Historically, appreciation for the roles of resource gradients in biology has fluctuated inversely to the popularity of genetic mechanisms. Nevertheless, in microbiology specifically, widespread recognition of the multicellular lifestyle has recently brought new emphasis to the importance of resource gradients. Most microorganisms grow in assemblages such as biofilms or spatially constrained communities with gradients that influence, and are influenced by, metabolism. In this Review, we discuss examples of gradient formation and physiological differentiation in microbial assemblages growing in diverse settings. We highlight consequences of physiological heterogeneity in microbial assemblages, including division of labour and increased resistance to stress. Our impressions of microbial behaviour in various ecosystems are not complete without complementary maps of the chemical and physical geographies that influence cellular activities. A holistic view, incorporating these geographies and the genetically encoded functions that operate within them, will be essential for understanding microbial assemblages in their many roles and potential applications.
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Lyons, N. A. & Kolter, R. On the evolution of bacterial multicellularity. Curr. Opin. Microbiol. 24, 21–28 (2015).
Caldwell, D. E. & Hirsch, P. Growth of microorganisms in two-dimensional steady-state diffusion gradients. Can. J. Microbiol. 19, 53–58 (1973).
Lappin-Scott, H. M. in Microbial biosystems: new frontiers: proceedings of the 8th International Symposium on Microbial Ecology Vol. 1 (eds Bell, C., Brylinski, M. & Johnson-Green, P. C.) 1–6 (Atlantic Canada Society for Microbial Ecology, 1999).
Babcsányi, I., Meite, F. & Imfeld, G. Biogeochemical gradients and microbial communities in Winogradsky columns established with polluted wetland sediments. FEMS Microbiol. Ecol. 93, fix089 (2017).
Child, C. M. Patterns and Problems of Development [by] C.M. Child Vol. 820 (Univ. of Chicago Press, 1941).
Blackstone, N. W. Charles Manning Child (1869–1954): the past, present, and future of metabolic signaling. J. Exp. Zool. B Mol. Dev. Evol. 306, 1–7 (2006).
Gurdon, J. B. & Bourillot, P. Y. Morphogen gradient interpretation. Nature 413, 797–803 (2001).
Fankhauser, C. & Christie, J. M. Plant phototropic growth. Curr. Biol. 25, R384–R389 (2015).
Moor, A. E. & Itzkovitz, S. Spatial transcriptomics: paving the way for tissue-level systems biology. Curr. Opin. Biotechnol. 46, 126–133 (2017).
Niethammer, P. Wound redox gradients revisited. Semin. Cell Dev. Biol. 80, 13–16 (2018).
Krejci, A. & Tennessen, J. M. Metabolism in time and space — exploring the frontier of developmental biology. Development 144, 3193–3198 (2017).
Flemming, H.-C. et al. Biofilms: an emergent form of bacterial life. Nat. Rev. Microbiol. 14, 563–575 (2016).
Flemming, H.-C. et al. Who put the film in biofilm? The migration of a term from wastewater engineering to medicine and beyond. NPJ Biofilms Microbiomes 7, 10 (2021).
Dragoš, A. & Kovács, Á. T. The peculiar functions of the bacterial extracellular matrix. Trends Microbiol. 25, 257–266 (2017).
Flemming, H.-C. & Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 8, 623–633 (2010).
Rasmussen, K. & Lewandowski, Z. Microelectrode measurements of local mass transport rates in heterogeneous biofilms. Biotechnol. Bioeng. 59, 302–309 (1998).
Costerton, J. W., Stewart, P. S. & Greenberg, E. P. Bacterial biofilms: a common cause of persistent infections. Science 284, 1318–1322 (1999).
Stewart, P. S. Diffusion in biofilms. J. Bacteriol. 185, 1485–1491 (2003).
Stewart, P. S. & Franklin, M. J. Physiological heterogeneity in biofilms. Nat. Rev. Microbiol. 6, 199–210 (2008).
Nealson, K. H. & Conrad, P. G. Life: past, present and future. Philos. Trans. R. Soc. Lond. B Biol. Sci. 354, 1923–1939 (1999).
Giri, S., Waschina, S., Kaleta, C. & Kost, C. Defining division of labor in microbial communities. J. Mol. Biol. 431, 4712–4731 (2019).
Evans, C. R., Kempes, C. P., Price-Whelan, A. & Dietrich, L. E. P. Metabolic heterogeneity and cross-feeding in bacterial multicellular systems. Trends Microbiol. 28, 732–743 (2020).
