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Fungal Biofilms and Polymicrobial Diseases

Department of Clinical Analysis, School of Pharmaceutical Sciences, São Paulo State University (UNESP), Araraquara SP 14800-903, Brazil
Department of Physiological Sciences, Piracicaba Dental School, University of Campinas (UNICAMP), Piracicaba SP 13414-018, Brazil
Author to whom correspondence should be addressed.
J. Fungi 2017, 3(2), 22;
Received: 22 February 2017 / Revised: 19 April 2017 / Accepted: 4 May 2017 / Published: 10 May 2017
(This article belongs to the Special Issue Fungal Biofilms)


Biofilm formation is an important virulence factor for pathogenic fungi. Both yeasts and filamentous fungi can adhere to biotic and abiotic surfaces, developing into highly organized communities that are resistant to antimicrobials and environmental conditions. In recent years, new genera of fungi have been correlated with biofilm formation. However, Candida biofilms remain the most widely studied from the morphological and molecular perspectives. Biofilms formed by yeast and filamentous fungi present differences, and studies of polymicrobial communities have become increasingly important. A key feature of resistance is the extracellular matrix, which covers and protects biofilm cells from the surrounding environment. Furthermore, to achieve cell–cell communication, microorganisms secrete quorum-sensing molecules that control their biological activities and behaviors and play a role in fungal resistance and pathogenicity. Several in vitro techniques have been developed to study fungal biofilms, from colorimetric methods to omics approaches that aim to identify new therapeutic strategies by developing new compounds to combat these microbial communities as well as new diagnostic tools to identify these complex formations in vivo. In this review, recent advances related to pathogenic fungal biofilms are addressed.

Graphical Abstract

1. Introduction

Biofilm formation by microorganisms, particularly bacteria, has been widely studied in recent years. This form of growth prevails in nature compared to planktonic or free cells, and is a cause of concern mainly in the clinic because of increased resistance to antimicrobials and environmental conditions [1,2,3,4]. Biofilms formed by pathogenic fungi have gained attention in recent years and several species among filamentous, yeast, and dimorphic fungi have been described as capable of developing into communities [4,5,6,7,8]. This review aims to discuss the development of biofilms formed by yeast and filamentous fungi, interactions among polymicrobial communities, resistance to commercially available antifungals, and aspects of in vitro and in vivo methodologies and models.

2. Yeasts and Filamentous Fungi Biofilms

Biofilms are sessile microbial communities that strongly adhere to surfaces and to each other and are protected by a polymeric extracellular matrix (ECM) composed primarily of polysaccharides [9,10,11,12]. Cells here exhibit increased resistance and different phenotypes compared to planktonic or free cells and are associated with the persistence of infections [4,13].
Pathogenic fungi can also adhere to abiotic surfaces such as prostheses and catheters; in particular, yeasts take advantage of this condition to gain access to blood circulation, reaching the internal organs of patients. This is alarming, as disseminated fungal infections have a high mortality rate [14].
Both yeast and filamentous fungi can form biofilms; however, studies of filamentous fungal biofilms are limited compared to those of yeasts [12,15]. According to Harding et al. [12], this is because for some time the biofilms formed by filamentous fungi did not fit the previous definitions of biofilms related to bacteria. Thus, the authors proposed a model for biofilm formation by filamentous fungi, suggesting that, despite the distinct morphology, this model was similar to bacterial and yeast biofilm development. The stages of development of filamentous fungi biofilms are described in Figure 1a and include propagule adsorption (I), involving contact of spores, hyphal fragments, or sporangia to a surface; active adhesion (II), in which adhesins are secreted by spores during germination and other reproductive structures; first microcolony formation (III), which involves elongation and hyphal branching, forming a monolayer with the production of extracellular matrix; second microcolony formation or initial maturation (IV), in which compact hyphae networks form in three dimensions, covering by an extracellular matrix, and formation of water channels; final maturation (V), in which fruiting bodies and other survivor structures are formed depending of the fungi; and, finally, the dispersion or planktonic phase (VI), in which conidia and/or hyphae fragments are released, beginning a new cycle. Another peculiarity of filamentous fungi is the secretion of small proteins known as hydrophobins. These proteins are involved in the adhesion of hyphae to hydrophobic surfaces and may be involved in biofilm formation [12,16].
Regarding yeasts, Candida albicans is the most studied model of biofilm formation and shows distinct phases of development that are similar to those of bacterial biofilms [3,12,17]. The development process involves fewer stages of development compared to filamentous fungi and includes the adsorption of yeast cells to a surface (i); followed by initial adhesion (ii), formation of basal layers of yeast with early development of hyphae and extracellular matrix (iii); biofilm maturation containing a significant number of yeast, hyphae, pseudohyphae, extracellular matrix, and water channels that allow the movement of nutrients (iv), and cell dispersion (v) (Figure 1b) [12].
In recent years, studies correlated to fungal biofilms have increased considerably and several species have shown the ability to form these communities. Paracoccidioides brasiliensis is a dimorphic fungus responsible for paracoccidioidomycosis, a systemic mycosis endemic in Latin America. Sardi et al. [8] characterized the biofilms formed by this fungi in the yeast phase and found that in vitro community formation was associated with increased gene expression of adhesins and enzymes such as GP43, enolase, GAPDH, and aspartyl proteinase and decreased in phospholipase expression.
Histoplasma capsulatum biofilm was first described by Pitangui et al. [6]. This fungus also features thermal dimorphism and is the cause of histoplasmosis, a respiratory and systemic mycosis whose evolution depends on the survival and replication of yeast in alveolar macrophages. Therefore, the authors investigated the biofilm formation of two clinical isolates in vitro, as well as their adhesion and internalization to pneumocytes.
Dermatophytes are fungi that invade keratinized tissues producing dermatophytosis, one of the most common dermatomycoses in human and animals [19,20]. Among dermatophytosis, onychomycosis often relapses and involves long, sometimes ineffective treatment. Given this context and the hypothesis of Burkhart et al. [21], which states that biofilm formation by dermatophytes can explain dermatophytomas, Costa-Orlandi et al. [7] confirmed in vitro biofilm formation by two of the most prevalent species worldwide: Trichophyton rubrum and T. mentagrophytes.
With respect to Histoplasma, Paracoccidioides, and Trichophyton, other studies are being carried to characterize these biofilms, either to correlate the communities to the greater resistance to antimicrobials or select probable biomarkers using “omics” approaches (unpublished data).
Additionally, with respect to pathogenic fungi, biofilms formed by Candida spp. have been studied since the mid-1990s. In vitro experiments [3,18,22,23,24,25,26,27,28] are predominant compared to in vivo experiments [29,30] and confirmed the heterogeneity of these biofilms composed of dense layers of yeast blastopores, hyphal, pseudohyphae, and ECM [31,32]. Several genes are involved in the adhesion, ECM production, quorum sensing, and morphogenesis of biofilms, particularly in C. albicans [15,33,34]. In addition, genetic analysis confirmed that both yeasts and hyphae have unique roles in biofilm formation by this species [34]. Paramonova et al. [35] showed that most filamentation is directly related to increasing the compressive force of biofilms, which makes them more resistant to adverse conditions such as vortexing and sonication. In an study of non-Candida albicans Candida, Silva et al. [36] analyzed the differences regarding the formation, morphology, and composition of the ECM of biofilms formed by C. glabrata, C. parapsilosis, and C. tropicalis. Regarding the morphology, some biofilms of C. parapsilosis were composed of both yeast and pseudohyphae, although biofilms formed by other isolates were composed of only yeast cells. Finally, in C. tropicalis, most biofilms were composed only of yeast cells, with few exceptions showing long hyphal filaments, while C. glabrata biofilms contained only yeast cells. With respect to the matrix composition, biofilms showed different amounts of carbohydrates and proteins in the three species tested.
Aspergillus spp. are saprophytic and opportunistic fungi involved in several biotechnological processes because they secrete enzymes, proteins, and metabolites and are involved in severe superficial and systemic pathologies [37,38,39]. Aspergillosis is considered the second major cause of nosocomial infection after C. albicans and shows a high mortality rate [39,40]. In immunocompromised or immunocompetent individuals with previous pulmonary cavities, these fungi may cause aspergilloma, invasive pulmonary aspergillosis, allergic bronchopulmonary aspergillosis, and even systemic dissemination [38,39,41]. Aspergilloma is a fungal mass showing characteristics of biofilms [40]. As with Candida biofilms, the biofilms formed by these filamentous fungi have been extensively studied in recent years and can develop on abiotic surfaces [5,40,42]. A study by Mowat et al. [5] showed that these biofilms reached maturation in 24 h. At maturation, the biomass density was increased and channels developed between hyphae to allow the passage of fluids and nutrients [40,43]. The ECM is composed of α-1,3-glucans, melanin, hydrophobins, galactomannan, monosaccharides, polyols, and antigens [40,44].
Additionally, biofilms formed by several other fungi have been studied including those formed by Cryptococcus spp. [45,46,47,48]; Malassezia spp. [49]; Trichosporon spp. [50]; Fusarium spp. [51,52,53]; Scedosporium spp., Lomentospora prolificans [54]; and Coccidioides spp. [55], among others.

