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Cellulose-mediated floc formation by the activated sludge bacterium Shinella zoogloeoides ATCC 19623

BMC Microbiology volume 22, Article number: 104 (2022) Cite this article

Abstract

Background

Bacterial floc formation plays a central role in the activated sludge (AS) process. The formation of AS flocs has long been known to require exopolysaccharide biosynthesis. We had demonstrated that both expolysaccharides and PEP-CTERM (a short C-terminal domain includes a near-invariant motif Pro-Glu-Pro (PEP)) proteins were required for floc-forming in Zoogloea resiniphila MMB, a dominant AS bacterium. However, the PEP-CTERM proteins are not encoded in the genome of AS bacterium Shinella zoogloeoides ATCC 19623 (formerly known as Zoogloea ramigera I-16-M) and other sequenced AS bacteria strains. The mechanism underlying floc formation of Shinella and related AS bacteria remained largely unclear.

Results

In this study, we have sequenced and annotated the complete genome of S. zoogloeoides ATCC 19623 (aka I-16-M), previously isolated in USA and treated as the neotype for the AS floc-forming bacterium Zoogloea ramigera I-16-M, and another AS strain XJ20 isolated in China. Mariner transposon mutagenesis had been conducted to isolate floc-forming-deficient mutants in the strain ATCC 19623 as previously performed by using Tn5 transposon three decades ago. The transposon insertional sites of multiple mutants were mapped to the gene cluster for bacterial cellulose synthesis (bcs) and secretion, and the role played by these genes in floc-formation had been further confirmed by genetic complementation. Interestingly, the restriction map of this bcs locus-flanking region was highly similar to that of the previously identified DNA fragment required for floc-formation in 1980s. Cellulase treatment abolished the floc-forming phenotype of S. zoogloeoides ATCC 19623 but not that of Z. resiniphila MMB strain. The FTIR spectral analyses revealed that the samples extracted from S. zoogloeoides ATCC 19623 were cellulose polymer.

Conclusion

Our results indicated that we have largely reproduced and completed the unfinished pioneering work on AS floc-formation mechanism, demonstrating that the floc-formation and flocculating capability of Shinella were mediated by extracellular cellulose polymers.

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Background

The AS process has been widely applied for municipal sewage and industrial wastewater treatment worldwide and has played a central role in water purification for over a century. Zoogloea ramigera and related floc-forming bacteria are required for formation of AS flocs key to gravitational effluent-and-sludge separation and AS recycling, two key elements for AS process. Butterfield et al. isolated a floc-forming organism from activated sludge and tentatively identified it as Z. ramigera, based on the phenotypes that the bacterial culture was capable of forming flocs similar to normal activated sludge and also exhibited oxidation characteristics similar to those of normal activated sludge [1]. Bacteria of the genus Zoogloea have historically been differentiated from other obligately aerobic, Gram-negative, non-spore-forming, rod-shaped bacteria that grow in aquatic habitats on the basis of a characteristic gelatinous matrix, often called a “zoogloeal matrix”, that surrounds clumps of cells when they are found in their aquatic habitat or in unshaken liquid culture in the laboratory [2]. However, the specimen had not been preserved and the taxonomy and microbiology of the Z. ramigera species were incomplete without a type strain. Therefore, three neotype strains for floc-forming Z. ramigera had been isolated from activated sludge and experimentally characterized during the 1960s [3,4,5]. However, these three strains obviously did not belong to the same species and one strain, ATCC 19544 (aka N.C. Dondero 106 strain), directly isolated by microdissection of a natural branched zoogloeal floc [5], has been recognized taxonomically as the neotype strain for the species of Z. ramigera Itzigsohn [6, 7]. The other two floc-forming strains, ATCC 19623 (aka I-16-M) and ATCC 25935 (aka P.R. Dugan 115 strain), were reclassified as novel species S. zoogloeoides of Rhizobiales and Duganella zoogloeoides of Burkholderiales, respectively, based on the phenotypic and 16S rDNA-based phylogenetic analyses [8,9,10]. In addition, another name Crabtreella saccharophila gen. nov., sp. nov. was proposed for strain ATCC 19623 (aka I-16-M) based on phylogenetic and chemotaxonomic analyses in March, 2006 [11]. Because S. zoogloeoides was proposed before Crabtreella saccharophila, the latter name is not a valid name according to the rules of the International Code of Nomenclature of Bacteria. All three could flocculate and each strain produces a different extracellular polysaccharide [12]. S. zoogloeoides ATCC 19623 strain produces a cellulose fibrillar polysaccharide [12], Z. ramigera ATCC 19544 (aka 106) produces an exopolymer reportedly containing amino sugars [13], and Duganella zoogloeoides ATCC 25935 (aka 115) produces a weakly acidic extracellular polysaccharide composed of D-glucose, D-galactose and pyruvic acid in an approximate molar ratio 11:3:1.5 [14, 15]. Anthony J. Sinskey et al. pioneered the study of genetics and molecular mechanism underlying the bacterial floc formation using S. zoogloeoides ATCC 19623 (still classified as Z. ramigera I-16-M at that time) as a model strain [16]. By using Tn5 transposon mutagenesis, they isolated the Tn5 insertion mutants deficient in EPS production by screening for the absence of fluorescence on plates containing the dye Calcofluor. They further determined whether a specific chromosomal DNA segment was required for exopolysaccharide biosynthesis and floc formation based on the planktonic cell growth (turbid cell culture) of the mutant and recovery of floc-forming phenotype and gravitational settling property of the mutants by genetic complementation using a genetic library, constructed in a broad-host-range cosmid vector, and introduced into the mutants via conjugation [16]. However, due to lack of easy sequencing methods in those days, the relevant genes had not been identified and the chromosomal DNA inserts of complementing recombinant plasmids had not been sequenced. In recent years, we have employed the similar methodology and techniques to identify the genes and gene clusters required for Z. resiniphila and other predominant floc-forming bacteria [17,18,19]. Our research has been greatly facilitated by high-throughput genome sequencing and more powerful genetic manipulation tools. Our previous researches had revealed that the formation of AS flocs requires both exopolysaccharide biosynthesis and expression of the PEP-CTERM domain proteins in Z. resiniphila, a dominant AS bacterium [17, 19]. The PEP-CTERM protein-sorting domain is usually a 25-residue carboxyl terminal domain that includes a near-invariant Proline-Glutamate-Proline (PEP) motif, a thirteen residue strongly hydrophobic sequence likely to span the membrane, and a five-residue strongly basic motif that often contains four arginine residues [20]. This kind of proteins also contain an amino terminal secretion signal peptide and the asparagine (N) residue is overrepresented and could be used for N-linked glycosylation [20]. A member of a wide-spread family of high copy number-per-genome PEP-CTERM genes were transcriptionally regulated by the RpoN (sigma54) sigma factor and PrsK-PrsR two-component system. Without PrsK or PrsR, Z. resiniphila cells were planktonic rather than aggregated into the floc structures that allow gravitational sludge settling and recycling. Moreover, the biosynthesis and secretion of exopolysaccharides remained unaffected in the prsK and prsR mutants. But the synthesized exopolysaccharides were released into the growth broth in soluble form. Overexpression of PEP-CTERM A protein (PepA) could circumvent the requirement of PrsK and PrsR for floc-formation [19]. We had also demonstrated that one of the four RpoN paralogues, RpoN1, regulated the floc formation but not exopolysaccharide biosynthesis in a floc-forming Aquincola tertiaricarbonis RN12 strain [18] and Azoarcus halotolerans [21]. The synthesized exopolysaccharide chains were not tightly bound to the bulk of cell aggregates but were released into the cultivation broth. A highly conserved large gene cluster, including two asparagine synthetase genes and other genes involved in exopolysaccharide biosynthesis and secretion, was encoded in the genome of Z. resiniphila MMB and A. tertiaricarbonis RN12 [17, 18]. Interestingly, both PEP-CTERM proteins and exopolysaccharide biosynthesis pathway were also encoded in the genome of many other AS bacteria including comammox strains of Nitrospira and polyphosphate-accumulating organisms Candidatus Accumulibacter phosphatis [17, 19].

