Regular Paper
A Novel Strain of the Cyanobacterial Growth-promoting Bacterium, Rhodococcus sp. AF2108, Enhances the Growth of Synechococcus elongatus
2024 年 39 巻 4 号 論文ID: ME24050
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2024 年 39 巻 4 号 論文ID: ME24050
To enhance the growth of the cyanobacterium Synechococcus elongatus, the present study conducted direct screening for cyanobacterium growth-promoting bacteria (CGPB) using co-cultures. Of the 144 strains obtained, four novel CGPB strains were isolated and phylogenetically identified: Rhodococcus sp. AF2108, Ancylobacter sp. GA1226, Xanthobacter sp. AF2111, and Shewanella sp. OR151. A co-culture of S. elongatus with the most effective CGPB strain, Rhodococcus sp. AF2108, achieved a 8.5-fold increase in the chlorophyll content of cyanobacterial cells over that in a monoculture. A flow cytometric analysis showed a 3.9-fold increase in the number of S. elongatus cells in the co-culture with Rhodococcus sp. AF2108. These results were attributed to increases in forward scattering and chlorophyll fluorescence intensities. The new Rhodococcus strain appears to be one of the most effective CGPBs described to date.
Cyanobacteria, as photoautotrophic prokaryotes, require only sunlight, carbon dioxide (CO2), water, and minimal nutrients for growth, thereby eliminating the need for expensive carbon sources and complex growth media (Singh and Dhar, 2019). Cyanobacteria have been employed to capture and convert CO2 into other compounds, such as ethanol (Liang et al., 2018), isopropanol (Hirokawa et al., 2015), and fatty acids (Ruffing, 2014). However, further enhancements in the productivity of cyanobacteria for economic feasibility are currently limited by their growth rates. Therefore, strategies that enhance cyanobacterial proliferation have potential in the further development of cyanobacteria-based bioprocesses.
Co-cultivations with symbiont bacteria may promote the proliferation and metabolic synthesis of cyanobacteria via the exchange of metabolites, molecules, or nutrients through complex mechanisms. A previous study reported that the cyanobacterium Synechocystis sp. PCC 6803 degraded phenanthrene and also increased the cyanobacterial content of chlorophyll 8-fold when co-cultured for 20 days with the aerobic heterotrophic bacterium Pseudomonas sp. GM41 (Abed, 2010). These findings indicate the utility of technique for bioremediating oil-polluted sites that circumvents the high costs associated with the use of organic and inorganic fertilizers (Abed, 2010). Symbiont co-cultivation may also reduce the use of exogeneous nutrients because a cyanobacterium and its symbiont working together may efficiently exchange remineralized compounds, thereby increasing nutrient cycling and reducing the discharge of wastewater (Wu et al., 2023). Therefore, a co-culture approach may be an eco-friendly alternative for enhancing biomass production.
In the present study, we performed symbiont co-culture screening for the cyanobacterium S. elongatus using bacterial isolates derived from agricultural wastewater. We obtained, taxonomically identified, and characterized novel cyanobacterium growth-promoting bacteria (CGPBs).
