Environmental Microbiology

Adaptive Responses of Shewanella decolorationis to Toxic Organic Extracellular Electron Acceptor Azo Dyes in Anaerobic Respiration

Yun Fang, Jun Liu, Guannan Kong, Xueduan Liu, Yonggang Yang, Enze Li, Xingjuan Chen, Da Song, Xuejiao You, Guoping Sun, Jun Guo, Meiying Xu
Harold L. Drake, Editor
DOI: 10.1128/AEM.00550-19

ABSTRACT

Bacterial anaerobic respiration using an extracellular electron acceptor plays a predominant role in global biogeochemical cycles. However, the mechanisms of bacterial adaptation to the toxic organic pollutant as the extracellular electron acceptor during anaerobic respiration are not clear, which limits our ability to optimize the strategies for the bioremediation of a contaminated environment. Here, we report the physiological characteristics and the global gene expression of an ecologically successful bacterium, Shewanella decolorationis S12, when using a typical toxic organic pollutant, amaranth, as the extracellular electron acceptor. Our results revealed that filamentous shift (the cells stretched to fiber-like shapes as long as 18 μm) occurred under amaranth stress. Persistent stress led to a higher filamentous cell rate and decolorization ability in subcultural cells compared to parental strains. In addition, the expression of genes involved in cell division, the chemotaxis system, energy conservation, damage repair, and material transport in filamentous cells was significantly stimulated. The detailed roles of some genes with significantly elevated expressions in filamentous cells, such as the outer membrane porin genes ompA and ompW, the cytochrome c genes arpC and arpD, the global regulatory factor gene rpoS, and the methyl-accepting chemotaxis proteins genes SHD_2793 and SHD_0015, were identified by site-directed mutagenesis. Finally, a conceptual model was proposed to help deepen our insights into both the bacterial survival strategy when toxic organics were present and the mechanisms by which these toxic organics were biodegraded as the extracellular electron acceptors.

IMPORTANCE Keeping toxic organic pollutants (TOPs) in tolerable levels is a huge challenge for bacteria in extremely unfavorable environments since TOPs could serve as energy substitutes but also as survival stresses when they are beyond some thresholds. This study focused on the underlying adaptive mechanisms of ecologically successful bacterium Shewanella decolorationis S12 when exposed to amaranth, a typical toxic organic pollutant, as the extracellular electron acceptor. Our results suggest that filamentous shift is a flexible and valid way to solve the dilemma between the energy resource and toxic stress. Filamentous cells regulate gene expression to enhance their degradation and detoxification capabilities, resulting in a strong viability. These novel adaptive responses to TOPs are believed to be an evolutionary achievement to succeed in harsh habitats and thus have great potential to be applied to environment engineering or synthetic biology if we could picture every unknown node in this pathway.

INTRODUCTION

Bacteria, with an overwhelming quantity in the biosphere, play a vital role in the control of environmental pollution by degradation or element transformation. The bacterial adaptive response to environmental stress is one of the most fundamental processes in bioremediation. For decades, researchers have attempted to investigate the secret of the ecologically successful bacteria as they can quickly respond to environmental changes with their strong functional and phenotypic plasticity (1–3). Bacterial responses to environmental pollutions may vary from species to species, and the successive colonization of a bacterium depends on its ability to sense the changes and to adapt suitably to ensure the best odds of survival. Some environmental pollutants could be irreversibly toxic or even lethal to bacteria, such as antibiotics and cyanides (4). Under sublethal stress, bacteria are allowed to develop an efficient strategy to survive; otherwise, they lose. Generally, (i) passively avoiding damage by reducing permeability and damage repair and (ii) actively removing pollutants by decomposition, transformation, insulation, and efflux are two major strategies to vent the pressure (5, 6). In many cases, however, toxic environmental pollutants may serve as nutrients or energy materials (e.g., carbon source, electron donors, or electron acceptors) as well.

Under hypoxic or anoxic conditions, bacteria can employ various toxic organic pollutants as electron acceptors for anaerobic respiration, which plays key roles in the biogeochemical cycle of elements (7–12), such as dimethyl sulfoxide (DMSO) respiration for the sulfur cycle in deep sea (8) and trimethylamine N-oxide (TMAO) and azo dye respiration for the nitrogen cycle in marine and river sediment (10, 12). DMSO and TMAO respirations have been extensively characterized, particularly in Escherichia coli, marine Shewanella spp., the purple phototrophic bacteria, and Rhodobacter sphaeroides and R. capsulatus (13, 14). Most azo dyes can only be used as extracellular electron acceptors during anaerobic respiration since they hardly permeate into microbial membrane due to their large molecular mass, and then they become more toxic daughter products such as aromatic amines, which might force bacteria to evolve special adaptive strategies to survive (12). Toxic organic pollutants such as DMSO, TMAO, nitrobenzene, and tetrachloroethene have been reported to be degraded by bacteria as intracellular or extracellular electron acceptors during anaerobic respiration (8–11). Some specific or nonspecific elements of electron chains for these organic pollutants have been found. However, more work is needed to illuminate how bacteria respond to toxic organic pollutants since the mechanism is still not very clear (8–12).

We hypothesize that bacteria would actively modify the gene expressions involved in energy conservation, metabolism, damage repair, and cell growth when sensing the TOPs to enhance their detoxification abilities and meanwhile would modulate their phenotype and conserve as much energy as possible from degrading the TOPs to survive or even grow in harsh habitats. To validate these hypotheses, we analyzed the responses of azo respiration bacterium Shewanella decolorationis S12 to amaranth, a typical azo dye, as the extracellular electron acceptor (15–17). This study provided new insights into our understanding of bacterial responses to environmental stress and laid a foundation for the efficient removal of TOPs in an anaerobic environment.