3. Polymicrobial Biofilms

Microbes rarely exist in single-species planktonic forms [56]. Most microorganisms live in complex communities, known as polymicrobial biofilms [11,57]. Similarly to most communities, biofilms are multicultural and well-engineered [58]. Interactions within these biofilms can be mutualistic, commensalistic, or antagonistic and microorganisms have evolved highly defined responses to sense and adapt to neighboring species [59,60].
In terms of human health, polymicrobial biofilms are prevalent throughout the human body, both during healthy and disease conditions [58]. However, the clinical concern regarding the synergies of polymicrobial biofilms is that the infection will be more severe and recalcitrant to treatment [61]. Microbial synergy is a cooperative interaction between two or more species that produces an effect not achieved by an individual species alone [56,62,63,64,65]. These synergistic interactions are more severe than infections with individual microorganisms [62], leading to increased antimicrobial resistance and prolonging the time necessary for host recovery [58].
Genetic diversity of biofilm communities increases the fitness of the residing community, making the species better able to survive to environmental pressures [61,66], resulting in accelerated growth [67], increased stress resistance [68,69], immune evasion [70,71,72,73], passive resistance [74], and metabolic cooperation [75,76]. In bacteria, it was demonstrated that this occurs because of an expanded gene pool that can be shared within the residents of the biofilm community [66,77,78].
Infections related to polymicrobial biofilms are most frequently observed in the urinary tract, lung, inner ear, urinary tract, oral cavity, wounds, and abiotic devices [66,79]. Biofilms at these sites can potentiate infection and induce a chronic inflammatory state, resulting in collateral damage to host tissue. This fact and the structure of biofilms help to protect microbes from antimicrobials, host immunity, and environmental factors [66].
In recent years, through the development of more sophisticated technologies, the understanding of the importance of polymicrobial infection in human fungal disease has increased. Fungal–polymicrobial interactions are important in a variety of disease states and niches including respiratory system infections, formation of dental plaque, invasive disease, skin and mucosal infections, and bloodstream infections [80].
Recent studies have shown that Candida rarely exists as monospecies and can colonize mucosal surfaces and prosthetic materials, such as dentures and catheters, throughout the human body. In addition, polymicrobial communities consisting of aggregates of other fungi and bacteria are highly prevalent and clinically important [81,82].
Oral candidiasis is one of the most well-defined fungal biofilm infections and is characterized by complex biofilms, which interact with bacteria and the host [81,83,84]. The relationship between Candida and streptococci is generally considered to be synergistic, where a streptococcal infection interacts with the hyphal filaments of Candida via cell surface adhesin SspB interacting with the hyphal cell wall protein Als3 [82,85,86,87]. Some studies showed that bacteria can enhance biofilm formation and the pathogenicity of C. albicans [82,88]. In this interaction, streptococci provide Candida with nutrients from the salivary pellicle [89], while Candida promote the survival of streptococci by lowering oxygen tension levels to those more acceptable for streptococcal growth and providing nutrients to stimulate bacterial growth [81].
Biofilms composed of Staphylococcus aureus and Candida albicans have been widely studied, as these two organisms are often found together in different types of infections, where they show enhanced virulence and resistance upon co-infection of hosts [64]. Staphylococcus aureus and Candida spp. are two of the most prevalent bloodstream pathogens and are responsible for severe morbidity and mortality in hospitalized patients. There is some evidence that they are commonly associated as co-infecting organisms [90,91,92,93]. In addition to the bloodstream, C. albicans and S. aureus have been co-isolated from various mucosal surfaces including the vaginal and oral mucosa in a biofilm mode of growth [91,94,95,96]. According to Peters et al. [91], a proteomics approach to identify proteins upregulated during a C. albicans–S. aureus interaction demonstrated that both species could induce a stress response upon their initial interaction, particularly when Candida was still in the yeast form. However, during biofilm maturation, some genes may be downregulated as a survival strategy, enabling survival within the host.
Moreover, although poorly studied, there are some reports of Candida–Candida mixed biofilms. Coco et al. [97] reported the isolation of C. albicans and C. glabrata co-infection from patients with severe inflammation and hypothesized that pathogenic synergy occurred. Further studies confirmed this synergy, in which C. albicans appeared to assist C. glabrata in invading in vitro reconstituted epithelium [98]. In another model corresponding to the human vaginal epithelium, C. glabrata in combination with C. albicans caused significant tissue damage compared to C. glabrata alone [99].
Recently, Martins et al. [26] reported the in vitro formation of a mixed biofilm containing C. albicans and C. rugosa, an emerging fungal pathogen found in Latin America, particularly in Brazil. Candida rugosa shows a lower susceptibility to fluconazole, amphotericin B, and echinocandins and is frequently found in elderly patients with a capacity to form biofilm [100,101,102,103,104,105]. Kirkpatrick et al. [106] also described mixed biofilms formed by C. albicans and C. dubliniensis and showed that C. albicans had a distinct competitive advantage over C. dubliniensis under planktonic growth conditions, while under biofilm growing conditions, C. dubliniensis was able to better withstand the rigorous competitive pressures from C. albicans.
Other fungi species were also described in vitro as part of polymicrobial biofilms. Aspergillus fumigatus and other species of Aspergillus are commonly found to co-colonize with Pseudomonas aeruginosa in the lungs of cystic fibrosis (CF) patients, open skin wounds, and cardiac implants [107]. Manavathu et al. [108] demonstrated in vitro that polymicrobial CF patient airway infection with P. aeruginosa and A. fumigatus produced mixed microbial biofilm with structural and functional characteristics differing from those of monomicrobial biofilms, which is a serious clinical problem in CF patients and other patient groups prone to airway infection with P. aeruginosa and A. fumigatus. Zheng et al. [107] demonstrated that phenazine-derived metabolites produced by P. aeruginosa can act as signals that affect A. fumigatus and A. nidulans development, shifting from weak vegetative growth to induced asexual sporulation (conidiation) along a decreasing phenazine gradient, affecting biofilm formation.
Numerous studies have described co-infections of fungi and bacteria in different diseases. As an example, the cystic fibrosis lung is a major site of polymicrobial infections, with bacteria such as P. aeruginosa, S. aureus, Burkholderia cepacia, Acinetobacter baumanii, Haemophilus influenzae mixed with C. albicans, A. fumigatus, and Scedosporium species [80]. However, additional studies are needed to understand how these interactions occur and to determine the involvement of biofilm communities in these infections.
At other sites of infections, different polymicrobial interactions between fungi and bacteria have been described. As an example, Candida interacting with Streptococcus and Lactobacilli; Porphyromonas gingivalis at oral sites; Candida, Aspergillus, Mucorales, and Fusarium with Pseudomonas and Staphylococcus in burn wounds and trauma sites; Candida and Cryptococcus with a wide range of Gram-negative and Gram-positive bacteria in the lower reproductive tract; Candida with Gram-positive and Gram-negative bacteria (typically Staphylococcus spp.) and interactions between dermatophyte species in the cutaneous site and vascular catheters, Enterobacteriaceae and Enterococcus spp. with Candida spp. in intra-abdominal site and Pseudomonas spp. and, finally, Enterobacteriaceae, Escherichia coli and Enterobacter faecalis with Candida spp. in the urinary tract [80,109]. Gastrointestinal tract and gut interactions between Candida spp. and E. coli, Helicobacter, Serratia marcescens, and Salmonella enterica subsp. enterica serovar Typhimurium have also been reported [80,110].
The increasing number of fungal infections associated with the increase in descriptions of fungal co-infections with bacteria and other microorganisms reveal the importance of further studies of the mechanisms and consequences of polymicrobial biofilm formation.