CGPBs were isolated from agriculture wastewater derived from cattle urine by a simplified activated sludge treatment. Agriculture waste has been used as a liquid fertilizer via phytohormone-like responses in Arabidopsis thaliana (Kato and Konishi, 2024). With the aim of increasing the variety of isolated species, six different isolation media were prepared for use in the present study: JCM medium number 520 (JCM 520) agar (2.0% [w/v] agar) (Biebl and Pfennig, 1981), consisting of 0.5 g L–1 KH2PO4, 0.25 g L–1 CaCl2·2H2O, 3.0 g L–1 MgSO4·7H2O, 0.68 g L–1 NH4Cl, 20 g L–1 NaCl, 3.0 g L–1 sodium malate, 3.0 g L–1 sodium pyruvate, 0.4 g L–1 yeast extract, 5 mg L–1 ferric citrate, 2 mg L–1 vitamin B12, 70 μg L–1 ZnCl2·5H2O, 100 μg L–1 MnCl2·4H2O, 60 μg L–1 H3BO3, 200 μg L–1 CoCl2·6H2O, 20 μg L–1 CuCl2·2H2O, 20 μg L–1 NiCl2·6H2O, and 40 μg L–1 Na2MoO4·H2O, pH adjusted to 6.8, and Ormerod agar (2.0% [w/v] agar) (Ormerod et al., 1961), composed of 2.0 g L–1 sodium malate, 0.1 g L–1 yeast extract, 0.2 g L–1 MgSO4·7H2O, 0.08 g L–1 CaCl2·H2O, 0.01 g L–1 FeSO4·7H2O, 0.9 g L–1 K2HPO4, 0.6 g L–1 KH2PO4, 1.25 g L–1 (NH4)2SO4, 0.02 g L–1 ethylenediaminetetraacetic acid sodium salt (EDTA-Na), 0.028 g L–1 H3BO3, 0.021 g L–1 MnSO4·4H2O, 0.075 g L–1 Na2MoO4·2H2O, 0.0024 g L–1 ZnSO4·2H2O, and 0.01 g L–1 Cu(NO3)2·3H2O, pH adjusted to 7.0. Agricultural wastewater, directly used as a medium component, was sterilized by autoclaving or filtration and was then used to prepare agar (20 g L–1) or gellan gum (0.8 g L–1) plates. No cations were added to agricultural wastewater for the preparation of gellan gum plates because wastewater contained a sufficient amount of cations to gelate the plates. Series-diluted agricultural wastewater was then spread on the plates and incubated aerobically at 24°C. Single colonies appearing on plates were selected and re-cultured in 20 mL of a rich medium containing 10 g L–1 peptone, 5 g L–1 NaCl, and 6 g L–1 yeast extract at 30°C for 48 h with rotary shaking at 168 rpm. Cultured bacterial cells were preserved at –80°C as 25% (w/v) glycerol stock before use. Isolated strains were named by alphabetical letters and numbers, with JM, OR, AF, AA, GF, and GA indicating the following isolation media: JCM520 agar, Ormerod agar, filtrated agricultural wastewater agar, autoclaved agricultural waste agar, filtrated agricultural wastewater gellan gum, and autoclaved agricultural wastewater gellan gum, respectively.
Screening of CGPBsSynechococcus elongatus PCC 7942 purchased from the Pasteur Culture Collection of Cyanobacteria (Institute Pasteur, France) was cultivated at 25°C for 7 days in BG11 medium. To select CGPBs, all bacterial strains and S. elongatus were cultured using a co-culture medium (Liu et al., 2021). To prepare a seed culture of S. elongatus, the cyanobacterial strain was cultivated in a 100-mL Erlenmeyer flask containing 40 mL of the co-culture medium at 30°C for 7 days with rotary shaking at 120 rpm in a thermostatic shaking incubator (Bioshaker BR-43FL; Taitec) under continuous 115–120 μmol m–2 s–1 photosynthetic photon flex density (PPFD) illumination (white LED irradiation unit, LC-450EXP; Taitec). Isolates from the glycerol stock (50-μL aliquots) were cultured in 1 mL of co-culture medium in 96-deep well plates at 30°C for 48 h with shaking at 1,200 rpm in a Deep Well Maximizer Bioshaker (M·BR-022UP; Taitec). The prepared S. elongatus (adjusted as 0.05 of OD730 Unit) and bacterial cells (adjusted as 0.01 of OD600 Unit) were inoculated into 200 μL of the co-culture medium in a 96-well microplate (Violamo Plate; As One). The microplate was incubated at 30°C with shaking at 600 rpm using a microplate mini shaker (PSU-2T; Biosan) in a growth chamber (MLR-352; PHCbi) under continuous 115–120 μmol m–2 s–1 PPFD illumination for 6 days, which was shorter than the normal flask cultivation of S. elongatus. To evaluate the growth-promoting effects of each bacterial strain, the intensity of chlorophyll fluorescence was measured at excitation and emission wavelengths of 488 and 683 nm, respectively, as an index of the growth of S. elongatus using a fluorescence plate reader (Varioskan LUX; Thermo Fisher Scientific). The fluorescence intensity of monocultured bacterial cells was also measured as the control.
