Reserve Flux Capacity in the Pentose Phosphate Pathway Enables Escherichia coli’s Rapid Response to Oxidative Stress

Construction of Escherichia coli K‐12 in‐frame, single‐gene knockout mutants: the Keio collection

Baba, T., Ara, T., Hasegawa, M., Takai, Y., Okumura, Y., Baba, M., Datsenko, K. A., Tomita, M., Wanner, B. L., & Mori, H. (2006) · Molecular Systems Biology · doi:10.1038/msb4100050 · Cited by 7,858

Free radicals, antioxidants, and nutrition

Fang, Y.-Z., Yang, S., & Wu, G. (2002) · Nutrition · doi:10.1016/s0899-9007(02)00916-4 · Cited by 2,607

Invited review: Oxidation of biological systems: Oxidative stress phenomena, antioxidants, redox reactions, and methods for their quantification

Kohen, R., & Nyska, A. (2002) · Toxicologic Pathology · doi:10.1080/01926230290166724 · Cited by 2,178

The molecular mechanisms and physiological consequences of oxidative stress: lessons from a model bacterium

Imlay, J. A. (2013) · Nature Reviews Microbiology · doi:10.1038/nrmicro3032 · Cited by 1,749

Oxidant signals and oxidative stress

Finkel, T. (2003) · Current Opinion in Cell Biology · doi:10.1016/s0955-0674(03)00002-4 · Cited by 1,393

Complete set of ORF clones of Escherichia coli ASKA library (A Complete Set of E. coli K-12 ORF Archive): Unique Resources for Biological Research

Kitagawa, M., Ara, T., Arifuzzaman, M., Ioka-Nakamichi, T., Inamoto, E., Toyonaga, H., & Mori, H. (2006) · DNA Research · doi:10.1093/dnares/dsi012 · Cited by 1,388

Inhibition of Pyruvate Kinase M2 by Reactive Oxygen Species Contributes to Cellular Antioxidant Responses

Anastasiou, D., Poulogiannis, G., Asara, J. M., Boxer, M. B., Jiang, J., Shen, M., Bellinger, G., Sasaki, A. T., Locasale, J. W., Auld, D. S., et al. (2011) · Science · doi:10.1126/science.1211485 · Cited by 1,218

The world of protein acetylation

Drazic, A., Myklebust, L. M., Ree, R., & Arnesen, T. (2016) · Biochimica et Biophysica Acta (BBA) - Proteins and Proteomics · doi:10.1016/j.bbapap.2016.06.007 · Cited by 847

The quantitative and condition-dependent Escherichia coli proteome

Schmidt, A., Kochanowski, K., Vedelaar, S. R., Ahrné, E., Volkmer, B., Callipo, L., Knoops, K., Bauer, M., Aebersold, R. H., & Heinemann, M. (2015) · Nature Biotechnology · doi:10.1038/nbt.3418 · Cited by 759

Glucose‐6‐phosphate dehydrogenase, NADPH, and cell survival

Stanton, R. C. (2012) · IUBMB Life · doi:10.1002/iub.1017 · Cited by 706

The pentose phosphate pathway in health and disease

TeSlaa, T. A., Ralser, M., Fan, J., & Rabinowitz, J. D. (2023) · Nature Metabolism · doi:10.1038/s42255-023-00863-2 · Cited by 714

Reactive oxygen species in pathogen clearance: The killing mechanisms, the adaption response, and the side effects

Li, H., Zhou, X., Huang, Y., Liao, B., Cheng, L., & Ren, B. (2021) · Frontiers in Microbiology · doi:10.3389/fmicb.2020.622534 · Cited by 284

Switching to the cyclic pentose phosphate pathway powers the oxidative burst in activated neutrophils

Britt, E. C., Lika, J., Giese, M. A., Schoen, T. J., Seim, G. L., Huang, Z., Lee, P. Y., Huttenlocher, A., & Fan, J. (2022) · Nature Metabolism · doi:10.1038/s42255-022-00550-8 · Cited by 237

Reconfiguration of metabolic fluxes in Pseudomonas putida as a response to sub-lethal oxidative stress