4. In Vitro Methods to Study Biofilms

4.1. Conventional Methods

Biofilm-associated infections are a serious public health problem because this microenvironment can reduce the efficacy and susceptibility to antifungal agents and avoid the host immune response. Thus, the development of biofilms has been extensively studied. Several in vitro methods are used to evaluation biofilm progression. The main characteristics assessed are cell adhesion, production of ECM, biofilm architecture, mechanism of drug resistance to antifungal agents, cell phenotypes, and for certain fungal species such as Candida albicans, yeast-to-hyphae transition [17,111,112]. The methods described below are employed in studies of both biofilms formed by filamentous fungi and those formed by yeasts.
Colorimetric assays are commonly used in studies of the development and susceptibility of biofilms to antifungal drugs. Hawser and Douglas [24] were pioneers in studies of fungal biofilm formation employing the methyltetrazolium assay (MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-iphenyltetrazolium bromide)). MTT is a yellow soluble salt; in the presence of metabolic activity, the salt is reduced to an insoluble purple formazan crystal. The same authors concluded that the converted MTT was highly correlated with biofilm dry weights and could be applied to evaluate fungal biomass. Since then, this technique has been widely accepted [111,113,114,115]. A disadvantage of this technique was reported by Manavathu et al. [108] when using MTT to determine the effects of antimicrobials on polymicrobial biofilms of A. fumigatus and P. aeruginosa. According to the authors, although this method was useful for monitoring monospecies biofilms of A. fumigatus, it was difficult to differentiate the contribution of each microorganism in the reduction of MTT compound when mixed biofilm was evaluated.
XTT (2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-5-[(phenylamino)carbonyl]-2H-tetrazolium hydroxide) is another tetrazolium salt employed to analyze biofilm development and drug susceptibility. This salt is converted to water-soluble colored formazan salt in the presence of metabolic activity by cellular effectors, such as mitochondrial dehydrogenases [111,116]. Among colorimetric methods, XTT has been the most widely used in recent years [5,6,7,8,46,48,117,118]. XTT is the method of choice employed in susceptibility tests [114]. Compared to MTT, the XTT technique is advantageous because the amount of formazan obtained as a product can be measured directly in the supernatant, while the MTT requires another step involving cell lysis, in which cells must be treated with dimethyl sulfoxide before optical density measurement [111]. However, there are some disadvantages to the use of XTT. Studies of the growth and metabolism of planktonic cells and biofilms of Candida reported that although there the colorimetric signal is proportional to the number of cells, there may be variations when comparing different strains of Candida. The authors stated that there may not be a linear relationship between the number of microorganisms and colorimetric signal, suggesting that this quantification is only valid after constructing a standard curve for each concentration of tetrazolium salt used. In addition, they also stated that significant salt retention may occur when comparing microorganisms in planktonic or biofilm forms.
In addition to MTT and XTT, other assays in microtiter plates for susceptibility testing and biofilm characterization have been explored, such as Alamar blue/resazurin [119,120,121]; safranin [7,42,122]; crystal violet [7,31,119]; Alician blue [123,124], and DMMB (1,9-dimethyl methylene blue) [119,125] (Table 1). Furthermore, scanning and transmission electron microscopes and confocal microscopy have been used to study and detect biofilm cells and ECM [6,7,8,126,127,128].
Scanning electron microscopy (SEM) is a technique in which a sample is prepared by fixation, dehydration, and drying, and the image is processed after coating the samples with a conductor such as gold or carbon under a high vacuum. However, drying and dehydration can alter biofilm morphology because of ECM collapse. Furthermore, artifacts can alter the images. Alternatively, environmental SEM has emerged as a method of choice because the biofilm can be observed without fixation and dehydration and the vacuum is moderate, preserving the morphology and structures of the surfaces [128,129].
Transmission electron microscopy (TEM) is also employed to visualize biofilm architecture. The preparation for TEM is similar to that of SEM. However, in this method, the biofilm is embedded in a resin that allows the ECM to remain stable, unlike in SEM. One disadvantage of TEM is that it is not possible to visualize the biofilm topography [128].
Confocal laser scanning microscopy (CLSM) is another tool used to analyze the biofilm three-dimensional (3D) architecture and thickness. In addition, it is possible to verify the presence of macromolecular compounds, such as polysaccharides, proteins, nucleic acids, and lipids [111,127]. Another advantage is that CLSM can be employed with or without fluorescence or with fluorescence in situ hybridization (FISH) to evaluate alterations of specific compounds of fungal populations over time and spatial relationships [130].
Among other microscopy techniques, which are often costly, scanning transmission X-ray microscopy is associated with near-edge X-ray absorption spectroscopy [127]; CLSM in combination with Raman microscopy (RM) [131]; episcopic differential interference contrast microscopy with and without fluorescence; Hoffman modulation contrast microscopy; and atomic force microscopy [132] have been used to examine biofilms in situ.

4.2. High-Throughput “Omics” Technologies in Biofilms Research

Microbial biofilms involve complex regulatory systems that can be elucidated using “omics” technologies. “Omics” approaches are powerful tools for quantifying differentially global variations between two different biological conditions on a transcriptomic, proteomic, and/or metabolomics scale and targeting for the discovery of novel therapeutics and/or biomarkers in the cell host–fungi interaction [135].
In general, these high-throughput “omics” tools compare two biological systems based on the abundance of messenger RNA transcripts (transcriptomic), proteins (proteomic), and other biomolecular components (metabolomics). These tools began in the so-called post-genomic era and were developed to overcome the limitations associated with previous genomic investigations of organisms in question. Genomic sequencing reflects genetic information through techniques that identify the correct nucleotide sequence in the organism's genome. In this context, aspects such as sequence, number, and syntenia of the genes contained in the nucleus of a cell remain static during the cell cycle; however there is a dynamic equilibrium between: (a) gene transcription, (b) protein translation, and (c) production of metabolic byproducts according to the biological situation to which a cell is subjected, thus defining different transcriptomes, proteomes, and metabolomes for the same cell throughout its cellular differentiation [136].
Therefore, transcriptomics, proteomics, and metabolomics analyses can be employed to determine differences between the transcriptional, translational, and metabolic signatures of a microorganism in biofilms and in planktonic growth. This field of research revealed new concepts; in this context, Azevedo et al. [137] suggested a new trend, so-called “biofomics,” an omics approach to the field of biofilms. The goal of this approach is to gather in an on-line database a large set of omics data generated from studies of a microorganism's ability to adhere to surfaces, communicate with its neighbors, and form biofilms. Such collected data would be freely available to the scientific community and identify a unique biofilm “signature” including important information such as environmental, physiological, and mutational factors that affect the ability of a microorganism to develop biofilms. This may positively impact systems biology and consequently the development of a new diagnostic tool and/or therapeutic for resistance development.
Recent studies using high-throughput omics methods to evaluate biofilms have revealed important information for the development of new therapies for biofilm-based fungi infections. For instance, a recent study showed that C. albicans constitutes the most prevalent and pathogenic species among all Candida species related to Candida bloodstream infections because of the species’ ability to form robust biofilms. Rajendran et al. [138] showed that clinical isolates of candidemia may be stratified as high- or low-biofilm formers (high biofilm formers (HBF) and low biofilm formers (LBF), respectively), resulting in a heterogeneous biofilm phenotype; according to its classification, this directly determines clinical outcomes and mortality. These authors found significant differential gene expression between C. albicans LBF and HBF by comparative RNA-Seq analysis on pre-characterized clinical isolates and emphasized the importance of the aspartate aminotransferase pathway in biofilm formation, which may be exploited as a potential target.
Concomitantly, according to Otto et al. [139], modern proteomic techniques provide a detailed description of the protein inventory of a cell and consequent changes in protein levels through relative or absolute quantification. In this sense, pathogenic fungi such as Candida species and C. neoformans have differential proteomes between planktonic and biofilm cultures as established by several authors [28,140,141,142,143]. However, features of biofilms formed by other pathogenic fungi, such as H. capsulatum, P. brasiliensis, and dermatophytes remain unknown. The findings of our group revealed a substantial difference in the protein profile between H. capsulatum yeasts structured in biofilms and planktonic growth, with more than 40 differentially expressed proteins identified by mass spectrometry. Such proteins are involved in the metabolism of amino acids, nuclear proteins, and translated proteins [10] and should be explored for the development of safer and more effective drugs. In addition, Pires et al. [28] used a proteomics approach to compare the protein expression of C. orthopsilosis planktonic and biofilm forms. According to these findings, differentially expressed proteins were linked to several suitable mechanisms that adjust the catalytic properties of the enzymes.
Additionally, an analysis of the metabolic signature of C. albicans during biofilm development has been published by Zhu et al. [27] who conducted gas chromatography-mass spectrometry and identified 31 differentially produced metabolites between the biofilm and planktonic cells. This study showed even that trehalose is involved in the formation of C. albicans biofilms, as the lack of this metabolite resulted in abnormal biofilm formation and increased the susceptibility of biofilms to the antifungal amphotericin B and miconazole.
In fact, advances in high-throughput DNA and RNA sequencing and mass spectrometric quantification of proteins and metabolites, combined with computational tools, have enabled researchers to obtain large amounts of data generated in these “omics” projects. Various research groups have integrated “omics” datasets. This combination represents much more than the sum of each “omics” analysis, generating data related to interactions that can occur among all classes of molecules in a cell. Thus, an understanding of interactions on different levels, such as genomic, epigenomic, transcriptional, proteomic, post-translational modification, and metabolic, enables the mapping and elucidation of peculiar information regarding the behavior of a cell during the same biological process [144].
Muszkieta et al. [135] integrated three different “omics” methods, microarray, RNA sequencing, and proteomics analysis, to compare the different transcriptomics and proteomics signatures of A. fumigatus biofilms. According to the authors, although utilization of several integrated “omics” methods remains challenging, such a combination provides potential information that answers important biological questions. Therefore, comparison and integration of “omics” approaches in different levels can provide exciting information for characterizing the microbial biofilm signature, such as formation potential, physiological activity, and structure.
In conclusion, although there are challenges in biofilms research using “omics” technologies, these approaches, whether isolated or integrated, provide promising results for future studies targeting novel potential therapeutics and/or biomarkers for the diagnosis of fungi biofilm-associated infections.