Since a series of researches such as the biosynthesis of polyhydrobutyrate (PHB), one of the bioplastic materials, have also been intensively conducted in S. zoogloeoides ATCC 19623 [22], but the genome data of S. zoogloeoides ATCC 19623 has not yet been published at the time of writing. In the present study, we sequenced and annotated the complete genome of the two S. zoogloeoides strains and conducted comparative genomics and molecular genetics analyses on the floc formation using mariner transposon mutagenesis to reproduce and complete the unfinished pioneering work on AS floc-formation mechanism. Our results could shed more light on the molecular mechanism underlying formation of activated sludge flocs and bioflocculation.

Results

Complete genome sequencing, annotation and comparative analysis

The complete genome sequences of both S. zoogloeoides ATCC 19623 and XJ20 strains were sequenced by using both Illumina HiSeq and Pacific Biosciences platforms. The complete genome of S. zoogloeoides ATCC 19623 was 4,848,065 base pairs (bp) in size with the G + C content of 64.43%, containing 9 rRNAs operons, and 53 tRNAs. On the other hand, the genome of S. zoogloeoides XJ20 was 5,141,032 bp in size with the G + C content of 63.89%, and also contained 9 rRNAs operons, and 53 tRNAs. Interestingly, the S. zoogloeoides XJ20 strain encoded a higher number of protein-coding genes (5090) than that of S. zoogloeoides ATCC 19623 (4687), which was consistent with its larger genome size. Moreover, there were 6 plasmids in XJ20 and 4 plasmids in S. zoogloeoides ATCC 19623, respectively (Table 1). The single circular genome maps of the two S. zoogloeoides genomes were shown in Fig. 1. Using a Venn diagram of two strains, the majority of homologous gene groups and unique gene groups were identified. The unique gene groups accounted for 880 and 1194 gene groups in S. zoogloeoides ATCC 19623 and S. zoogloeoides XJ20, respectively (Supplemental Fig. 1). The average genomic nucleotide identity (ANI) between S. zoogloeoides XJ20 and S. zoogloeoides ATCC 19623 was 93.88%, which was lower than the species delineation thresholds (95–96%), suggesting that S. zoogloeoides XJ20 might not belong to the same species sensu stricto [23]. We conducted comparative genomics analysis of the two strains. These two chromosome genomes were aligned with Progressive Mauve using default parameters [24]. The genome synteny and collinearity between strain S. zoogloeoides ATCC 19623 and S. zoogloeoides XJ20 was high (Supplemental Fig. 2), though there were still some rearrangements and translocation. However, the large gene cluster for exopolysaccharide biosynthesis and secretion, PEP-CTERM proteins and the PrsK-PrsR two-component system regulating the expression of PEP-CTERM proteins, which are required for the floc-forming of Z. resiniphila MMB strain and A. tertaricarbonis RN12 strain [17,18,19], were not encoded in the genome of S. zoogloeoides ATCC 19623, S. zoogloeoides XJ20 and other sequenced Shinella strains. It was strongly suggested that the floc-forming mechanism in Shinella species was different from that of Zoogloea species, one of the predominant floc-forming bacteria in activated sludge.

