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Keywords:

  • biofilm;
  • resistance;
  • attachment;
  • development;
  • inhibition;
  • dispersion

Abstract

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

Biofilms are complex microbial communities consisting of microcolonies embedded in a matrix of self-produced polymer substances. Biofilm cells show much greater resistance to environmental challenges including antimicrobial agents than their free-living counterparts. The biofilm mode of life is believed to significantly contribute to successful microbial survival in hostile environments. Conventional treatment, disinfection and cleaning strategies do not proficiently deal with biofilm-related problems, such as persistent infections and contamination of food production facilities. In this review, strategies to control biofilms are discussed, including those of inhibition of microbial attachment, interference of biofilm structure development and differentiation, killing of biofilm cells and induction of biofilm dispersion.


Introduction of microbial biofilms

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

Bacteria form surface attached biofilm communities as one of the most important survival strategies in nature (Costerton et al., 1995). Biofilms consist of water, bacterial cells and a wide range of self-generated extracellular polymeric substances (EPS) referred to as the matrix. Microbial biofilms affect world economy at the level of billions of dollars with regard to equipment damage, product contamination, energy losses and infections. Conventional methods that would otherwise lead to eradication of non-attached, non-aggregated (planktonic) microbes are often ineffective to the microbial populations inside the biofilms due to their particular physiology and physical matrix barriers (Stewart, 2002). Therefore, novel strategies based on a more fulfilling understanding of the biofilm phenomenon are urgently needed.

In vitro studies reveal that biofilm development is a dynamic and complicated process that involves many different components and various group activities. For example, a representative diagram of biofilm development on vacant glass surfaces in a continuously irrigated flow chamber by the opportunistic pathogen Pseudomonas aeruginosa is depicted in Fig. 1. Pseudomonas aeruginosa cells attach to the glass surfaces or substratum by means of surface appendages such as type IV pili and flagellum (O'Toole & Kolter, 1998). Shortly after initial attachment, non-motile subpopulation of P. aeruginosa cells starts microcolony formation, which requires both Pel and Psl extracellular polysaccharides as well as biosurfactant (Pamp & Tolker-Nielsen, 2007; Yang et al., 2011). Quorum sensing systems and iron signalling are highly induced in the microcolonies, which favour release of extracellular DNA (eDNA), an important EPS material (Hentzer et al., 2005; Allesen-Holm et al., 2006). Motile subpopulation of P. aeruginosa cells then moves to the microcolonies formed by the non-motile subpopulation via flagellum-mediated chemotaxis and binds to the eDNA through type IV pili (Barken et al., 2008; Yang et al., 2009ab). The association between non-motile and motile subpopulations of P. aeruginosa cells leads to the formation of mushroom-shaped biofilm structures with distinct physiological states (such as tolerance to treatment by different antibiotics) (Bjarnsholt et al., 2005; Haagensen et al., 2007; Yang et al., 2007; Pamp et al., 2008). Under stressful conditions (Webb et al., 2003; Banin et al., 2006; Barraud et al., 2006; Haagensen et al., 2007), P. aeruginosa biofilm cells will become activated and cause dispersion of the biofilms. A summary of strategies to combat biofilms is described in Fig. 1 and will be discussed in details in the following text.

image

Figure 1. Summary of biofilm development stages and biofilm combating strategies. Biofilm development stages: 1, individual planktonic cells attach to the surface; 2, attached cells form microcolonies; 3, subpopulations interact with each other during biofilm structure development;4, macrocolonies are formed in mature biofilms; 5, dead cells accumulate under stressful conditions; 6, cells are released from the biofilm macrocolonies. Red dots represent eDNA; blue and yellow cycles represent cells of the non-motile and motile biofilm subpopulations respectively; cyan peripheral cycles represent the Pel and Psl polysaccharides (adapted from Yang et al., 2011).

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Strategies to inhibit microbial attachment

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

Microbial attachment to a surface is a universal phenomenon in nature and is essential for biofilm formation. In recent years, a series of different approaches have been developed to reduce microbial attachment, including biochemical approaches, physicochemical approaches and biological approaches.

