Formation of the biofilm matrix induces a unique environment for bacteria that allows the dynamic biofilm mode of life. Biofilms, and the resulting lifestyle, are built in specific, defined steps, producing a bacterial community that is heterogeneous in space and time.
Extracellular polymeric substances (EPS) immobilize biofilm cells, keeping them in long-term close proximity and, thus, allowing intense interactions to occur, including cell–cell communication, horizontal gene transfer and the formation of synergistic microconsortia.
Owing to the retention of extracellular enzymes in the matrix, a versatile external digestive system is generated: dissolved and particulate nutrients imported through the water phase of the matrix can be sequestered, accumulated and utilized. The matrix acts as an ultimate recycling yard, keeping all the components of lysed cells available, including DNA, and possibly therefore serving as a large genetic archive. Gradient formation creates a wide range of very different habitats, contributing to biodiversity in biofilms.
The matrix protects organisms in the biofilm from desiccation, biocides, antibiotics, heavy metals, ultraviolet radiation, host immune defences and many protozoan grazers.
Eventually, EPS can serve as a nutrient source, but — as for many other structural polymers in biology — some EPS components are only slowly biodegradable. The vast variety of EPS components means that their complete degradation requires a wide range of enzymes.
Ecologically, competition and cooperation in the confined space of the EPS matrix, and competition for the limited nutrients in particular, lead to constant adaptation of population fitness.
The microorganisms in biofilms live in a self-produced matrix of hydrated extracellular polymeric substances (EPS) that form their immediate environment. EPS are mainly polysaccharides, proteins, nucleic acids and lipids; they provide the mechanical stability of biofilms, mediate their adhesion to surfaces and form a cohesive, three-dimensional polymer network that interconnects and transiently immobilizes biofilm cells. In addition, the biofilm matrix acts as an external digestive system by keeping extracellular enzymes close to the cells, enabling them to metabolize dissolved, colloidal and solid biopolymers. Here we describe the functions, properties and constituents of the EPS matrix that make biofilms the most successful forms of life on earth.
This is a preview of subscription content, access via your institution
Open Access articles citing this article.
BMC Oral Health Open Access 30 September 2023
“Sharing the matrix” – a cooperative strategy for survival in Salmonella enterica serovar Typhimurium
BMC Microbiology Open Access 23 August 2023
Future Journal of Pharmaceutical Sciences Open Access 05 June 2023
Subscribe to this journal
Receive 12 print issues and online access
$209.00 per year
only $17.42 per issue
Rent or buy this article
Prices vary by article type
Prices may be subject to local taxes which are calculated during checkout
Wingender, J., Neu, T. & Flemming, H.-C. in Microbial Extracellular Polymeric Substances (eds Wingender, J., Neu, T. & Flemming, H.-C.) 1–19 (Springer, Heidelberg, 1999).
Karatan, E. & Watnik, P. Signals, regulatory networks, and materials that build and break bacterial biofilms. Microbiol. Mol. Biol. Rev. 73, 310–347 (2009). An excellent paper on aspects of the regulation of the biofilm matrix.
Xavier, J. B. & Foster, K. R. Cooperation and conflict in microbial biofilms. Proc. Natl Acad. Sci. USA 104, 876–881 (2007). An important and inspiring discussion on microbial interactions, including the role of the matrix.
Flemming, H. C., Neu, T. R. & Wozniak, D. The EPS matrix: the house of biofilm cells. J. Bacteriol. 189, 7945–7947 (2007).
Allison, D. G., Sutherland, I. W. & Neu, T. R. in Biofilm Communities: Order from Chaos? (eds McBain, A. et al.) 381–387 (BioLine, Cardiff, 1998).
Zogaj, X., Nimtz, M., Rohde, M., Bokranz, W. & Römling, U. The multicellular morphotypes of Salmonella typhimurium and Escherichia coli produce cellulose as the second component of the extracellular matrix. Mol. Microbiol. 39, 1452–1463 (2001).
Schooling, S. R. & Beveridge, T. J. Membrane vesicles: an overlooked component of the matrices of biofilms. J. Bacteriol. 188, 5945–5957 (2006).
