Surfing in the storm: how Paraburkholderia xenovorans thrives under stress during biodegradation of toxic aromatic compounds and other stressors

Abstract

The adaptive mechanisms of Burkholderiales during the catabolism of aromatic compounds and abiotic stress are crucial for their fitness and performance. The aims of this report are to review the bacterial adaptation mechanisms to aromatic compounds, oxidative stress, and environmental stressful conditions, focusing on the model aromatic-degrading Paraburkholderia xenovorans LB400, other Burkholderiales, and relevant degrading bacteria. These mechanisms include (i) the stress response during aromatic degradation, (ii) the oxidative stress response to aromatic compounds, (iii) the metabolic adaptation to oxidative stress, (iv) the osmoadaptation to saline stress, (v) the synthesis of siderophore during iron limitation, (vi) the proteostasis network, which plays a crucial role in cellular function maintenance, and (vii) the modification of cellular membranes, morphology, and bacterial lifestyle. Remarkably, we include, for the first time, novel genomic analyses on proteostasis networks, carbon metabolism modulation, and the synthesis of stress-related molecules in P. xenovorans. We analyzed these metabolic features in silico to gain insights into the adaptive strategies of P. xenovorans to challenging environmental conditions. Understanding how to enhance bacterial stress responses can lead to the selection of more robust strains capable of thriving in polluted environments, which is critical for improving biodegradation and bioremediation strategies.

Keywords: Paraburkholderia; adaptation; aromatic compound; chaperone; proteostasis; stress.

Figure 1.

Molecular oxidative stress responses of P. xenovorans during exposure to oxidizing agents and 4‐hydroxyphenylacetate. (A) Response to paraquat and H₂O₂. (B) Response during 4‐hydroxyphenylacetate (4-HPA) catabolism in the presence of FldX1. 4‐HPA enters the cell through a transmembrane protein and is catabolized via the homoprotocatechuate and homogentisate degradation pathways. The preferred route, homogentisate, generates ROS via the MhaAB enzyme during an enzymatic reaction facilitated by FldX1, which serves as an electron shuttle. This is reflected by a downregulation of the antioxidant response (antioxidant proteins and genes), enhancing bacterial fitness. The long‐chain flavodoxin FldX1 acts as an electron shuttle between sources of reducing power and metabolic pathways. Green and red arrows indicate the upregulation and downregulation of genes, proteins, or metabolites, respectively. Created in https://BioRender.com.

Figure 2.

(Chloro)biphenyl, benzoate, and catechol catabolic pathways and formation of toxic metabolic intermediates. The enzymes of the upper (chloro)biphenyl pathway are encoded by the bph locus of P. xenovorans LB400. Toxic intermediates are shown in gray: (chloro)2,3-dihydro-2,3-dihydroxybiphenyl and (chloro)2,3-dihydroxybiphenyl. Enzymes: BphA1A2A3A4, biphenyl-2,3-dioxygenase; BphB, 2,3-dihydro-2,3-dihydroxybiphenyl-2,3-dehydrogenase; BphC, cis-2,3-dihydrobiphenyl-2,3-diol dehydrogenase; BphD, 2-Hydroxy-6-oxo-6-phenylhexa-2,4-dienoate hydrolase; BphH, 2-hydroxypenta-2,4-dienoate hydratase; BphI, 4-hydroxy-2-oxovalerate aldolase; BphJ, acetaldehyde dehydrogenase; BenA, benzoate 1,2-dioxygenase α subunit; BenB, benzoate 1,2-dioxygenase ß subunit; BenC, benzoate 1,2-dioxygenase ferredoxin reductase component; BenD, benzoate diol dehydrogenase; DH, dehydrogenase; CatA, catechol 1,2 dioxygenase; and MCI, muconate cycloisomerase. Formation of ROS by BphA1A2A3A4 and BphC is indicated.

Figure 3.