Biochemical approaches

Antimicrobial agents immobilized on surfaces can kill attaching organisms. Various methods are used to generate antimicrobial surfaces. Non-covalently binding, covalently immobilization and polymer matrix loading of antimicrobial agents are routinely used approaches for this purpose. For example, antimicrobial peptides (AMPs) were loaded on micro-porous calcium phosphate (CaP)-coated titanium surface up to 9 μg cm−2 using a simple soaking technique, and this surface exhibited antimicrobial activity against both Gram-positive (Staphylococcus aureus) and Gram-negative (P. aeruginosa) bacteria (Kazemzadeh-Narbat et al., 2010). However, surfaces coated with such ‘conventional’ antimicrobials are usually considered short-term with respect to ‘life-time’. New methods that would enable a long-term coating of antimicrobials are under development. Synthesis of permanent, non-leaching antibacterial surfaces represents a new trend of coating technology. As an example of that, single-walled carbon nanotubes (SWNTs) were reported to have strong antimicrobial activities against microbes (Vecitis et al., 2010). Electrospun polymer mats with incorporated narrow diameter SWNTs were found to significantly reduce bacterial colonization and subsequent biofilm formation (Schiffman & Elimelech, 2011).

Besides microbicidal agents, non-microbicidal agents are also used to block microbial attachment. For example, pathogens often bind human cell surface through pili and form biofilm in vivo (Tsui et al., 2003; Okahashi et al., 2011). A 12-mer peptide (RQERSSLSKPVV), which binds to the structural protein PilS of the type IVB pili of Salmonella Typhi, was isolated by using a ribosome display system and shown to inhibit adhesion to or invasion of human monocytic THP-1 cells by piliated S. Typhi (Wu et al., 2005). This group also identified high-affinity single-stranded RNA aptamer [S-PS(8.4)] as a type IVB pilus-specific ligand and further showed that the aptamer [S-PS(8.4)] could significantly inhibit the entry of the piliated S. Typhi into human THP-1 cells (Pan et al., 2005). Bovine lactoferrin was also shown to interact with cable pili of Burkholderia cenocepacia and efficiently inhibit invasion of alveolar epithelial cells by free-living bacteria or biofilm (Ammendolia et al., 2010).

Physicochemical approaches

Increasing efforts have been put on development of modified surfaces with anti-adhesive properties by means of physicochemistry. For example, electropolished stainless steel was shown to significantly reduce attachment and biofilm formation by bacterial cells than the sand-blasted and sanded stainless steel surfaces (Arnold & Bailey, 2000). Raulio et al. (2008) reported that hydrophilic or hydrophobic coated stainless steel by diamond-like carbon or certain fluoropolymers could reduce or almost eliminate adhesion and biofilm formation by Staphylococcus epidermidis, Deinococcus geothermalis, Meiothermus silvanus and Pseudoxanthomonas taiwanensis (Raulio et al., 2008). A robust peptide-based coating technology for modifying the surface of titanium (Ti) metal through non-covalent binding was introduced by Khoo et al. (2009). In their study, a short HKH tripeptide motif containing peptide (e.g. SHKHGGHKHGSSGK) possessing affinity for Ti was identified by means of a phage display based screening and amino acid substitution study. Based on this peptide, a PEGylated analogue was found to rapidly coat Ti and efficiently block the adsorption of fibronectin and attachment of S. aureus (Khoo et al., 2009).

Anti-adhesive properties and microbicidal properties are combined by researchers when designing novel surfaces. In a recent study, Yuan et al. (2011) immobilized lysozyme to the chain ends of poly(ethylene glycol) branches of the grafted poly(ethylene glycol) monomethacrylate (PEGMA) polymer after PEGMA was coated to stainless steel surfaces (Yuan et al., 2011). This coating strategy is efficient in preventing bovine serum albumin adsorption, bacterial adhesion and killing of Escherichia coli and S. aureus cells.

