Medema, antiSMASH 4.0-improvements in chemistry prediction and gene cluster boundary identification, Nucleic Acids Res, (2017). [63] M. Lechner, S. Findeiss, L. Steiner, M. Marz, P.F. Stadler, S.J. Prohaska, Proteinortho: detection of (co-)orthologs in large-scale analysis, BMC bioinformatics, 12 (2011) 124. [64] E. Pruesse, J. Peplies, F.O. Glöckner, SINA: accurate high-throughput multiple sequence alignment of ribosomal RNA genes, Bioinformatics, 28 (2012) 1823-1829. [65] A. Stamatakis, RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies, Bioinformatics, 30 (2014) 1312-1313. [66] J. Bondoso, J. Harder, O.M. Lage, rpoB gene as a novel molecular marker to infer phylogeny in Planctomycetales, Antonie van Leeuwenhoek, 104 (2013) 477-488.

Jo

ur na

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re

-p

ro of

[67] F. Sievers, A. Wilm, D. Dineen, T.J. Gibson, K. Karplus, W. Li, R. Lopez, H. McWilliam, M. Remmert, J. Söding, Fast, scalable generation of high‐quality protein multiple sequence alignments using Clustal Omega, Molecular systems biology, 7 (2011) 539. [68] J. Wittmann, B. Dreiseikelmann, C. Rohde, M. Rohde, J. Sikorski, Isolation and characterization of numerous novel phages targeting diverse strains of the ubiquitous and opportunistic pathogen Achromobacter xylosoxidans, PloS one, 9 (2014) e86935. [69] W.F. Tivol, A. Briegel, G.J. Jensen, An improved cryogen for plunge freezing, Microsc Microanal, 14 (2008) 375-379. [70] A. Rigort, F.J. Bauerlein, E. Villa, M. Eibauer, T. Laugks, W. Baumeister, J.M. Plitzko, Focused ion beam micromachining of eukaryotic cells for cryoelectron tomography, Proc Natl Acad Sci U S A, 109 (2012) 4449-4454. [71] M. Schaffer, J. Mahamid, B.D. Engel, T. Laugks, W. Baumeister, J.M. Plitzko, Optimized cryo-focused ion beam sample preparation aimed at in situ structural studies of membrane proteins, J Struct Biol, (2016). [72] X. Li, P. Mooney, S. Zheng, C.R. Booth, M.B. Braunfeld, S. Gubbens, D.A. Agard, Y. Cheng, Electron counting and beam-induced motion correction enable near-atomic-resolution single-particle cryo-EM, Nat Meth, 10 (2013) 584-590. [73] J.R. Kremer, D.N. Mastronarde, J.R. McIntosh, Computer visualization of threedimensional image data using IMOD, J Struct Biol, 116 (1996) 71-76. [74] S. Nickell, F. Forster, A. Linaroudis, W.D. Net, F. Beck, R. Hegerl, W. Baumeister, J.M. Plitzko, TOM software toolbox: acquisition and analysis for electron tomography, J Struct Biol, 149 (2005) 227-234. [75] A. Martinez-Sanchez, I. Garcia, S. Asano, V. Lucic, J.J. Fernandez, Robust membrane detection based on tensor voting for electron tomography, J Struct Biol, 186 (2014) 49-61. [76] P. Rast, I. Glockner, C. Boedeker, O. Jeske, S. Wiegand, R. Reinhardt, P. Schumann, M. Rohde, S. Spring, F.O. Glockner, C. Jogler, M. Jogler, Three Novel Species with Peptidoglycan Cell Walls form the New Genus Lacunisphaera gen. nov. in the Family Opitutaceae of the Verrucomicrobial Subdivision 4, Front Microbiol, 8 (2017) 202. [77] L.T. Miller, Single derivatization method for routine analysis of bacterial whole-cell fatty acid methyl esters, including hydroxy acids, J Clin Microbiol, 16 (1982) 584-586. [78] L.D. Kuykendall, M.A. Roy, apos, J.J. Neill, T.E. Devine, Fatty Acids, Antibiotic Resistance, and Deoxyribonucleic Acid Homology Groups of Bradyrhizobium japonicum, Int J Syst Evol Microbiol, 38 (1988) 358-361. [79] P. Kämpfer, R.M. Kroppenstedt, Numerical analysis of fatty acid patterns of coryneform bacteria and related taxa, Can J Microbiol, 42 (1996) 989-1005. [80] S.F. Altschul, T.L. Madden, A.A. Schaffer, J. Zhang, Z. Zhang, W. Miller, D.J. Lipman, Gapped BLAST and PSI-BLAST: a new generation of protein database search programs., Nucleic Acids Res, 25 (1997) 3389-3402. [81] Y. Bao, V. Chetvernin, T. Tatusova, Improvements to pairwise sequence comparison (PASC): a genome-based web tool for virus classification, Arch Virol, 159 (2014) 3293-3304. [82] R. Aghnatios, M. Drancourt, Gemmata species: Planctomycetes of medical interest, Future Microbiol, 11 (2016) 659-667. [83] P. Hirsch, M. Muller, Methods and sources for the enrichment and isolation of budding, nonprosthecate bacteria from freshwater, Microb Ecol, 12 (1986) 331-341. [84] P. Hirsch, M. Müller, Planctomyces limnophilus sp. nov., a stalked and budding bacterium from freshwater, Syst Appl Microbiol, 6 (1985) 276-280. [85] G. Reintjes, C. Arnosti, B.M. Fuchs, R. Amann, An alternative polysaccharide uptake mechanism of marine bacteria, ISME J, (2017).