Douglas, A. E. The microbial exometabolome: ecological resource and architect of microbial communities. Philos. Trans. R. Soc. Lond. B Biol. Sci. 375, 20190250 (2020).
Fritts, R. K., McCully, A. L. & McKinlay, J. B. Extracellular metabolism sets the table for microbial cross-feeding. Microbiol. Mol. Biol. Rev. 85, e00135-20 (2021).
Chen, J. & Strous, M. Denitrification and aerobic respiration, hybrid electron transport chains and co-evolution. Biochim. Biophys. Acta 1827, 136–144 (2013).
Konhauser, K. O. Introduction to Geomicrobiology (Wiley, 2009).
Fenchel, T. & Finlay, B. Oxygen and the spatial structure of microbial communities. Biol. Rev. Camb. Philos. Soc. 83, 553–569 (2008).
Bishop, P. L. & Yu, T. A microelectrode study of redox potential change in biofilms. Water Sci. Technol. 39, 179–185 (1999).
Jo, J., Cortez, K. L., Cornell, W. C., Price-Whelan, A. & Dietrich, L. E. An orphan cbb3-type cytochrome oxidase subunit supports Pseudomonas aeruginosa biofilm growth and virulence. eLife 6, e30205 (2017). This paper shows that phenazine production affects the redox gradient within P. aeruginosa biofilms and that mutations that alter ETC composition affect this gradient, implicating specific ETC complexes in phenazine reduction.
Dal, Co,A., van Vliet, S. & Ackermann, M. Emergent microscale gradients give rise to metabolic cross-feeding and antibiotic tolerance in clonal bacterial populations. Philos. Trans. R. Soc. Lond. B Biol. Sci. 374, 20190080 (2019).
Cole, J. A., Kohler, L., Hedhli, J. & Luthey-Schulten, Z. Spatially-resolved metabolic cooperativity within dense bacterial colonies. BMC Syst. Biol. 9, 15 (2015). This paper describes a model in which acetate exchange supports metabolic cross-feeding in E. coli biofilms.
Wolfsberg, E., Long, C. P. & Antoniewicz, M. R. Metabolism in dense microbial colonies: 13C metabolic flux analysis of E. coli grown on agar identifies two distinct cell populations with acetate cross-feeding. Metab. Eng. 49, 242–247 (2018). Together with Cole et al. (2015), this study tests the model of acetate exchange supporting metabolic cross-feeding in E. coli biofilms.
Díaz-Pascual, F. et al. Spatial alanine metabolism determines local growth dynamics of Escherichia coli colonies. eLife 10, e70794 (2021).
Decho, A. W., Norman, R. S. & Visscher, P. T. Quorum sensing in natural environments: emerging views from microbial mats. Trends Microbiol. 18, 73–80 (2010).
Wolpert, L. in Current Topics in Developmental Biology Vol. 117 (ed. Wassarman, P. M.) 597–608 (Academic, 2016).
Redfield, R. J. Is quorum sensing a side effect of diffusion sensing? Trends Microbiol. 10, 365–370 (2002).
Hense, B. A. et al. Does efficiency sensing unify diffusion and quorum sensing? Nat. Rev. Microbiol. 5, 230–239 (2007).
West, S. A., Winzer, K., Gardner, A. & Diggle, S. P. Quorum sensing and the confusion about diffusion. Trends Microbiol. 20, 586–594 (2012).
Lee, J.-H., Wood, T. K. & Lee, J. Roles of indole as an interspecies and interkingdom signaling molecule. Trends Microbiol. 23, 707–718 (2015).
Zarkan, A., Liu, J., Matuszewska, M., Gaimster, H. & Summers, D. K. Local and universal action: the paradoxes of indole signalling in bacteria. Trends Microbiol. 28, 566–577 (2020).
Kumar, A. & Sperandio, V. Indole signaling at the host–microbiota–pathogen interface. mBio 10, e01031-19 (2019). This study uses measurements of indole concentrations in the gut lumen and gut epithelial cells to infer the presence of an indole gradient and shows that the indole concentration influences virulence in gut-colonizing bacteria.
Comolli, J. C. & Donohue, T. J. Differences in two Pseudomonas aeruginosa cbb3 cytochrome oxidases. Mol. Microbiol. 51, 1193–1203 (2004).
Kawakami, T., Kuroki, M., Ishii, M., Igarashi, Y. & Arai, H. Differential expression of multiple terminal oxidases for aerobic respiration in Pseudomonas aeruginosa. Environ. Microbiol. 12, 1399–1412 (2010).