Table 1 General characteristics of the Shinella zoogloeoides two strains genome
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Fig. 1
figure 1

Circular genome maps of Shinella zoogloeoides ATCC 19623 (a) and XJ20 (b) from inside to outside indicate the following: Circle 1: G + C skew; green, GC skew>0; orange, GC skew<0; Circle 2: G + C content (median represents the above average content, the outer circle is greater than the average content (red), and the inner circle is less than the average content (blue)); Circle 3: rRNA and tRNA genes distribution represented in scaffold sequence; Circle 4 and 5: open reading frame (ORF) distribution, minus and plus strand, different colors represent the functional classification of COG with different CDS; Circle 6: indicator of genome size

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Identification of a bacterial cellulose synthesis (bcs) gene cluster required for floc formation

Previously the Tn5 transposon mutagenesis had been conducted on the floc-forming activated sludge bacterium S. zoogloeoides ATCC 19623 (still classified as Z. ramigera I-16-M at that time) strain to identify the genes required for biosynthesis of extracellular polysaccharides and floc formation [16]. A series of mutants deficient in exopolysaccharide biosynthesis were isolated and plasmids from a genomic library, containing at least 14 kilobases of common insert DNA, could restore the floc forming phenotype of the mutants after entering the cells via conjugation. However, the DNA sequence and relevant genes had not been determined due to the technical limitation at that time. In the present study, the transposon mutagenesis was conducted on the S. zoogloeoides ATCC 19623 strain by using mariner transposon as previously described [25]. Sixty-one floc-forming-deficient mutants had been isolated and the transposon insertional site had been mapped in nineteen mutants (Supplemental Table S1). As presented in Fig. 2 a5 and b5, these mutants do not form the flocs in the agitated liquid culture or the thick pellicle on the surface of the static liquid culture. Microscopic observation shows that the bacterial cultures of those mutants are usually composed of single cells and tiny cell aggregates (Fig. 2 a2, 2b2), but not large flocs as previously shown [17,18,19]. For example, one of the mutants has interrupted bcsA and the other two bcsB. Furthermore, the plasmid-borne bcsA gene (pBBR1MCS-2-bcsA) and bcsB gene (pBBR1MCS-2-bcsB) restored the floc formation phenotype to these two mutants in genetic complementation analyses (Fig. 2a, b). These mutants were mapped to a bacterial cellulose synthesis (bcs) gene cluster (locus tags K8M09_02915- K8M09_02940). This gene cluster was composed of bcsN, bcsA, bcsB and bcsK (Fig. 2c).

Fig. 2
figure 2

The genetic complementation analysis of the bcsA and bcsB genes in transposon mutant SZM19, SZM6 and SZM25. a The genetic complementation analysis of the bcsA in trans in the mutants SZM19. Plasmid-borne wild-type bcsA gene restored the phenotype of the SZM19 mutant to floc formation, similar to that of the wild-type strain carrying the empty vector. (1) Wild-type strain with the pBBR1MCS-2 empty vector, visualized under a microscope after staining with crystal violet solution. (2) SZM19 mutant with pBBR1MCS-2. (3) SZM19 mutant with pBBR1MCS-2-bcsA construct. (4) Photograph of agitated bacterial cultures (culture tubes contain bacteria from panels (1) to (3), from left to right). (5) Photograph of settled bacterial cultures. b The genetic complementation analysis of the bcsB in trans in mutants SZM6. (1) Wild-type strain with the pBBR1MCS-2 empty vector, visualized under a microscope. (2) SZM6 mutant with pBBR1MCS-2. (3) SZM6 mutant with pBBR1MCS-2-bcsB construct. (4) Photograph of agitated culture (culture tubes contain bacteria from panels (1) to (3), from left to right). (5) Photograph of settled bacterial cultures. c The cellulose biosynthesis gene cluster of Shinella zoogloeodies ATCC 19623 strain and the genes disrupted by insertions of the mariner transposon, mapped in the transposon mutants deficient for bacterial floc formation

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The restriction map of bcs gene cluster-flanking region is highly similar to that of previously identified exopolysaccharide biosynthesis gene region

We constructed a digital restriction map of the identified bcs gene cluster-flanking region, which was highly similar to that of the 32 kilobases exopolysaccharide biosynthesis gene region previously identified from the complementing cosmids of gene library by Easson, Sinskey and Peoples [16]. The counterpart of most of the EcoR I restriction fragments could be found in the digital restriction map and the measured size of these fragments were close to their real length, suggesting that the quality of the original restriction map was good (Fig. 3). The inconsistency between the two maps might be due to the technical limitation of restriction mapping. Some small restriction fragments could not be readily identified by the agarose gel electrophoresis and ethidium bromide staining methods and were missed in the original restriction map. Nevertheless, the resolution of agarose gel electrophoresis was not good enough to differentiate the bands containing two or more restriction fragments of similar sizes. These results indicated that we have largely reproduced and further confirmed the previous findings of Easson, Sinskey and Peoples [16].