Taxonomical analysis of CGPBsThe genomic DNAs of CGPB strains were prepared using the alkaline-SDS method, with some modifications (Sambrook et al., 1989). CGPB culture broth (1 mL) was centrifuged at 10,000×g at 4°C for 1 min. Each cell pellet was resuspended in 560 μL Tris-EDTA buffer (pH 8.0), and 10 μL of 10 mg mL–1 proteinase K (Wako Pure Chemical), and 30 μL of 10% (w/v) SDS was then added. The suspension was incubated at 37°C for 1 h. Genomic DNA was twice extracted with phenol/chloroform/isoamyl alcohol (48:24:1) and purified by isopropanol precipitation. The partial 16S ribosomal RNA (16S rRNA) gene was amplified with the primers 27F (5′-AGRGTTYGATYMTGGCTCAG-3′) and 1492R (5′-RGYTACCTTGTTACGACTT-3′). Amplification was performed on a T-100 thermocycler (Bio-Rad) using the following protocol: initial denaturation at 95°C for 3 min, followed by 30 cycles at 95°C for 30 s, 55°C for 30 s, and 72°C for 2 min, with a final elongation at 72°C for 6 min. Amplified products were purified using the Wizard SV Gel and PCR Clean-Up system (Promega). Sequences were generated from the purified products with BigDye Terminator Cycle Sequencing kit v3.1 (Thermo Fisher Scientific) and the following primers: 27F (5′-AGRGTTYGATYMTGGCTCAG-3′), 341F (5′-CAATGGRSGVRASYCTGAHS-3′), 909F (5′-AAACTYAAARRAATTGACGG-3′), P699D (5′-YAACGAGCGMRACCC-3′), 1492R (5′-RGYTACCTTGTTACGACTT-3′), P699R (5′-GGGTYKCGCTCGTTR-3′), 518R (5′-ATTACCGCGGCTGCTGG-3′), and 338R (5′-TGCTGCCTCCCGTAGGAGT-3′) (Klindworth et al., 2013). The resulting sequences were analyzed on an ABI Prism 3130 DNA analyzer (Applied Biosystems Hitachi). After a BLAST search analysis against the NCBI GenBank database (http://www.ncbi.nlm.nih.gov/), sequences were aligned in ClustalW v2.0 and subjected to a phylogenetic analysis by the neighbor-joining method (Larkin et al., 2007). A phylogenetic tree was generated using TreeView v1.6.6 (Page, 1996). DNA sequences were deposited in the DDBJ database under accession numbers LC731267–LC731270. To examine morphological characteristics, CGPBs were cultured in AccuDiaTM Standard Method Agar (Shimadzu Diagnostics Corporation) at 30°C for 72 h, and were observed under a microscope (Nikon Ts2R, Nikon) with a ×100 magnitude lens using the difference interface method. Gram staining was performed using a common procedure. Microscopic images were obtained by a digital camera system (Ti2; Nikon). Cell sizes were measured by imaging software (NIS-Elements, Nikon).
Flask cultivationInocula of isolated strains and S. elongatus were prepared by 100-mL shake flask cultivation in the described rich medium and BG11, respectively. The inoculation ratio and number of days of the CGPB pre-culturation used for flask cultivation were those that exerted the strongest growth-promoting effects on S. elongatus in preliminary investigations. CGPB seed cultures were prepared by shake flask cultivation in 100-mL Erlenmeyer flasks containing 40 mL of the co-culture medium at 30°C with rotary shaking at 168 rpm. Co-cultures, which were conducted using the same culture medium as in the screening experiment, were performed in 100-mL Erlenmeyer flasks containing 40 mL of the co-culture medium at 30°C for 7 days with rotary shaking at 120 rpm.
Chlorophyll contents were measured as described in a previous study (Broddrick et al., 2016). Following cultivation, 1-mL samples of culture broth were collected and centrifuged at 15,000×g at 4°C for 7 min. The supernatant was removed, and cell pellets were resuspended in 1 mL of chilled methanol. To extract chlorophyll from cells, samples were incubated at 4°C for 1 h in the absence of light. The absorbance of each sample at 665 and 720 nm was measured using a Thermo Scientific Multiskan Sky spectrometer (Thermo Fisher Scientific), with methanol as the blank for calibration purposes. The concentration of chlorophyll a was calculated using the following equation:
Chl a (μg mL–1)=12.9447(A665–A720)
S. elongatus cell numbers and forward scattering and chlorophyll fluorescence intensity per cell were measured using a CyFlow Cube 8 flow cytometer (Sysmex) equipped with a 488-nm laser. The voltages of forward scattering (FSC-H), side scattering (SSC-H), and chlorophyll fluorescence (FL2-H filter; cut-off of 675 nm) were set at 200.0, 275.0, and 675.0 V, respectively. To detect S. elongatus, detection gating was set by the fluorescence intensity of chlorophyll in flow cytometry software. We herein confirmed that background fluorescence derived from the isolated strains may be ignored in the analysis. The morphological assay of S. elongatus using a microscope was performed using the above described methods.