Decho, A. W. Microbial exopolymer secretions in ocean environments: their role(s) in food webs and marine processes. Oceanogr. Mar. Biol. Annu. Rev. 28, 73–153 (1990). A classic review of microbial interactions, including the role of EPS.
Decho, A. W. Microbial biofilms in intertidal systems: an overview. Cont. Shelf Res. 20, 1257–1273 (2000).
Flemming, H.-C. & Wingender, J. in Encyclopedia of Environmental Microbiology (ed. Bitton, G.) 1223–1231 (Wiley, New York, 2002).
Decho, A. W., Visscher, P. T. & Reid, R. P. Production and cycling of natural microbial exopolymers (EPS) within a marine stromatolite. Paleogeogr. Paleoclimatol. Paleoecol. 219, 71–86 (2005).
Ortega-Morales, B. O. et al. Characterization of extracellular polymers synthesized by tropical intertidal biofilm bacteria. J. Appl. Microbiol. 102, 254–264 (2006).
Sutherland, I. W. The biofilm matrix – an immobilized but dynamic microbial environment. Trends Microbiol. 9, 222–227 (2001).
De Beer, D., Stoodley, P., Roe, F. & Lewandowski, Z. Effects of biofilm structures on oxygen distribution and mass transport. Biotechnol. Bioeng. 43, 1131–1138 (1994). A classic paper introducing microelectrode techniques to biofilm research.
Lawrence, J. R., Swerhone, G. D. W., Kuhlicke, U. & Neu, T. R. In situ evidence for microdomains in the polymer matrix of bacterial microcolonies. Can. J. Microbiol. 53, 450–458 (2007). The use of lectin-staining analysis for characterizing target structures in the EPS matrix.
Wagner, M., Ivleva, N. P., Haisch, C., Niessner, R. & Horn, H. Combined use of confocal laser scanning microscopy (CLSM) and Raman microscopy (RM): investigations on EPS-matrix. Water Res. 43, 63–76 (2009).
Watnik, P. I. & Kolter, R. Steps in the development of a Vibrio cholerae El Tor biofilm. Mol. Microbiol. 34, 586–595 (1999).
Danese, P. N., Pratt, L. A. & Kolter, R. Exopolysaccharide production is required for development of Escherichia coli K-12 biofilm architecture. J. Bacteriol. 182, 3593–3596 (2000).
Branda, S. S., Chu, F., Kearns, D. B., Losick, R. & Kolter, R. A major protein component of the Bacillus subtilis biofilm matrix. Mol. Microbiol. 59, 1229–1238 (2006).
Lux, R., Li, Y., Lu, A. & Shi, W. Detailed three-dimensional analysis of structural features of Myxococcus xanthus fruiting bodies using confocal laser scanning microscopy. Biofilms 1, 293–303 (2004).
Tielen, P., Strathmann, M., Jaeger, K. E., Flemming, H.-C. & Wingender, J. Alginate acetylation influences initial surface colonization by mucoid Pseudomonas aeruginosa. Microbiol. Res. 160, 165–176 (2005).
Franklin, M. J. & Ohman, D. E. Identification of algF in the alginate biosynthetic gene cluster of Pseudomonas aeruginosa which is required for alginate acetylation. J. Bacteriol. 175, 5057–5065 (1993).
Wozniak, D. et al. Alginate is not a significant component of the extracellular polysaccharide matrix of PA14 and PAO1 Pseudomonas aeruginosa biofilms. Proc. Natl Acad. Sci. USA 100, 7907–7912 (2003).
Körstgens, V., Flemming, H.-C., Wingender, J. & Borchard, W. Influence of calcium ions on the mechanical properties of a model biofilm of mucoid Pseudomonas aeruginosa. Water Sci. Technol. 43, 49–57 (2001).
Nielsen, P. H. & Jahn, A. in Microbial Extracellular Polymeric Substances (eds Wingender, J., Neu, T. & Flemming, H.-C.) 49–72 (Springer, Heidelberg, 1999).
Tapia, J. M. et al. Extraction of extracellular polymeric substances from the acidophilic bacterium Acidophilium. Water Sci. Technol. 59, 1959–1967 (2009).