RESULTS

Filamentous cell formed under amaranth condition.Morphological transitions were observed in S. decolorationis S12 cells under azo-reducing conditions. In the anaerobic incubation with amaranth, S12 cells stretched to fiber-like shapes as long as 18 μm (Fig. 1A), much longer than the normal rod size (∼2 μm, Fig. 1B). When transferred to a fresh medium without amaranth (both aerobic and anaerobic), these filamentous cells tended to restore the normal sizes (Fig. 1C), suggesting that the filamentous shift was one of the S12 adaptive responses to amaranth. To validate this, we analyzed more morphological shifts under different concentrations of amaranth using viability staining and laser scanning confocal microscopy (LSCM) (Fig. 1D). The results showed that filamentous cells were elongated cells with multiple nucleoids and maintained high metabolic activity (colored green). Furthermore, we analyzed the cell size distribution and decolorization efficiency under different concentrations of amaranth (0, 0.1, 0.2, 0.4, 0.5, 1.0, 2.0, 3.0, 4.0, and 5.0 mM) in the presence or absence of oxygen over time (Fig. 1E; see Fig. S1 in the supplemental material). For anaerobic conditions without amaranth, no filamentous cells were found in 32 h. When the amaranth was in a range of 0 to 2.0 mM, filamentous cells appeared at 8 h, and the filamentous rate increased over time and peaked at 24 h, with a maximum filamentous rate of 62%; it then decreased at 32 h. For the anaerobic condition with 3.0 to 5.0 mM amaranth, the filamentous cell occurred at 8 h, and the filamentous cell rate fluctuated in 32 h, with an average filamentous rate of 20%. For the aerobic condition, no obvious decolorization of amaranth was observed, and no filamentous cells were detected with 0 to 2.0 mM amaranth, and yet filamentous cells appeared with an average rate of 20% when the concentration of amaranth was more than 3.0 mM. Notably, we analyzed the anaerobic growth of cells in the presence of 0.5 mM amaranth. As shown in Fig. S1C, the cell number significantly increased over the time course of the experiment, while the concentration of amaranth showed an opposite trend. The initial inoculum was approximately 1 × 107 CFU/ml. In 8 h, the cell number doubled, that is, increased to 2.0 × 107 CFU/ml and reached a maximum of 3.78 × 107 CFU/ml in a 12-h reaction. However, for the control without amaranth, the cell count was 1.19 × 107 CFU/ml in 8 h and was approximately 1.51 × 107 CFU/ml in a 12-h reaction. These data showed that amaranth could serve as terminal electron acceptor under the anaerobic condition. Similar filamentous cells were detected for other azo dyes, such as acid red 13 and β-naphthol violet (Fig. S2).

FIG 1
FIG 1

Filamentous cell count and observation of S. decolorationis S12 cells under different azo dye conditions. (A) TEM images of filamentous cells with amaranth; (B) TEM images of normal cells without amaranth; (C) SEM images of cells from subculture of filamentous cells without amaranth; (D) LSCM images of cells under gradient concentrations of amaranth. Green and red represent the live and dead cells, respectively. Am, amaranth. (E) Length distributions of cells under different concentrations of amaranth.

Higher decolorization ability in filamentous cells.Serial batch cultivations were employed to examine the responses of S12 to persistent azo-dye stress because S12 cells were allowed to adapt or evolve in the daily serial transfer to fresh media containing amaranth. It turned out the derivatives performed better in amaranth degradation than the parental strains, which could be considered an advantage of the adaptation (Fig. 2A). Furthermore, a positive relationship (R2 = 0.928, P = 0.008) was observed between the decolorization efficiency and the filamentous cell rate, implying that the cells with filamentous structure somehow have advantages in reducing azo dyes (Fig. 2B).

FIG 2
FIG 2

Decolorization of azo dye and the filamentous cell rate of clones from ancestral and subcultural S. decolorationis S12. (A) Decolorization of azo dye; (B) filamentous cell rate. A t test was used to test the difference between ancestral and subcultural S. decolorationis S12 (**, P < 0.01).

Overall review of transcriptional responses of strain S12 to azo dye.In order to understand the mechanisms of S. decolorationis S12 response to azo dyes and to probe the key functional genes involved in extracellular azo respiration, RNA sequencing (RNA-Seq) technology was used to profile the global gene expressions of strain S12 exposed to 0.5 mM amaranth for 0.5 h (normal cells) or 8 h (filamentous cells appeared at this time point). The statistical results of transcriptomic data are shown in Table S1, and the expression of 27 genes from four different operons was analyzed to evaluate the data (Fig. S3). The patterns of gene expression within the same operon were consistent, suggesting a solid reliability of the transcriptomic data. Among the total open reading frames in the genome, ∼20% were upregulated and ∼16% were downregulated after azo incubation. Based on COG functional classification (18), genes with significantly changed expressions were mainly involved in the process of energy production and conversion, translation, posttranslational modification, protein turnover, chaperone functions, inorganic ion transport and metabolism, amino acid metabolism and transport, signal transduction, coenzyme metabolism, and cell motility (Fig. 3), especially signal transduction, the energy process, damage repair, and transcriptional regulation.

FIG 3
FIG 3

Heatmap of significantly affected gene COG categories responding to azo dye. The numbers of gene members of each COG category altered (up- or downregulated) are summarized in the heatmap. The COG category is listed on the left y axis, and the total number of genes comprising that category is provided (labeled as n). B, chromatin structure and dynamics; C, energy production and conversion; D, cell cycle control and mitosis; E, amino acid metabolism and transport; F, nucleotide metabolism and transport; G, carbohydrate metabolism and transport; H, coenzyme metabolism; I, lipid metabolism; J, translation; K, transcription; L, replication and repair; M, cell wall/membrane/envelope biogenesis; N, cell motility; O, posttranslational modification, protein turnover, chaperone functions; P, inorganic ion transport and metabolism; Q, secondary metabolites biosynthesis, transport and catabolism; R, general functional prediction only; S, function unknown; T, signal transduction; U, intracellular trafficking and secretion; V, defense mechanisms. The percentage of all gene members of the COG category altered is provided on the right side. The heatmap is sorted by this percentage.