Fig. 3
figure 3

Comparison of the two EcoR I restriction maps. a EcoR I restriction maps of pPS25, pPS27, pPS30, pPS48, and the composite region identified from the complementing cosmids of gene library by Easson, Sinskey and Peoples [16]; b The digital restriction map of the identified bcs gene cluster-flanking region. Sizes of each fragment are shown in kilobases. Illustrator for Biological Sequences (IBS1.0.3) and Adobe Illustrator CS6 programs were used to draw the maps

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Cellulase supplement prevented the floc formation of Shinella strains but not Z. resiniphila

Spontaneous flocculation of Escherichia coli cells could be induced by overexpression of the bcsB gene and those flocs were sensitive to proteinase K and NaOH, but not to cellulase, indicating that the main component of the flocs was proteinaceous [26]. In this study, we added the commercial cellulase, proteinase K or NaOH solution to the bacterial cultures of Z. resiniphila MMB and S. zoogloeoides ATCC 19623. Our results showed that the flocs of Shinella could be disrupted by addition of cellulase, while the floc formation of the Z. resiniphila MMB strain was not affected (Fig. 4). Both proteinase K and NaOH could not de-flocculate the flocs of Shinella, only Z. resiniphila MMB exhibited slight sensitivity to NaOH solution. These results further verified that the extracellular polymers required for floc formation of Shinella were the cellulose materials vulnerable to cellulase degradation.

Fig. 4
figure 4

Treatment of Shinella zoogloeoides ATCC 19623 and Zoogloea resiniphila MMB cell flocs with cellulase, proteinase K and NaOH solution

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Characterization of bacterial exopolymers

The involvement of cellulose in the formation of flocs has been confirmed by molecular genetics. According to the extraction methods of cellulose from other strains, FTIR analysis was used to prove that the substance extracted from S. zoogloeoides ATCC 19623 was cellulose. The FTIR spectra of all the cellulose could provide the information about their functional groups and the state of bonds in their structure (Fig. 5). The FTIR spectrum of bacterial cellulose sample was highly comparable with that of the commercial cellulose product used as the control. The hydroxyl groups were observed at ~3400–3440 cm−1, which indicated (−OH) stretching vibrations. The broader peak bands indicate intermolecular hydrogen bonding in the cellulose molecule, and also these hydroxyl groups are responsible for facilitating substitution in the molecule, consistent with the previously reported results [27]. The peak at ~2800–2900 cm−1 represents symmetric (−CH) stretching vibrations indicating the presence of methyl/methylene functional group. The peak at ~1620–1640 cm−1 reveals a water molecule associated with cellulose (absorbed H2O) bending vibrations. The carbonyl groups (C=O) are observed at the ~1420–1440 cm−1. The peak at~1040–1068 cm−1 indicates the presence of (C–O) stretching vibration in the polymer. These significant peaks further confirmed that the extracted sample was a cellulose polymer.

Fig. 5
figure 5

FTIR spectra of bacteria cellulose from Shinella zoogloeoides ATCC 19623 and commercial cellulose. NICOLET 5700 FTIR Spectrometer was used with diamond ATR spectrophotometer for 24 scans in the range of 4000–400 cm−1 with a resolution of 4 cm−1. Bacteria cellulose extracted from Shinella zoogloeoides ATCC 19623, commercial cellulose purchased from Sangon Biotech, Shanghai, China

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Discussion

Based on the core community of activated sludge [28] and our own isolation of AS bacteria, many predominate AS bacteria exhibited floc-forming phenotypes, such as small flocs and less strengths, similar to those of S. zoogloeoides. Those key AS bacteria such as Acidovorax, Comamonas, Diaphorobacter and Hydrogenophaga actually lack the PEP-CTERM genes and EPS biosynthesis genes previously described for Z. resiniphila, instead, they encode cellulose biosynthesis pathway similar to that of Shinella described in this study. The mechanism underlying the floc formation of Shinella is different from that of Zoogloea and other predominant activated sludge bacteria in which both the PEP-CTERM proteins and extracellular polysaccharides are required for floc formation [17,18,19].

The Shinella genus is phylogenetically close to the Agrobacterium and Rhizobium genera of Hyphomicrobiales, and S. zoogloeodies ATCC 19623 exhibits a floc-forming phenotype, which requires bacterial cellulose synthesis as shown previously [16] and in this study. The comparative genomic analysis showed that S. zoogloeodies ATCC 19623 was highly collinear to S. zoogloeodies XJ20 isolated from Xinjiang (Supplemental Fig. 2). Sixty-one floc-forming-deficient mutants had been isolated and the transposon insertional site has been mapped in nineteen mutants in the S. zoogloeodies ATCC 19623 strain. Genetic complementation was conducted to confirm whether the identified genes are required for floc-formation and the insertion site of three mutants was mapped to a bacterial cellulose synthesis (bcs) gene cluster. This gene cluster was composed of bcsN, bcsA, bcsB and bcsK (Fig. 2), which could be found in other bacteria, like Agrobacterium fabrum C58 and Azospirillum lipoferum 4B [29]. Omadjela et al. reported that BcsA and BcsB are necessary and sufficient for the formation of the polysaccharide chain in vitro in most studied bacteria [30]. The protein encoded by bcsA contains the catalytically active subunit with a PilZ domain, which is responsive to c-di-GMP. BcsB is located in the periplasm and is anchored in the membrane by a single transmembrane helix [31]. The functions of BcsK are probably the same as those of AlgK and the N-terminal part of BcsC, i.e., interacting with the peptidoglycan and organizing the entire cellulose secretion complex [29]. The functions of BcsN remain unclear. Our results clearly demonstrated that bacterial cellulose biosynthesis was required for floc formation of S. zoogloeoides ATCC 19623 strain. The FTIR analysis showed the sample extracted from S. zoogloeoides ATCC 19623 was cellulose polymer (Fig. 5). Biosynthesis and regulatory pathways of cellulose are also highly conserved among these bacteria of Hyphomicrobiales [32]. Shinella has been frequently found in the activated sludge samples, but it may not be one of the predominant genera because the Shinella genus is not among the 28 taxa of global core community of activated sludge [28]. On the other hand, the floc-forming bacteria including Zoogloea, Thauera and Dechloromonas are among the core community and encode the exopolysaccharide biosynthesis gene clusters and PEP-CTERM domain containing proteins. The fluorescent probe complementary to the 16S ribosomal RNAs of the S. zoogloeoides ATCC 19623 strain bound to only a few cells of activated sludge samples [33]. But this kind of small and unstable flocs, which as minor players of high species diversity in AS, still facilitates integration of these bacteria into AS flocs.