Brown, M. J. & Lester, J. N. Comparison of bacterial extracellular polymer extraction methods. Appl. Environ. Microbiol. 40, 179–185 (1980).
Frølund, B., Palmgren, R., Keiding, K. & Nielsen, P.-H. Extraction of extracellular polymers from activated sludge using a cation exchange resin. Water Res. 30, 1749–1758 (1996). The original description of one of the most frequently used and most successful methods of isolating EPS.
Wingender. J., Strathmann, M., Rode, A., Leis, A. & Flemming, H.-C. Isolation and biochemical characterization of extracellular polymeric substances from Pseudomonas aeruginosa. Methods Enzymol. 336, 302–314 (2001).
Römling, U. Molecular biology of cellulose production in bacteria. Res. Microbiol. 153, 205–212 (2002).
Ude, S., Arnold, D. L., Moon, C. D., Timms-Wilson, T. & Spiers, A. J. Biofilm formation and cellulose expression among diverse environmental Pseudomonas isolates. Environ. Microbiol. 8, 1997–2011 (2006).
Wang, X. et al. Impact of biofilm matrix components on interaction of commensal Escherichia coli with the gastrointestinal cell line HT-29. Cell. Mol. Life Sci. 63, 2352–2363 (2007).
Sutherland, I. W. in Comprehensive Glycoscience Vol. 2 (ed. Kamerling, J. P.) 521–558 (Elsevier, Doordrecht, 2007). The best and most comprehensive overview of the polysaccharide moiety of EPS.
Götz, F. Staphylococcus and biofilms. Mol. Microbiol. 43, 1367–1378 (2002).
Jefferson, K. K. in Bacterial Polysaccharides. Current Innovations and Future Trends (ed. Ullrich, M.) 175–186 (Caister Academic, Norfolk, UK, 2009).
Vaningelgem, F. et al. Biodiversity of exopolysaccharides produced by Streptococcus thermophilus strains is reflected in their production and their molecular and functional characteristics. Appl. Environ. Microbiol. 70, 900–912 (2004).
Ryder, C., Byrd, M., Wozniak, D. J. Role of exopolysaccharides in Pseudomonas aeruginosa biofilm development. Curr. Opin. Microbiol. 10, 644–648 (2007).
Byrd, M. S. et al. Genetic and biochemical analyses of the Pseudomonas aeruginosa Psl exopolysaccharide reveal overlapping roles for polysaccharide synthesis enzymes in Psl and LPS production. Mol. Microbiol. 73, 622–638 (2009).
Ma, L. et al. Assembly and development of the Pseudomonas aeruginosa biofilm matrix. PLoS Pathog. 5, e1000354 (2009).
Skillman, L., Sutherland, I. W. & Jonse, M. V. The role of exopolysaccharides in dual species biofilm development. J. Appl. Microbiol. 85, S13–S18 (1999).
Conrad, A. et al. Fatty acid lipid fractions in extracellular polymeric substances of activated sludge flocs. Lipids 38, 1093–1105 (2003).
Jahn, A. & Nielsen, P. H. Cell biomass and exopolymer composition in sewer biofilms. Water Sci. Technol. 37, 17–24 (1998).
Wingender, J., Jaeger, K.-E. & Flemming, H.-C. in Microbial Extracellular Polymeric Substances (eds Wingender, J., Neu, T. & Flemming, H.-C.) 231–251 (Springer, Heidelberg, 1999).
Shimao, M. Biodegradation of plastics. Curr. Opin. Biotechnol. 12, 242–247 (2001).
Busalmen, J. P., Vázquez, M. & de Sánchez, S. R. New evidences on the catalase mechanism of microbial corrosion. Electrochim. Acta 47, 1857–1865 (2002).
Wingender, J. & Jaeger, K.-E. in Encyclopedia of Environmental Microbiology (ed. Bitton, G.) 1207–1223 (Wiley, New York, 2002).
Mayer, C. et al. The role of intermolecular interactions studies on model systems for bacterial biofilms. Int. J. Biol. Macromol. 26, 3–16 (1999).
Zhang, X. & Bishop, P. Biodegradability of biofilm extracellular polymeric substances. Chemosphere 50, 63–69 (2003).
