Developing Techniques for the Utilization of Planctomycetes As Producers of Bioactive Molecules

Abstract

Planctomycetes are conspicuous, ubiquitous, environmentally important bacteria. They can attach to various surfaces in aquatic habitats and form biofilms. Their unique FtsZ-independent budding cell division mechanism is associated with slow growth and doubling times from 6 h up to 1 month. Despite this putative disadvantage in the struggle to colonize surfaces, Planctomycetes are frequently associated with aquatic phototrophic organisms such as diatoms, cyanobacteria or kelp, whereby Planctomycetes can account for up to 50% of the biofilm-forming bacterial population. Consequently, Planctomycetes were postulated to play an important role in carbon utilization, for example as scavengers after phototrophic blooms. However, given their observed slow growth, such findings are surprising since other faster- growing heterotrophs tend to colonize similar ecological niches. Accordingly, Planctomycetes were suspected to produce antibiotics for habitat protection in response to the attachment on phototrophs. Recently, we demonstrated their genomic potential to produce non-ribosomal peptides, polyketides, bacteriocins, and terpenoids that might have antibiotic activities. In this study, we describe the development of a pipeline that consists of tools and procedures to cultivate Planctomycetes for the production of antimicrobial compounds in a chemically defined medium and a procedure to chemically mimic their interaction with other organisms such as for example cyanobacteria. We evaluated and adjusted screening assays to enable the hunt for planctomycetal antibiotics. As proof of principle, we demonstrate antimicrobial activities of planctomycetal extracts from Planctopirus limnophila DSM 3776, Rhodopirellula baltica DSM 10527, and the recently isolated strain Pan216. By combining UV/Vis and high resolution mass spectrometry data from high-performance liquid chromatography fractionations with growth inhibition of indicator strains, we were able to assign the antibiotic activity to candidate peaks related to planctomycetal antimicrobial compounds. The MS analysis points toward the production of novel bioactive molecules with novel structures. Consequently, we developed a large scale cultivation procedure to allow future structural elucidation of such compounds. Our findings might have implications for the discovery of novel antibiotics as Planctomycetes represent a yet untapped resource that could be developed by employing the tools and methods described in this study.

Keywords: Planctomycetes; Planctopirus limnophila; Rhodopirellula baltica; antibiotics; natural products; screening; secondary metabolites.

FIGURE 1

Growth of Planctopirus limnophila, Rhodopirellula baltica and strain Pan216 under different cultivation conditions. Logarithmic OD₆₀₀ values as measure of growth over 96 h for (A) P. limnophila, (B) R. baltica and 255 h for (C) strain Pan216 under different cultivation conditions: M1 and M2: complex culture media containing yeast extract, peptone and glucose (green curves). Maintain medium 1 or Maintain medium 2 (MM1 and MM2) are chemically defined but lack a carbon source and thus, unsupplemented, do not allow for bacterial growth of Planctomycetes (red curves). MM1 and MM2 were supplemented with glucose (blue curves), dextran (orange curves) or N- acetyl-D-glucosamine (NAG; pink curves) to allow growth of P. limnophila, R. baltica and strain Pan216. Each time point represents the mean of triplicate measurements.

FIGURE 2

High performance liquid chromatography (HPLC) analysis of R. baltica and P. limnophila extracts. (A) HPLC-analysis of extracts generated from P. limnophila cultivated in MM1 supplemented with glucose (green curve) and NAG (orange curve). Feeding glucose resulted in higher major peaks than NAG. Three minor peaks were exclusively produced if glucose was added (black asterisk), while one major peak was unique to NAG cultivation (red asterisk). (B) HPLC-analysis of extracts generated from R. baltica cultivated in MM2 supplemented with glucose (green curve) or NAG (orange curve). The intensity of the secondary metabolite peaks was higher in extracts from R. baltica cultivated with NAG as carbon source compared to extracts from cultivation with glucose. Extracts from glucose cultivation showed five peaks after 12 and 40–56 min with intensity only up to 2.2 × 10⁵ Au, while same signal peaks from extracts from NAG cultivation showed intensity up to 10⁶ Au. Three of the major peaks were significantly higher (red asterisk), while one showed less signal intensity (red arrow). The HPLC spectrum contains two additional minor peaks (black arrows) while one minor peak vanished (black asterisk). (C) Comparison of HPLC- chromatograms of XAD extracts from P. limnophila cultivated in MM1 (blue curve) and R. baltica in MM2 (red curve) supplemented with glucose as sole carbon source. Secondary metabolites signals from P. limnophila extracts were detected after a retention time of 7 and 26–38 min with an intensity of 5 × 10⁴ up to 3.8 × 10⁵ Au, while for R. baltica a signal was detected after 10 min with an intensity of 2.5 × 10⁵ Au and additional signals were detected after 44–57 min with an intensity of 1.8 × 10⁵ up to 2.4 × 10⁵ Au. (D) Comparison of HPLC- chromatograms of XAD extracts from P. limnophila cultivated in MM1 (blue curve) and R. baltica in MM2 (red curve) supplemented with NAG as sole carbon source. Secondary metabolite signals from P. limnophila were detected with intensity up to 3 × 10⁵ Au after a retention time of 25–38 min while secondary metabolite peaks from R. baltica extracts after 12 and 40–56 min with intensity up to 10⁶ Au.