As compared to the highly diverse exopolysaccharides, cellulose is more degradable and vulnerable because many microbes synthesize and secrete cellulase. Our results have also shown that the flocs of Shinella could be disrupted by addition of cellulase (Fig. 4). Bacteria cellulose biosynthesis is regulated by the expression of bcs genes, which appear to be expressed in response to the second messenger cyclic-di-GMP [34]. The c-di-GMP is synthesized by diguanylate cyclases (DGCs) that carry GGDEF domains and degraded by phosphodiesterases (PDEs) that carry either EAL or HD-GYP domains, all these domains are essential for enzymatic activity [35]. Indeed, even though the GGDEF and EAL domain-containing proteins have opposing activities, these two domains are often found coupled in the same proteins, which are referred to as hybrid proteins because they carry both domains [36]. We found eight hybrid proteins containing GGDEF and EAL domain, six stand-alone c-di-GMP PDEs and four stand-alone DGCs are present in the genome. These multiple genes together control the intracellular c-di-GMP levels. Easson et al. described spontaneous mutations and genetic instability of the strain Shinella in the region encoding the cellulose biosynthetic cluster [16]. Genome instability can be mediated by genetic elements, homologous and illegitimate recombination. A number of mobile elements (including insertion sequences, miniature inverted-repeat transposable elements, repetitive extragenic palindromic sequences and bacterial interspersed mosaic elements, transposons, transposable bacteriophages and genomic islands), introns and integrons play a role in genomic instability [37]. The number of gene islands, prophage and insertion sequences in the genome of S. zoogloeoides ATCC 19623 was 20, 4, 11, respectively. It remains unknown whether other genes are required for the floc formation of S. zoogloeoides ATCC 19623, in addition to the bcsA and bcsB genes involved in biosynthesis of extracellular cellulose polymers. Shinella strains may serve as a good bioflocculant of wide-range of applications.

Conclusions

In this study we further explored the mechanism underlying the floc formation of S. zoogloeoides ATCC 19623 (previously Zoogloea ramigera I-16-M) and related activated sludge bacteria, and sequenced and annotated the complete genome of S. zoogloeoides ATCC 19623 and XJ20 strains. The comparative genomic analysis showed that the chromosomes of S. zoogloeoides ATCC 19623 and XJ20 strain were highly collinear, although they were isolated from activated sludge of United States and China, respectively. Our results clearly demonstrate that bacterial cellulose biosynthesis gene bcsA and bcsB are required for floc formation of S. zoogloeoides ATCC 19623 and the flocs could be disrupted by addition of cellulase. This mechanism may account for the smaller and less stable flocs formed by highly diverse bacteria in activated sludge. These results indicate that we have largely reproduced and completed the unfinished pioneering research on the genes required for floc-forming in S. zoogloeoides ATCC 19623 by Easson, Sinskey and Peoples three decades ago.

Methods

Bacterial strains, plasmids, and culture conditions

The bacterial strains and plasmids used in this study are listed in Supplemental Table S2. The S. zoogloeoides ATCC 19623 strain was obtained from China Center for Type Culture Collection (CCTCC) and the XJ20 strain was isolated from the activated sludge of Municipal Sewage Treatment Plant, Xinjiang Uygur Autonomous Region, China, collected by Yaqi Liu in September, 2019. Bacterial strains were cultured in the Luria-Bertani broth (5 g/L yeast extract, 10 g/L tryptone, 10 g/L sodium chloride (NaCl, pH 7.0)/plates (supplemented with 15–30 μg/ml of gentamycin or 50 μg/ml of kanamycin and 50 μg/ml of diaminopimelic acid, when necessary), and Zoogloea medium (ZM) [19] at 28 °C.

Genome sequencing, annotation and comparative genomic analysis

Two complete genomes of S. zoogloeoides were sequenced using a combination of PacBio RS and Illumina sequencing platforms. The Illumina data was used to evaluate the complexity of the genome and correct the PacBio long reads. Firstly, we used the SOAP de novo version 2.21 package to do genome assembly with multiple-Kmer parameters (21–41) and got the optimal results of the assembly. Secondly unicycler was used to assemble the PacBio corrected long reads. Finally, GapCloser software was subsequently applied to fill up the remaining local inner gaps and correct the single base polymorphism for the final assembly results. The complete circle of the genome was drawn with Circos v0.64. Genome annotation was performed using RAST (Rapid Annotation using Subsystem Technology). The rRNA genes were predicted by RNAmmer [38], and tRNAscan-SE [39] was used to identify the tRNA genes. The interactive web-based software system GC-Profile [40] was used to investigate the genomic GC content. The genomes alignment was performed by Progressive Mauve [24]. Nucleotide and protein sequences were retrieved from NCBI database by using BLAST searches at the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/BLAST). Homologous gene analysis was performed using OrthoMCL program. The relationship between sequenced strains was determined by Average Nucleotid Identity (ANI) calculation with web tool ANI calculator [41] (https://www.ezbiocloud.net/tools/ani).