Nikel, P. I., Fuhrer, T., Chavarría, M., Sánchez-Pascuala, A., Sauer, U., & de Lorenzo, V. (2021) · The ISME Journal · doi:10.1038/s41396-020-00884-9 · Cited by 156

The Role of Reactive Species on Innate Immunity

Andrés, C., Pérez de la Lastra, J., Juan, C., Plou, F., & Pérez-Lebeña, E. (2022) · Vaccines · doi:10.3390/vaccines10101735 · Cited by 142

The GAPDH redox switch safeguards reductive capacity and enables survival of stressed tumour cells

Talwar, D., Miller, C. G., Großmann, J. L., Szyrwiel, Ł., Schwecke, T., Demichev, V., Mikecin, A., Mayakonda, A., Lutsik, P., Veith, C., et al. (2023) · Nature Metabolism · doi:10.1038/s42255-023-00781-3 · Cited by 137

Mismatch-CRISPRi Reveals the Co-varying Expression-Fitness Relationships of Essential Genes in Escherichia coli and Bacillus subtilis

Hawkins, J. S., Silvis, M. R., Koo, B., Peters, J. M., Osadnik, H., Jost, M., Hearne, C. C., Weissman, J. S., Todor, H., & Gross, C. A. (2020) · Cell Systems · doi:10.1016/j.cels.2020.09.009 · Cited by 126

Awakening a latent carbon fixation cycle in Escherichia coli

Satanowski, A., Dronsella, B., Noor, Ε., Vögeli, B., He, H., Wichmann, P., Erb, T. J., Lindner, S. N., & Bar‐Even, A. (2020) · Nature Communications · doi:10.1038/s41467-020-19564-5 · Cited by 115

Allosteric Feedback Inhibition Enables Robust Amino Acid Biosynthesis in E. coli by Enforcing Enzyme Overabundance

Sander, T., Farke, N., Diehl, C., Kuntz, M. A., Glatter, T., & Link, H. (2019) · Cell Systems · doi:10.1016/j.cels.2018.12.005 · Cited by 94

Corrigendum: Reactive Oxygen Species in Pathogen Clearance: The Killing Mechanisms, the Adaption Response, and the Side Effects

Li, H., Zhou, X., Huang, Y., Liao, B., Cheng, L., & Ren, B. (2021) · Frontiers in Microbiology · doi:10.3389/fmicb.2021.685133 · Cited by 91

Modular (de)construction of complex bacterial phenotypes by CRISPR/nCas9-assisted, multiplex cytidine base-editing

Volke, D. C., Martino, R. A., Kozaeva, E., Smania, A. M., & Nikel, P. I. (2022) · Nature Communications · doi:10.1038/s41467-022-30780-z · Cited by 89

Multi-omics Analysis of CRISPRi-Knockdowns Identifies Mechanisms that Buffer Decreases of Enzymes in E. coli Metabolism

Donati, S., Kuntz, M. A., Pahl, V., Farke, N., Beuter, D., Glatter, T., Gomes‐Filho, J. V., Randau, L., Wang, C., & Link, H. (2020) · Cell Systems · doi:10.1016/j.cels.2020.10.011 · Cited by 83

High‐Z‐Sensitized Radiotherapy Synergizes with the Intervention of the Pentose Phosphate Pathway for In Situ Tumor Vaccination

Wang, Y., Chen, J., Duan, R., Gu, R., Wang, W., Wu, J., Lian, H., Hu, Y., & Yuan, A. (2022) · Advanced Materials · doi:10.1002/adma.202109726 · Cited by 83

Sod1 integrates oxygen availability to redox regulate NADPH production and the thiol redoxome

Montllor-Albalate, C., Kim, H.-J., Thompson, A. E., Jonke, A. P., Torres, M. P., & Reddi, A. R. (2021) · Proceedings of the National Academy of Sciences · doi:10.1073/pnas.2023328119 · Cited by 84

Investigation of the methylerythritol 4-phosphate pathway for microbial terpenoid production through metabolic control analysis

Volke, D. C., Rohwer, J. M., Fischer, R., & Jennewein, S. (2019) · Microbial Cell Factories · doi:10.1186/s12934-019-1235-5 · Cited by 85

Redox rebalance against genetic perturbations and modulation of central carbon metabolism by the oxidative stress regulation