Jo

ur na

lP

re

-p

ro of

[86] J.C. Orr, V.J. Fabry, O. Aumont, L. Bopp, S.C. Doney, R.A. Feely, A. Gnanadesikan, N. Gruber, A. Ishida, F. Joos, R.M. Key, K. Lindsay, E. Maier-Reimer, R. Matear, P. Monfray, A. Mouchet, R.G. Najjar, G.K. Plattner, K.B. Rodgers, C.L. Sabine, J.L. Sarmiento, R. Schlitzer, R.D. Slater, I.J. Totterdell, M.F. Weirig, Y. Yamanaka, A. Yool, Anthropogenic ocean acidification over the twenty-first century and its impact on calcifying organisms, Nature, 437 (2005) 681-686. [87] R.G. Wetzel, 11 - THE INORGANIC CARBON COMPLEX, in: Limnology (Third Edition), Academic Press, San Diego, 2001, pp. 187-204. [88] E. Stackebrandt, J. Ebers, Taxonomic parameter revisited: tarnished gold standards, Microbiol Today, 33 (2006) 152-155. [89] M. Kim, H.-S. Oh, S.-C. Park, J. Chun, Towards a taxonomic coherence between average nucleotide identity and 16S rRNA gene sequence similarity for species demarcation of prokaryotes, Int J Syst Evol Microbiol, 64 (2014) 346-351. [90] L.G. Wayne, D.J. Brenner, R.R. Colwell, P.A.D. Grimont, O. Kandler, M.I. Krichevsky, L.H. Moore, W.E.C. Moore, R.G.E. Murray, E. Stackebrandt, M.P. Starr, H.G. Truper, Report of the Ad Hoc Committee on Reconciliation of Approaches to Bacterial Systematics, Int J Syst Evol Microbiol, 37 (1987) 463-464. [91] B.J. Tindall, R. Rosselló-Móra, H.-J. Busse, W. Ludwig, P. Kämpfer, Notes on the characterization of prokaryote strains for taxonomic purposes, Int J Syst Evol Microbiol, 60 (2010) 249-266. [92] H. Brüssow, C. Canchaya, W.D. Hardt, Phages and the evolution of bacterial pathogens: from genomic rearrangements to lysogenic conversion, Microbiol Mol Biol Rev, 68 (2004) 560-602, table of contents. [93] A.A. Ivanova, D.G. Naumoff, K.K. Miroshnikov, W. Liesack, S.N. Dedysh, Comparative Genomics of Four Isosphaeraceae Planctomycetes: A Common Pool of Plasmids and Glycoside Hydrolase Genes Shared by Paludisphaera borealis PX4T, Isosphaera pallida IS1BT, Singulisphaera acidiphila DSM 18658T, and Strain SH-PL62, Front Microbiol, (2017). [94] R. Uebe, D. Schuler, C. Jogler, S. Wiegand, Reevaluation of the Complete Genome Sequence of Magnetospirillum gryphiswaldense MSR-1 with Single-Molecule Real-Time Sequencing Data, Genome Announc, 6 (2018). [95] C. Jogler, M. Kube, S. Schubbe, S. Ullrich, H. Teeling, D.A. Bazylinski, R. Reinhardt, D. Schuler, Comparative analysis of magnetosome gene clusters in magnetotactic bacteria provides further evidence