Transposon mutagenesis and genetic complementation

The mariner transposon mutant libraries were generated and screened as previously described [17, 42, 43], with Escherichia coli WM3064 strains carrying the transposon delivery suicide plasmids pFAC (courtesy by Dr. John Mekalanos, Harvard Medical School) as the donor and the S. zoogloeoides ATCC 19623 strain as the recipient for biparental conjugation [18]. The transposon insertion site in each mutant was mapped as previously described [17]. For genetic complementation analyses, the target genes were PCR amplified and cloned into the pBBR1MCS-2 [44]. The primers for PCR reactions are listed in the Supplemental Table S2. The resultant constructs and empty vector were transferred into the S. zoogloeoides wild type strain and mutant strains via conjugation using WM3064 as a donor strain.

Assays of flocs sensitivity to enzyme treatment

Floc formation was studied in the ZM medium [19]. Z. resiniphila MMB has been used as negative control. Each tube contained 3 ml of medium, incubation was carried out on a rotary shaker at 28 °C for 36 h After the flocs were settled and washed three times with water in the same test tube by decantation. The collected flocs were treated with 2 mg/ml cellulase (Trichoderma reesei ATCC 26921, Sangon Biotech, Shanghai, China) in 2.5 mM 2-morpholinoethanesulfonic acid buffer (pH 5.5), 2 mg/ml proteinase K (Tritirachium album limber, Sangon Biotech, Shanghai, China) in 50 mM Tris buffer (pH 8.0), or 0.5 M NaOH solution at 37 °C for 2 h [28].

Extracellular polymer extraction, purification and spectrometry analyses

Aliquots of 15 ml bacteria culture were inoculated into a flask containing 1.5 L of ZM medium. Later the conical flasks were left for static incubations at 28 °C. Cells were harvested by centrifuge after a week of incubation and cell pellets were washed with deionized water to remove the residual medium. The cell pellets were then boiled in 1% NaOH at 80 °C for 90 min to dissolve bacterial cell mass and rinsed with deionized water until the filtrate reached neutral pH value. The purified sample was then dried by freeze-drying. Dried polymers were analyzed for both chemical and structural features with commercial cellulose as a reference [43]. FTIR spectral analyses were performed to ascertain the similarity between the functional groups of the sample and the reference molecule.

Availability of data and materials

The complete genome sequencing datasets of Shinella zoogloeoides ATCC 19623 and Shinella zoogloeoides XJ20 used in this study were deposited in GenBank under accession CP086610 for ATCC 19623 chromosome, CP086611 - CP086614 for ATCC 19623 plasmids, CP093528 for XJ20 chromosome, CP093529 - CP093534 for XJ20 plasmids.

References

  1. Butterfield CT, Ruchhoft CC, McNamee PD. Studies of sewage purification VI Biochemical oxidation by sludges developed by pure cultures of bacteria isolated from activated sludge. Public Health Rep (1896–1970). 1937;52(14):387–412.

    Article  CAS  Google Scholar 

  2. Dugan PR. The Genus Zoogloea. In: Starr MP, Stolp H, Trüper HG, Balows A, Schlegel HG, editors. The Prokaryotes. Berlin: Springer; 1981. p. 764–70.

    Chapter  Google Scholar 

  3. Dugan PR, Lundgren DG. Isolation of the floc-forming organism Zoogloea-ramigera and its culture in complex and synthetic media. Appl Microbiol. 1960;8(6):357–61.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Crabtree K, Mccoy E. Zoogloea ramigera Itzigsohn, identification and description. Int J Syst Bacteriol. 1967;17(1):1–10.

    Article  Google Scholar 

  5. Unz RF, Dondero NC. Predominant bacteria in natural zoogloeal colonies .II. Physiology and nutrition. Can J Microbiol. 1967;13(12):1683–94.

    Article  CAS  PubMed  Google Scholar 

  6. Skerman VBD, McGowan V, Sneath PHA, editors. Approved lists of bacterial names. Washington (DC): ASM Press; 1989.

    Google Scholar 

  7. Unz RF. Neotype strain of Zoogloea ramigera Itzigsohn. Int J Syst Bacteriol. 1971;21:91–9.

    Article  Google Scholar 

  8. Shin YK, Hiraishi A, Sugiyama J. Molecular systematics of the genus Zoogloea and emendation of the genus. Int J Syst Bacteriol. 1993;43(4):826–31.

    Article  CAS  PubMed  Google Scholar 

  9. Hiraishi A, Shin YK, Sugiyama J. Proposal to reclassify Zoogloea ramigera IAM 12670 (P. R. Dugan 115) as Duganella zoogloeoides gen. nov., sp. nov. Int J Syst Bacteriol. 1997;47(4):1249–52.

    Article  CAS  PubMed  Google Scholar 

  10. An DS, Im WT, Yang HC, Lee ST. Shinella granuli gen. nov., sp. nov., and proposal of the reclassification of Zoogloea ramigera ATCC 19623 as Shinella zoogloeoides sp. nov. Int J Syst Evol Microbiol. 2006;56(Pt 2):443–8.