Arai, H. et al. Enzymatic characterization and in vivo function of five terminal oxidases in Pseudomonas aeruginosa. J. Bacteriol. 196, 4206–4215 (2014).
Dietrich, L. E. P. et al. Bacterial community morphogenesis is intimately linked to the intracellular redox state. J. Bacteriol. 195, 1371–1380 (2013).
Eschbach, M. et al. Long-term anaerobic survival of the opportunistic pathogen Pseudomonas aeruginosa via pyruvate fermentation. J. Bacteriol. 186, 4596–4604 (2004).
Glasser, N. R., Kern, S. E. & Newman, D. K. Phenazine redox cycling enhances anaerobic survival in Pseudomonas aeruginosa by facilitating generation of ATP and a proton-motive force. Mol. Microbiol. 92, 399–412 (2014).
Jo, J., Price-Whelan, A., Cornell, W. C. & Dietrich, L. E. P. Interdependency of respiratory metabolism and phenazine-associated physiology in Pseudomonas aeruginosa PA14. J. Bacteriol. 202, e00700-19 (2020).
Lin, Y.-C. et al. Phenazines regulate Nap-dependent denitrification in Pseudomonas aeruginosa biofilms. J. Bacteriol. 200, e00031-18 (2018).
Schiessl, K. T. et al. Phenazine production promotes antibiotic tolerance and metabolic heterogeneity in Pseudomonas aeruginosa biofilms. Nat. Commun. 10, 762 (2019). This paper describes the effects of phenazine production on metabolic heterogeneity across depth and provides evidence for cross-feeding between fermentative and respiratory subpopulations in P. aeruginosa biofilms, and also shows that phenazines differentially influence the responses of biofilm subpopulations to antibiotic exposure.
Greenhagen, B. T. et al. Crystal structure of the pyocyanin biosynthetic protein PhzS. Biochemistry 47, 5281–5289 (2008).
Sakhtah, H. et al. The Pseudomonas aeruginosa efflux pump MexGHI-OpmD transports a natural phenazine that controls gene expression and biofilm development. Proc. Natl Acad. Sci. USA 113, E3538–E3547 (2016).
Bellin, D. L. et al. Electrochemical camera chip for simultaneous imaging of multiple metabolites in biofilms. Nat. Commun. 7, 10535 (2016).
Lambden, P. R. & Guest, J. R. Mutants of Escherichia coli K12 unable to use fumarate as an anaerobic electron acceptor. Microbiology 97, 145–160 (1976).
Kiley, P. J. & Beinert, H. Oxygen sensing by the global regulator, FNR: the role of the iron–sulfur cluster. FEMS Microbiol. Rev. 22, 341–352 (1998).
Myers, K. S. et al. Genome-scale analysis of Escherichia coli FNR reveals complex features of transcription factor binding. PLoS Genet. 9, e1003565 (2013).
Crofts, A. A. et al. Enterotoxigenic E. coli virulence gene regulation in human infections. Proc. Natl Acad. Sci. USA 115, E8968–E8976 (2018).
Lin, Y.-C., Cornell, W. C., Jo, J., Price-Whelan, A. & Dietrich, L. E. P. The Pseudomonas aeruginosa complement of lactate dehydrogenases enables use of d- and l-lactate and metabolic cross-feeding. mBio 9, e00961-18 (2018).
Okegbe, C. et al. Electron-shuttling antibiotics structure bacterial communities by modulating cellular levels of c-di-GMP. Proc. Natl Acad. Sci. USA 114, E5236–E5245 (2017).
Fong, G.-H. & Takeda, K. Role and regulation of prolyl hydroxylase domain proteins. Cell Death Differ. 15, 635–641 (2008).
Beebout, C. J. et al. Respiratory heterogeneity shapes biofilm formation and host colonization in uropathogenic Escherichia coli. mBio 10, e02400-18 (2019).
Arnaouteli, S. et al. Bifunctionality of a biofilm matrix protein controlled by redox state. Proc. Natl Acad. Sci. USA 114, E6184–E6191 (2017).
Worlitzsch, D. et al. Effects of reduced mucus oxygen concentration in airway Pseudomonas infections of cystic fibrosis patients. J. Clin. Invest. 109, 317–325 (2002).
Kolpen, M. et al. Nitrous oxide production in sputum from cystic fibrosis patients with chronic Pseudomonas aeruginosa lung infection. PLoS ONE 9, e84353 (2014).