5. In Vivo Models to Study Fungal Biofilms

Biofilm formation provides protection to fungal cells (stress, immune system, antifungal drugs). Recently, different techniques based on morphologic and biochemical characterization have become available for evaluating biofilms in vitro. However, biofilms must be studied using complex organisms. The use of in vivo models provides important information regarding the influence of host factors on biofilm formation. Some host factors have been found to affect biofilm formation. (1) Flow conditions: some fungi species can form biofilm under static conditions, while other species produce biofilm under different flow conditions, requiring the production of a large amount of ECM, reflecting increased antifungal resistance [145,146,147]. (2) Substrate: surfaces with certain topography and hydrophobicity conditions favor biofilm formation to provide good adhesion conditions. Medical devices are carefully designed to resist microbe adhesion; however, molecules present in host fluids can promote biofilm formation in these devices. Moreover, biotic surfaces can present receptors that facilitate the cell-microorganism link [147,148,149]. (3) Nutrient conditions: the availability of sugar, proteins, and metal ions impact biofilm characteristics [147,150,151]. (4) Immune system: leukocytes and mononuclear cells interact with biofilm [149,152]; different antibodies also interact with biofilm [153].
The use of medical devices (catheters, dentures, and subcutaneous implants) permits the adhesion of fungal cells and formation of biofilms, making the eradication of infection difficult [154]. Candida spp. are among the most studied yeasts in the context of biofilms. This yeast can grow as biofilm on biotic surfaces during oral, oropharyngeal, and vulvovaginal infection [149,155]. Aspergillus fumigatus can form biofilm during different clinical presentations like sinusitis, pulmonary aspergillosis, and aspergilloma [38]. Based on this, models using medical devices to study in vivo biofilm were developed to better understand biofilm life cycle and treatment. Models using only biotic surfaces have also been described.
The vascular catheter model is one of the most widely used approaches for evaluating in vivo biofilm formation. This model simulates biofilm clinical infections and host conditions such as flow, nutrition, and immune system. In contrast, the surgical process for implanting the catheter is laborious and invasive [147,156]. This in vivo model can be performed in different animal such as mice, rats, and rabbits. The advantage of using mice is their low cost compared to other organisms. However, the surgical procedures are more difficult to execute because of the diameter of the vessels. The rabbit model is easier to manipulate, but more costly [156]. Microorganisms can access vascular catheters through the skin, fluids, medicines, and other avenues, forming biofilm on the intraluminal or extraluminal side of this device [157]. This model was also described in rats and reproduced architectural structures as in vitro models [158]. Martinez et al. [159] used a vascular catheter model in rats to assess the effect of catheters coated with chitosan, a crustacean exoskeleton polymer. Promising results were obtained for the inhibition of Candida spp. biofilm formation and cell viability decrease. The vascular catheter model using rabbits was first described by Schinabeck et al. [160] in a study of biofilm formation and treatment. Amphotericin B liposomal was found to be effective for eliminating biofilms and treating C. albicans infection associated with the catheter. Using the same model, Ghannoum et al. [157] tested the efficacy of micafungin against Candida biofilms, revealing the ability of this antifungal to eradicate the infection and inhibit biofilm growth.
Subcutaneous in vivo biofilm models possess advantages such as the ability to implant more than one device per animal. Compared to other models, the procedure is fast, less aggressive, and requires a shorter anesthesia period [29,147]. In addition, this system can mimic joint prostheses; however, the model suffers from nutrient deprivation because of inconsistent fluid irrigation [161]. A subcutaneous rat model was developed to study C. albicans biofilm; this avascular location of the catheters reduced the experimental procedure and the biofilm extracellular matrix was observed just two days after infection [29]. Candida glabrata cannot produce hypha, which is considered an essential phenomenon for C. albicans biofilm formation [15]. Based on this, Schinabeck et al. [160] demonstrated in a rabbit subcutaneous catheter model that C. glabrata formed biofilm in vivo with half the thickness of the C. albicans biofilm, despite that C. glabrata in vivo biofilm was susceptible to echinocandins but not to fluconazole.
Mucosal tissues exhibit satisfactory conditions for biofilm growth, mainly because they provide a nutrient-rich environment for microbial communities and because of the unique host–microbial interactions that can affect both the host responses and biofilm development, an advantage of this model compared to abiotic surfaces [149]. However, the recognition of many tissue infections as biofilms remains a critical process [149]. Oral candidiasis was found to commonly affect immunosuppressed patients [162]; moreover, denture stomatitis related to polymicrobial biofilm including Candida spp. induces mucosal biofilms. Thus, immunosuppressed animals are used to simulate this model, inoculating the fungi using a swab on the tongue or sublingually [163]. A rat model was demonstrated to be suitable for analysis of in vivo mucosal device-associated infections, particularly given the cost of the animal; moreover, rats show great potential for mimicking denture stomatitis as well as for evaluating mixed microorganism biofilm and the immune response [164]. Mouse experimental oral candidiasis was used to evaluate the effect of photodynamic therapy [165]. In another study, induction of oral candidiasis was performed in mice using a bioluminescent strain of C. albicans to evaluate the action of the polyphenol lichochalcone-A. Longitudinal imaging and histological analysis of mice infected with the bioluminescent strain and after treatment revealed the antifungal efficacy of lichochalcone-A [166]. An oropharyngeal candidiasis model in mouse [163] was used to demonstrate the in vivo efficacy of miltefosine, which was previously used in vitro and efficiently inhibited biofilm formation and for oropharyngeal treatment [167].
The incidence of urinary catheter-associated infection is high during hospital confinement. To study the involvement of Candida spp. biofilm in this infection, a urinary in vivo biofilm model was developed. Rat and mice were used as models to study Candida spp. biofilm characteristics as well as the importance of the biofilm in the persistence of the candiduria, biofilm antifungal efficacy, mutant phenotypes, and new anti-adhesive materials for catheters [168,169].
Fusarium spp. can form biofilms on contact lenses, which is a risk factor for the development of keratitis. Sun et al. [170] developed an in vivo model for contact lenses containing Fusarium spp. biofilm, which were applied to the mouse cornea. Important features of Fusarium spp. biofilm formation in vivo and the immune response were elucidated. Taking a different approach, Pinnock et al. [171] used ex vivo corneas from both rabbits and humans to study C. albicans and F. solani biofilms as alternatives to in vitro or in vivo models to study keratitis. Another approach was developed to simulate Fusarium spp. keratitis by direct application to the cornea without contact lenses in a murine model using fluorescent staining. This simple method enables detection of infection in early stages [172].
Despite the use of these mammalian models for studying in vivo biofilm formation and treatment, the use of invertebrate models to determine biofilm characteristics has recently been encouraged because of ethical issues related to mammalians models. The use of Galleria mellonella, an insect model, revealed information regarding the virulence of Cryptococcus spp. planktonic and biofilm cells, demonstrating that cells originating from biofilm killed G. mellonella larvae faster than planktonic cells [173]. The effectiveness of acetylcholine for inhibiting C. albicans biofilm was studied in the G. mellonella model [174]. In addition, the in vivo virulence of clinical strains of C. albicans capable of forming biofilms was evaluated in G. mellonella, revealing the strong virulence of biofilm-producing strains [175].
Significant advances have been made in understanding biofilms and in their therapy by using in vivo models. However, many aspects of biofilms remain unclear and in vivo studies are fundamental for understanding biofilm–host interactions and developing anti-biofilm compounds/strategies.