Genes for cell division proteins.RNA-Seq data showed that although no significant change was observed in the expression of ftsZ (SHD_0544, cell division protein FtsZ), a series of cell division genes were downregulated when exposed to amaranth (Table 1), such as zapA (SHD_3759, cell division protein ZapA), zapB (SHD_0848, cell division protein ZapB), ftsN (SHD_1891, sporulation domain-containing protein FtsN), merB (SHD_2830, rod shape-determining protein MreB), and merC (SHD_2829, rod shape-determining protein MreC). These results suggested that the formation of filamentous cells could be a result of stalled cell division.

View this table:
TABLE 1

Expression and differential expression pattern of key genes in response to azo dye

Genes for chemotaxis and signal transduction systems.Chemotaxis and signal transduction systems mediate the response of motile bacteria to a chemical stimulus. Among the 77 genes involved in chemotaxis in S. decolorationis S12, most (43/77, 56%) were stimulated in the filamentous cells, with the highest being the four methyl-accepting chemotaxis protein (MCP) genes SHD_2793 (3.5-log2-fold induction, P < 0.001), SHD_1488 (2.49-log2-fold induction, P < 0.001), SHD_1981 (2.48-log2-fold induction, P < 0.001), and SHD_0015 (2.46-log2-fold induction, P < 0.001) (Table S2). Two gene clusters encoding chemotaxis proteins (CheABDRWY) were also upregulated in filamentous cells, especially the signal transduction histidine kinase gene cheA1 (SHD_0014, 2.85-log2-fold induction, P < 0.001) and the response regulator receiver protein gene cheY1 (SHD_0012, 2.88-log2-fold induction, P < 0.001) (Fig. S4). Meanwhile, the expressions of the genes involved in cell motility were also increased in the filamentous cells, including SHD_1332 (fimbrial biogenesis outer membrane usher protein, 1.52-log2-fold induction), SHD_1333 (pilus assembly C-terminal domain protein, 1.57-log2-fold induction), and the gene cluster SHD_2104-2111 (flgF, 1.31; flgG, 1.45; flgH, 1.75; flgI, 1.47; flgJ, 1.57; flgK, 1.46; flgL, 1.48; SHD_2111, 0.6-log2-fold induction). It is of much interest to examine the mechanism of chemotaxis to azo dyes in S. decolorationis S12. Still, we do not know which exact signal(s) triggered the transduction pathway, but some outer membrane porin proteins give us cues, such as OmpW and OmpA. The gene ompW was significantly induced (SHD_2431, 3.57-log2-fold induction, P < 0.001) and the FPKM value (i.e., the fragments per kilobase of transcript per million mapped reads) was very high (855.4 for normal cells and 10,178.4 for filamentous cells). Another gene, ompA, was also induced (SHD_3106, 1.51-log2-fold induction, P < 0.001). A knockout mutant carrying an in-frame deletion of ompA or ompW was constructed (ΔompA and ΔompW). An obvious decrease in amaranth decolorization rate occurred for these mutants, especially for the ΔompA mutant, in comparison to the wild-type S12 (Fig. 4), and the decolorization rate of their complemented strains returned to normal levels (Fig. S5). Similar results were observed with other azo dyes, such as methyl orange and methyl red (data not shown). These results indicated that in order to use amaranth as electron acceptor, S. decolorationis S12 upregulated the expressions of signal transduction genes and changed the chemotactic strategies.

FIG 4
FIG 4

Anaerobic reduction of amaranth by wild-type (WT) and mutant strains of S. decolorationis S12.

Genes for cytochromes in energy conservation.Electron transport/transfer from electron donors inside bacteria to extracellular acceptors (azo bond) lead to the reduction of azo dye. The cytochrome c family was a group of electron transfer proteins, which were vital for energy conservation during that process. Our results showed that the genes encoding the cytochrome c biogenesis protein (SHD_0398-0401) and the heme exporter protein (SHD_0392-0396) were significantly upregulated (Table 1). According to the gene expression profiles, the electron transfer pathway was put forward as follows (Fig. 5). The gene operons SHD_2062-2063 and SHD_3465-3466 encoding cytochrome bd ubiquinol oxidase for ubiquinol (or menaquinoe) oxidization were significantly induced in the filamentous cells. The gene SHD_1345 encoding CymA, a cytoplasmic-membrane-bound cytochrome c for transporting electrons from the quinol pool to a number of reductases, was induced. Two periplasmic tetraheme cytochrome c protein-encoding genes, SHD_2946/cctA (cytochrome c3, 1.20-log2-fold induction) and SHD_3291/cytC (tetraheme cytochrome c, 1.76-log2-fold induction), were upregulated. Meanwhile, operon SHD_2525-2531 containing two outer membrane protein genes, mtrB and mtrE, and five cytochrome c protein genes, mtrC, mtrF, mtrA, mtrD, and omcA, showed at least a log2-fold induction of 1.37. In particular, another operon SHD_1874-1876 containing one outer membrane protein gene, arpB, and two cytochrome c protein genes, arpC and arpA, as well as the decaheme cytochrome c gene SHD_2419/arpD showed a log2-fold induction ranging from 3.17 up to 8.1. This explains why slight decreases in the decolorization rate were observed in ΔarpC and ΔarpD mutants compared to the wild-type S12 (Fig. 4). Products of SHD_1874/arpC and SHD_2419/arpD potentially function as an extracellular azoreductase in S. decolorationis S12. Further experimental evidences are required to identify their exact biological roles.