    Article  CAS  PubMed  Google Scholar 

  11. Xie CH, Yokota A. Zoogloea oryzae sp nov., a nitrogen-fixing bacterium isolated from rice paddy soil, and reclassification of the strain ATCC 19623 as Crabtreella saccharophila gen. nov., sp nov. Int J Syst Evol Microbiol. 2006;56(Pt 3):619–24.

    Article  CAS  PubMed  Google Scholar 

  12. Dugan PR. Bioflocculation, and the accumulation of chemicals by floc-forming organisms. Final Report: United States, N.P., Environmental Protection Technology Series; 1975. EPA-600/2–75-032.

    Google Scholar 

  13. Farrah SR, Unz RF. Isolation of exocellular polymer from Zoogloea strains mp6 and 106 and from activated-sludge. Appl Environ Microbiol. 1976;32(1):33–7.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Ikeda F, Shuto H, Saito T, Fukui T, Tomita K. An extracellular polysaccharide produced by Zoogloea-ramigera 115. Eur J Biochem. 1982;123(2):437–45.

    Article  CAS  PubMed  Google Scholar 

  15. Franzen LE, Norberg AB. Structural investigation of the acidic polysaccharide secreted by Zoogloea ramigera 115. Carbohydr Res. 1984;128:111–7.

    Article  CAS  Google Scholar 

  16. Easson DD, Sinskey AJ, Peoples OP. Isolation of Zoogloea ramigera I-16-M exopolysaccharide biosynthetic genes and evidence for instability within this region. J Bacteriol. 1987;169:4518–24.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  17. An W, Guo F, Song Y, Gao N, Bai S, Dai J, et al. Comparative genomics analyses on EPS biosynthesis genes required for floc formation of Zoogloea resiniphila and other activated sludge bacteria. Water Res. 2016;102:494–504.

    Article  CAS  PubMed  Google Scholar 

  18. Yu D, Xia M, Zhang L, Song Y, Duan Y, Yuan T, et al. RpoN (sigma54) is required for floc formation but not for extracellular polysaccharide biosynthesis in a floc-forming Aquincola tertiaricarbonis strain. Appl Environ Microbiol. 2017;83(14):e00709–17.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Gao N, Xia M, Dai J, Yu D, An W, Li S, et al. Both widespread PEP-CTERM proteins and exopolysaccharides are required for floc formation of Zoogloea resiniphila and other activated sludge bacteria. Environ Microbiol. 2018;20(5):1677–92.

    Article  CAS  PubMed  Google Scholar 

  20. Haft DH, Paulsen IT, Ward N, Selengut JD. Exopolysaccharide-associated protein sorting in environmental organisms: the PEP-CTERM/EpsH system. Application of a novel phylogenetic profiling heuristic. BMC Biol. 2006;4:29.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  21. Li S, Zhao L, Han J, Liu S, Dai J, Fu G, et al. Azoarcus halotolerans sp. nov., a novel member of Rhodocyclaceae isolated from activated sludge collected in Hong Kong. Int J Syst Evol Microbiol. 2020;70(11):5799–805.

    Article  CAS  PubMed  Google Scholar 

  22. Fukui T, Yoshimoto A, Matsumoto M, Hosokawa S, Saito T. Enzymatic synthesis of poly-beta-hydroxybutyrate in Zoogloea ramigera. Arch Microbiol. 1976;110(23):149–56.

    Article  CAS  PubMed  Google Scholar 

  23. Kim M, Oh HS, Park SC, Chun J. Towards a taxonomic coherence between average nucleotide identity and 16S rRNA gene sequence similarity for species demarcation of prokaryotes. Int J Syst Evol Microbiol. 2014;64(Pt 2):346–51.

    Article  CAS  PubMed  Google Scholar 

  24. Darling AE, Mau B, Perna NT. progressiveMauve: Multiple Genome Alignment with Gene Gain, Loss and Rearrangement. PLoS One. 2010;5(6):e11147.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Bouhenni R, Gehrke A, Saffarini D. Identification of genes involved in cytochrome c biogenesis in Shewanella oneidensis, using a modified mariner transposon. Appl Environ Microbiol. 2005;71(8):4935–7.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Ojima Y, Minh Hong N, Yajima R, Taya M. Flocculation of Escherichia coli Cells in Association with Enhanced Production of Outer Membrane Vesicles. Appl Environ Microbiol. 2015;81(17):5900–6.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  27. Guzel M, Akpinar O. Production and characterization of bacterial cellulose from citrus peels. Waste Biomass Valori. 2018;10:2165–75.

    Article  CAS  Google Scholar 

  28. Wu LW, Ning DL, Zhang B, Li Y, Zhang P, Shan XY, et al. Global diversity and biogeography of bacterial communities in wastewater treatment plants. Nat Microbiol. 2019;4(7):1183–95.

    Article  CAS  PubMed  Google Scholar 

  29. Romling U, Galperin MY. Bacterial cellulose biosynthesis: diversity of operons, subunits, products, and functions. Trends Microbiol. 2015;23(9):545–57.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Omadjela O, Narahari A, Strumillo J, Melida H, Mazur O, Bulone V, et al. BcsA and BcsB form the catalytically active core of bacterial cellulose synthase sufficient for in vitro cellulose synthesis. Proc Natl Acad Sci U S A. 2013;110(44):17856–61.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Morgan JLW, Strumillo J, Zimmer J. Crystallographic snapshot of cellulose synthesis and membrane translocation. Nature. 2013;493(7431):181–6.

    Article  CAS  PubMed  Google Scholar 

  32. Robledo M, Rivera L, Jiménez-Zurdo JI, et al. Role of Rhizobium endoglucanase CelC2 in cellulose biosynthesis and biofilm formation on plant roots and abiotic surfaces. Microb Cell Factories. 2012;11:125.