Cowley, E. S., Kopf, S. H., LaRiviere, A., Ziebis, W. & Newman, D. K. Pediatric cystic fibrosis sputum can be chemically dynamic, anoxic, and extremely reduced due to hydrogen sulfide formation. mBio 6, e00767 (2015). This study shows that sputum from persons with cystic fibrosis contains steep chemical gradients, describes models for the distribution of O2 in the cystic fibrosis airway and provides a comprehensive overview of metabolisms that could support microbial growth and survival in this setting.
Quinn, R. A. et al. Biogeochemical forces shape the composition and physiology of polymicrobial communities in the cystic fibrosis lung. mBio 5, e00956–13 (2014).
Cendra, M. D. M., Blanco-Cabra, N., Pedraz, L. & Torrents, E. Optimal environmental and culture conditions allow the in vitro coexistence of Pseudomonas aeruginosa and Staphylococcus aureus in stable biofilms. Sci. Rep. 9, 16284 (2019).
DePas, W. H. et al. Exposing the three-dimensional biogeography and metabolic states of pathogens in cystic fibrosis sputum via hydrogel embedding, clearing, and rRNA labeling. mBio 7, e00796-16 (2016). This study reveals the extreme structural heterogeneity in sputum from persons with cystic fibrosis, and applies an in situ hybridization chain reaction in combination with standard fluorescence in situ hybridization to obtain insight into the growth rates of microorganisms present in sputum samples.
Flynn, J. M., Niccum, D., Dunitz, J. M. & Hunter, R. C. Evidence and role for bacterial mucin degradation in cystic fibrosis airway disease. PLoS Pathog. 12, e1005846 (2016).
Ley, R. E. et al. Unexpected diversity and complexity of the Guerrero Negro hypersaline microbial mat. Appl. Environ. Microbiol. 72, 3685–3695 (2006).
Prieto-Barajas, C. M., Valencia-Cantero, E. & Santoyo, G. Microbial mat ecosystems: structure types, functional diversity, and biotechnological application. Electron. J. Biotechnol. 31, 48–56 (2018).
Dupraz, C. et al. Processes of carbonate precipitation in modern microbial mats. Earth-Sci. Rev. 96, 141–162 (2009).
Hoehler, T. M., Bebout, B. M. & Des Marais, D. J. The role of microbial mats in the production of reduced gases on the early Earth. Nature 412, 324–327 (2001).
Bosak, T., Knoll, A. H. & Petroff, A. P. The meaning of stromatolites. Annu. Rev. Earth Planet. Sci. 41, 21–44 (2013).
Wagner, K., Besemer, K., Burns, N. R., Battin, T. J. & Bengtsson, M. M. Light availability affects stream biofilm bacterial community composition and function, but not diversity. Environ. Microbiol. 17, 5036–5047 (2015).
Mobberley, J. M. et al. Organismal and spatial partitioning of energy and macronutrient transformations within a hypersaline mat. FEMS Microbiol. Ecol. 93, fix028 (2017). This study comprehensively describes physicochemical and biological features of a highly stratified cyanobacterial mat.
Airs, R. L. & Keely, B. J. A high resolution study of the chlorophyll and bacteriochlorophyll pigment distributions in a calcite/gypsum microbial mat. Org. Geochem. 34, 539–551 (2003).
Ohkubo, S. & Miyashita, H. A niche for cyanobacteria producing chlorophyll f within a microbial mat. ISME J. 11, 2368–2378 (2017).
Abed, R. M. M., Kohls, K., Leloup, J. & de Beer, D. Abundance and diversity of aerobic heterotrophic microorganisms and their interaction with cyanobacteria in the oxic layer of an intertidal hypersaline cyanobacterial mat. FEMS Microbiol. Ecol. 94, fix183 (2018).
Cypionka, H., Widdel, F. & Pfennig, N. Survival of sulfate-reducing bacteria after oxygen stress, and growth in sulfate-free oxygen-sulfide gradients. FEMS Microbiol. Lett. 31, 39–45 (1985).
Canfield, D. E. & Des Marais, D. J. Aerobic sulfate reduction in microbial mats. Science 251, 1471–1473 (1991).
Santegoeds, C. M., Ferdelman, T. G., Muyzer, G. & de Beer, D. Structural and functional dynamics of sulfate-reducing populations in bacterial biofilms. Appl. Environ. Microbiol. 64, 3731–3739 (1998).