FIG 5
FIG 5

Conceptual cellular model of S. decolorationis S12 responses to amaranth. Red or blue fonts indicate increased or decreased gene expression, respectively. (A) P, phosphate. (B) Solid lines represent material transformation or transport, and dashed lines represent electron transports. DPs, degradation products; Q, ubiquinone; QH2, ubiquinol. (C) Perpendicular line represents negative regulation. (D) Arrow represents positive regulation. CRP, cyclic AMP receptor protein; AHR, alkyl hydroperoxide reductase; ThiR, thioredoxin.

Genes for damage repair.It is inevitable to confront stress and maintain redox homeostasis for cells under harsh conditions. The data showed that genes involved in DNA repair, protein refolding, antioxidant activity, and stress response protein were upregulated in the filamentous cells (Table 1). Expression of the DNA repair genes SHD_1650, SHD_2003, SHD_2561, SHD_3077/radC, and SHD_3320 was induced. Two pairs of chaperone genes, dnaJ/dnaK and groEL/groES, which may play a vital role in protein refolding for injured proteins, were almost 2.0 log2-fold induced. Two heat shock protein genes, SHD_0443/ibpA (HSP20, 3.66-log2-fold induction, P < 0.001) and SHD_0530/htpG (HSP90, 2.36-log2-fold induction, P < 0.001), were also significantly upregulated. Furthermore, several genes encoding proteases and peptidases were induced in the filamentous cells: SHD_1787 encoding peptidase S8/S53 with serine-type endopeptidase activity was induced by 3.9-log2 fold, SHD_1862-1863 encoding proteases HslU and HslV with ATP-dependent peptidase activity was induced by 2.2- and 1.4-log2 fold, and SHD_2978/clpB encoding ATP-dependent protease was induced by 3.3-log2 fold. Most of the genes involved in redox homeostasis were also induced in the filamentous cells, such as SHD_3266 and SHD_4043 (encoding alkyl hydroperoxide reductase), SHD_4071 (encoding thioredoxin), and SHD_3569-3570 (encoding a glutathione/cysteine transporter) (Table 1). These results suggested that the antioxidant was highly associated with azo respiration because its synthesis and transport were enhanced in the filamentous cells more than in the normal S12 cells.

Genes for transcriptional regulation.The sigma factor is essential for transcription initiation. Six sigma factor-encoding genes—rpoD (σ70), rpoE (σ24), rpoF (σ28), rpoH (σ32), rpoS (σ38), and rpoN (σ54)—were detected in this study (Table S3). The expression of rpoD was highly enhanced (FPKM > 500) in both filamentous and normal cells since it is a housekeeping sigma factor gene and keeps essential genes and pathways operating (19). The extracytoplasmic heat stress sigma factor gene, rpoE, was slightly induced in the filamentous cell (log2-fold induction = 0.58, P = 0.015). Interestingly, rpoS (σ38) showed an unexpected expression profile between filamentous and normal cells, with a log2-fold change of 1.45 (P = 0.002) in the former. The deletion of rpoS caused the significant decrease in the decolorization rate of strain S12, suggesting its crucial role in regulation of azo respiration (Fig. 4). In this study, rpoS may actively regulate the two-component system narQP since DNA binding sites of σ38 were found upstream of narQP, and the expression of narQP was induced in the filamentous cells (1.69- and 1.15-log2-fold induction for narQ and narP, respectively). The crp gene, induced in the filamentous cells, may serve as a regulator for narQP, ccmABCDE, cydAB, mcp, and ompW. Moreover, another gene, dksA, facilitating the binding of σ38 to the promoter was also upregulated (log2-fold induction = 1.56, P = 0.0002).

DISCUSSION

Bacteria comprise the largest part of the Earth’s biomass in terms of number of species, performing essential functions in natural and artificial environments, as well as on and inside living organisms. With the development and application of the stable isotope labeling technology, high-throughput molecular biology techniques, and other new analytical detection techniques, considerable studies showed that functional microorganisms in the polluted environment could utilize toxic pollutants as their own nutrients. These functional microorganisms are indispensable in the process of environmental purification, and they provide good biological resources for bioremediation. Pollutants with high redox potentials are ubiquitous and can be either electron acceptors for those able to tolerate and reduce them or stress for those that cannot. The bacterial adaptive responses discovered in this study involved morphological switch and function enhancement, which we believe were driven by the modulation of the original gene expression network.

Filament response and decolorization enhancement.One hypothesis is that phenotype modulation, especially morphological shift, is a survival strategy to promote pollutant degradation and detoxification. In this study, bacterial morphological change, from rod-shaped to filamentous, was observed, with a toxic organic pollutant being the extracellular electron acceptor. Bacteria are known to undergo filamentous shift when coping with other environmental stress, such as cold, UV radiation, chromium, and BTEX (benzene, toluene, ethylbenzene, and o-xylene) (1, 20–22). It is believed that filamentation has been implicated in bacterial survival under environmental stresses with several benefits (23–26). First, stretched growth of a cell increases its uptake-proficient surface without changing its surface/volume ratio appreciably. Second, filamentation increases specific surface area in direct contact with the solid medium and may benefit cells attached to a surface. Third, the fiber-like filamentous cell body allows cells to access more energy materials that would be out of reach by rod-shaped ones and connects the nutrient-rich and -deficient regions via intracellular transfer. Finally, delayed cell division could conserve energy for survival activities. Previous studies showed that this phenomenon is a temporary response to suboptimal environment, and yet no evidence has been proposed to identify whether it could be passed on and enhanced in derivatives. In our work, subculture of strain S12 showed that the cells experiencing long-time exposure to azo dyes could adapt to amaranth with higher decolorization ability and filamentous rate. We inferred that induced transcriptional and/or epigenetic regulations and even adaptive evolution in genotype might occur during long-time stress exposure.