Indole-3-acetic acid (IAA) and siderophore assaysIAA production was estimated using a colorimetric method based on Salkowski reagent (Ehmann, 1977). Strains were cultivated in IAA production medium containing 10 g L–1 peptone, 6.0 g L–1 yeast extract, 1.0 g L–1 L-tryptophan, and 5.0 g L–1 NaCl at 30°C for 2 days. After cultivation, the supernatant was isolated by centrifugation at 3,000×g at 4°C for 15 min. Two hundred microliters of the supernatant, 10 μL of 3.5 mM phosphate buffer (pH 7.0), and 400 μL of Salkowski reagent were mixed on a microplate and incubated for 1 h in the dark. Absorbance at 535 nm was measured using a microplate reader. IAA concentrations were calculated by a standard curve. Pseudomonas simiae OLi (DSM 18861), which was obtained from the German Collection of Microorganism and Cell Cultures GmbH (DSMZ, German), was used as the positive control in the IAA assay. Colorimetric measurements were conducted in triplicate. Siderophore production was analyzed using the chromo azurol S (CAS) blue agar assay (Schwyn and Neilands, 1987). P. fluorescens NBRC 14160, which was obtained from the National Biological Resource Center (NBRC), the National Institute of Technology and Evaluation, Japan, was used as a siderophore positive control.
Whole-cell hydrogenase assayA hydrogenase assay was performed as described in a previous study (Sebastiampillai et al., 2022). Isolated strains were cultured in modified tryptone-yeast extract-Tris (TYET) including 10 g L–1 tryptone, 5 g L–1 yeast extract, 50 mM Tris-HCl (pH 7.5), 4 g L–1 glucose 30 mM formate, 1 μM sodium selenite, and 1 μM sodium molybdate, which had been prepared through filtration with 0.22-μm polyethersulfone membrane. Cells of the isolated strain with 0.1 OD630 unit were inoculated into 200 μL of TYET broth in a 96-well plastic plate (Violamo Plate; As One Corporation) and then incubated at 30°C for 6 h. After the incubation, OD630 was measured using a plate reader (Varioskan LUX; Thermo Fisher Scientific). A developing solution (20 μL), including 10 mg mL–1 benzyl viologen and 250 mM sodium formate in 20 mM Tris-HCl buffer (pH 7.5), was added to each well. Changes in A630 were monitored by the plate reader every 30 s for 5 min. Increases in A630 per OD630 for 1 min were calculated from the time course.
In the present study, we isolated and screened 144 potential CGPB strains for their effects on the growth of S. elongatus. Due to equipment limitations, the screening experiment was conducted in two batches. In the first CGPB screening, 33 strains increased chlorophyll fluorescence when co-cultured with S. elongatus (Fig. 1). Strains AF2108, GA1226, AF2111, and OR151 enhanced the growth of co-cultured S. elongatus by 7.5-, 5.3-, 4.7-, and 3.9-fold, respectively, from that in the monoculture. These strains were selected for subsequent experiments.
The co-culture combination with Synechococcus elongatus PCC 7942 in the co-culture screening experiment in which the chlorophyll fluorescence intensity of S. elongatus PCC 7942 was more than 1.0-fold higher than that in the mono-culture of S. elongatus PCC 7942 on day 6. Bar graphs (A) and (B) show two batches of the screening experiment conducted in the present study. Error bars indicate standard deviations (n=3).
According to the phylogenetic analysis of their 16S rRNA gene sequences, the four isolated CGPB strains belonged to three classes: Actinomycetia, Alphaproteobacteria, and Gammaproteobacteria (Fig. 2). Strain AF2108 was closely related (99.95% sequence similarity) to the Gram-positive actinomycete Rhodococcus cerastii C5 (Kämpfer et al., 2013). Morphological observations (Fig. S1) showed that AF2108 was a Gram-positive, non-motile, and short-rod (length of 1.09–3.48 μm and width of 0.39–0.98) strain. Colonies were round, convex, and yellowish orange in color. These morphological characteristics corresponded to those of the type strain R. cerastii C5 (Kämpfer et al., 2013). Therefore, strain AF2108 was identified as Rhodococcus sp., which was closely related to R. cerastii. In the phylogenetic tree (Fig. 2), strains GA1226 and AF2111 belonged to the Xanthobacteraceae family of Gammaproteobacteria and were closely related to Ancylobacter rudongensis AS1.1761T and Xanthobacter flavus NBRC 14759T, with 99.56 and 100.0% sequence similarities, respectively. GA1226 was a Gram-negative, non-motile, curved-rod (length of 1.55–3.65 μm and width of 0.45–0.94 μm) strain (Fig. S1). Colonies were white, round, convex, and opaque, which corresponded to the type strain A. rudongensis (Xin et al., 2004). AF2111 was a Gram-negative, rod-shape (length of 1.00–4.56 μm and width of 0.58–1.02 μm) strain. Colonies were yellow, round, entire, convex, and opaque. These morphological characteristics corresponded to those of the type strain X. flavus (Malik and Claus, 1979). Therefore, strains GA1226 and AF2111 were assigned as Ancylobacter sp. and Xanthobacter sp., respectively. The phylogenetic analysis placed strain OR151 within the genus Shewanella. OR151 was a Gram-negative, motile, rod-shaped (length of 1.04–2.43 μm and width of 0.39–0.97 μm) strain. Colonies were beige, circular, and convex. In the phylogenetic tree, strain OR151 was the most closely related to Shewanella putrefaciens and Shewanella profunda; however, the strain was not assigned to either species based on the partial 16S rRNA sequence and, thus, was designated as Shewanella sp. OR151. According to the phylogenetic analysis, the four CGPB strains were widely distributed across bacterial phyla.