6. Physical and Molecular Resistance in Fungal Biofilms

Among the defining characteristics of biofilms are their high resistance to antimicrobial agents and production of ECM. The matrix protects and envelops the entire biofilm, providing an ideal structure for cell cohesion and adhesion [176]. The ECM also retains water and nutrients derived from matrix materials hydrolyzed by enzymes produced by microorganisms [176].
The most medically relevant function of the extracellular matrix is its ability to provide a physical barrier between biofilm cells and immune system and often drugs used for treatment [9,154,177]. Fungus biofilms were reported to be up to 1000-fold more resistant to antifungal agents than planktonic cells, but the mechanism of this resistance remains unclear [10,123].
Antifungal resistance is very complex and multifactorial. It may be inducible in response to a drug, biochemical alterations, or an irreversible genetic change resulting from prolonged exposure (Table 2). Specifically, alterations or overexpression of target molecules, active extrusion through efflux pumps, limited diffusion, tolerance and cell density (quorum sensing), and the ECM are all characterized mechanisms used by fungi to combat the effects of antifungal treatment (Figure 2).
Planktonic cells generally depend on irreversible genetic changes to maintain a resistant phenotype, while biofilms persist because of their physical presence and population density, providing a nearly inducible resistant phenotype regardless of the defined genetic changes [10,187,188]. Further, factors including pH, temperature, oxygen availability, and other environmental stresses alter the biofilm architecture and possibly antifungal susceptibility [189].
A defining characteristic of a biofilm is its ECM, which is self-produced and may contain proteins, polysaccharides, lipids, nucleic acids, and other molecules [176,190] that can interact with each other and with the cell surface to form a robust and protective network [191]. ECM composition varies across species and even growth conditions [176]; however, the ECM composition of many biofilms remains unknown [176]. Functionally, the ECM can serve as a protective barrier against chemical and biological antimicrobial agents, including many prescribed antifungal drugs [178,190]. In some instances, the ECM can contribute to antifungal resistance by binding to antifungals, thereby preventing access to their intended target at the surface or within fungal cells [192].
Quorum sensing is related to the mechanism by which microorganisms communicate and coordinate their behavior through the secretion of signaling molecules [193,194]. Cells respond to these quorum-sensing molecules (QSMs) through the expression or repression of quorum-dependent target genes [10,195]. QSMs play a role in several mechanisms, including biofilm development, morphogenesis, and limitation of cell population, among others. They are also important during the infectious process, particularly for dissemination [194,196]. Farnesol was described for the first time in C. albicans by Hornby et al. [197] as a QSM. Contact between C. albicans and exogenous farnesol results in several responses, including activation of genes involved in drug resistance (CaFCR1 and CaPDR16) [189,198]. Another study conducted by Sharma et al. [199] demonstrated that farnesol modulated the action of drugs in C. albicans planktonic cells. In addition, Ramage et al. [200] concluded that farnesol inhibits hyphae development during the initial phase of biofilm formation, compromising the structure. Farnesol also affects many other microorganisms such as S. aureus, S. cerevisiae, Aspergillus spp., P. brasiliensis, and Mycobacterium smegmatis [201,202]. The detection of QSMs is of fundamental importance, as they have specific roles in biofilm physiology (Table 3).
Persisters represent a small subpopulation of cells that spontaneously enter a dormant, non-dividing state. When a population is treated with an antimicrobial, normal cells die, while persisters survive. When therapy is discontinued, the persistent cells can restore the biofilms, thus explaining why biofilm infections are recurrent [203,204].
The molecular mechanisms that promote antifungal resistance in fungal biofilms are not completely understood. Some studies have shown that efflux pumps contribute to azole resistance only during the early phase of biofilm formation. In addition, membrane sterol composition contributes to azole resistance, but this occurs during the intermediate and mature phases [205]. According to Soto [206], the upregulation of drug efflux pumps also causes drug resistance in several biofilm-forming microorganisms. These pumps are divided into two groups; the first is linked to ATP-binding cassette transporters encoded by CDR-genes, while the second is composed of the major facilitator superfamily encoded by MDR-genes [178]. Activation of drug efflux pumps occurs through the “expulsion” of antifungal drugs after contact with the biofilm. The increases the expression of the CDR1 and MDR1 genes, in which has been correlated to the resistance of yeast to azole drugs. Some data regarding biofilm-associated resistance have shown that the expression of efflux pump genes is increased during the first hours of biofilm formation [178].

7. Conclusions

Biofilm formation by saprophytic and pathogenic fungi is of great concern. Despite recent advances in the study of biofilms formed by species in the genus Candida, Cryptococcus, and Aspergillus, reports of biofilm formation by other species are increasing. Therefore, an understanding of the molecular mechanisms and key factors involved in establishing these infections is necessary. In addition, interactions between biofilms of polymicrobial origin and the host should be prioritized in studies, particularly those of recently described biofilms. Finally, the discovery of new treatment alternatives capable of controlling or destroying these microbial communities is essential.


We thank the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) 150261/2016-0 (CBCO), the Fundação de Apoio à Pesquisa do Estado de São Paulo-FAPESP [2013/10917-9 (LS), 2015/03700-9 (MJSMG), 2015/14023-8 (HCO), 2016/11836-0 (AMFA)], the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES), and the Programa de Apoio ao Desenvolvimento Científico (PADC-FCFAR UNESP) for financial support.