Bacterial filamentation is due to the lack of septation (typically caused by DNA damage or cell wall damage) during cell division and results in a stretched cell body. Under anaerobic conditions, two aromatic amines, 1-aminenaphthylene-4-sulfonic acid and 1-aminenaphthylene-2-hydroxy-3,6-disulfonic acid, are the main biodegradation products of amaranth (11). Amaranth and its degradation products had considerable toxicity to S. decolorationis S12, but the latter were much more dangerous because of their elevated hydrophobicity and smaller molecules, which allow them to easily penetrate the cytoplasmic membrane and to accumulate inside the cytosol (11, 27). Cell division is a complex process involving more than 30 proteins in Escherichia coli, the model strain for prokaryotic study. In general, FtsZ, FtsA, and ZipA assemble to form the basal proto-ring complex anchoring to the cytoplasmic membrane (28, 29); FtsK, FtsQ, FtsB, and FtsL then adhere to the divisome, the complex interacting with the nucleoid and the periplasmic space (30, 31). Next follows the incorporation of five proteins (FtsW, FtsI, FtsN, MreB, and MreC) related to the synthesis of divisive peptidoglycans (32, 33) and another five (ZapA, ZapB, MinC, MinD, and MinE) related to the segregation of the nucleoid (34–37), which ultimately results in cell wall invagination and the formation of two daughter cells. Study on the divisome in the Shewanella genus is limited, but the genes potentially encoding the proteins mentioned above could also be located in the S12 genome. The gene expression profiling indicated that fstN (encoding a sporulation domain-containing protein) had a curial role in morphological determination under aromatic stress. FtsN is the last protein in the hierarchy of divisome assembly, and its C-terminal peptidoglycan-binding domain (SROP) may contribute to its midcell localization via specific recognition of a septal signal (38–40). E. coli cells expressing less FtsN stopped dividing, while their mean cell length continued increasing, and formed a filamentous cell body (31, 41). Khandige et al. previously suggested that another SPOR domain-containing-protein-encoding gene, damX, controlled the filamentous morphology in E. coli, and yet it was not differentially expressed in rod and filamentous S12 cells (42). The precise location of the degraded amaranth and proto-ring components during filamentation and the role that cell divisome proteins (e.g., FstN, MreB, MreC, ZapA, and ZapB) play in this scheme remain subjects for future study.

For another, under aerobic conditions, no filamentous cell appeared without adding amaranth, indicating that amaranth and its degradation products might trigger filamentous phenomena. Moreover, it was observed that a significantly higher concentration of amaranth (≥3 mM) was needed to induce filamentation of cells under aerobic conditions in comparison to that (≥0.5 mM) under anaerobic conditions, which might be attributed to two reasons: (i) oxygen and amaranth competed for electrons, and a higher concentration of amaranth (≥3 mM) was more competitive and (ii) different compositions of biodegradation products might be produced with different concentrations of amaranth under anaerobic and aerobic conditions (11), and a higher concentration of amaranth (≥3 mM) and its degradation products might have more cytotoxicity for triggering cell filamentation. Accordingly, there may be different induction mechanisms of cell filamentation when amaranth was biodegraded under anaerobic and aerobic conditions. These inferences provide a basis for future research.

Transcriptional modulations concerning amaranth degradation and detoxification.Our second hypothesis is that the expression of functional genes would respond to azo stresses, especially those related to energy conservation, metabolism, damage repair, and cell growth. In order to survive and grow under such an environment, bacteria must keep adjusting their metabolism and behavior after sensing the specific cues with dedicated receptors. The best-characterized signal transduction system in bacteria is that of E. coli involved in chemotaxis (43, 44). At the molecular level, it comprises a two-component signal transduction pathway in which sensory information perceived by dedicated receptors (such as E. coli Tsr, Tar, Aer, Trg, and Tap) is relayed to the flagellar motor(s) via a series of phosphorylation events initiated at the chemotactic kinase CheA, which is coupled to the receptors via CheW and modulates the phosphorylation state of the CheY response regulator. Phosphorylated CheY has an increased affinity for flagellar motor proteins, and their combination results in a directed flagellar rotation against the stress, probably via conformational spread. In our study, the expression of S12 chemotaxis-involved genes is vigorous. We postulate that signal recognized by receptors is relayed to CheY-1 by CheA-1. CheY-1 undergoes a conformational change and becomes able to bind to the motor switch, causing a change in the direction of flagellar rotation to approach amaranth. Although the chemotactic signaling pathway is well conserved among motile bacteria, the numbers and types of methyl-accepting chemotactic proteins (MCPs; the sensory transducers), which are typically membrane-bound proteins responsible for the detection of extracellular signals, vary greatly among species. The presence of large MCP families in bacteria is thought to reflect their metabolic versatility and their ability to adapt to the complex environment in which they live. For example, S. decolorationis S12 has 43 MCPs, Pseudomonas species have on average 33, and E. coli only has 5 (16, 45, 46). Only a few MCPs have been functionally characterized to detect aromatic pollution. For instance, Aer2 could detect phenylacetic acid and PcaY could detect aromatic acid in P. putida F1, and MCP2211 could detect 4-hydroxybenozate in Comamonas testosteroni (47–49). It is reported that MCPs could either directly detect chemoattractants that enter the periplasm (e.g., PcaY senses aromatic acid) (48) or indirectly detect metabolic intermediates of chemoattractants. For example, Aer2 senses the flavin adenine dinucleotide (FAD)/reduced FAD (FADH2) ratio and MCP2211 senses succinic acid resulting from the tricarboxylic acid cycle, which indicate the levels of phenylacetic acid and 4-hydroxybenozate individually (47, 49). Nevertheless, no study has reported MCPs being the electron transporters for azo dyes. In this study, four MCP genes, SHD_2793, SHD_1488, SHD_1981, and SHD_0015, were significantly induced under azo stress, and the deletion of SHD_2793 and SHD_0015 caused an obvious decrease in the decolorization efficiency of strain S12 (Fig. 4), suggesting that these MCPs were likely to relay the amaranth signal. Currently, it is not clear how the degradation products of amaranth traverse the outer membrane, but the outer membrane porins seem to facilitate their transport. Amaranth was degraded extracellularly, and we supposed that degradation products could enter the periplasm and were recognized by specific MCPs to switch on a chemotaxis response. Several upregulated MCPs (products of SHD_2793, SHD_1488, SHD_1981, and SHD_0015) may serve as receptors for chemoattractants and the elevated outer membrane porins such as OmpW and OmpA may be employed to transport these degradation products. This idea was supported by Pilsl et al. (50) and Foulds et al. (51), who reported that OmpW was associated with the formation of colicin S4 porin, and OmpA functioned as a receptor for colicin L in E. coli. Interestingly, an operon SHD_2710-2713 was induced by over 3-log2-fold during amaranth decolorization, which we believed was an adaptive response of S12 to azo stress. The product of SHD_2710-2713 was an efflux transporter of the RND family (refers to resistance, nodulation, and cell division), which was reported to be a multidrug transporter (52). It may detoxify by expelling the toxic degradation products outside the cell.