16S rRNA-based phylogenetic reconstruction of four aerobic heterotrophic bacterial strains used in the present study. Bacillus licheniformis ATCC 14580T was set as the outgroup. Bootstrap value ≥50%.
After optimizing seed culture periods and CGPB inoculation rates in multi-well plate cultures (Fig. S2), we performed flask co-cultures to confirm the growth-promoting activity of the CGPB strains in more detail. The seed culture periods and inoculation rates of Rhodococcus sp. AF2108, Ancylobacter sp. GA1226, Xanthobacter sp. AF2111, and Shewanella sp. OR151 were set to 1 day and OD600=0.02, 1 day and OD600=0.04, 2 days and OD600=0.02, and 2 days and OD600=0.02, respectively. As shown in Fig. 3, we monitored the content of chlorophyll a over time as a cyanobacterial growth index during flask-based cyanobacteria–CGPB co-cultivation and cyanobacterial monocultivation. When Rhodococcus sp. AF2108 was co-cultured with S. elongatus, the content of chlorophyll a derived from S. elongatus was 8.5-fold higher after 168 h than that of the monocultured cyanobacterium. At each corresponding time point, the other CGPBs were found to exert weaker growth-promoting effects. The highest fold change in growth observed in the co-culture of Ancylobacter sp. GA1226 with S. elongatus from that in the monoculture with the cyanobacterium alone, 2.8-fold, was recorded after 72 h. When co-cultured with S. elongatus, Xanthobacter sp. AF2111 and Shewanella sp. OR151 increased cyanobacterial growth by 1.3- and 1.7-fold, respectively, after 168 h. However, the growth-promoting effects of Xanthobacter sp. AF2111 and Shewanella sp. OR151 co-cultured with S. elongatus in flasks were weaker than those obtained using multi-well microplates. These differences in cyanobacterial growth promotion were caused by variations in physical conditions (other than temperature), such as the aeration level (agitation rate) and light intensity, between the two experiments due to differences in vessel sizes. Regardless of these differences between culture scales, Rhodococcus sp. AF2108 exhibited highly stable cyanobacterial growth-promoting activity. According to flow cytometric data, the numbers of S. elongatus cells counted in co-cultures with Rhodococcus sp. AF2108, Ancylobacter sp. GA1226, and Shewanella sp. OR151 were 3.9-, 2.0-, and 1.5-fold higher than that in the corresponding monoculture (Table 1). In contrast, co-culturing S. elongatus with Xanthobacter sp. AF2111 did not significantly increase the number of cyanobacterial cells counted. The intensities of forward and side scattering as the index of cell size were measured in the present study. Even though the intensity of side scattering was 0.9-fold weaker in both co-cultures with Rhodococcus sp. AF2108 and Shewanella sp. OR151, forward scattering was 1.5- and 1.7-fold larger, respectively, in co-cultures with the aforementioned strains than that of monocultured cyanobacteria. On the other hand, no significant differences were observed between co-cultures and the monoculture in the case of Ancylobacter sp. GA1226 and Xanthobacter sp. AF2111 (Table 1). To confirm cell morphology during co-cultures, microscope observations were performed (Table S1). The cell width of cyanobacterial cells markedly increased in cases of Rhodococcus sp. AF2108 and Shewanella sp. OR151. The observed cell area in cases of Rhodococcus sp. AF2108 and Shewanella sp. OR151 also corresponded to the results of forward scattering by flow cytometry. According to microscopic observations, Ancylobacter sp. GA1226 and Xanthobacter sp. AF2111 did not appear to affect cyanobacterial cell sizes. Cell size depends on the basic processes of cell physiology, and changes in cell size have a profound impact on metabolic flux, biosynthetic capacity, and nutrient exchange (Marshall et al., 2012). In a previous study, cell enlargement was noted following supplementation with metal ions (Nishino et al., 2018). We also measured the chlorophyll fluorescence intensity per cell of S. elongatus under each cultivation condition (Table 1). After co-culturing with Rhodococcus sp. AF2108 or Shewanella sp. OR151, chlorophyll fluorescence intensities per cell were 2.0- and 1.3-fold higher, respectively, than that of the cyanobacterium in the monoculture; therefore, co-culturing with Ancylobacter sp. GA1226 or Xanthobacter sp. AF2111 did not significantly affect chlorophyll fluorescence intensity per cell. These results revealed that co-cultivations with different partner strains led to not only in differences in the total amount of chlorophyll a, but also the total number of cells, cell size, and chlorophyll fluorescence intensity per cell of S. elongatus.