FIGURE 3

Measurement of biomass production compared to optical density and glucose consumption of P. limnophila and R. baltica at constant pH. (A) Biomass production of P. limnophila compared to optical density at 600 nm (OD₆₀₀) and glucose consumption at constant pH over 6 days in MM1 supplemented with glucose in a 10 l fermenter. Glucose concentration decreased from 250 to 100 mg/l (blue curve), while the OD₆₀₀ increased from 0.15 up to 0.99 (purple curve) and correlating with the OD₆₀₀ also the dry weight increased from 0.1 g/l up to 1.8 g/l (red curve) during cultivation. The pH was 7.2 (green curve). (B) Biomass production of R. baltica compared to, OD₆₀₀-value and glucose consumption at constant pH over 6 days in MM2 supplemented with glucose in a 10 l fermenter. Glucose concentration decreased from 600 to 450 mg/l (blue curve), while the OD₆₀₀ increased from 0.2 to 1.170 (purple curve) and correlating with the OD₆₀₀ also the dry weight increased from 0.18 g/l and to 2.18 g/l (red curve) during cultivation. The pH was stabilized at 7.5 (green curve).

FIGURE 4

Activity assays of R. baltica methanol extracts. (A) Agar plate diffusion assay of R. baltica methanol cell extracts against Bacillus subtilis. Disks are coated with 90 μl methanol (MeOH), R. baltica cell or XAD extract respectively and incubated over night on soft-agar plates inoculated with B. subtilis. The clear areas around the disks treated with XAD extract show a distinct zone of B. subtilis growth inhibition, while methanol or cell extract show no effect. (B) Dual culture assay of R. baltica and E. coli. E. coli grew best with the maximal chosen distance of 15 mm to R. baltica. E. coli cells grew weaker when the distance was reduced to 10 mm and weakest at 2 mm (white arrows). (C) Dilution minimum inhibitory concentration (MIC) assay with R. baltica cell and XAD extracts. Growth inhibitory effect of cell and XAD methanol extracts on: treated E. coli TolC deletion mutant, S. aureus DSM 346 and M. luteus DSM 1790 cells. XAD: serial dilution column with XAD extract; Cell: serial dilution column with cell extract; C-: serial dilution column with methanol as negative control; C+: respectively untreated cells as positive control. Red circles highlight inhibited cells. If the same methanol concentration also inhibited the growth, the effect of the extracts was not taken into account (see C first two wells of XAD extract). The last well in which no bacteria grow contains the antibiotic at its minimal inhibitory concentration.

FIGURE 5

Fractionation analysis of planctomycetal crude extracts. (A) Combined chromatogram of HPLC fractionations, UV/Vis, mass- and tandem mass (MS/MS) spectrometry to identify the active compound, which causes the growth inhibition of P. limnophila crude extract #2 against B. subtilis on a 96 well plate. Fractionation of 15 μL extract for P. limnophila replicate #2 led to reproducible inhibition of B. subtilis growth in well ‘B6’, which had been collected from 23 to 23.5 min. The chromatogram shows a major peak at this retention time. The same peak was observed at a retention time of 22.4 min in the HRESIMS system. Its molecular formula C₂₆H₅₁NO₆ was deduced from its [M+H]⁺ peak at m/z 474.3880 as well as its [M+Na]⁺, [2M+H]⁺, and [2M+Na]⁺ peaks at m/z 496.3608, 947.7514, and 969.7325, respectively (Table 1). The assignment of the molecular ion cluster was further confirmed by the observation of the [M+CO₂H]^- peak at m/z 518.3 in the ESIMS spectrum measured in negative mode. (B) Combined chromatogram of HPLC fractionations, UV/Vis, mass spectrometry to identify three active compounds, which causes growth inhibition of B. subtilis of Pan216 crude extract #1 in a 96 well plate. In case of strain Pan216 both replicates resulted in different inhibition patterns as well. Inhibition of wells B7, C7, and G7 was observed that correspond to three major peaks of the slightly lipophilic region of the chromatogram. These three peaks were observed in HRESIMS spectrum at 25.1, 25.8, and 27.6 min; molecular formulae of C₁₈H₃₀O₂, C₂₂H₃₂O₂, and C₂₂H₃₄O₂ were assigned from their [M+H]⁺ and [M+Na]⁺ peak pairs (Table 1). These assignments were further confirmed by [M+H]^+/ [M-H]^- pairs in positive/negative mode ESIMS spectra. Wells A12 and H12 are negative controls treated with tetracycline and crude extract respectively.