Biological approaches

Biological approaches have great potential in alleviating microbial attachments. Microbial species coexist and interact extensively with each other in natural biofilms. The competition for substrates serves as one of the major evolutionary driving forces in these multiple-species biofilms (Xavier & Foster, 2007; Xavier et al., 2009). Thus, many bacteria are capable of synthesizing and excreting chemicals that inhibit biofilm formation by other species. For example, biosurfactants are synthesized and excreted by many bacteria, which inhibit attachment by their competitors (Zeraik & Nitschke, 2010; Luna et al., 2011). Thus, biosurfactants producing probiotic bacteria are proposed as potential biofilm control agents (Rodrigues et al., 2004; Falagas & Makris, 2009).

Biological approaches for controlling biofilms are well studied in dental plaque biofilms. The oral microbial flora contains many beneficial species that are able to halt the progression of oral disease. Probiotic strain Lactobacillus acidophilus was shown to reduce the biofilm formation of Streptococcus mutans, one of primary dental cariogen, through inhibiting attachment (Tahmourespour & Kermanshahi, 2011). The early dental plaque colonizer Streptococcus gordonii secretes proteases that reduce subsequent colonization of S. mutans by inactivating its competence-stimulating peptide signalling system (Wang et al., 2010). In a recent study, Ogawa et al. (2011) identified exo-beta-d-fructosidase from the culture supernatants of Streptococcus salivarius as an active substance to inhibit S. mutans biofilm formation (Ogawa et al., 2011).

Strategies to inhibit biofilm structure development and differentiation

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

Young biofilms are often more susceptible to antimicrobial agents than mature biofilms (Drenkard & Ausubel, 2002; Mukherjee et al., 2003; Allesen-Holm et al., 2006; Ito et al., 2009). The large amounts of EPS in the mature biofilms can act as a diffusion barrier to antimicrobial agents (Hoyle et al., 1992; Souli & Giamarellou, 1998; Anderl et al., 2000). The high cell density in the mature biofilms can induce cell-to-cell communication (quorum sensing) systems, which up-regulate expression of genes contributing to antibiotic resistance (De Kievit et al., 2001; Bjarnsholt et al., 2005) and release of protecting DNA (Hunt et al., 1995; Allesen-Holm et al., 2006). Also, competition for nutrients can lead to subpopulation differentiation and spatial physiological heterogeneity in the mature biofilms, which further cause antibiotic resistance (Xu et al., 1998; Walters et al., 2003). Thus, strategies for interfering structure development and differentiation of biofilms are being developed by many research groups.

Disruption of biofilm EPS matrix

Enzymatically and chemically disrupting biofilm EPS matrix is proved to be an efficient approach to arrest biofilm structure development. Longhi et al. (2008) reported that protease treatment of Listeria monocytogenes cells digests surface protein Ami4b autolysin, internalinB, and ActA and reduces both invasion ability and biofilm formation of L. monocytogenes (Longhi et al., 2008). Biofilm formation by S. epidermidis and S. aureus requires surface protein (Aap and SasG) that contain sequence repeats known as G5 domain (Rohde et al., 2005; Corrigan et al., 2007; Geoghegan et al., 2010). Dimerization of the G5 domains in the presence of Zn2+ is essential for these proteins to function as intercellular adhesin (Conrady et al., 2008). Zn2+ chelation was shown to specifically prevent biofilm formation by S. epidermidis and methicillin-resistant S. aureus, which was proposed as a potential approach for combating biofilm-related infections (Conrady et al., 2008). Antiparasitic drug nitazoxanide and its active metabolite, tizoxanide, were reported to inhibit S. epidermidis biofilm formation possibly by targeting the zinc-dependent adhesin Aap (Tchouaffi-Nana et al., 2010).

Polysaccharide intercellular adhesin (PIA) synthesized by the icaADBC operon of Staphylococci is one of the best understood EPS components and is essential for Staphylococci biofilm development. Thus the ica genes represent potential targets for biofilm inhibitors. Oduwole et al. (2010) reported that the antibacterial agent povidone-iodine at sub-inhibitory concentrations has anti-biofilm activity against S. epidermidis by activating the icaR transcriptional repressor in S. epidermidis and reducing the transcription of the icaADBC operon (Oduwole et al., 2010). More recently, the organosulfur compound from garlic, allicin, was shown to inhibit PIA biosynthesis and biofilm development by S. epidermidis (Cruz-Villalon & Perez-Giraldo, 2011). Sulfhydryl compounds such as dithiothreitol, beta-mercaptoethanol or cysteine were also shown to reduce S. aureus biofilm formation by inhibiting PIA biosynthesis probably through metabolic interventions (Wu et al., 2011ab).