When S12 reached amaranth by chemotaxis, an electron transport chain from cytoplasm to the extracellular amaranth was established and cytochrome c was sure to play a key role in it. That explains why in filamentous cells the transcription of cytochrome c-related genes (e.g., cymA, cctA, mtrCAB-omcA-mtrFDE, and arpABCD) was maintained at high levels. The mtrCAB-omcA operon encodes two decaheme c-type cytochromes and an outer membrane protein linked to metal reduction such as Fe(III), Mn(IV), U(VI), and Cr(VI), which could also be found in other Shewanella species (53). Bencheikh-Latmani et al. (54) reported that the expression of mtrCAB-omcA was upregulated under metal-reducing conditions in S. oneidensis MR-1, suggesting that the increased expression detected in the presence of azo dyes is nonspecific. The operon mtrCAB-omcA-mtrFDE was also involved in the extracellular reduction of textile dyes in S. oneidensis (55, 56). The high expression of CymA (a periplasmic tetraheme cytochrome c anchored in the inner membrane) in the presence of azo dyes is consistent with that in the presence of nitrate, fumarate, Fe(III), and Mn(IV), all of which require CymA during the reduction pathway in S. oneidensis MR-1 (53, 57, 58). Knockout of cymA resulted in a substantial loss of azo reduction ability in S. oneidensis (53). cctA was another gene involved in electron transfer during Fe(III) reduction in S. frigidimarina (59) and S. oneidensis (60). Decaheme cytochrome c (SHD_2419, ArpD) in S. decolorationis S12 has an 82% similarity with the corresponding protein (SO_1659) in S. oneidensis, a surface-bound decaheme cytochrome c lipoprotein belonging to the OmcA/MtrC family. Octahame cytochrome c (SHD_1874, ArpC) in S. decolorationis S12 also has a high sequence similarity to the corresponding proteins in S. frigidimarina (Sfri_3556, 81%) and S. oneidensis (SO_4144, 97%), which were reported to be the reductases for tetrathionate, nitrite, and hydroxylamine (60, 61). It is proved that E. coli could tolerate and reduce azo dyes, but there was no evidence showing any specifically induced cytochrome located in the outer membrane (62), suggesting different pathways of azo reduction for E. coli K-12 and S. decolorationis S12.

As we mentioned above, filamentous cells confronted more stress than the normal cells did. Pleiotropic regulations with respect to damage repair and transcriptional regulation network were implemented to relieve the pressure. Heat shock proteins (HSPs) act as molecular chaperones, and their functions included prevention of protein aggregation, protein degradation, protein trafficking, and maintenance of signaling proteins in a ready-to-activate conformation (63, 64). HSP20 was reported to form large heterooligomeric aggregates to protect other proteins against stress-induced denaturation and aggregation, and HSP90 can regulate a specific subset of cellular signaling proteins (65, 66). Crabbé et al. (67) also found a pronounced induction of the HSP IbpA and recombinase RecA in the filamentous cells of Pseudomonas putida KT2440. It was found that alkyl hydroperoxide reductase (AHR), usually referred to as peroxiredoxin, as well as glutathione and thioredoxin was an important antioxidant in bacteria preventing damage to important cellular components caused by reactive oxygen species such as free radicals and peroxides (68–70). High expression of alkyl hydroperoxide reductase was also observed under Co(III)-, Mn(VI)-, thiosulfate-, and nitrate-reducing conditions in S. oneidensis (71). RpoS is a central sigma factor in E. coli involved in gene expression under a large number of stress conditions (72). In this research, the ΔrpoS mutant lost almost 50% of decolorization capacity compared to the wild-type strain. A regulative gene network was put forward according to bioinformatic tools, and much experimental work was needed to certify it. According to the data from Regprecise (http://regprecise.lbl.gov/), transcriptional factor NarP, regulated by RpoS, potentially regulated MtrCBA, MtrFED, OcmA, and OmpW, as well as a global regulator CRP (cyclic AMP receptor protein) in Shewanella spp., since the DNA binding sites where NarP activates their expressions were localized upstream of the coding regions. In addition, although azo dyes are toxic to S12, we did not find significant effects of the genes encoding transcriptional regulators of the MarR family, which determine the tolerance of E. coli to antibiotics, organic solvents, and oxidative stress, as well as azo dyes (62, 73). It seems therefore that different regulative systems existed in S. decolorationis S12 and E. coli.