Central carbon metabolism and metabolic pathways for stress-related molecules in P. xenovorans LB400. Pathways are shown for: (A) catabolism of aromatic compounds, and (B) metabolism of stress-related molecules. Intermediates in bold are anabolic precursors for the synthesis of stress-related molecules. 1, Citrate synthase (GltA1 and GltA2). 2, Aconitate hydratase (AcnA1 and AcnA2). 3, Isocitrate dehydrogenase (Idh1, Idh2, and Idh3). 4, 2-Ketoglutarate dehydrogenase (SucA). 5, Succinyl-CoA synthase (SucC). 6, Succinate dehydrogenase (SdhC). 7, Fumarate hydratase (FumCB). 8, Malate dehydrogenase (Mdh). 9, Isocitrate lyase (MtbI). 10, Malate synthase (AceB or GlcG). 11, Malate dehydrogenase (oxaloacetate decarboxylating) (MaeB1, MaeB2, and MaeB3). 12, Oxaloacetate decarboxylase (Oad). 13, Phosphoenolpyruvate (PEP) carboxykinase (PckG). 14, Fructose-1,6-biphosphate (FBP) aldolase (CbbA1, CbbA2, and CbbA3). 15, FBP 1,6-biphosphatase (Fbp1 and Fbp2). 16, Glucose-6-phosphate isomerase (Pgi). 17, Glucose-6-phosphate dehydrogenase (Zwf1, Zwf2, and Zwf3). 18, Phosphogluconolactonase (Pgl). 19, 6-Phosphogluconate dehydrogenase (Pgdh1 and Pgdh2). 20, 6-Phosphogluconate dehydratase (Edd). 21, 2-Keto-3-deoxy-6-phosphogluconate (KDPG) aldolase (Eda). 22, Proton translocating transhydrogenase (PntAB). 23, NADP-dependent ferredoxin oxidoreductase (Fpr). G3P, glycerate-3-phosphate. GAP, glyceraldehyde-3-phosphate. 6-PGL, 6-phosphogluconolactone. PhaZ, poly(3-hydroxybutyrate) (P(3HB)) depolymerase. PhaY, P(3HB) oligomer hydrolase. P(3HB), poly(3-hydroxybutyrate). (R)-3HB, (R)-3-hydroxybutyrate. NRPS, nonribosomal peptide synthase. SerA, d-3-phosphoglycerate dehydrogenase. SerC, phosphoserine aminotransferase. SerB, phosphoserine phosphatase. GdhA, l-glutamate dehydrogenase. ArgJ, arginine synthesis bifunctional protein. NagK, acetylglutamate kinase. ArgC, N-acetyl-γ-glutamylphosphate reductase. ArgD, acetylornithine aminotransferase. ArgG, argininosuccinate synthase. ArgH, argininosuccinate lyase. AdiA, Arginine decarboxylase. SpeB, agmatinase. MbaA, MbaB, nonribosomal peptide synthases (NRPS). MbaC, l-ornithine-5-monooxygenase (Vargas-Straube et al. 2016). PhaA, 2-ketothiolase. PhaB, (R)-3-hydroxybutyryl-CoA reductase. PhaC, polyhydroxyalkanoate (PHA) synthase (Urtuvia et al. 2018). PPO, PEP, pyruvate, oxaloacetate node. EMP, Embden–Meyerhof–Parnas. ED, Entner–Doudoroff. Non-OxPP, the nonoxidative branch of pentose–phosphate (PP) pathway. Ox-PP, oxidative branch of the PP pathway. NADPH formation is highlighted in the corresponding enzymatic reactions. Central metabolic pathways were evaluated in P. xenovorans LB400 through the BlastKO software of the Kyoto Encyclopedia of Genes and Genomes (KEGG) database. Enzymes were corroborated through the bidirectional best-hit approach using the BlastP tool against the Swiss-Prot curated database considering sequences identities >30% and alignment coverage >70% of the query sequence. Sequences with empirical evidence at the transcript or protein level were considered as references. Gene context was also evaluated with the KEGG genome database.

Figure 4.