Interference with group activities

Biofilm formation involves many ‘social’ activities including those of quorum sensing, iron siderophore and biosurfactant production (Davies et al., 1998; Davey et al., 2003; Banin et al., 2005; Alhede et al., 2009). Interference of these group activities can affect biofilm architecture and antibiotic resistance.

Quorum sensing is widely used by microorganisms to coordinate communal behaviours such as bioluminescence, swarming and production of virulence (Rasmussen et al., 2000; DeLisa et al., 2001; Miller et al., 2002). Quorum sensing regulation is achieved by synthesizing and releasing small signal molecules by many denoted autoinducers (AIs), a word inspired from their positive feedback effect on expression of bioluminescense. The structures of AIs and their receptors have been extensively characterized (Shaw et al., 1997; Vannini et al., 2002; Bottomley et al., 2007). Thus, a diverse set of close analogues have been identified as quorum sensing inhibitors (QSIs) and transcriptomic analysis suggest that they exert their effects by binding to the quorum sensing receptor proteins by displacing the cognate AIs (Hentzer et al., 2003; Glansdorp et al., 2004; Rasmussen et al., 2005). Recently, several crystal structures of the quorum-sensing regulatory proteins with their cognate AIs have been reported (Vannini et al., 2002; Bottomley et al., 2007; De Silva et al., 2007; Kim et al., 2010), and in line with that computational modelling approaches have been employed to design potential QSIs. Yang et al. (2009a b) applied molecular docking and virtual screening and identified three recognized drugs, salicylic acid, nifuroxazide, and chlorzoxazone, as QSIs of P. aeruginosa (Yang et al., 2009a b). AI structurally unrelated QSIs were discovered by Soulere et al. (2010) through docking-based screening on a 2344 chemical compounds library (Soulere et al., 2010). Besides docking, structure-activity relationship methods are also applied to design and identify novel QSIs (Steenackers et al., 2010; Brackman et al., 2011).

Over the past few years, researchers have identified quorum-quenching enzymes from many prokaryotic and eukaryotic organisms, which degrade quorum-sensing signal molecules (Dong et al., 2007). Bacillus spp. produces a N-acyl-homoserine lactone lactonase that hydrolyses this major group quorum sensing AI in Gram-negative bacteria (Augustine et al., 2010). Mammalian cells was shown to produce paraoxonases (PON1, PON2, and PON3) that hydrolytically inactivate quorum sensing signal N-(3-oxododecanoyl)-l-homoserine lactone from P. aeruginosa (Teiber et al., 2008). Recently, metagenomic approaches are widely applied to identify novel enzymes from nature. Bijtenhoorn et al. (2011) isolated and biochemically characterized a novel N-acyl-homoserine lactone hydrolase, BpiB05, from the soil metagenome (Bijtenhoorn et al., 2011). BpiB05 is not distantly related to any of the currently known N-acyl-homoserine lactone hydrolases and strongly reduces motility, pyocyanin synthesis and biofilm formation by P. aeruginosa (Bijtenhoorn et al., 2011). Quorum-quenching enzymes have been immobilized on surfaces and applied as anti-biofilm agents (Kim et al., 2011; Ng et al., 2011).

Metabolic intervention

Secondary metabolites may serve as intercellular pathogenic signals, which regulate numerous phenomena including biofilm formation (Dufour & Rao, 2011). Thus, metabolic intervention can be used to affect development and differentiation of biofilms.

The green tea epigallocatechin gallate was shown to reduce both quorum sensing and biofilm development of P. aeruginosa through inhibiting the enoyl-acyl carrier protein reductase from the type II fatty acid synthesis pathway (Yang et al., 2010). A cyclopropane-containing fatty acid, lyngbyoic acid, from the marine cyanobacterium was shown to directly inhibit LasB enzymatic activity and reduce the production of pyocyanin and elastase in P. aeruginosa (Kwan et al., 2011).