Shimizu, K., & Matsuoka, Y. (2019) · Biotechnology Advances · doi:10.1016/j.biotechadv.2019.107441 · Cited by 85

Redox Systems Biology: Harnessing the Sentinels of the Cysteine Redoxome

Held, J. M. (2019) · Antioxidants and Redox Signaling · doi:10.1089/ars.2019.7725 · Cited by 78

Increase in antioxidant capacity associated with the successful subclone of hypervirulent carbapenem-resistant Klebsiella pneumoniae ST11-KL64

Wang, R., Zhang, A., Sun, S., Yin, G., Wu, X., Ding, Q., Wang, Q., Chen, F., Wang, S., van Dorp, L., et al. (2024) · Nature Communications · doi:10.1038/s41467-023-44351-3 · Cited by 79

Metabolic Modelling as a Framework for Metabolomics Data Integration and Analysis

Volkova, S., Matos, M. R. A., Mattanovich, M., & de Mas, I. M. (2020) · Metabolites · doi:10.3390/metabo10080303 · Cited by 77

Impact of ROS-Induced Damage of TCA Cycle Enzymes on Metabolism and Virulence of Salmonella enterica serovar Typhimurium

Noster, J., Persicke, M., Chao, T., Krone, L., Heppner, B., Hensel, M., & Hansmeier, N. (2019) · Frontiers in Microbiology · doi:10.3389/fmicb.2019.00762 · Cited by 74

Protein allocation and utilization in the versatile chemolithoautotroph Cupriavidus necator

Jahn, M., Crang, N., Janasch, M., Hober, A., Forsström, B., Kimler, K., Mattausch, A., Chen, Q., Asplund‐Samuelsson, J., & Hudson, E. P. (2021) · eLife · doi:10.7554/elife.69019 · Cited by 77

The enzymes of the oxidative phase of the pentose phosphate pathway as targets of reactive species: consequences for NADPH production

Fuentes-Lemus, E., Reyes, J. S., Figueroa, J. D., Davies, M. J., & López‐Alarcón, C. (2023) · Biochemical Society Transactions · doi:10.1042/bst20231027 · Cited by 74

Building kinetic models for metabolic engineering

Foster, C., Wang, L., Dinh, H. V., Suthers, P. F., & Maranas, C. D. (2020) · Current Opinion in Biotechnology · doi:10.1016/j.copbio.2020.11.010 · Cited by 70

Reserve Flux Capacity in the Pentose Phosphate Pathway by NADPH Binding Is Conserved across Kingdoms

Christodoulou, D., Kuehne, A., Estermann, A., Fuhrer, T., Lang, P., & Sauer, U. (2019) · iScience · doi:10.1016/j.isci.2019.08.047 · Cited by 69

Rewiring Central Carbon Metabolism Ensures Increased Provision of Acetyl-CoA and NADPH Required for 3-OH-Propionic Acid Production

Qin, N., Li, L., Ji, X., Li, X., Zhang, Y., Larsson, C., Chen, Y., Nielsen, J., & Liu, Z. (2020) · ACS Synthetic Biology · doi:10.1021/acssynbio.0c00264 · Cited by 60

Facile synthesis of copper oxide nanoparticles (CuONPs) using green method to promote photocatalytic and biocidal applications

Rehman, K. U., Khan, A. U., Tahir, K., Nazir, S., Albalawi, K. M., Hassan, H. M. A., Alabbad, E. A., Refat, M. S. E.-D., Al‐Shehri, H. S., & Aldawsari, A. M. (2022) · Journal of Molecular Liquids · doi:10.1016/j.molliq.2022.119453 · Cited by 56

Increased CO2 fixation enables high carbon-yield production of 3-hydroxypropionic acid in yeast

Qin, N., Li, L., Wan, X., Ji, X., Chen, Y., Li, C., Liu, P., Zhang, Y., Yang, W., Jiang, J., et al. (2024) · Nature Communications · doi:10.1038/s41467-024-45557-9 · Cited by 60

Metabolism of non-growing bacteria

Lempp, M., Lubrano, P., Bange, G., & Link, H. (2020) · Biological Chemistry · doi:10.1515/hsz-2020-0201 · Cited by 53