Prediction of adaptive responses to saline stress in P. xenovorans LB400. Under high osmolarity conditions, compatible solutes accumulate at high concentrations within the cell. After controlling turgor imbalance, large conductance mechanosensitive channels (MscL) rapidly release edible solutes and larger molecular metabolites. The ATP-dependent K⁺ transporter (KdpFABC) actively drives potassium into the cell under low intracellular K⁺ concentration conditions, contributing to the regulation of cellular osmotic pressure. The ectoine/5-hydroxyectoine transporter (EhuABD) allows the uptake of these compatible solutes, which can be metabolized in the P. xenovorans cell as a carbon and energy source through the expression of the doeABDC genes, yielding l-aspartate as the final product, and acetate, 2-oxoglutarate, and glutamate as metabolic intermediates. The transporters for compatible solute precursors (ProU, ProP, and OusA) enter glycine–betaine and l-proline and their precursors, such as choline/betaine, into the cell, which are dehydrogenated by the enzymes BetA and BetB to form glycine-betaine. BetI is the negative transcriptional regulator of the betAB genes, activated by increased glycine–betaine osmoprotectant concentration. ProU directly imports the osmoprotectant. Trehalose plays a dual role in cellular regulation under saline stress, acting either as a compatible solute (trehalose) in response to osmotic pressure fluctuations or as a carbon and energy source when it is metabolized into glucose and glucose-6-phosphate. The identified determinants associated with compatible solutes in the genome of P. xenovorans are highlighted in bold. Protein sequences obtained from the NCBI (National Center for Biotechnology Information) platform were selected and cross-referenced with the Uniprot KB–Swiss Prot database to determine molecular determinants associated with salinity using experimentally validated sequences as a filter. The cutoff values for positive alignments were set at identity and coverage higher than 40% and 70%, respectively. Created in https://BioRender.com.

Figure 5.

Evolutionary relationships of the bacterium P. xenovorans with other relevant aromatic-degrading bacterial strains, and abundance of genes encoding chaperones and proteases in their genomes. (A) Phylogenetic tree of aromatic-degrading Burkholderiales and other bacteria. Evolutionary phylogenetic tree of bacterial 16S rRNA genes were constructed with MUSCLE alignment (Edgar 2004) and Maximum-likelihood clustering (1,000 bootstrap). (B) The genomes of P. xenovorans LB400, C. metallidurans CH34, B. cepacia ATCC 25416, P. putida (NBRC 1416 and KT2440), A. evansii KB740, R. jostii RHA1, S. enterica serovar Typhimurium LT2, and two strains of E. coli (O157:H7 Sakai and BW25113) were evaluated. The heatmap indicates the number of gene copies. Genome sequences were obtained from the National Center for Biotechnology Information (NCBI) database. The strains and genome accession codes are listed in Table S1. Assessment of classical cytoplasmic chaperones (TF, DnaK, DnaJ, DjlA, CbpA GrpE, GroEL, GroES, HtpG, ClpB, HscAB, and MsrAB), membrane and periplasmic chaperones (HtrA, FstH, SurA, Skp, YidC, Spy, and HdeAB), stress response holdase chaperones (Hsp33, Hsp20, Hsp31, SlyD, SlpA, CnoX, and RidA), and proteolytic systems (ClpX/ClpA/ClpC/ClpP/ClpY/ClpQ, Lon) were performed by sequence comparative NCBI Blast tools (https://blast.ncbi.nlm.nih.gov/Blast.cgi). Conserved domains/motifs and ATP-binding sites were confirmed using CDART (Conserved Domain Architecture Retrieval Tool) and CDD (Conserved Domains) tools from NCBI portal (https://www.ncbi.nlm.nih.gov/cdd).

Figure 6.

Schematic representation of the predicted proteostasis network in P. xenovorans. Newly synthesized proteins emerge from the ribosome and are assisted by trigger factor (TF) and chaperone systems such as DnaK/J, GrpE, and GroEL/ES for proper folding. Unfolded proteins are managed by a network of chaperones and proteases that mediate refolding or degradation. Numbers next to each factor correspond to the number of copies identified in the genome of P. xenovorans. Created in https://BioRender.com.

Figure 7.

Comprehensive stress response mechanisms of P. xenovorans to aromatic compounds and abiotic stressors. The diagram illustrates the metabolic, physiological, and morphological adaptations of P. xenovorans to chemicals (aromatic compounds, PCBs, and oxidizing compounds) and abiotic stressors (high salinity and nutrient scarcity). Adaptive metabolic responses include the increase of reductive power, synthesis/degradation of P(3HB), synthesis of siderophores. Adaptive physiological responses include the upregulation of foldases and holdases, and ROS-scavenging enzymes. Adaptive morphological responses include membrane and cell morphology changes, along with lifestyle changes (planktonic and biofilm). Genetic and metabolic engineering approaches highlight potential enhancements of bacterial stress tolerance, wherein antioxidants play a role in mitigating oxidative stress. Overcoming stress strategies play a key role in enhancing the biodegradation of pollutants and bacterial tolerance to abiotic stress. Created in https://BioRender.com.