Iron metabolism and signalling play essential roles on biofilm formation and regulate production of virulence factors (Bollinger et al., 2001; Banin et al., 2005; Johnson et al., 2005; Sonnleitner et al., 2011). In 2002, Singh et al. reported that iron chelator lactoferrin stimulates twitching motility and prevents biofilm formation by P. aeruginosa (Singh et al., 2002). Iron-binding compounds were also reported to reduce biofilm formation of P. aeruginosa under anaerobic conditions (O'May et al., 2009). The transition metal gallium (Ga3+) is chemically similar to iron and was found to efficiently interfere iron uptake and biofilm formation by P. aeruginosa (Kaneko et al., 2007).

Strategies to kill biofilm cells

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

Conventional antimicrobial therapy to eradicate biofilm-related infections is frequently ineffective. The resistance mechanisms of biofilm cells to antimicrobial agents are rather complicated and vary greatly among biofilms in different stages (Stewart, 2002; Davies, 2003). Novel anti-biofilm strategies have been extensively proposed and tested in recent years.

Novel anti-biofilm agents

Two-component regulatory systems are involved in biofilm formation by many bacterial species (Li et al., 2002; Hancock & Perego, 2004; Tomaras et al., 2008; Petrova & Sauer, 2010). Qin et al. (2006) identified novel inhibitors of the S. epidermidis YycG histidine kinase through structure-based virtual screening and further showed that five of these inhibitors display bactericidal effects on both planktonic and biofilm cells of S. epidermidis (Qin et al., 2006). Addition of exogenous competence-stimulating peptide beyond the levels necessary for competence was shown to induce S. mutans cell death in both planktonic and biofilm cultures though the ComDE two-component signal transduction systems (Qi et al., 2005).

Siderophore-mediated iron uptake and signalling are required for biofilm structure development and maturation (Banin et al., 2005; Johnson et al., 2005; Yang et al., 2009a). Siderophore-antibiotic conjugates are used as ‘Trojan Horses’ to combat pathogenic bacteria (Miller et al., 1991; Budzikiewicz, 2001). Banin et al. (2008) reported that the desferrioxamine-gallium (DFO-Ga) conjugate kills planktonic cells and blocks biofilm formation by P. aeruginosa (Banin et al., 2008). They also showed that a combination of DFO-Ga and gentamicin causes massive killing of cells in mature P. aeruginosa biofilms (Banin et al., 2008).

Recently, AMPs are proposed to be promising agents against biofilms (Batoni et al., 2011). AMPs combined with antibiotics were shown to rapidly kill most of the cells in biofilms formed by pathogenic bacteria (Pamp et al., 2008; Herrmann et al., 2010). However, AMPs have undesirable properties such as nonspecific toxicity and low stability, which limit their application. Thus, numerous approaches are applied to modify the structures of AMPs and obtain novel peptides or peptidomimetics. AMP mimetics were reported by different research groups to be highly active against biofilms (Flemming et al., 2009; Hua et al., 2010).

Enhancement of antimicrobial penetration

The activity of antimicrobial agents against biofilms is largely hindered by the dense microcolonies embedded in the EPS matrix. Approaches to enhance antimicrobial penetration in biofilms have been evaluated by different research groups. Alipour et al. (2009) reported that co-administration of DNase and alginate lyase significantly enhance activity of certain aminoglycosides in reducing biofilm growth and cystic fibrosis sputum bacterial counts of P. aeruginosa (Alipour et al., 2009). Lipopeptide biosurfactant produced by Bacillus licheniformis was shown to significantly enhance the efficacy of antibiotics in killing E. coli biofilms (Rivardo et al., 2011). Micelle-encapsulated antibiotics and antibiotic-encapsulated biodegradable polymeric nanoparticles are also reported to efficiently kill biofilm cells (Jones, 2005; Cheow et al., 2010). Efflux pump systems are involved in biofilm formation and antimicrobial resistance (Pamp et al., 2008; Zhang & Mah, 2008). Inactivation of efflux systems by efflux pump inhibitors was reported to abolish bacterial biofilm formation or enhance antimicrobial activity against biofilms (Kvist et al., 2008; Liu et al., 2010).