Increased energy demand from anabolic-catabolic processes drives β-lactam antibiotic lethality

Lobritz, M. A., Andrews, I. W., Braff, D., Porter, C. B. M., Gutierrez, A., Furuta, Y., Cortes, L. B. G., Ferrante, T. C., Bening, S. C., Wong, F. J., et al. (2022) · Cell chemical biology · doi:10.1016/j.chembiol.2021.12.010 · Cited by 55

Extensive regulation of enzyme activity by phosphorylation in Escherichia coli

Schastnaya, E., Nakic, Z. R., Gruber, C., Doubleday, P. F., Krishnan, A., Johns, N. I., Park, J., Wang, H. H., & Sauer, U. (2021) · Nature Communications · doi:10.1038/s41467-021-25988-4 · Cited by 50

Large-scale kinetic metabolic models of Pseudomonas putida KT2440 for consistent design of metabolic engineering strategies

Tokić, M. S., Mišković, L., & Vassily Hatzimanikatis, A. (2020) · Biotechnology for Biofuels · doi:10.1186/s13068-020-1665-7 · Cited by 48

Persistence of Intracellular Bacterial Pathogens—With a Focus on the Metabolic Perspective

Eisenreich, W., Rudel, T., Heesemann, J., & Goebel, W. (2021) · Frontiers in Cellular and Infection Microbiology · doi:10.3389/fcimb.2020.615450 · Cited by 49

Cofactor Specificity of Glucose-6-Phosphate Dehydrogenase Isozymes in Pseudomonas putida Reveals a General Principle Underlying Glycolytic Strategies in Bacteria

Volke, D. C., Olavarría, K., & Nikel, P. I. (2021) · mSystems · doi:10.1128/msystems.00014-21 · Cited by 42

Transomics data-driven, ensemble kinetic modeling for system-level understanding and engineering of the cyanobacteria central metabolism

Nishiguchi, H., Hiasa, N., Uebayashi, K., Liao, J. C., Shimizu, H., & Matsuda, F. (2019) · Metabolic Engineering · doi:10.1016/j.ymben.2019.01.004 · Cited by 43

Escherichia coli metabolism under short-term repetitive substrate dynamics: adaptation and trade-offs

Vasilakou, E., van Loosdrecht, M. C. M., & Wahl, A. (2020) · Microbial Cell Factories · doi:10.1186/s12934-020-01379-0 · Cited by 43

Genomic diversity across the Rickettsia and ‘Candidatus Megaira’ genera and proposal of genus status for the Torix group

Davison, H., Pilgrim, J., Wybouw, N., Parker, J., Pirro, S., Hunter-Barnett, S., Campbell, P. M., Blow, F., Darby, A. C., Hurst, G. D. D., et al. (2022) · Nature Communications · doi:10.1038/s41467-022-30385-6 · Cited by 45

CRISPR interference screens reveal growth–robustness tradeoffs inSynechocystissp. PCC 6803 across growth conditions

Miao, R., Jahn, M., Shabestary, K., Peltier, G., & Hudson, E. P. (2023) · The Plant Cell · doi:10.1093/plcell/koad208 · Cited by 47

Microbial physiological engineering increases the efficiency of microbial cell factories

Liu, H., Qi, Y., Zhou, P., Ye, C., Gao, C., Chen, X., & Liu, L. (2021) · Critical Reviews in Biotechnology · doi:10.1080/07388551.2020.1856770 · Cited by 41

Ribulose-1,5-Bisphosphate Carboxylase/Oxygenase (RubisCO) Is Essential for Growth of the Methanotroph Methylococcus capsulatus Strain Bath

Henard, C. A., Wu, C., Xiong, W., Henard, J. M., Davidheiser‐Kroll, B., Orata, F. D., & Guarnieri, M. T. (2021) · Applied and Environmental Microbiology · doi:10.1128/aem.00881-21 · Cited by 40

Hostile Takeover: How Viruses Reprogram Prokaryotic Metabolism

Jacobson, T. B., Callaghan, M. M., & Amador‐Noguez, D. (2021) · Annual Review of Microbiology · doi:10.1146/annurev-micro-060621-043448 · Cited by 40

Relationship between myo-inositol synthesis and carbohydrate metabolism changes in Mozambique tilapia (Oreochromis mossambicus) under acute hypersaline stress