Author Contributions

Caroline B. Costa-Orlandi, Janaina C. O. Sardi, Nayla S. Pitangui, Haroldo C. de Oliveira, Liliana Scorzoni, Mariana C. Galeane, Kaila P. Medina-Alarcón, Wanessa C. M. A. Melo, Mônica Y. Marcelino, Jaqueline D. Braz, Ana Marisa Fusco-Almeida and Maria José S. Mendes-Giannini contributed substantially to the manuscript.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Lopez-Ribot, J.L. Candida albicans biofilms: More than filamentation. Curr. Biol. 2005, 15, R453–R455. [Google Scholar] [CrossRef] [PubMed]
  2. Donlan, R.M. Biofilms: Microbial life on surfaces. Emerg. Infect. Dis. 2002, 8, 881–890. [Google Scholar] [CrossRef] [PubMed]
  3. Chandra, J.; Kuhn, D.M.; Mukherjee, P.K.; Hoyer, L.L.; McCormick, T.; Ghannoum, M.A. Biofilm formation by the fungal pathogen Candida albicans: Development, architecture, and drug resistance. J. Bacteriol. 2001, 183, 5385–5394. [Google Scholar] [CrossRef] [PubMed]
  4. Fanning, S.; Mitchell, A.P. Fungal biofilms. PLoS Pathog. 2012, 8, e1002585. [Google Scholar] [CrossRef] [PubMed]
  5. Mowat, E.; Butcher, J.; Lang, S.; Williams, C.; Ramage, G. Development of a simple model for studying the effects of antifungal agents on multicellular communities of Aspergillus fumigatus. J. Med. Microbiol. 2007, 56, 1205–1212. [Google Scholar] [CrossRef] [PubMed]
  6. Pitangui, N.S.; Sardi, J.C.; Silva, J.F.; Benaducci, T.; Moraes da Silva, R.A.; Rodriguez-Arellanes, G.; Taylor, M.L.; Mendes-Giannini, M.J.; Fusco-Almeida, A.M. Adhesion of Histoplasma capsulatum to pneumocytes and biofilm formation on an abiotic surface. Biofouling 2012, 28, 711–718. [Google Scholar] [CrossRef] [PubMed]
  7. Costa-Orlandi, C.B.; Sardi, J.C.; Santos, C.T.; Fusco-Almeida, A.M.; Mendes-Giannini, M.J. In vitro characterization of Trichophyton rubrum and T. mentagrophytes biofilms. Biofouling 2014, 30, 719–727. [Google Scholar] [CrossRef] [PubMed]
  8. Sardi Jde, C.; Pitangui Nde, S.; Voltan, A.R.; Braz, J.D.; Machado, M.P.; Fusco Almeida, A.M.; Mendes Giannini, M.J. In vitro Paracoccidioides brasiliensis biofilm and gene expression of adhesins and hydrolytic enzymes. Virulence 2015, 6, 642–651. [Google Scholar] [CrossRef] [PubMed]
  9. Costerton, J.W.; Stewart, P.S.; Greenberg, E.P. Bacterial biofilms: A common cause of persistent infections. Science 1999, 284, 1318–1322. [Google Scholar] [CrossRef] [PubMed]
  10. Sardi Jde, C.; Pitangui Nde, S.; Rodriguez-Arellanes, G.; Taylor, M.L.; Fusco-Almeida, A.M.; Mendes-Giannini, M.J. Highlights in pathogenic fungal biofilms. Rev. Iberoam. Micol. 2014, 31, 22–29. [Google Scholar] [CrossRef] [PubMed]
  11. Donlan, R.M.; Costerton, J.W. Biofilms: Survival mechanisms of clinically relevant microorganisms. Clin. Microbiol. Rev. 2002, 15, 167–193. [Google Scholar] [CrossRef] [PubMed]
  12. Harding, M.W.; Marques, L.L.; Howard, R.J.; Olson, M.E. Can filamentous fungi form biofilms? Trends Microbiol. 2009, 17, 475–480. [Google Scholar] [CrossRef] [PubMed]
  13. Martinez, L.R.; Fries, B.C. Fungal biofilms: Relevance in the setting of human disease. Curr. Fungal Infect. Rep. 2010, 4, 266–275. [Google Scholar] [CrossRef] [PubMed]
  14. Verstrepen, K.J.; Klis, F.M. Flocculation, adhesion and biofilm formation in yeasts. Mol. Microbiol. 2006, 60, 5–15. [Google Scholar] [CrossRef] [PubMed]
  15. Blankenship, J.R.; Mitchell, A.P. How to build a biofilm: A fungal perspective. Curr. Opin. Microbiol. 2006, 9, 588–594. [Google Scholar] [CrossRef] [PubMed]
  16. Wosten, H.A. Hydrophobins: Multipurpose proteins. Annu. Rev. Microbiol. 2001, 55, 625–646. [Google Scholar] [CrossRef] [PubMed]
  17. Nett, J.; Andes, D. Candida albicans biofilm development, modeling a host-pathogen interaction. Curr. Opin. Microbiol. 2006, 9, 340–345. [Google Scholar] [CrossRef] [PubMed]
  18. Pires, R.H.; Montanari, L.B.; Martins, C.H.; Zaia, J.E.; Almeida, A.M.; Matsumoto, M.T.; Mendes-Giannini, M.J. Anticandidal efficacy of cinnamon oil against planktonic and biofilm cultures of Candida parapsilosis and Candida orthopsilosis. Mycopathologia 2011, 172, 453–464. [Google Scholar] [CrossRef] [PubMed]
  19. Weitzman, I.; Summerbell, R.C. The dermatophytes. Clin. Microbiol. Rev. 1995, 8, 240–259. [Google Scholar] [CrossRef]
  20. Costa-Orlandi, C.B.; Magalhães, G.M.; Oliveira, M.B.; Taylor, E.L.; Marques, C.R.; de Resende-Stoianoff, M.A. Prevalence of dermatomycosis in a Brazilian tertiary care hospital. Mycopathologia 2012, 174, 489–497. [Google Scholar] [CrossRef] [PubMed]
  21. Burkhart, C.N.; Burkhart, C.G.; Gupta, A.K. Dermatophytoma: Recalcitrance to treatment because of existence of fungal biofilm. J. Am. Acad. Dermatol. 2002, 47, 629–631. [Google Scholar] [CrossRef] [PubMed]
  22. Martins, M.; Henriques, M.; Lopez-Ribot, J.L.; Oliveira, R. Addition of DNase improves the in vitro activity of antifungal drugs against Candida albicans biofilms. Mycoses 2012, 55, 80–85. [Google Scholar] [CrossRef] [PubMed]
  23. Kuhn, D.M.; George, T.; Chandra, J.; Mukherjee, P.K.; Ghannoum, M.A. Antifungal susceptibility of Candida biofilms: Unique efficacy of amphotericin B lipid formulations and echinocandins. Antimicrob. Agents Chemother. 2002, 46, 1773–1780. [Google Scholar] [CrossRef] [PubMed]
  24. Hawser, S.P.; Douglas, L.J. Biofilm formation by Candida species on the surface of catheter materials in vitro. Infect. Immun. 1994, 62, 915–921. [Google Scholar] [PubMed]
  25. Marcos-Zambrano, L.J.; Gomez-Perosanz, M.; Escribano, P.; Zaragoza, O.; Bouza, E.; Guinea, J. Biofilm production and antibiofilm activity of echinocandins and liposomal amphotericin B in echinocandin-resistant yeast species. Antimicrob. Agents Chemother. 2016, 60, 3579–3586. [Google Scholar] [CrossRef] [PubMed]
  26. Martins, C.H.; Pires, R.H.; Cunha, A.O.; Pereira, C.A.; Singulani, J.L.; Abrão, F.; Moraes, T.; Mendes-Giannini, M.J. Candida/candida biofilms. First description of dual-species Candida albicans/C. rugosa biofilm. Fungal Biol. 2016, 120, 530–537. [Google Scholar] [CrossRef] [PubMed]
  27. Zhu, Z.; Wang, H.; Shang, Q.; Jiang, Y.; Cao, Y.; Chai, Y. Time course analysis of Candida albicans metabolitesduring biofilm development. J. Proteome Res. 2013, 12, 2375–2385. [Google Scholar] [CrossRef] [PubMed]
  28. Pires, R.H.; Cataldi, T.R.; Franceschini, L.M.; Labate, M.V.; Fusco-Almeida, A.M.; Labate, C.A.; Palma, M.S.; Soares Mendes-Giannini, M.J. Metabolic profiles of planktonic and biofilm cells of Candida orthopsilosis. Future Microbiol. 2016, 11, 1299–1313. [Google Scholar] [CrossRef] [PubMed]