Phage activity

In recent years, phages are suggested as alternatives to antibiotics for the treatment of biofilms. Phages are inexpensive and specific against a host or host range, and will not affect the normal microflora of the environment where they are applied. A T7-like lytic phage against P. aeruginosa isolated from Pavana river water has been shown to prevent and disperse biofilms of P. aeruginosa (Ahiwale et al., 2011). Carson et al. (2010) reported that lytic bacteriophages could eradicate established biofilms of Proteus mirabilis and E. coli, and impregnation of hydrogel-coated catheter sections with these lytic bacteriophages could prevent biofilm formation on catheter biomaterials (Carson et al., 2010).

Some phages also possess polysaccharide-degrading enzymes that can rapidly destroy the integrity of biofilms (Suthereland et al., 2004). A P. aeruginosa-specific phage was isolated and shown to produce alginase to depolymerize the alginate capsule from the mucoid cystic fibrosis isolates of P. aeruginosa (Glonti et al., 2010). This alginase might accelerate phagocytic uptake of bacteria and perturb bacterial biofilms of patients with cystic fibrosis. An engineered bacteriophage which expresses a biofilm-degrading enzyme during infection was reported to simultaneously attack the biofilm cells and the EPS matrix (Lu & Collins, 2007). A cell-wall-degrading enzyme SAL-2 from a new podoviridae S. aureus bacteriophage (SAP-2) was cloned and expressed by Son et al. (2010). The SAL-2 enzyme has specific lytic activity against S. aureus with a minimum inhibitory concentration of about 1 μg mL−1 and can efficiently remove S. aureus biofilms (Son et al., 2010).

Phages are also reported to improve the conventional antimicrobial treatment to biofilm related infections. Verma et al. (2010) reported that the depolymerase-mediated structural changes caused by a lytic bacteriophage (KPO1K2) enhance ciprofloxacin activity against mature biofilms of Klebsiella pneumonia (Verma et al., 2010). They also reported that combination of bacteriophage and ciprofloxacin efficiently kills K. pneumonia biofilm cells and restricts the formation of resistant variants when compared with individual treatments (Verma et al., 2009).

Strategies to induce biofilm dispersion

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

It is well known that environmental cues such as oxygen and carbon substrate concentration can trigger biofilm dispersion (Applegate & Bryers, 1991; Thormann et al., 2005; Gjermansen et al., 2010; Newell et al., 2011). Biofilm dispersion often coincides with alteration of the biofilm EPS components. Understanding the modulation of biofilm EPS components and transduction of the dispersion signals would greatly facilitate the development of dispersion-based strategies to control biofilm formation.

Genetic regulators of biofilm formation and dispersion

In recent years, genetic regulators and signal transduction pathways for biofilm dispersion have been identified from a number of microorganisms. Gjermansen et al. (2010) reported that overexpression of a plasmid-born EAL domain protein reduces intracellular c-di-GMP level and activates the LapG cysteine proteinase in biofilms formed by Pseudomonas putida (Gjermansen et al., 2010). The activated LapG proteinase can digest the LapA protein, which functions both as a surface adhesin and as a biofilm matrix component, and cause dispersion of P. putida biofilms (Gjermansen et al., 2010). Three two-component systems, BfiSR, BfmSR and MifSR, are reported to be essential for regulating P. aeruginosa biofilm development (Petrova & Sauer, 2009). Inhibiting the expression of bfiS, bfmR and mifR genes in mature biofilms leads to biofilm architectural collapse and biomass loss (Petrova & Sauer, 2009). Boles & Horswill (2008) reported that activation of the agr quorum-sensing system of S. aureus by autoinducing peptide addition or glucose depletion can trigger biofilm dispersion via a protease-mediated mechanism (Boles & Horswill, 2008).