Fike, D. A., Gammon, C. L., Ziebis, W. & Orphan, V. J. Micron-scale mapping of sulfur cycling across the oxycline of a cyanobacterial mat: a paired nanoSIMS and CARD-FISH approach. ISME J. 2, 749–759 (2008).
Lee, J. Z. et al. Fermentation couples Chloroflexi and sulfate-reducing bacteria to Cyanobacteria in hypersaline microbial mats. Front. Microbiol. 5, 61 (2014).
Harris, J. K. et al. Phylogenetic stratigraphy in the Guerrero Negro hypersaline microbial mat. ISME J. 7, 50–60 (2013).
Ruff, S. E. et al. Global dispersion and local diversification of the methane seep microbiome. Proc. Natl Acad. Sci. USA 112, 4015–4020 (2015).
Drewniak, L. et al. Physiological and metagenomic analyses of microbial mats involved in self-purification of mine waters contaminated with heavy metals. Front. Microbiol. 7, 1252 (2016).
Lin, S. et al. Biological sulfur oxidation in wastewater treatment: a review of emerging opportunities. Water Res. 143, 399–415 (2018).
Dworkin, M. Sergei Winogradsky: a founder of modern microbiology and the first microbial ecologist. FEMS Microbiol. Rev. 36, 364–379 (2012).
Jørgensen, B. B. & Revsbech, N. P. Colorless sulfur bacteria, Beggiatoa spp. and Thiovulum spp., in O2 and H2S microgradients. Appl. Environ. Microbiol. 45, 1261–1270 (1983).
Nelson, D. C., Wirsen, C. O. & Jannasch, H. W. Characterization of large, autotrophic Beggiatoa spp. abundant at hydrothermal vents of the guaymas basin. Appl. Environ. Microbiol. 55, 2909–2917 (1989).
Kreutzmann, A.-C. & Schulz-Vogt, H. N. Oxidation of molecular hydrogen by a chemolithoautotrophic Beggiatoa strain. Appl. Environ. Microbiol. 82, 2527–2536 (2016).
Pasulka, A. et al. SSU-rRNA gene sequencing survey of benthic microbial eukaryotes from Guaymas Basin hydrothermal vent. J. Eukaryot. Microbiol. 66, 637–653 (2019).
Treude, T. et al. Consumption of methane and CO2 by methanotrophic microbial mats from gas seeps of the anoxic Black Sea. Appl. Environ. Microbiol. 73, 2271–2283 (2007).
Vlamakis, H., Aguilar, C., Losick, R. & Kolter, R. Control of cell fate by the formation of an architecturally complex bacterial community. Genes Dev. 22, 945–953 (2008).
Kolodkin-Gal, I. et al. Respiration control of multicellularity in Bacillus subtilis by a complex of the cytochrome chain with a membrane-embedded histidine kinase. Genes Dev. 27, 887–899 (2013).
Serra, D. O. & Hengge, R. Stress responses go three dimensional — the spatial order of physiological differentiation in bacterial macrocolony biofilms. Environ. Microbiol. 16, 1455–1471 (2014).
Okegbe, C., Price-Whelan, A. & Dietrich, L. E. P. Redox-driven regulation of microbial community morphogenesis. Curr. Opin. Microbiol. 18, 39–45 (2014).
Zacharia, V. M. et al. Genetic network architecture and environmental cues drive spatial organization of phenotypic division of labor in Streptomyces coelicolor. mBio 12, e00794-21 (2021).
Váchová, L. & Palková, Z. How structured yeast multicellular communities live, age and die? FEMS Yeast Res. 18, 1–9 (2018).
Hengge, R. Linking bacterial growth, survival, and multicellularity — small signaling molecules as triggers and drivers. Curr. Opin. Microbiol. 55, 57–66 (2020).
Klauck, G., Serra, D. O., Possling, A. & Hengge, R. Spatial organization of different sigma factor activities and c-di-GMP signalling within the three-dimensional landscape of a bacterial biofilm. Open. Biol. 8, 180066 (2018). This study uses fluorescent reporters of gene expression to characterize the growth states of subpopulations in E. coli biofilms, revealing an unexpectedly complex pattern across depth.
Serra, D. O., Klauck, G. & Hengge, R. Vertical stratification of matrix production is essential for physical integrity and architecture of macrocolony biofilms of Escherichia coli. Environ. Microbiol. 17, 5073–5088 (2015).
Serra, D. O., Richter, A. M. & Hengge, R. Cellulose as an architectural element in spatially structured Escherichia coli biofilms. J. Bacteriol. 195, 5540–5554 (2013).