The growth of Synechococcus elongatus co-cultured with and without CGPB strains (AF2108, GA1226, AF2111, and OR151) in the flask scale co-culture experiment. The growth of S. elongatus was monitored by measuring the content of chlorophyll a. Error bars indicate standard deviations (n=3).
Flow cytometry specifically analyzed Synechococcus elongatus on day 7 after the co-culture with CGPBs and the monoculture.
| Number of cells counted (×108 mL–1) |
Forward scatter intensity per cell (×102) |
Side scatter intensity per cell (×10) |
Chlorophyll fluorescence intensity cell–1 (×102) |
|
|---|---|---|---|---|
| Co-culture with AF2108 | 17.8±0.8* | 13.9±0.4* | 4.95±0.37* | 1.94±0.18* |
| Co-culture with GA1226 | 9.3±0.3* | 8.1±0.0* | 5.48±0.95 | 1.21±0.10 |
| Co-culture with AF2111 | 5.8±0.6 | 9.6±0.4 | 6.07±0.50 | 1.46±0.25 |
| Co-culture with OR151 | 6.7±0.3* | 14.9±0.3* | 4.96±0.12* | 1.24±0.10* |
| Monoculture of S. elongatus | 4.6±0.1 | 9.1±0.1 | 5.64±0.06 | 0.98±0.09 |
Note: Values are means±standard deviations (n=3). Asterisks indicate significant differences (P<0.05) from S. elongatus (monoculture). The significance of differences was calculated by the Tukey-Kramer method.
To estimate the cyanobacterial growth promoting factors of the CGPBs obtained, siderophore production, IAA production, and whole-cell hydrogenase activity were measured (Table S2). Siderophore production by Rhodococcus sp. AF2108 and Shewanella sp. OR151 was positive in the CAS blue agar assay, with P. fluorescens as the positive control, whereas that by Ancylobacter sp. GA1226 and Xanthobacter sp. AF2111 was only weakly positive. All CGPB strains produced IAA; however, the amount of IAA was less than 18% of the positive control, P. simiae. Whole-cell hydrogenase activity was observed for Shewanella sp. OR151, but not for the other strains.
In the present study, CGPBs were successfully isolated and efficiently enhanced the growth of the cyanobacterium S. elongatus. Table 2 summarizes the results of various studies on the enhancing effects of co-cultures on the growth of cyanobacteria. Isolates of Rhodococcus sp., Ancylobacter sp., Xanthobacter sp., and Shewanella sp. were described as CGPBs in the present study. In a previous study, Pseudomonas sp. GM41, which was identified as the most effective CGPB analyzed (Abed, 2010), induced an 8-fold increase in the cyanobacterial content of chlorophyll during a 20-day co-cultivation with the cyanobacterium Synechocystis PCC6803 in the presence of hexadecane. In the present study, Rhodococcus sp. AF2108 increased the content of chlorophyll a in S. elongatus by 8.5-fold during a 7-day co-cultivation (Fig. 3). This result indicates that Rhodococcus sp. AF2108 exerted stronger growth-promoting effects over a shorter period than Pseudomonas sp. GM41; therefore, to the best of our knowledge, Rhodococcus sp. AF2108 is the most effective CGPB described to date. Synechococcus is a promising host for producing biomass and metabolites, such as those generated by photosynthesis from atmospheric carbon dioxide via synthetic metabolic pathways, and may be genetically modified (Oliver and Atsumi, 2015). Nevertheless, the low proliferation rate of Synechococcus limits production efficiency. Difficulties are associated with enhancing the growth rate through improvements in the flux of total metabolic pathways via metabolic engineering. Even though the mechanisms responsible for the observed growth promotion are presently unknown, the CGPBs isolated in the present study may contribute to a new Synechococcus growth improvement strategy. A CGPB–Synechococcus co-culture system may be applied to various bioprocesses using autotrophic Synechococcus metabolic pathways. We noted that co-culturing with Rhodococcus sp. AF2108 increased cyanobacterial growth, as assessed by the content of chlorophyll and cell number indexes, by 8.5-fold (Fig. 3) and 3.9-fold (Table 1), respectively, over that in the cyanobacterial monocultivation.