    Article  CAS  Google Scholar 

  33. Rossellomora RA, Wagner M, Amann R, Schleifer KH. The abundance of Zoogloea ramigera in sewage-treatment plants. Appl Environ Microbiol. 1995;61(2):702–7.

    Article  CAS  Google Scholar 

  34. Ross P, Mayer R, Weinhouse H, Amikam D, Huggirat Y, Benziman M, et al. The cyclic diguanylic acid regulatory system of cellulose synthesis in Acetobacter xylinum - chemical synthesis and biological-activity of cyclic-nucleotide dimer, trimer, and phosphothioate derivatives. J Biol Chem. 1990;265(31):18933–43.

    Article  CAS  PubMed  Google Scholar 

  35. Hengge R. Principles of c-di-GMP signalling in bacteria. Nat Rev Microbiol. 2009;7(4):263–73.

    Article  CAS  PubMed  Google Scholar 

  36. Seshasayee ASN, Fraser GM, Luscombe NM. Comparative genomics of cyclic-di-GMP signalling in bacteria: post-translational regulation and catalytic activity. Nucleic Acids Res. 2010;38(18):5970–81.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Darmon E, Leach DR. Bacterial genome instability. Microbiol Mol Biol Rev. 2014;78(1):1–39.

    Article  PubMed  PubMed Central  Google Scholar 

  38. Lagesen K, Hallin P, Rodland EA, Staerfeldt HH, Rognes T, Ussery DW. RNAmmer: consistent and rapid annotation of ribosomal RNA genes. Nucleic Acids Res. 2007;35(9):3100–8.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Todd MI, Sean RE. tRNAscan-SE: a program for improved detection of transfer RNA genes in genomic sequence. Nucleic Acids Res. 1997;25(5):955–64.

    Article  Google Scholar 

  40. Gao F, Zhang CT. GC-Profile: a web-based tool for visualizing and analyzing the variation of GC content in genomic sequences. Nucleic Acids Res. 2006;34(Web Server issue):W686–91.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Yoon S-H, Ha S-M, Lim J, Kwon S, Chun J. A large-scale evaluation of algorithms to calculate average nucleotide identity. Antonie Van Leeuwenhoek. 2017;110(10):1281–6.

    Article  CAS  PubMed  Google Scholar 

  42. Wong SM, Mekalanos JJ. Genetic footprinting with mariner-based transposition in Pseudomonas aeruginosa. Proc Natl Acad Sci U S A. 2000;97(18):10191–6.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Kovach ME, Elzer PH, Hill DS, Robertson GT, Farris MA, Roop RM 2nd, et al. Four new derivatives of the broad-host-range cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes. Gene. 1995;166(1):175–6.

    Article  CAS  PubMed  Google Scholar 

  44. Gopu G, Govindan S. Production of bacterial cellulose from Komagataeibacter saccharivorans strain BC1 isolated from rotten green grapes. Prep Biochem Biotechnol. 2018;48(9):842–52.

    Article  CAS  PubMed  Google Scholar 

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Acknowledgements

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Funding

This work was supported by the grant from National Key R&D Program of China (No. 2019YFA0905500), the Strategic Priority Research Program of the Chinese Academy of Sciences Grant (No. XDA23040401), and the Anhui Academy of Agricultural Sciences Wetland Ecology and Application Technology Innovation Team Project (2021YL055).

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Authors and Affiliations

  1. Institute of Hydrobiology, Chinese Academy of Sciences, Wuhan, 430072, China

    Na Gao, Jingcheng Dai, Yaqi Liu, Shuyang Li, Jing Wang & Dongru Qiu

  2. Key Laboratory of Freshwater Aquaculture and Enhancement of Anhui Province, Fisheries Research Institute, Anhui Academy of Agricultural Sciences, Hefei, 230031, China

    Na Gao & Wenxuan Lu

  3. University of Chinese Academy of Sciences, Beijing, 100049, China

    Na Gao, Jingcheng Dai, Shuyang Li, Jing Wang & Dongru Qiu

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Contributions

D.Q. and N.G. designed the experiments; N.G., J.D., Y.L., S. L., and J.W. did the data curation and formal analysis; N.G. and D.Q. wrote the original manuscript; W.L. and D.Q. reviewed and edited the manuscript. All authors have read and approved the manuscript.

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Correspondence to Dongru Qiu.

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Supplementary Information

Additional file 1: Supplemental Figure 1.

Venn diagram showing the number of genes shared by both the strains and unique genes in Shinella zoogloeoides ATCC19623 and XJ20 strain. Supplemental Figure 2. Nucleotide-based alignment of the choromosome genomes from the two strains. Homologous blocks are shown as identically colored regions and linked across the genomes. Regions that are inverted relative to ATCC19623 are shown below the central axis of each sequence. These two chromosome genomes were aligned with Progressive Mauve using default parameters. Supplemental Table S1. Transposon mutants of Shinella zoogloeoides ATCC 19623 defective in floc formation, mapped insertional sites and genetic complementation ATCC19623. Supplemental Table S2. Bacterial strains, plasmids and primers used in this study.

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Gao, N., Dai, J., Liu, Y. et al. Cellulose-mediated floc formation by the activated sludge bacterium Shinella zoogloeoides ATCC 19623. BMC Microbiol 22, 104 (2022). https://doi.org/10.1186/s12866-022-02516-y

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BMC Microbiology

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