Pérez-Osorio, A. C., Williamson, K. S. & Franklin, M. J. Heterogeneous rpoS and rhlR mRNA levels and 16S rRNA/rDNA (rRNA gene) ratios within Pseudomonas aeruginosa biofilms, sampled by laser capture microdissection. J. Bacteriol. 192, 2991–3000 (2010).
Williamson, K. S. et al. Heterogeneity in Pseudomonas aeruginosa biofilms includes expression of ribosome hibernation factors in the antibiotic-tolerant subpopulation and hypoxia-induced stress response in the metabolically active population. J. Bacteriol. 194, 2062–2073 (2012).
Santoro, C., Arbizzani, C., Erable, B. & Ieropoulos, I. Microbial fuel cells: from fundamentals to applications. A review. J. Power Sources 356, 225–244 (2017).
Walter, X. A. et al. From the lab to the field: self-stratifying microbial fuel cells stacks directly powering lights. Appl. Energy 277, 115514 (2020).
Chadwick, G. L., Jiménez Otero, F., Gralnick, J. A., Bond, D. R. & Orphan, V. J. NanoSIMS imaging reveals metabolic stratification within current-producing biofilms. Proc. Natl Acad. Sci. USA 116, 20716–20724 (2019).
Lebedev, N., Strycharz-Glaven, S. M. & Tender, L. M. Spatially resolved confocal resonant Raman microscopic analysis of anode-grown Geobacter sulfurreducens biofilms. ChemPhysChem 15, 320–327 (2014).
Renslow, R. et al. Metabolic spatial variability in electrode-respiring Geobacter sulfurreducens biofilms. Energy Environ. Sci. 6, 1827–1836 (2013).
Beauregard, P. B., Chai, Y., Vlamakis, H., Losick, R. & Kolter, R. Bacillus subtilis biofilm induction by plant polysaccharides. Proc. Natl Acad. Sci. USA 110, E1621–E1630 (2013).
Pereira, F. C. & Berry, D. Microbial nutrient niches in the gut. Environ. Microbiol. 19, 1366–1378 (2017).
Rudrappa, T., Biedrzycki, M. L. & Bais, H. P. Causes and consequences of plant-associated biofilms. FEMS Microbiol. Ecol. 64, 153–166 (2008).
Stacy, A., Fleming, D., Lamont, R. J., Rumbaugh, K. P. & Whiteley, M. A commensal bacterium promotes virulence of an opportunistic pathogen via cross-respiration. mBio 7, e00782-16 (2016).
Davis, K. M., Mohammadi, S. & Isberg, R. R. Community behavior and spatial regulation within a bacterial microcolony in deep tissue sites serves to protect against host attack. Cell Host Microbe 17, 21–31 (2015). In this study, the authors use gene expression as a read-out for NO and detect a gradient in Y. tuberculosis microcolonies growing within the spleens of a murine host; they use mutants in both the mouse host background and the background of the bacterial pathogen to show that NO produced by host cells is consumed by bacteria at the periphery of the microcolony, thereby creating the gradient.
von Ohle, C. et al. Real-time microsensor measurement of local metabolic activities in ex vivo dental biofilms exposed to sucrose and treated with chlorhexidine. Appl. Environ. Microbiol. 76, 2326–2334 (2010).
Mark Welch, J. L., Dewhirst, F. E. & Borisy, G. G. Biogeography of the oral microbiome: the site-specialist hypothesis. Annu. Rev. Microbiol. 73, 335–358 (2019).
Xiao, J. et al. Biofilm three-dimensional architecture influences in situ pH distribution pattern on the human enamel surface. Int. J. Oral. Sci. 9, 74–79 (2017).
Bowen, W. H., Burne, R. A., Wu, H. & Koo, H. Oral biofilms: pathogens, matrix, and polymicrobial interactions in microenvironments. Trends Microbiol. 26, 229–242 (2018).
Kim, D. et al. Spatial mapping of polymicrobial communities reveals a precise biogeography associated with human dental caries. Proc. Natl Acad. Sci. USA 117, 12375–12386 (2020). This paper reports several findings important for understanding how biofilm communities can promote development of dental caries (tooth decay), and identifies a specific type of biofilm architecture in which community stratification supports pH gradient formation and contributes to the robustness of the pathogenic population.
Donaldson, G. P., Lee, S. M. & Mazmanian, S. K. Gut biogeography of the bacterial microbiota. Nat. Rev. Microbiol. 14, 20–32 (2016).