Effects of co-culturing cyanobacteria with bacteria or microalgae reported from various studies.
| Co-culture | Culture time (day) | Product | Fold change | Ref. | |
|---|---|---|---|---|---|
| Cyanobacteria | Co-culture microbes | ||||
| Synechococcus elongatus PCC 7942 | Rhodococcus sp. AF2108 | 7 | Biomass (Chlorophyll a) | 8.5 | This study |
| Ancylobacter sp. GA1226 | 2.8 | ||||
| Xanthobacter sp. AF2111 | 1.3 | ||||
| Shewanella sp. OR151 | 1.7 | ||||
| Synechococcus elongatus PCC 7942 | Rhodotorula glutinis ATCC 204091 | 30 | Lipid yield | 1.4–1.6 | Li et al., 2017 |
| Synechocystis sp. PCC6803 | Pseudomonas sp. GM41 | 20 | Biomass (Chlorophyll a) | 8 | Abed, 2010 |
| Synechocystis salina LEGE 06079 | Chlorella vulgaris CCAP 211/11B | 7 | Biomass (dry cell weight) |
1.62 | Gonçalves et al., 2016 |
| Pseudokirchneriella subcapitata CCAP 278/4 | 1.07 | ||||
| Microcystis aeruginosa LEGE 9134 | 1.40 | ||||
| Spirulina platensis UTEX 1926 | Rhodotorula glutinis 2.541 | 5 | Lipid production | 3.92 | Xue et al., 2010 |
In the present study, we did not examine the mechanisms underlying cyanobacterial growth enhancements. Based on previous findings on plant growth-promoting bacteria (PGPBs) and microalgae growth-promoting bacteria (MGPBs), four hypotheses have been proposed to explain how CGPBs enhance cyanobacterial growth: (1) by exchanging nutrients between CGPBs and cyanobacteria, (2) by enhancing nitrogen fixation via CGPBs, (3) by signaling via phytohormones or phytohormone-like factors, and (4) by enhancing the uptake of insoluble materials by siderophores.
Cyanobacteria exude dissolved organic matter, which becomes available for bacteria. In return, bacteria remineralize sulfur, nitrogen, and phosphorus to support the further growth of cyanobacteria (Buchan et al., 2014). In specific interactions, bacteria supply vitamin B group derivatives as organic cofactors or produce siderophores to bind iron; siderophores increase the bioavailability of iron for cyanobacteria, while cyanobacteria, in return, provide dissolved organic carbon for bacteria (Croft et al., 2005; Amin et al., 2012). In a previous study, Rhodococcus sp. increased the biomass of Chlamydomonas reinhardtii cc124 by 46% over that of the monocultured microalgal strain (Lakatos et al., 2014). In addition, Rhodococcus sp. promoted microalgal growth and hydrogen production during co-cultivation, which indicated that oxygen elimination is the most crucial factor for algal hydrogen production and that efficient bacterial respiration is essential for the activation of algal Fe-hydrogenase. At the same time, increased carbon dioxide emissions from bacteria may enhance carbon fixation by microalgae. In the present study, co-culturing with Rhodococcus sp. AF2108 resulted in cyanobacterial growth that was 8.5-fold (Fig. 3) and 3.9-fold (Table 1) higher, according to the content of chlorophyll and cell number indexes, respectively, than that of cyanobacteria in a monoculture. These fold changes in growth were larger than that in a previous study, which reported a 1.46-fold increase in C. reinhardtii cc124 biomass (Lakatos et al., 2014). Although the cyanobacterial growth-promoting mechanism of Rhodococcus sp. AF2108 is unknown, the marked increase observed in the growth of S. elongatus cannot be solely explained—unlike in the previous study (Lakatos et al., 2014)—by enhanced proliferation and respiration due to decreases and increases in the levels of dissolved oxygen and carbon dioxide, respectively.
Xanthobactor autotrophicus is a bacterial species that fixes nitrogen by simultaneously expressing hydrogenase and nitrogenase. The genus Xanthobacter is also exploited for sustainable ammonia and biofertilizer production (Liu et al., 2017). However, Xanthobacter sp. AF2111 did not exhibit whole-cell hydrogenase activity (Table S2). Based on the present results, nitrogen fixation by Xanthobacter sp. AF2111 did not appear to promote cyanobacterial growth.