Zhu, J., Wang, X., Bu, X., Wang, C., Pan, J., Li, E., Shi, Q., Zhang, M., Qin, J. G., & Chen, L. (2020) · Aquaculture · doi:10.1016/j.aquaculture.2020.736005 · Cited by 39

Recent advances in constraint and machine learning-based metabolic modeling by leveraging stoichiometric balances, thermodynamic feasibility and kinetic law formalisms

Suthers, P. F., Foster, C., Sarkar, D., Wang, L., & Maranas, C. D. (2020) · Metabolic Engineering · doi:10.1016/j.ymben.2020.11.013 · Cited by 39

Quantitative modeling of pentose phosphate pathway response to oxidative stress reveals a cooperative regulatory strategy

Hurbain, J., Thommen, Q., Anquez, F., & Pfeuty, B. (2022) · iScience · doi:10.1016/j.isci.2022.104681 · Cited by 42

Functional decomposition of metabolism allows a system-level quantification of fluxes and protein allocation towards specific metabolic functions

Mori, M., Cheng, C., Taylor, B. R., Okano, H., & Hwa, T. (2023) · Nature Communications · doi:10.1038/s41467-023-39724-7 · Cited by 36

ROS-induced stress promotes enrichment and emergence of antibiotic resistance in conventional activated sludge processes

Manna, B., Zhou, X., & Singhal, N. (2025) · Water Research · doi:10.1016/j.watres.2025.123366 · Cited by 37

Milligrams to kilograms: making microbes work at scale

Cordell, W. T., Avolio, G., Takors, R., & Pfleger, B. F. (2023) · Trends in biotechnology · doi:10.1016/j.tibtech.2023.05.002 · Cited by 36

Dynamic Metabolomics for Engineering Biology: Accelerating Learning Cycles for Bioproduction

Vavricka, C. J., Hasunuma, T., & Kondo, A. (2019) · Trends in biotechnology · doi:10.1016/j.tibtech.2019.07.009 · Cited by 35

Expanding the search for small-molecule antibacterials by multidimensional profiling

Ortmayr, K., de la Cruz, R., & Zampieri, M. (2022) · Nature Chemical Biology · doi:10.1038/s41589-022-01040-4 · Cited by 35

Why is manganese so valuable to bacterial pathogens?

Čapek, J., & Večerek, B. (2023) · Frontiers in Cellular and Infection Microbiology · doi:10.3389/fcimb.2023.943390 · Cited by 32

Global insights into intermediate metabolites: Signaling, metabolic divergence and stress response modulation in plants

Samsami, H., & Maali-Amiri, R. (2024) · Plant Physiology and Biochemistry · doi:10.1016/j.plaphy.2024.108862 · Cited by 32

Eat in or Take out? Metabolism of Intracellular Salmonella enterica

Kehl, A., Noster, J., & Hensel, M. (2020) · Trends in Microbiology · doi:10.1016/j.tim.2020.03.005 · Cited by 30

Microbes with higher metabolic independence are enriched in human gut microbiomes under stressPreprint

Veseli, I., Chen, Y. T., Schechter, M. S., Vanni, C., Fogarty, E. C., Watson, A. R., Jabrì, B., Blekhman, R., Willis, A. D., Yu, M., et al. (2023) · eLife · doi:10.7554/elife.89862 · Cited by 33

From Escherichia coli mutant 13C labeling data to a core kinetic model: A kinetic model parameterization pipeline

Foster, C., Gopalakrishnan, S., Antoniewicz, M. R., & Maranas, C. D. (2019) · PLoS Computational Biology · doi:10.1371/journal.pcbi.1007319 · Cited by 29

Kinetic modeling of metabolism: Present and future

Islam, M. M., Schroeder, W. L., & Saha, R. (2021) · Current Opinion in Systems Biology · doi:10.1016/j.coisb.2021.04.003 · Cited by 28

Elucidating yeast glycolytic dynamics at steady state growth and glucose pulses through kinetic metabolic modeling

Lao‐Martil, D., Schmitz, J. P. J., Teusink, B., & van Riel, N. A. W. (2023) · Metabolic Engineering · doi:10.1016/j.ymben.2023.03.005 · Cited by 27