Conceptual cellular model of S. decolorationis S12 responses to azo dye.In this study, S. decolorationis S12 was used to demonstrate the bacterial responses and adaptive mechanism under azo dye stress. Considering all of the experimental results and our general knowledge together, a conceptual cellular model is proposed (Fig. 5). Stress sensation marks the initiation of all cascade responses. The initial signal of azo dye is recognized by porins OmpA/OmpW and then relayed to chemoreceptors (MCPs). After a series of signal transduction and transcriptional regulations, chemotactic proteins (CheABDRWY) are finally activated, which trigger the chemotactic system to function and S12 cells begin to approach azo dyes. Another benefit from the transcriptional regulations is a morphological switch from rod-shaped to filamentous that occurs to S12 cells, along with enhanced decolorization ability and an extended access to azo dyes. However, based on all clues collected so far, we could not identify which appears earlier, the chemotaxis or the morphological switch. When in direct contact with azo dyes, S12 cells accelerate their electron transfer to reduce them. In our study, amaranth has problems entering the cytoplasm and thus is reduced extracellularly as the final electron acceptor. Cytochrome c proteins may play central roles in this process. Three clusters of them (MtrABC-OmcA, MtrDEF-OmcA, and ArpABCD) are potential azoreductases, and the other three (CymA, CytC, and CttA) are periplasmic proteins. The degradation products of amaranth are more toxic, partially because they are smaller molecules with higher hydrophobicity, making them easier to penetrate the cytoplasmic membrane and accumulate inside. In response to that, S12 ceases cell division to conserve energy for survival activities, including the exclusion of toxic products via RND efflux pumps and the enhanced biosynthesis of molecular chaperons (e.g., Hsp90, Hsp20, and AHR) to repair damaged DNA and proteins. We also found a positive feedback in that the internal toxic reactants would stimulate S12 to switch on the regulation factor σ38 to further activate genes involved in chemotaxis and electron transfer, all of which eventually enhance the viability of S. decolorationis S12 in anaerobic and polluted habitats.

MATERIALS AND METHODS

Chemicals, organism, media, and cultivation.Amaranth, acid red 13, and β-naphthol violet were purchased from Sigma (St. Louis, MO); their molecular structures are depicted in Fig. S6. S. decolorationis S12 was isolated from the activated-sludge of a textile-printing wastewater treatment plant, Guangzhou, China (15). S. decolorationis S12 was cultivated by transferring a single clone to a 150-ml conical flask containing 50 ml of LB medium (10 g/liter peptone, 5 g/liter yeast extract, 10 g/liter NaCl) and was then incubated in a shaker (160 rpm, 30°C). The cells were harvested in the middle of the exponential growth phase (∼8 h) by centrifugation, washed twice, and resuspended with phosphate buffer (0.1 M [pH 7.4]) prior to inoculation. The cells were inoculated into the revised lactate mineral (LM) medium (pH 7.0) containing 7.64 g/liter Na2HPO4, 3.00 g/liter KH2PO4, 0.50 g/liter NH4Cl, 1.00 g/liter NaCl, 5.00 mmol/liter sodium lactate, 0.50 g/liter yeast extract, and 0.50 mmol/liter amaranth and were statically cultivated at 30°C in an anaerobic workstation (BugBox; Ruskinn Technology). The initial inoculum was approximately 1 × 107 CFU/ml. A standard anaerobic technique was used throughout the study, as described previously, to guarantee the oxygen depletion (17). All batch experiments were conducted in 100-ml serum bottles containing 40 ml of LM medium under anaerobic conditions. Each experiment was independently performed in triplicate.

Azo dye response experiment of S. decolorationis S12.Stress response experiments were conducted to examine the effect of persistent azo dye stress on S. decolorationis S12. Three replicates of monoculture were set up following the protocol modified from Lawrence et al. (74) in Fig. S7. Every 24 h, cells from each microcosm were passed on to a new batch into 40 ml of fresh LM medium, and a total of 5 batches over 5 days were obtained. The bacteria were isolated from the final batch by plating on LB medium agar, picking up single colonies, and resuspending them in LB medium. All six subcultural colonies (e101 to e106) were inoculated into LM medium to compare the decolorization activity with six ancestral culture colonies (a001 to a006). Frozen isolates from the ancestral culture were first grown in LB medium, and then an aliquot was sampled to start the assay cultures. The optical densities at 600 nm (OD600s) of all the stock cultures were measured before inoculation to ensure the similar initial cell concentrations. The decolorization rates of the cells from each batch were monitored every 2 h, and the cell morphologies were observed in 24 h. Serial dilutions of amaranth (0, 0.1, 0.2, 0.4, 0.5, 1.0, 2.0, 3.0, 4.0, and 5.0 mmol/liter) were used to examine the effect of azo dye on cell morphology in the presence or absence of oxygen.

Microexamination of cell morphology.In order to monitor the morphology of strain S12 during decolorization, the cells were stained with crystal violet and then immobilized on a glass slide. Image capture and analysis were performed by a Leica DM LB light microscope and a Leica Qwin V3 image analysis system (Leica, Inc., Solms, Germany). Normal S12 cell width is ∼0.8 μm, and the length is ∼2 μm. In this study, cells longer than 5 μm are defined as filamentous cells. Filamentous cell rate was calculated as follows: filamentous cell rate (%) = n/N × 100%, where n is the filamentous cell number, and N is the total cell number. Approximately 200 cells were measured for each culture. Observation of the surface morphology with a high resolution was conducted by a transmission electron microscope (TEM) and scanning electron microscopy (SEM).