Russell, R. R. B. in Bacterial Polysaccharides. Current Innovations and Future Trends (ed. Ullrich, M.) 143–156 (Caister Academic, Norfolk, UK, 2009).
Laue, H. et al. Contribution of alginate and levan production to biofilm formation by Pseudomonas syringae. Microbiology 152, 2909–2918 (2006).
Sauer, K. et al. Characterization of nutrient-induced dispersion in Pseudomonas aeruginosa PAO1 biofilm. J. Bacteriol. 186, 7312–7326 (2004).
Gjermansen, M. Ragas, P., Sternberg, C., Molin, S. & Tolker-Nielsen, T. Characterization of starvation-induced dispersion in Pseudomonas putida biofilms. Environ. Microbiol. 7, 894–906 (2005).
Kaplan, J. B. et al. Genes involved in the synthesis and degradation of matrix polysaccharide in Actinobacillus actinomycetemcomitans and Actinobacillus pleuropneumoniae biofilms. J. Bacteriol. 186, 8213–8220 (2004).
Lynch, D. J., Fountain, T. L. & Mazurkiewicz, Banas, J. A. Glucan-binding proteins are essential for shaping Streptococcus mutans biofilm architecture. FEMS Microbiol. Lett. 268, 158–165 (2007).
Higgins, M. J. & Novak, J. T. Characterization of exocellular protein and its role in bioflocculation. J. Environ. Eng. (New York) 123, 479–485 (1997).
Mora, P., Rosconi, F., Franco Fraguas, L. & Castro-Sowinski, S. Azospirillum brasilense Sp7 produces an outer-membrane lectin that specifically binds to surface-exposed extracellular polysaccharide produced by the bacterium. Arch. Microbiol. 189, 519–524 (2008).
Tielker, D. et al. Pseudomonas aeruginosa lectin LecB is located in the outer membrane and is involved in biofilm formation. Microbiology 151, 1313–1323 (2005).
Diggle, S. P. et al. The galactophilic lectin, LecA, contributes to biofilm development in Pseudomonas aeruginosa. Environ. Microbiol. 8, 1095–1104 (2006).
Johansson, E. M. V. et al. Inhibition and dispersion of Pseudomonas aeruginosa biofilms by glycopeptide dendrimers targeting the fucose-specific lectin LecB. Chem. Biol. 15, 1249–1257 (2008).
Lasa, I. & Penadés, J. R. Bap: a family of surface proteins involved in biofilm formation. Res. Microbiol. 157, 99–107 (2006). A description of the role of non-enzymatic proteins in the biofilm matrix.
Otzen, D. & Nielsen, P. H. We find them here, we find them there: functional bacterial amyloid. Cell. Mol. Life Sci. 65, 910–927 (2007).
van Schaik, E. J. et al. DNA binding: a novel function of Pseudomonas aeruginosa type IV pili. J. Bacteriol. 187, 1455–1464 (2005).
Izano, E. A., Amarante, M. A., Kher, W. B. & Kaplan, J. B. Differential roles of poly-N-acetylglucosamine surface polysaccharide and extracellular DNA in Staphylococcus aureus and Staphylococcus epidermidis biofilms. Appl. Environ. Microbiol. 74, 470–476 (2008).
Molin, S. & Tolker-Nielsen, T. Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilisation of the biofilm structure. Curr. Opin. Biotechnol. 14, 255–261 (2003).
Watanabe, M. et al. Growth and flocculation of a marine photosynthetic bacterium Rhodovulum sp. Appl. Microbiol. Biotechnol. 50, 682–691 (1998).
Yang, L. et al. Effects of iron on DNA release and biofilm development by Pseudomonas aeruginosa. Microbiology 153, 1318–1328 (2007).
Whitchurch, C. B., Tolker-Nielsen, T., Ragas, P. S. & Mattick, J. S. Extracellular DNA required for bacterial biofilm formation. Science 295, 1487 (2002). The first report on the functional relevance of eDNA to biofilms.
Vilain, S., Pretorius, J. M., Theron J. & Broezel, V. S. DNA as an adhesion: Bacillus cereus requires extracellular DNA to form biofilms. Appl. Environ. Microbiol. 75, 2861–2868 (2009).