Metabolomics and Microbial Metabolism: Toward a Systematic Understanding

Holbrook‐Smith, D., Trouillon, J., & Sauer, U. (2023) · Annual Review of Biophysics · doi:10.1146/annurev-biophys-030722-021957 · Cited by 29

Dark accumulation of downstream glycolytic intermediates initiates robust photosynthesis in cyanobacteria

Tanaka, K., Shirai, T., Vavricka, C. J., Matsuda, M., Kondo, A., & Hasunuma, T. (2022) · PLANT PHYSIOLOGY · doi:10.1093/plphys/kiac602 · Cited by 27

Inside the phagosome: A bacterial perspective

Hampton, M. B., & Dickerhof, N. (2023) · Immunological Reviews · doi:10.1111/imr.13182 · Cited by 28

Fundamental Differences in Inactivation Mechanisms of Escherichia coli O157:H7 Between Chlorine Dioxide and Sodium Hypochlorite

Bridges, D. F., Lacombe, A., & Wu, V. C. H. (2022) · Frontiers in Microbiology · doi:10.3389/fmicb.2022.923964 · Cited by 26

Time-resolved, deuterium-based fluxomics uncovers the hierarchy and dynamics of sugar processing by Pseudomonas putida

Volke, D. C., Gurdo, N., Milanesi, R., & Nikel, P. I. (2023) · Metabolic Engineering · doi:10.1016/j.ymben.2023.07.004 · Cited by 25

Redefining the Clostridioides difficile σ B Regulon: σ B Activates Genes Involved in Detoxifying Radicals That Can Result from the Exposure to Antimicrobials and Hydrogen Peroxide

Boekhoud, I. M., Michel, A., Corver, J., Jahn, D., & Smits, W. K. (2020) · mSphere · doi:10.1128/msphere.00728-20 · Cited by 25

Metabolic plasticity enables lifestyle transitions of Porphyromonas gingivalis

Moradali, M. F., & Davey, M. E. (2021) · npj Biofilms and Microbiomes · doi:10.1038/s41522-021-00217-4 · Cited by 26

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Homolak, J., Perhoč, A. B., Knezović, A., Kodvanj, I., Virag, D., Barilar, J. O., Riederer, P. F., & Šalković‐Petrišić, M. (2021) · Molecular Nutrition & Food Research · doi:10.1002/mnfr.202100400 · Cited by 25

A critical bioenergetic switch is regulated by IGF2 during murine cartilage development

Hollander, J. M., Li, L., Rawal, M., Wang, S. K., Shu, Y., Zhang, M., Nielsen, H. C., Rosen, C. J., & Zeng, L. (2022) · Communications Biology · doi:10.1038/s42003-022-04156-4 · Cited by 24

The antenna-like domain of the cyanobacterial ferrochelatase can bind chlorophyll and carotenoids in an energy-dissipative configuration

Pazderník, M., Mareš, J., Pilný, J., & Sobotka, R. (2019) · Journal of Biological Chemistry · doi:10.1074/jbc.ra119.008434 · Cited by 24

Design of microbial catalysts for two-stage processes

Shabestary, K., Klamt, S., Link, H., Mahadevan, R., Steuer, R., & Hudson, E. P. (2024) · Nature Reviews Bioengineering · doi:10.1038/s44222-024-00225-x · Cited by 23

Role of Premycofactocin Synthase in Growth, Microaerophilic Adaptation, and Metabolism of Mycobacterium tuberculosis

Gopinath, K., Kaiser, P., Constant, P., Abed, U. A., Schmid, M., Frese, C. K., Brinkmann, V., Daffé, M., & Kaufmann, S. H. E. (2021) · mBio · doi:10.1128/mbio.01665-21 · Cited by 24

Lactic acid fermentation: A maladaptive mechanism and an evolutionary throwback boosting cancer drug resistance

Gnocchi, D., Sabbà, C., & Mazzocca, A. (2023) · Biochimie · doi:10.1016/j.biochi.2023.01.005 · Cited by 23

Microbes with higher metabolic independence are enriched in human gut microbiomes under stress