Zheng, L., Kelly, C. J. & Colgan, S. P. Physiologic hypoxia and oxygen homeostasis in the healthy intestine. A Review in the Theme: Cellular Responses to Hypoxia. Am. J. Physiol. Cell Physiol. 309, C350–C360 (2015).
De Weirdt, R. & Van de Wiele, T. Micromanagement in the gut: microenvironmental factors govern colon mucosal biofilm structure and functionality. NPJ Biofilms Microbiomes 1, 15026 (2015).
Rivera-Chávez, F., Lopez, C. A. & Bäumler, A. J. Oxygen as a driver of gut dysbiosis. Free Radic. Biol. Med. 105, 93–101 (2017).
Brauner, A., Fridman, O., Gefen, O. & Balaban, N. Q. Distinguishing between resistance, tolerance and persistence to antibiotic treatment. Nat. Rev. Microbiol. 14, 320–330 (2016).
Anderson, G. G. & O’Toole, G. A. Innate and induced resistance mechanisms of bacterial biofilms. Curr. Top. Microbiol. Immunol. 322, 85–105 (2008).
Høiby, N. et al. The clinical impact of bacterial biofilms. Int. J. Oral. Sci. 3, 55–65 (2011).
Rocha-Granados, M. C., Zenick, B., Englander, H. E. & Mok, W. W. K. The social network: impact of host and microbial interactions on bacterial antibiotic tolerance and persistence. Cell. Signal. 75, 109750 (2020).
Königs, A. M., Flemming, H.-C. & Wingender, J. Nanosilver induces a non-culturable but metabolically active state in Pseudomonas aeruginosa. Front. Microbiol. 6, 395 (2015).
Borriello, G. et al. Oxygen limitation contributes to antibiotic tolerance of Pseudomonas aeruginosa in biofilms. Antimicrob. Agents Chemother. 48, 2659–2664 (2004).
Balaban, N. Q. et al. Definitions and guidelines for research on antibiotic persistence. Nat. Rev. Microbiol. 17, 441–448 (2019).
Yan, J. & Bassler, B. L. Surviving as a community: antibiotic tolerance and persistence in bacterial biofilms. Cell Host Microbe 26, 15–21 (2019).
Jennings, L. K. et al. Pseudomonas aeruginosa aggregates in cystic fibrosis sputum produce exopolysaccharides that likely impede current therapies. Cell Rep. 34, 108782 (2021).
Koo, H., Allan, R. N., Howlin, R. P., Stoodley, P. & Hall-Stoodley, L. Targeting microbial biofilms: current and prospective therapeutic strategies. Nat. Rev. Microbiol. 15, 740–755 (2017).
Hall, C. W. & Mah, T.-F. Molecular mechanisms of biofilm-based antibiotic resistance and tolerance in pathogenic bacteria. FEMS Microbiol. Rev. 41, 276–301 (2017).
Andersson, D. I. & Hughes, D. Microbiological effects of sublethal levels of antibiotics. Nat. Rev. Microbiol. 12, 465–478 (2014).
Høiby, N., Bjarnsholt, T., Givskov, M., Molin, S. & Ciofu, O. Antibiotic resistance of bacterial biofilms. Int. J. Antimicrob. Agents 35, 322–332 (2010).
Hermsen, R., Deris, J. B. & Hwa, T. On the rapidity of antibiotic resistance evolution facilitated by a concentration gradient. Proc. Natl Acad. Sci. USA 109, 10775–10780 (2012).
Zhang, Q. et al. Acceleration of emergence of bacterial antibiotic resistance in connected microenvironments. Science 333, 1764–1767 (2011).
Kowalski, C. H., Morelli, K. A., Schultz, D., Nadell, C. D. & Cramer, R. A. Fungal biofilm architecture produces hypoxic microenvironments that drive antifungal resistance. Proc. Natl Acad. Sci. USA 117, 22473–22483 (2020).
Quinn, R. A. et al. Niche partitioning of a pathogenic microbiome driven by chemical gradients. Sci. Adv. 4, eaau1908 (2018).
Cantor, M. D., van den Tempel, T., Hansen, T. K. & Ardö, Y. in Cheese 4th edn Ch. 37 (eds McSweeney, P. L. H., Fox, P. F., Cotter, P. D. & Everett, D. W.) 929–954 (Academic, 2017).
Ong, L. et al. in Cheese 4th edn Ch. 33 (eds McSweeney, P. L. H., Fox, P. F., Cotter, P. D. & Everett, D. W.) 829–863 (Academic, 2017).