Mulcahy, H., Charron-Mazenod, L. & Lewenza, S. Extracellular DNA chelates and induces antibiotic resistance in Pseudomonas aeruginosa biofilms. PLoS Pathog. 4, e1000213 (2008).
Allesen-Holm, M. et al. A characterization of DNA release in Pseudomonas aeruginosa cultures and biofilms. Mol. Microbiol. 59, 1114–1128 (2006).
Böckelmann, U. et al. Bacterial extracellular DNA forming a defined network-like structure. FEMS Microbiol. Lett. 262, 31–38 (2006).
Jurcisek, J. A. & Bakaletz, L. O. Biofilms formed by nontypeable Haemophilus influenzae in vivo contain both double-stranded DNA and type IV pilin protein. J. Bacteriol. 189, 3868–3875 (2007).
Steinberger, R. E. & Holden, P. A. Extracellular DNA in single- and multiple-species unsaturated biofilms. Appl. Environ. Microbiol. 71, 5404–5410 (2005).
Neu, T. R. & Poralla, K. An amphiphilic polysaccharide from an adhesive Rhodococcus strain. FEMS Microbiol. Lett. 49, 389–392 (1988).
Neu, T. R., Dengler, T., Jann, B. & Poralla, K. Structural studies of an emulsion-stabilizing exopolysaccharide produced by an adhesive, hydrophobic Rhodococcus strain. J. Gen. Microbiol. 138, 2531–2537 (1992).
Sand, W. & Gehrke, T. Extracellular polymeric substances mediate bioleaching/biocorrosion via interfacial processes involving iron(III) ions and acidophilic bacteria. Res. Microbiol. 157, 49–56 (2006).
Matsuyama, T. & Nakagawa, Y. Surface-active exolipids: analysis of absolute chemical structures and biological functions. J. Microbiol. Methods 25, 165–175 (1996).
Ron, E. Z. & Rosenberg, E. Natural role of biosurfactants. Environ. Microbiol. 3, 229–236 (2001).
Neu, T. Significance of bacterial surface-active compounds in interactions of bacteria with interfaces. Microbiol. Rev. 60, 151–166 (1996). A valuable and comprehensive overview of the concept and ecological role of biosurfactants as part of the EPS matrix.
Panilaitis, B., Castro, G. R., Solaiman, D. & Kaplan, D. L. Biosynthesis of emulsan biopolymers from agro-based feedstocks. J. Appl. Microbiol. 102, 531–537 (2007).
Leck, C. & Bigg, E. K. Biogenic particles in the surface microlayer and overlaying atmosphere in the central Arctic Ocean during summer. Tellus B 57, 305–316 (2005).
Davey, M. E., Cajazza, N. C. & O´Toole, G. A. Rhamnolipid surfactant production affects biofilm architecture in Pseudomonas aeruginosa PAO1. J. Bacteriol. 185, 1027–1036 (2003).
Boles, B. R., Thoendel, M & Singh, P. K. Self-generated diversity produces “insurance effects” in biofilms communities Proc. Natl Acad. Sci. USA 101, 16630–16635 (2004).
Pamp, S. J., Gjermansen, M. & Tolker-Nielsen, T. in The Biofilm Mode of Life. Mechanisms and Adaptations (eds Kjelleberg, S. & Givskov, M.) 37–69 (Horizon Bioscience, Norfolk, UK, 2007).
Or, D., Phutane, S. & Dechesne, A. Extracellular polymeric substances affecting pore-scale hydrologic conditions for bacterial activity in unsaturated soils. Vadose Zone J. 6, 298–305 (2007).
Tamaru, Y., Takami, Y., Yoshida, T. & Sakamoto, T. Crucial role of extracellular polysaccharides in desiccation and freezing tolerance in the terrestrial cyanobacterium Nostoc commune. Appl. Environ. Microbiol. 71, 7327–7333 (2005).
Roberson, E. B., Firestone, M. K. Relationship between desiccation and exopolysaccharide production in a soil Pseudomonas sp. Appl. Environ. Microbiol. 58, 1284–1291 (1992).
Potts, M. Desiccation tolerance of prokaryotes. Microbiol. Rev. 58, 755–805 (1994).