Veseli, I., Chen, Y. T., Schechter, M. S., Vanni, C., Fogarty, E. C., Watson, A. R., Jabrì, B., Blekhman, R., Willis, A. D., Yu, M., et al. (2025) · eLife · doi:10.1101/2023.05.10.540289 · Cited by 29

Engineering of Biosynthesis Pathway and NADPH Supply for Improved L-5-Methyltetrahydrofolate Production by Lactococcus lactis

Lu, C., Liu, Y., Li, J., Liu, L., & Du, G. (2020) · Journal of Microbiology and Biotechnology · doi:10.4014/jmb.1910.10069 · Cited by 22

Laboratory evolution, transcriptomics, and modeling reveal mechanisms of paraquat tolerance

Rychel, K., Tan, J., Patel, A., Lamoureux, C. R., Hefner, Y., Szubin, R., Johnsen, J., Mohamed, E. T., Phaneuf, P. V., Anand, A., et al. (2023) · Cell Reports · doi:10.1016/j.celrep.2023.113105 · Cited by 23

Yeast increases glycolytic flux to support higher growth rates accompanied by decreased metabolite regulation and lower protein phosphorylation

Chen, M., Xie, T., Li, H., Zhuang, Y., Xia, J., & Nielsen, J. (2023) · Proceedings of the National Academy of Sciences · doi:10.1073/pnas.2302779120 · Cited by 23

Ionizing Radiation-induced Proteomic Oxidation in Escherichia coli

Bruckbauer, S. T., Minkoff, B. B., Yu, D., Cryns, V. L., Cox, M. M., & Sussman, M. R. (2020) · Molecular & Cellular Proteomics · doi:10.1074/mcp.ra120.002092 · Cited by 23

Analogous Metabolic Decoupling in Pseudomonas putida and Comamonas testosteroni Implies Energetic Bypass to Facilitate Gluconeogenic Growth

Wilkes, R. A., Waldbauer, J., & Aristilde, L. (2021) · mBio · doi:10.1128/mbio.03259-21 · Cited by 21

Identifying metabolic features and engineering targets for productivity improvement in CHO cells by integrated transcriptomics and genome-scale metabolic model

Huang, Z., & Yoon, S. (2020) · Biochemical Engineering Journal · doi:10.1016/j.bej.2020.107624 · Cited by 20

Enzymes of glycolysis and the pentose phosphate pathway as targets of oxidants: Role of redox reactions on the carbohydrate catabolism

Fuentes-Lemus, E., Usgame, K., Fierro, A., & López‐Alarcón, C. (2025) · Redox Biochemistry and Chemistry · doi:10.1016/j.rbc.2025.100049 · Cited by 18

Dissolved iron released from nanoscale zero-valent iron (nZVI) activates the defense system in bacterium Pseudomonas putida, leading to high tolerance to oxidative stress

Yeap, C. S. Y., Nguyen, N. H. A., Špánek, R., Too, C. C., Beneš, V., Provazník, J., Černík, M., & Ševců, A. (2022) · Journal of Hazardous Materials · doi:10.1016/j.jhazmat.2022.129627 · Cited by 18

Growth-rate-dependent and nutrient-specific gene expression resource allocation in fission yeast

Kleijn, I. T., Martínez-Segura, A., Bertaux, F., Saint, M., Kramer, H., Shahrezaei, V., & Marguerat, S. (2022) · Life Science Alliance · doi:10.26508/lsa.202101223 · Cited by 16

Dynamics in redox metabolism, from stoichiometry towards kinetics

Verhagen, K. J. A., van Gulik, W. M., & Wahl, A. (2020) · Current Opinion in Biotechnology · doi:10.1016/j.copbio.2020.01.002 · Cited by 18

Beyond Resistance: Tolerance and Resilience of Bacteria to Photodynamic and Oxidative Stress

Rapacka-Zdończyk, A. (2025) · International Journal of Molecular Sciences · doi:10.3390/ijms26188908 · Cited by 19

Role of FruR transcriptional regulator in virulence of Listeria monocytogenes and identification of its regulon

Abdelhamed, H., Ramachandran, R., Narayanan, L., Islam, S., Ozan, O., Freitag, N. E., & Lawrence, M. L. (2022) · PLoS ONE · doi:10.1371/journal.pone.0274005 · Cited by 16

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