Abraham, S., Cachon, R., Colas, B., Feron, G. & De Coninck, J. Eh and pH gradients in Camembert cheese during ripening: measurements using microelectrodes and correlations with texture. Int. Dairy. J. 17, 954–960 (2007).
Monds, R. D. & O’Toole, G. A. The developmental model of microbial biofilms: ten years of a paradigm up for review. Trends Microbiol. 17, 73–87 (2009).
Jorth, P., Spero, M. A., Livingston, J. & Newman, D. K. Quantitative visualization of gene expression in mucoid and nonmucoid Pseudomonas aeruginosa aggregates reveals localized peak expression of alginate in the hypoxic zone. mBio 10, e02622-19 (2019).
Serra, D. O. & Hengge, R. A c-di-GMP-based switch controls local heterogeneity of extracellular matrix synthesis which is crucial for integrity and morphogenesis of Escherichia coli macrocolony biofilms. J. Mol. Biol. 431, 4775–4793 (2019).
Hansen, J. M., Jones, D. P. & Harris, C. The redox theory of development. Antioxid. Redox Signal. 32, 715–740 (2020).
Vander Heiden, M. G. & DeBerardinis, R. J. Understanding the intersections between metabolism and cancer biology. Cell 168, 657–669 (2017).
Miyazawa, H. & Aulehla, A. Revisiting the role of metabolism during development. Development 145, dev131110 (2018).
Carmeliet, P. et al. Role of HIF-1α in hypoxia-mediated apoptosis, cell proliferation and tumour angiogenesis. Nature 394, 485–490 (1998).
Stamatelos, S. K., Bhargava, A., Kim, E., Popel, A. S. & Pathak, A. P. Tumor ensemble-based modeling and visualization of emergent angiogenic heterogeneity in breast cancer. Sci. Rep. 9, 5276 (2019).
Hobson-Gutierrez, S. A. & Carmona-Fontaine, C. The metabolic axis of macrophage and immune cell polarization. Dis. Model. Mech. 11, dmm034462 (2018).
Lyssiotis, C. A. & Kimmelman, A. C. Metabolic interactions in the tumor microenvironment. Trends Cell Biol. 27, 863–875 (2017).
Gifford, R. M. & Evans, L. T. Photosynthesis, carbon partitioning, and yield. Annu. Rev. Plant. Physiol. 32, 485–509 (1981).
Weits, D. A., van Dongen, J. T. & Licausi, F. Molecular oxygen as a signaling component in plant development. N. Phytol. 229, 24–35 (2021).
Work in the Dietrich laboratory is supported by National Institutes of Health (NIH)/National Institute of Allergy and Infectious Diseases (NIAID) grant R01AI103369.
The authors declare no competing interests.
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Interacting assemblages of multiple species in a common location.
Assemblages of microbial cells encased in a self-produced matrix.
An adhesive, extracellular material composed of polysaccharides, DNA and/or proteins produced by microbial cells in biofilms.
The exchange of metabolites between cells.
- Oxygenic phototrophs
Organisms that use light as a source of energy and water as an electron donor, thereby producing molecular O2.
A signalling molecule that forms a concentration gradient across a developing organism and elicits concentration-dependent responses.
The stepwise reduction of nitrate to nitrogen gas, carried out by integral membrane and membrane-associated proteins. Some steps in denitrification pathways contribute to the proton motive force, and thereby the synthesis of ATP.
Redox-active compounds produced by Pseudomonas aeruginosa and other bacteria that can shuttle electrons between cells and distant oxidants.
Using an organic or inorganic chemical (as opposed to light) as a source of energy for metabolism.
Organisms that use an organic carbon source in metabolism.
- Intertidal mats
Biofilm communities dominated by cyanobacteria that form within the tidal range on seashores.
Using an inorganic carbon source in metabolism.
A (usually heritable) trait conferred, for example, by dedicated proteins that efflux or degrade an antimicrobial compound, allowing the producing organism to grow at antimicrobial concentrations that would otherwise inhibit growth.
A (heritable or non-heritable) trait exhibited by the majority of cells in a population that confers survival during transient exposure to an antimicrobial.
A non-heritable property exhibited by a minority of cells in a population that survives exposure to an antimicrobial; that is, tolerance that is restricted to a subpopulation.
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Jo, J., Price-Whelan, A. & Dietrich, L.E.P. Gradients and consequences of heterogeneity in biofilms. Nat Rev Microbiol 20, 593–607 (2022). https://doi.org/10.1038/s41579-022-00692-2
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