Flemming, H.-C. & Leis, A. in Encyclopedia of Environmental Microbiology (ed. Bitton, G.) 2958–2967 (Wiley, New York, 2002).
Van Hullebusch, E. D., Zandvoord, M. H. & Lens, P. N. L. Metal immobilization by biofilms: mechanisms and analytical tools. Rev. Environ. Sci. Biotechnol. 2, 9–33 (2004).
Wuertz, S. et al. A new method for extraction of extracellular polymeric substances from biofilms and activated sludge suitable for direct quantification of sorbed metals. Water Sci. Technol. 43, 25–34 (2001).
Schmitt, J., Nivens, D., White, D. C. & Flemming, H.-C. Changes of biofilm properties in response to sorbed substances — an FTIR-ATR-study. Water Sci. Technol. 32, 149–155 (1995).
Stoodley, P., Cargo, R., Rupp, C. J., Wilson, S. & Klapper, I. Biofilm material properties as related to shear-induced deformation and detachment phenomena. J. Ind. Microbiol. Biotechnol. 29, 361–367 (2003). A seminal article about the measurement and ecological role of biofilm stability.
Klausen, M. M., Thomsen, T. R., Nielsen, J. L., Mikkelsen, L. H. & Nielsen, P. H. Variations in microcolony strength of probe-defined bacteria in activated sludge flocs. FEMS Microbiol. Ecol. 50, 123–132 (2004).
Gerbersdorf, S. U., Jancke, T., Westrich, B. & Paterson, D. M. Microbial stabilization of riverine sediments by extracellular polymeric substances. Geobiology 6, 57–69 (2008).
Jaeger-Zuern, I. & Gruberg, M. Podostemaceae depend on sticky biofilms with respect to attachment to rocks in waterfalls. Int. J. Plant Sci. 161, 599–607 (2000).
Rupp, C. J., Fux, C. A. & Stoodley, P. Viscoelasticity of Staphylococcus aureus biofilms in response to fluid shear allows resistance to detachment and facilitates rolling migration. Appl. Environ. Microbiol. 71, 2175–2178 (2005).
Shaw, T., Winston, M., Rupp, C. J., Klapper, I. & Stoodley, P. Commonality of elastic relaxation times in biofilms. Phys. Rev. Let. 93, 098102 (2004).
Hohne, D. N., Younger, G. J. & Solomon, M. J. Flexible multifluidic device for mechanical property characterization of soft viscoelastic solids such as bacterial biofilms. Langmuir 25, 7743–7751 (2009).
Webb, J. et al. Cell death in Pseudomonas aeruginosa biofilm development. J. Bacteriol. 185, 4585–4592 (2003). A thorough discussion of the dynamics of the EPS matrix and the concept of a microbial analogue to programmed cell death that leads to alterations of the matrix architecture.
Borlee, B. R. et al. Pseudomonas aeruginosa uses a cyclic-di-GMP-regulated adhesin to reinforce the biofilm extracellular matrix. Mol. Microbiol. 75, 827–842 (2010).
De Kievit, T. R. Quorum sensing in Pseudomonas aeruginosa biofilms. Environ. Microbiol. 11, 279–288 (2009).
Cooksey, K. E. & Wigglesworth-Cooksey, B. in Encyclopedia of Environmental Microbiology (ed. Bitton, G.) 1051–1063 (Wiley, New York, 2002).
de Brouwer, J. F. C., Wolfstein, K., Ruddy, G. K., Jones, T. E. R. & Stal, L. J. Biogenic stabilization of intertidal sediments: the importance of extracellular polymeric substances produced by benthic diatoms. Microbiol. Ecol. 49, 501–512 (2005).
Dade, W. B. et al. Effects of bacterial exopolymer adhesion on the entrainment of sand. Geomicrobiol. J. 8, 1–16 (1990).
Domozych, D. S., Kort, S., Benton, S. & Yu, T. The extracellular polymeric substance of the green alga Penium margaritaceum and its role in biofilm formation. Biofilms 2, 129–144 (2005).
Baillie, G. S. & Douglas, L. J. Matrix polymers of Candida biofilms and their possible role in biofilm resistance to antifungal agents. J. Antimicrob. Chemother. 46, 397–403 (2000).