  29. Ricicová, M.; Kucharíková, S.; Tournu, H.; Hendrix, J.; Bujdáková, H.; Van Eldere, J.; Lagrou, K.; van Dijck, P. Candida albicans biofilm formation in a new in vivo rat model. Microbiology 2010, 156, 909–919. [Google Scholar] [CrossRef] [PubMed]
  30. Kucharikova, S.; Vande Velde, G.; Himmelreich, U.; van Dijck, P. Candida albicans biofilm development on medically-relevant foreign bodies in a mouse subcutaneous model followed by bioluminescence imaging. J. Vis. Exp. 2015, 52239. [Google Scholar] [CrossRef] [PubMed]
  31. Henriques, M.; Azeredo, J.; Oliveira, R. Candida albicans and Candida dubliniensis: Comparison of biofilm formation in terms of biomass and activity. Br. J. Biomed. Sci. 2006, 63, 5–11. [Google Scholar] [CrossRef] [PubMed]
  32. Pires, R.H.; Santos, J.M.; Zaia, J.E.; Martins, C.H.; Mendes-Giannini, M.J. Candida parapsilosis complex water isolates from a haemodialysis unit: Biofilm production and in vitro evaluation of the use of clinical antifungals. Memórias Inst. Oswaldo Cruz 2011, 106, 646–654. [Google Scholar] [CrossRef]
  33. Bonhomme, J.; d’Enfert, C. Candida albicans biofilms: Building a heterogeneous, drug-tolerant environment. Curr. Opin. Microbiol. 2013, 16, 398–403. [Google Scholar] [CrossRef] [PubMed]
  34. Finkel, J.S.; Mitchell, A.P. Genetic control of Candida albicans biofilm development. In Nature Reviews. Microbiology; Nature Publishing Group: London, UK, 2011; Volume 9, pp. 109–118. [Google Scholar]
  35. Paramonova, E.; Krom, B.P.; van der Mei, H.C.; Busscher, H.J.; Sharma, P.K. Hyphal content determines the compression strength of Candida albicans biofilms. Microbiology 2009, 155, 1997–2003. [Google Scholar] [CrossRef] [PubMed]
  36. Silva, S.; Henriques, M.; Martins, A.; Oliveira, R.; Williams, D.; Azeredo, J. Biofilms of non-Candida albicans Candida species: Quantification, structure and matrix composition. Med. Mycol. 2009, 47, 681–689. [Google Scholar] [CrossRef] [PubMed]
  37. Gonzalez-Ramirez, A.I.; Ramirez-Granillo, A.; Medina-Canales, M.G.; Rodriguez-Tovar, A.V.; Martinez-Rivera, M.A. Analysis and description of the stages of Aspergillus fumigatus biofilm formation using scanning electron microscopy. BMC Microbiol. 2016, 16, 243. [Google Scholar] [CrossRef] [PubMed]
  38. Müller, F.M.; Seidler, M.; Beauvais, A. Aspergillus fumigatus biofilms in the clinical setting. Med. Mycol. 2011, 49, S96–S100. [Google Scholar] [CrossRef] [PubMed]
  39. Kaur, S.; Singh, S. Biofilm formation by Aspergillus fumigatus. Med. Mycol. 2014, 52, 2–9. [Google Scholar] [PubMed]
  40. Ramage, G.; Rajendran, R.; Gutierrez-Correa, M.; Jones, B.; Williams, C. Aspergillus biofilms: Clinical and industrial significance. FEMS Microbiol. Lett. 2011, 324, 89–97. [Google Scholar] [CrossRef] [PubMed]
  41. Williams, C.; Rajendran, R.; Ramage, G. Aspergillus biofilms in human disease. Adv. Exp. Med. Biol. 2016, 931, 1–11. [Google Scholar] [PubMed]
  42. Seidler, M.J.; Salvenmoser, S.; Muller, F.M. Aspergillus fumigatus forms biofilms with reduced antifungal drug susceptibility on bronchial epithelial cells. Antimicrob. Agents Chemother. 2008, 52, 4130–4136. [Google Scholar] [CrossRef] [PubMed]
  43. Villena, G.K.; Fujikawa, T.; Tsuyumu, S.; Gutierrez-Correa, M. Structural analysis of biofilms and pellets of Aspergillus niger by confocal laser scanning microscopy and cryo scanning electron microscopy. Bioresour. Technol. 2010, 101, 1920–1926. [Google Scholar] [CrossRef] [PubMed]
  44. Beauvais, A.; Schmidt, C.; Guadagnini, S.; Roux, P.; Perret, E.; Henry, C.; Paris, S.; Mallet, A.; Prevost, M.C.; Latge, J.P. An extracellular matrix glues together the aerial-grown hyphae of Aspergillus fumigatus. Cell. Microbiol. 2007, 9, 1588–1600. [Google Scholar] [CrossRef] [PubMed]
  45. Ajesh, K.; Sreejith, K. Cryptococcus laurentii biofilms: Structure, development and antifungal drug resistance. Mycopathologia 2012, 174, 409–419. [Google Scholar] [CrossRef] [PubMed]
  46. Martinez, L.R.; Casadevall, A. Susceptibility of Cryptococcus neoformans biofilms to antifungal agents in vitro. Antimicrob. Agents Chemother. 2006, 50, 1021–1033. [Google Scholar] [CrossRef] [PubMed]
  47. Pettit, R.K.; Repp, K.K.; Hazen, K.C. Temperature affects the susceptibility of Cryptococcus neoformans biofilms to antifungal agents. Med. Mycol. 2010, 48, 421–426. [Google Scholar] [CrossRef] [PubMed]
  48. Martinez, L.R.; Mihu, M.R.; Han, G.; Frases, S.; Cordero, R.J.; Casadevall, A.; Friedman, A.J.; Friedman, J.M.; Nosanchuk, J.D. The use of chitosan to damage Cryptococcus neoformans biofilms. Biomaterials 2010, 31, 669–679. [Google Scholar] [CrossRef] [PubMed]
  49. Bumroongthai, K.; Chetanachan, P.; Niyomtham, W.; Yurayart, C.; Prapasarakul, N. Biofilm production and antifungal susceptibility of co-cultured Malassezia pachydermatis and Candida parapsilosis isolated from canine seborrheic dermatitis. Med. Mycol. 2016, 54, 544–549. [Google Scholar] [CrossRef] [PubMed]
  50. Iturrieta-Gonzalez, I.A.; Padovan, A.C.; Bizerra, F.C.; Hahn, R.C.; Colombo, A.L. Multiple species of trichosporon produce biofilms highly resistant to triazoles and amphotericin B. PLoS ONE 2014, 9, e109553. [Google Scholar] [CrossRef] [PubMed]
  51. Imamura, Y.; Chandra, J.; Mukherjee, P.K.; Lattif, A.A.; Szczotka-Flynn, L.B.; Pearlman, E.; Lass, J.H.; O’Donnell, K.; Ghannoum, M.A. Fusarium and Candida albicans biofilms on soft contact lenses: Model development, influence of lens type, and susceptibility to lens care solutions. Antimicrob. Agents Chemother. 2008, 52, 171–182. [Google Scholar] [CrossRef] [PubMed]
  52. Mukherjee, P.K.; Chandra, J.; Yu, C.; Sun, Y.; Pearlman, E.; Ghannoum, M.A. Characterization of fusarium keratitis outbreak isolates: Contribution of biofilms to antimicrobial resistance and pathogenesis. Investig. Ophthalmol. Vis. Sci. 2012, 53, 4450–4457. [Google Scholar] [CrossRef] [PubMed]
  53. Peiqian, L.; Xiaoming, P.; Huifang, S.; Jingxin, Z.; Ning, H.; Birun, L. Biofilm formation by Fusarium oxysporum f. Sp. Cucumerinum and susceptibility to environmental stress. FEMS Microbiol. Lett. 2013, 350, 138–145. [Google Scholar] [CrossRef] [PubMed]
  54. Mello, T.P.; Aor, A.C.; Goncalves, D.S.; Seabra, S.H.; Branquinha, M.H.; Santos, A.L. Assessment of biofilm formation by Scedosporium apiospermum, S. aurantiacum, S. minutisporum and Lomentospora prolificans. Biofouling 2016, 32, 737–749. [Google Scholar] [CrossRef] [PubMed]