for horizontal gene transfer, Environ Microbiol, (2009). [96] C. Jogler, W. Lin, A. Meyerdierks, M. Kube, E. Katzmann, C. Flies, Y. Pan, R. Reinhard, D. Schüler, Towards cloning the magnetotactic metagenome: Identification of magnetosome island gene clusters in uncultivated magnetotactic bacteria from different aquatic sediments, Appl Environ Microbiol in press, (2009). [97] N. Ward, J.T. Staley, J.A. Fuerst, S. Giovannoni, H. Schlesner, E. Stackebrandt, The Order Planctomycetales, Including the Genera Planctomyces, Pirellula, Gemmata and Isosphaera and the Candidatus Genera Brocadia, Kuenenia and Scalindua, in: M. Dworkin (Ed.) The Prokaryotes, Springer, New York, 2006, pp. 757-793. [98] K. Labutti, J. Sikorski, S. Schneider, M. Nolan, S. Lucas, T. Glavina Del Rio, H. Tice, J.F. Cheng, L. Goodwin, S. Pitluck, K. Liolios, N. Ivanova, K. Mavromatis, N. Mikhailova, A. Pati, A. Chen, K. Palaniappan, M. Land, L. Hauser, Y.J. Chang, C.D. Jeffries, B.J. Tindall, M. Rohde, M. Goker, T. Woyke, J. Bristow, J.A. Eisen, V. Markowitz, P. Hugenholtz, N.C. Kyrpides, H.P. Klenk, A. Lapidus, Complete genome sequence of Planctomyces limnophilus type strain (Mu 290), Stand Genomic Sci, 3 (2010) 47-56.

Jo

ur na

lP

re

-p

ro of

[99] E. Adriaenssens, J.R. Brister, How to Name and Classify Your Phage: An Informal Guide, Viruses, 9 (2017). [100] B.B. Hsu, T.E. Gibson, V. Yeliseyev, Q. Liu, L. Lyon, L. Bry, P.A. Silver, G.K. Gerber, Dynamic Modulation of the Gut Microbiota and Metabolome by Bacteriophages in a Mouse Model, Cell Host Microbe, 25 (2019) 803-814 e805. [101] L. Bai, C. Cao, C. Wang, H. Xu, H. Zhang, V.I. Slaveykova, H.L. Jiang, Towards Quantitative Understanding of the Bioavailability of Dissolved Organic Matter in Freshwater Lake during Cyanobacteria Blooming, Environ Sci Technol, (2017). [102] S. Yadav, R. Vaddavalli, S. Siripuram, R.V.V. Eedara, S. Yadav, O. Rabishankar, T. Lodha, S. Chintalapati, V. Chintalapati, Planctopirus hydrillae sp. nov., an antibiotic producing Planctomycete isolated from the aquatic plant Hydrilla and its whole genome shotgun sequence analysis, J Antibiot (Tokyo), 71 (2018) 575-583. [103] Z.M. Gao, Y. Wang, O.O. Lee, R.M. Tian, Y.H. Wong, S. Bougouffa, Z. Batang, A. AlSuwailem, F.F. Lafi, V.B. Bajic, P.Y. Qian, Pyrosequencing reveals the microbial communities in the Red Sea sponge Carteriospongia foliascens and their impressive shifts in abnormal tissues, Microb Ecol, 68 (2014) 621-632.