Chandra, J. et al. Biofilm formation by the fungal pathogen Candida albicans: development, architecture, and drug resistance. J. Bacteriol. 183, 5385–5394 (2001).
Verstrepen, K. J. & Klis, F. M. Flocculation, adhesion and biofilm formation in yeasts. Mol. Microbiol. 60, 5–15 (2006).
Zolghadr, B. et al. Appendage-mediated surface adherence of Sulfolobus solfataricus. J. Bacteriol. 192, 104–110 (2009).
Wrangstadh, M., Szewzyk, U., Östling, J. & Kjelleberg, S. Starvation-specific formation of peripheral exopolysaccharide by a marine Pseudomonas sp., strain S9. Appl. Environ. Microbiol. 56, 2065–2072 (1990).
Neu, T. R., Lawrence, J. R. in Microbial Extracellular Polymeric Substances (eds Wingender, J., Neu, T. & Flemming, H.-C.) 22–47 (Springer, Heidelberg, 1999).
Dow, J. M. et al. Biofilm dispersal in Xanthomonas campestris is controlled by cell–cell signalling and is required for full virulence to plants. Proc. Natl Acad. Sci. USA 100, 10995–11000 (2003).
Davies, D. G. & Marques, C. N. H. A fatty acid messenger is responsible for inducing dispersion in microbial biofilms. J. Bacteriol. 191, 1393–1403 (2009).
Kolodkin-Gal, I. et al. D-amino acids trigger biofilm disassembly. Science 328, 627–629 (2010).
Tait, K., Skillman, L. C. & Sutherland, I. W. The efficacy of bacteriophage as a method of biofilm eradication. Biofouling 18, 305–311 (2002).
Nardini, M., Lang, D. M., Liebeton, K., Jaeger, K.-E. & Dijkstra, B. W. Crystal structure of Pseudomonas aeruginosa lipase in the open conformation. The prototype for family i.1 of bacterial lipases. J. Biol. Chem. 275, 31219–31225 (2000).
Grobe, S., Wingender, J., Trüper, H. G. Characterization of mucoid Pseudomonas aeruginosa strains isolated from technical water systems. J. Appl. Bacteriol. 79, 94–102 (1995).
We are grateful for the inspiring cooperation with partners in the research group on 'Physico-chemistry of Biofilms': W. Borchard, K.-E. Jaeger, H. Kuhn, C. Mayer and W. Veeman. We also acknowledge financial support by the German Research Foundation to various EPS research projects. Furthermore, constructive, critical and stimulating comments and discussions with I. Sutherland are highly appreciated.
The authors declare no competing financial interests.
A loose definition for microbial aggregates that usually accumulate at a solid–liquid interface and are encased in a matrix of highly hydrated EPS. Included in this definition are cell aggregates such as flocs (floating biofilms) and sludge, which are not attached to an interface but which share the characteristics of biofilms. Multispecies biofilms can form stable microconsortia, develop physiochemical gradients, and undergo horizontal gene transfer and intense cell–cell communication, and these consortia therefore represent highly competitive environments.
- Extracellular polymeric substances
Hydrated biopolymers (including polysaccharides, proteins, nucleic acids and lipids) that are secreted by biofilm cells to encase and immobilize microbial aggregates. These biopolymers are responsible for the macroscopic appearance of biofilms, which are frequently referred to as 'slime'.
- Humic substance
A component of the natural organic matter in soil and water enviroments. Humic substances are mixtures of compounds that are formed by limited degradation and transformation of dead organic matter and that are resistant to complete biodegradation. They can be divided into three main fractions: humic acids, fulvic acids and humin. They usually include phenolic and polyaromatic compounds (containing peptide and carbohydrate moieties with carboxylic substituents), providing the acidic character.
A long, thin, helically shaped bacterial appendage that provides motility. A flagellum consists of several components and moves by rotation, much like a propeller. The motor is anchored in the cytoplasmic membrane and the cell wall.
A bacterial surface structure that is similar to a fimbria but is typically a longer structure, and that is present on the cell surface in one or two copies. Pili can be receptors for bacteriophages and also facilitate genetic exchange between bacterial cells during conjugation. Type IV pili mediate twitching motility, which is a flagella-independent form of bacterial translocation over surfaces, and can be involved in biofilm development.