In a previous study, plant-associated Rhodococcus qingshengii RL1, in the Actinomycetia clade in the phylogenetic tree (Fig. 2), exhibited the ability to produce the phytohormone, IAA (Kuhl et al., 2021). In the case of microalgae, extracellular IAA has been reported to enhance algal growth (Dao et al., 2018); however, it currently remains unclear whether IAA enhances the growth of Synechococcus. In a previous study in which cyanobacteria were exposed to chromium, exogenous supplementation with IAA and kinetin (a cytokinin-like synthetic phytohormone) attenuated the effects of chromium toxicity by decreasing chromium uptake (Tiwari et al., 2020). In the present study, all CGPBs produced a small amount of IAA from tryptophan (Table S2). Therefore, IAA produced by the strains during the co-culture may have stimulated cyanobacterial growth. However, it may not the sole factor promoting cyanobacterial growth because the ability to produce IAA was markedly weaker than that of the positive control species, P. simiae.
Members of the genus Shewanella decrease toxicity in the environment by reducing various electron acceptors (Dikow, 2011). In a previous study, a species of Shewanella rapidly protected Synechococcus sp. PCC7002 from Fe2+ toxicity before major oxidation induced cellular damage (Szeinbaum et al., 2021). Although iron plays a key role in cyanobacterial physiology, an excess of free intracellular iron is extremely harmful because it catalyzes the formation of reactive oxygen species (ROS) through Fenton reactions, leading to oxidative stress (Sauer et al., 2001). Therefore iron uptake and metabolism need to be tightly regulated in order to ensure a supply that maintains intracellular concentrations within non-toxic levels (González et al., 2012). In a previous study, S. putrefaciens W3-18-1 increased iron availability and provided more facile, energy-efficient mechanisms for iron acquisition by Synechococcus sp. PCC7002, a conclusion inferred from the broad and consistent decrease in the mRNA levels of Fe-regulated Synechococcus sp. PCC7002 genes during the co-cultivation (Beliaev et al., 2014). These findings showed that Shewanella may have expanded the upper tolerable Fe2+ concentration and increased the uptake of iron for the cyanobacterium, and also suggest that the growth of S. elongatus was promoted by siderophores produced by Shewanella sp. OR151. Although cyanobacteria produce siderophores, such as schizokinen and synechobactin A (Årstøl and Hohmann-Marriott, 2019), they still require high levels of iron for photosynthesis (Sunda and Huntsman, 2015). By chelating insoluble ferric ions, siderophores help increase iron uptake by cyanobacteria (Neilands, 1995). Siderophores, such as putrebactin, bisucaberin, and avaroferrin, have been identified in the genus Shewanella (Liu et al., 2022). In the present study, the CAS blue agar assay showed that siderophore production by Rhodococcus sp. AF2108 and Shewanella sp. OR151 was positive (Table S2), whereas that by the other strains was only weakly positive. Based on these results, siderophores produced by the two strains may have promoted cyanobacterial growth during the co-culture. Furthermore, the siderophore-positive strains, Rhodococcus sp. AF2108 and Shewanella sp. OR151, increased the cell size of S. elongatus. Although siderophores appeared to affect cyanobacterial cell size, further evidence is needed.
In the present study, four novel CGPB strains, Rhodococcus sp. AF2108, Ancylobacter sp. GA1226, Xanthobacter sp. AF2111, and Shewanella sp. OR151, were isolated and exerted growth-promoting effects on S. elongatus; these CGPBs increased the content of chlorophyll a in S. elongatus by 8.5-, 2.8-, 1.3-, and 1.7-fold, respectively. According to flow cytometry, the four CGPBs also increased the cell number, forward scattering intensity, and chlorophyll fluorescence intensity per cell of S. elongatus to varying extents. Rhodococcus sp. AF2108 exerted faster and stronger growth-promoting effects than bacteria in previous studies, which indicates that Rhodococcus sp. AF2108 is the most effective CGPB strain identified to date. These four newly isolated CGPBs will serve as important components of a new Synechococcus growth improvement strategy. The present results indicate that IAA and siderophores stimulate cyanobacterial growth. These CGPBs may be applied to various bioprocesses that rely on autotrophic Synechococcus metabolic pathways.
Tan, P. Y., Kato, Y., and Konishi, M. (2024) A Novel Strain of the Cyanobacterial Growth-promoting Bacterium, Rhodococcus sp. AF2108, Enhances the Growth of Synechococcus elongatus. Microbes Environ 39: ME24050.
https://doi.org/10.1264/jsme2.ME24050
We thank Mr. Masashi Ishida for his advice and support with experiments. This work was partly supported by the METI Monozukuri R&D Support Grant Program for SMEs (grant number JPJ005698) and JSPS KAKENHI Grant Numbers JP23H02115 and JP23K26808.