To estimate the viable and total counts of bacteria during the decolorization, cells were fixed and stained with fluorescence using a Live/Dead BacLight viability kit (Molecular Probes, Inc., Eugene, OR) according to the manufacturer’s protocol and observed under a laser scanning confocal microscope (LSM700, Zeiss, Braunschweig, German).

Transcriptional analysis of filamentous and normal cells. (i) RNA isolation, enrichment, and sequencing.To examine differences of the global gene expression between filamentous and normal cells, the RNAs of triplicate samples that were exposed to 0.5 mM amaranth for 0.5 h and another three exposed for 8 h were sequenced. Cell pellets from centrifugation (5,000 × g, 10 min) were suspended in 1 ml of RNAprotect bacterial reagent (Qiagen, Carlsbad, CA) for 5 min, harvested by centrifugation (5,000 × g, 10 min), and resuspended in 200 μl of RNase-free water before assay. The mixture was digested by lysozyme (2.5 mg/ml) for 10 min at 25°C and then protease K (1 mg/ml) for another 10 min at 25°C. RNA samples were purified by an RNeasy kit (Qiagen, Valencia, CA) and treated with RNase-free DNase I (Qiagen, Carlsbad, CA) to digest residual genomic DNA. RNA concentration was determined by spectrophotometry (BioSpecnano; Shimadzu, Kyoto, Japan). RNA quality was assessed using a nanochip on a bioanalyzer (model 2100; Agilent Technologies, Santa Clara, CA), and the RNA integrity numbers (RIN) were >8.5, indicating good qualities (Table S4). Equal amounts of six DNA-free RNA samples were pooled for further analysis. rRNA was removed by using a RiboZero rRNA removal kit (Epicenter, Madison, WI) according to the manufacturer’s protocols (75). mRNA was used to construct a 300-bp cDNA library and then sequenced on a HiSeq 2000 platform (Illumina, San Diego, CA) in the same lane with an index 101 PE method. A detailed description for RNA sequencing is included in the supplemental material.

(ii) RNA-Seq data processing.The raw data after sequencing went through a four-step quality control to construct a clean data set with high reliability. Sequences having any one of the four situations below were eliminated: (i) 3 or more “N” bases in a single read, (ii) adapter contamination, (iii) more than 36 bases with Q-score below 20, and (iv) duplicated reads generated in PCR. The clean data were mapped on the annotated genomic objects identified in the genome of S. decolorationis S12 using TopHat (v2.0.11) and SAMtools (v0.1.18.0), yielding BAM files (76, 77). Sorted and indexed BAM files were analyzed by Cufflinks (v2.0.0) (78) to calculate the number of fragments per kilobase of transcript per million mapped reads (FPKM) for all genes and thereby to detect differentially expressed genes. Based on the calculations made by Cuffdiff, genes with |log2-fold changes (filamentous verse normal cell)| > 1 and a false discovery rate (FDR) P value of <0.05 were marked as significant. Based on the relative expression (79), genes were grouped into six major categories: very high (FPKM > 500), high (200 < FPKM < 500), moderate (50 < FPKM < 200), modest (10 < FPKM < 50), low (2 < FPKM value< 10), and not expressed (FPKM < 2).

(iii) Sequence accession number.The transcriptomes of S. decolorationis S12 exposed to amaranth treatment are available at the NCBI Gene Expression Omnibus database under accession number GSE64532.

Construction of mutant and complementary strains.Deletion mutation strains of S. decolorationis S12 (ΔompA, ΔompW, ΔarpC, ΔarpD, ΔrpoS, ΔSHD_2793, and ΔSHD_0015) were individually constructed using the fusion PCR method (80). Primers used for the mutagenesis are listed in Table S5. pHG102 was used in the genetic complementation of mutation strains (80). The target DNA fragments harboring those genes were amplified with primer pairs listed in Table S6. A detailed description for the construction of mutant and complementary strains is included in the supplemental material.

UV analyses and statistical analysis.The maximum absorbance of amaranth solution occurred at 520 nm, and thus the decolorization of amaranth was measured by monitoring the decrease in A520 over time. Considering a linearity between the A520 and the concentration of amaranth, the decolorization rate was calculated as (A – B)/A × 100%, where A is the initial absorbance, and B is the observed absorbance after the reaction. All assays were done in triplicate. In addition, cell biomass was estimated by the OD600. Data processing and figure plotting were conducted by using Office Excel 2010, Origin V8.0, and SPSS V17.0 (SPSS, Inc. Chicago, IL). The differences between treatments were examined by t test, and those with P values of <0.05 were considered significant.

ACKNOWLEDGMENTS

This research was funded by the National Natural Science Foundation of China (91851202, 31600077, 51678163, U1701243, 41603074, and 41773132), the High-Level Leading Talent Introduction Program of GDAS (2016GDASRC-0208), a China Postdoctoral Science Foundation grant (2017M612622), the Guangdong Provincial Natural Science Foundation (2016A030306021), and the Science and Technology Planning Project of Guangzhou City (201707020021). The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Yinghua Cen from Guangdong Institute of Microbiology for suggestions during manuscript preparation and Ailin Zhang from Guangdong Institute of Microbiology for acquiring morphological data.

FOOTNOTES

    • Received 8 March 2019.
    • Accepted 29 May 2019.
    • Accepted manuscript posted online 7 June 2019.

  • Supplemental material for this article may be found at https://doi.org/10.1128/AEM.00550-19.

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KEYWORDS

Shewanella decolorationis S12
extracellular electron acceptor
filamentous shift
toxic organic pollutant
transcriptomic analysis

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