A filamentous structure composed of one or a few proteins that extends from the surface of a cell and can have diverse functions. Fimbriae are involved in attachment to both animate and inanimate surfaces and in the formation of pellicles and biofilms. They assist in the disease process of some pathogens, such as S. enterica, Neisseria gonorrhoea and Bordetella pertussis.
- Membrane vesicle
A vesicle that is formed from the outer membrane of Gram-negative bacteria, is secreted from the cell surface and contains extracellular enzymes and nucleic acids. These vesicles may represent mobile elements in the EPS matrix.
A discrete polysaccharide (sometimes also protein) layer that is firmly attached to the surface of a bacterial cell, closely surrounding it, in contrast to less compact, amorphous slime that is shed into the more distant extracellular environment.
A protein or glycoprotein of plant, animal or microbial origin that binds to carbohydrates with a characteristic specificity. Fluorescently labelled lectins can be used as probes to investigate EPS composition, enabling the microscopic in situ detection of EPS and their distribution in biofilms.
- Raman microscopy
A spectroscopic technique based on inelastic light scattering (Raman scattering) of monochromatic laser light in the near-ultraviolet range, revealing vibrational, rotational and other low-frequency modes in a system. The technique is used for the analysis of chemical bonds and is suitable for very small volumes, allowing spectra and chemical information to be obtained for the molecules present in that volume.
- Matrix void
A pore or channel in the biofilm matrix that contains liquid water and is not filled with hydrated EPS molecules.
A laminated microbial mat that is typically built from layers of filamentous cyanobacteria and other microorganisms that become fossilized. Stromatolites are the oldest records of life on Earth, dating back 3.5 billion years.
- Surface-active property
The ability of a molecule to alter the interface of two different phases. Substances with surface-active properties (surfactants) are amphipatic molecules with both hydrophilic and hydrophobic (generally hydrocarbon) moieties. They partition preferentially at the interface between fluid phases with different degrees of polarity and hydrogen bonding, such as oil–water interfaces.
A substance that is synthesized by living cells (mostly bacteria and yeasts) and that is surface active. Biosurfactants reduce surface tension, stabilize emulsions, promote foaming and are generally non-toxic and biodegradable. When grown on hydrocarbon substrates as a carbon source, microorganisms can synthesize a wide range of biosurfactants, such as glycolipids and phospholipids. These chemicals are apparently synthesized to emulsify the hydrocarbon substrate and facilitate its transport into the cells. In some bacterial species, such as P. aeruginosa, biosurfactants are also involved in a group movement behaviour called swarming motility.
- Hydraulic decoupling
The formation of areas that have virtually no exchange of water content with their environment. An example is a desiccated EPS layer that covers an area with a high water content but has very low water transport through the layer, retaining the water underneath.
- Elasticity modulus
The tendency of an object or material to reversibly develop an elastic force in response to deformation. Mathematically, the elasticity modulus is the proportionality factor between the force and the deformation, or, in other words, the slope on a plot of stress versus strain in the elastic deformation region. Stiff materials have a higher elasticity modulus, whereas soft materials have a lower one.
- Stress relaxation
A deviation from the ideal elastic behaviour of a material due to an internal relief of stress under constant strain. Some materials, when put under mechanical tension, undergo internal flow processes (termed 'creep') that are at least partially irreversible and lead to a constant deformation of the test specimen.
About this article
Cite this article
Flemming, HC., Wingender, J. The biofilm matrix. Nat Rev Microbiol 8, 623–633 (2010). https://doi.org/10.1038/nrmicro2415
This article is cited by
BMC Oral Health (2023)
Future Journal of Pharmaceutical Sciences (2023)
Media component bovine serum albumin facilitates the formation of mycobacterial biofilms in response to reductive stress
BMC Microbiology (2023)
Micafungin effect on Pseudomonas aeruginosa metabolome, virulence and biofilm: potential quorum sensing inhibitor
AMB Express (2023)
Novel biological aqua crust enhances in situ metal(loid) bioremediation driven by phototrophic/diazotrophic biofilm