Genetically engineered regulators are used to manipulate biofilm formation and dispersion. Uppuluri et al. (2010) demonstrated that modulation of NRG1 gene expression affects biofilm formation and dispersion by Candida albicans (Uppuluri et al., 2010). Hong et al. (2010a) used random mutagenesis to obtain variants of the global transcriptional regulator Hha, which controls biofilm formation of E. coli probably by activation of proteases (Hong et al., 2010a). One of the obtained Hha variants, Hha13D6 (D22V, L40R, V42I and D48A), causes nearly complete biofilm dispersion by increasing apoptosis (Hong et al., 2010a). The same authors also engineered another global regulator H-NS of E. coli to control its biofilm formation (Hong et al., 2010a b).

Dispersion-inducing agents

Analysis of signal transduction molecules involved in biofilm dispersion has led to identification of a series of biofilm dispersion-inducing agents. The plant pathogen Xanthomonas campestris pathovar campestris (Xcc) regulates the production of virulence factors via a small molecule diffusible signal factor (DSF) (Dow et al., 2003). Addition of DSF can activate an extracellular enzyme, single endo-beta-1,4-mannanase, which disrupts the extracellular polysaccharide xanthan and triggers the dispersion of the Xcc biofilms (Dow et al., 2003). A DSF structurally related short-chain fatty acid signalling molecule, cis-2-decenoic acid, was identified from P. aeruginosa cultures and found to induce the dispersion of established biofilms formed by many bacterial species, such as P. aeruginosa, E. coli, K. pneumoniae, P. mirabilis, Streptococcus pyogenes, Bacillus subtilis, S. aureus, and C. albicans (Davies & Marques, 2009).

Barraud et al. (2006) reported that the anaerobic respiration processes are involved in P. aeruginosa biofilm dispersion, and nitric oxide (NO) can cause dispersion of P. aeruginosa biofilms (Barraud et al., 2006). They further showed that the NO donor sodium nitroprusside efficiently disperses P. aeruginosa biofilms and greatly enhances the activity of conventional antimicrobial compounds against P. aeruginosa biofilms. Ginseng extract was recently shown to disperse P. aeruginosa biofilms by facilitating twitching and swimming motility, which further enhance the activity of conventional antimicrobial compounds against P. aeruginosa biofilms (Wu et al., 2011a). 2-aminoimidazole-derived anti-biofilm agents are extensively studied and are shown to enhance the activity of conventional antibiotics against biofilms (Richards & Melander, 2008; Richards et al., 2008; Rogers et al., 2010; Rogers et al., 2011).

Agents targeting the EPS components are frequently reported to induce biofilm dispersion. Bacillus licheniformis secretes an extracellular DNase (NucB) that rapidly disperses the biofilms formed by both Gram-positive and Gram-negative bacteria (Nijland et al., 2010). D-amino acids treatment was shown to cause the release of amyloid fibers that link cells in biofilms at nanomolar concentrations and disperse biofilms formed by S. aureus and P. aeruginosa (Kolodkin-Gal et al., 2010). Johansson et al. (2008) screened combinatorial libraries of multivalent fucosyl-peptide dendrimers and identified high-affinity ligands of the fucose-specific lectin (LecB) of P. aeruginosa (Johansson et al., 2008). They showed these dendrimers can completely disperse biofilms formed by the wild-type strain and several clinical P. aeruginosa isolates.

Conclusion

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References

There is an urgent need to develop novel strategies to control biofilms in industrial and clinical settings. A wide range of promising approaches have been evaluated in different biofilm model systems. However, dealing with natural biofilms formed by multi-species is more complicated than the biofilms formed by single-species in our model systems since the mechanisms of multi-species biofilm formation is not well investigated. More reliable techniques for investigating biofilms and better model systems for evaluating control strategies are still required.

References

  1. Top of page
  2. Abstract
  3. Introduction of microbial biofilms
  4. Strategies to inhibit microbial attachment
  5. Strategies to inhibit biofilm structure development and differentiation
  6. Strategies to kill biofilm cells
  7. Strategies to induce biofilm dispersion
  8. Conclusion
  9. Acknowledgement
  10. References