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Figure Legends

Fig. 1. Sampling location and sponge. Map showing the location of Lake Constance in the border region of Germany (DE), Switzerland (CH) and Austria (AT) a, the sampled Ephydatia sp. sponge in situ b and ex-situ c. Bar 1 cm. Planctopirus ephydatiae spb1 T Planctopirus limnophila Mü290T

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Planctopirus hydrillae JC280T

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Rubinisphaera brasiliensisDSM5305T

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Planctomicrobium piriforme P3T

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Gimesia maris 534-30T 55 100

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Rhodopirellula & Rubripirellula Roseimaritima ulvae UC8T

Mariniblastus fucicola FC18T

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Blastopirellula marina SH106T 100

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Blastopirellula cremea LHWP2T

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Fuerstia marisgermanicae NH11T 58

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Schlesneria paludicola PL7T 99

Pirellula staleyi ATCC27377T

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Bythopirellula goksoyri Pr1dT 100

Thermophilic group

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Gemmata obscuriglobus UQM2246T

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Gemmata massiliana IIL30 T

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Fimbriiglobus ruber SP5T Telmatocola sphagniphila SP2T

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Zarvazinella formosa A10 T Aquisphaera giovannonii OJF2T

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Singulisphaera

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Paludisphaera borealis PX4T Isosphaera pallida IS1BT Phycisphaerae Anammox

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Fig. 2. Phylogenetic tree of the phylum Planctomycetes. Phylogenetic position of strain spb1T (bold) within the phylum Planctomycetes based on 16S rRNA gene sequence comparison with the maximum likelihood algorithm. Bootstrap values after 1000 resamplings are given at the nodes. Selected species of the phylum Verrucomicrobia served as outgroup. The depicted phylogenetic tree displays described planctomycetal strains but was calculated in the presence of all sequenced isolates recently published by Wiegand et al.[58].

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Fig. 3. Light microscopy of strain spb1T and P. limnophila DSM 3776T. Phase-contrast images of type strain P. limnophila DSM 3776T (a,b) compared to the strain spb1T (c,d). Aggregates (a,c), single cells (b,d). Bar 1µm.

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Fig. 4. Scanning electron micrographs of strain spb1T and P. limnophila DSM 3776T. Type strain P. limnophila DSM 3776T (a-c) compared to strain spb1T (d-f). Aggregates with few (a,b) and lots of fiber-like structures (d,e). Crateriform pits (c,f, white arrowheads). Bar 1µm.

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Fig. 5. Transmission electron micrographs of strain spb1T and P. limnophila DSM 3776T. TEM analysis of negative stained P. limnophila DSM 3776T (a) and spb1T (b) cells. Fiber-like structures are more prominent in strain spb1T. Bar 1µm.

rpoB

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Fig. 6. Phylogeny of the genus Planctopirus based on marker genes and genome analyses, illustrated as Venn-diagrams. The three strains of the genus Planctopirus are depicted as different colored circles (blue: P. limnophila DSM 3776T, grey: P. hydrillae JC280T, yellow: spb1T). For the two marker genes, 16S rRNA and rpoB, the identity is given for each strain compared to the two others (as strain spb1 possesses two different 16S rRNA genes, values might differ by 0.1%). For Average Nucleotide Identity (ANI), the corresponding value is given, while for the genome-genome comparison, the total numbers of shared genes are given.

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Fig. 7. Bacteriophages of P. limnophila DSM 3776T. Cells of P. limnophila DSM 3776T with associated bacteriophages (a,b, white arrowheads) in transmission electron microscopy and a selected slice of the tomogram obtained by cryo-electron tomography (c). Bar 200 nm.