  55. Davis, L.E.; Cook, G.; Costerton, J.W. Biofilm on ventriculo-peritoneal shunt tubing as a cause of treatment failure in coccidioidal meningitis. Emerg. Infect. Dis. 2002, 8, 376–379. [Google Scholar] [CrossRef] [PubMed]
  56. Peters, B.M.; Jabra-Rizk, M.A.; O’May, G.A.; Costerton, J.W.; Shirtliff, M.E. Polymicrobial interactions: Impact on pathogenesis and human disease. Clin. Microbiol. Rev. 2012, 25, 193–213. [Google Scholar] [CrossRef] [PubMed]
  57. Galván, E.M.; Mateyca, C.; Ielpi, L. Role of interspecies interactions in dual-species biofilms developed in vitro by uropathogens isolated from polymicrobial urinary catheter-associated bacteriuria. Biofouling 2016, 32, 1067–1077. [Google Scholar] [CrossRef] [PubMed]
  58. Stacy, A.; McNally, L.; Darch, S.E.; Brown, S.P.; Whiteley, M. The biogeography of polymicrobial infection. Nat. Rev. Microbiol. 2016, 14, 93–105. [Google Scholar] [CrossRef] [PubMed]
  59. Hibbing, M.E.; Fuqua, C.; Parsek, M.R.; Peterson, S.B. Bacterial competition: Surviving and thriving in the microbial jungle. Nat. Rev. Microbiol. 2010, 8, 15–25. [Google Scholar] [CrossRef] [PubMed]
  60. Willems, H.M.; Xu, Z.; Peters, B.M. Polymicrobial biofilm studies: From basic science to biofilm control. Curr. Oral Health Rep. 2016, 3, 36–44. [Google Scholar] [CrossRef] [PubMed]
  61. Wolcott, R.; Costerton, J.W.; Raoult, D.; Cutler, S.J. The polymicrobial nature of biofilm infection. Clin. Microbiol. Infect. 2013, 19, 107–112. [Google Scholar] [CrossRef] [PubMed]
  62. Murray, J.L.; Connell, J.L.; Stacy, A.; Turner, K.H.; Whiteley, M. Mechanisms of synergy in polymicrobial infections. J. Microbiol. 2014, 52, 188–199. [Google Scholar] [CrossRef] [PubMed]
  63. Rendueles, O.; Ghigo, J.M. Multi-species biofilms: How to avoid unfriendly neighbors. FEMS Microbiol. Rev. 2012, 36, 972–989. [Google Scholar] [CrossRef] [PubMed]
  64. Burmølle, M.; Ren, D.; Bjarnsholt, T.; Sørensen, S.J. Interactions in multispecies biofilms: Do they actually matter? Trends Microbiol. 2014, 22, 84–91. [Google Scholar] [CrossRef] [PubMed]
  65. Short, F.L.; Murdoch, S.L.; Ryan, R.P. Polybacterial human disease: The ills of social networking. Trends Microbiol. 2014, 22, 508–516. [Google Scholar] [CrossRef] [PubMed]
  66. Gabrilska, R.A.; Rumbaugh, K.P. Biofilm models of polymicrobial infection. Future Microbiol. 2015, 10, 1997–2015. [Google Scholar] [CrossRef] [PubMed]
  67. Jemielita, M.; Taormina, M.J.; Burns, A.R.; Hampton, J.S.; Rolig, A.S.; Guillemin, K.; Parthasarathy, R. Spatial and temporal features of the growth of a bacterial species colonizing the zebrafish gut. mBio 2014, 5. [Google Scholar] [CrossRef] [PubMed]
  68. Monier, J.M.; Lindow, S.E. Differential survival of solitary and aggregated bacterial cells promotes aggregate formation on leaf surfaces. Proc. Natl. Acad. Sci. USA 2003, 100, 15977–15982. [Google Scholar] [CrossRef] [PubMed]
  69. Connell, J.L.; Wessel, A.K.; Parsek, M.R.; Ellington, A.D.; Whiteley, M.; Shear, J.B. Probing prokaryotic social behaviors with bacterial “Lobster traps”. mBio 2010, 1, e00202–e00210. [Google Scholar] [CrossRef] [PubMed]
  70. Blanchette-Cain, K.; Hinojosa, C.A.; Akula Suresh Babu, R.; Lizcano, A.; Gonzalez-Juarbe, N.; Munoz-Almagro, C.; Sanchez, C.J.; Bergman, M.A.; Orihuela, C.J. Streptococcus pneumoniae biofilm formation is strain dependent, multifactorial, and associated with reduced invasiveness and immunoreactivity during colonization. mBio 2013, 4, e00745-13. [Google Scholar] [CrossRef] [PubMed]
  71. Guggenberger, C.; Wolz, C.; Morrissey, J.A.; Heesemann, J. Two distinct coagulase-dependent barriers protect Staphylococcus aureus from neutrophils in a three dimensional in vitro infection model. PLoS Pathog. 2012, 8, e1002434. [Google Scholar] [CrossRef] [PubMed]
  72. Kragh, K.N.; Alhede, M.; Jensen, P.; Moser, C.; Scheike, T.; Jacobsen, C.S.; Seier Poulsen, S.; Eickhardt-Sørensen, S.R.; Trøstrup, H.; Christoffersen, L.; et al. Polymorphonuclear leukocytes restrict growth of Pseudomonas aeruginosa in the lungs of cystic fibrosis patients. Infect. Immun. 2014, 82, 4477–4486. [Google Scholar] [CrossRef] [PubMed]
  73. 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 2015, 17, 21–31. [Google Scholar] [CrossRef] [PubMed]
  74. Weimer, K.E.; Juneau, R.A.; Murrah, K.A.; Pang, B.; Armbruster, C.E.; Richardson, S.H.; Swords, W.E. Divergent mechanisms for passive pneumococcal resistance to β-lactam antibiotics in the presence of haemophilus influenzae. J. Infect. Dis. 2011, 203, 549–555. [Google Scholar] [CrossRef] [PubMed]
  75. Elias, S.; Banin, E. Multi-species biofilms: Living with friendly neighbors. FEMS Microbiol. Rev. 2012, 36, 990–1004. [Google Scholar] [CrossRef] [PubMed]
  76. Fischbach, M.A.; Sonnenburg, J.L. Eating for two: How metabolism establishes interspecies interactions in the gut. Cell Host Microbe 2011, 10, 336–347. [Google Scholar] [CrossRef] [PubMed]
  77. Tuttle, M.S.; Mostow, E.; Mukherjee, P.; Hu, F.Z.; Melton-Kreft, R.; Ehrlich, G.D.; Dowd, S.E.; Ghannoum, M.A. Characterization of bacterial communities in venous insufficiency wounds by use of conventional culture and molecular diagnostic methods. J. Clin. Microbiol. 2011, 49, 3812–3819. [Google Scholar] [CrossRef] [PubMed]
  78. Ehrlich, G.D.; Hu, F.Z.; Shen, K.; Stoodley, P.; Post, J.C. Bacterial plurality as a general mechanism driving persistence in chronic infections. Clin. Orthop. Relat. Res. 2005, 20–24. [Google Scholar] [CrossRef]
  79. Kolenbrander, P.E.; Palmer, R.J.; Periasamy, S.; Jakubovics, N.S. Oral multispecies biofilm development and the key role of cell-cell distance. Nat. Rev. Microbiol. 2010, 8, 471–480. [Google Scholar] [CrossRef] [PubMed]
  80. Dixon, E.F.; Hall, R.A. Noisy neighbourhoods: Quorum sensing in fungal-polymicrobial infections. Cell. Microbiol. 2015, 17, 1431–1441. [Google Scholar] [CrossRef] [PubMed]
  81. O’Donnell, L.E.; Millhouse, E.; Sherry, L.; Kean, R.; Malcolm, J.; Nile, C.J.; Ramage, G. Polymicrobial Candida biofilms: Friends and foe in the oral cavity. FEMS Yeast Res. 2015, 15, fov077. [Google Scholar] [CrossRef] [PubMed]
  82. Jack, A.A.; Daniels, D.E.; Jepson, M.A.; Vickerman, M.M.; Lamont, R.J.; Jenkinson, H.F.; Nobbs, A.H. Streptococcus gordonii comcde (competence) operon modulates biofilm formation with Candida albicans. Microbiology 2015, 161, 411–421. [Google Scholar] [CrossRef] [PubMed]
  83. Rautemaa, R.; Ramage, G. Oral candidosis—Clinical challenges of a biofilm disease. Crit. Rev. Microbiol. 2011, 37, 328–336. [Google Scholar] [CrossRef] [PubMed]
  84. Dongari-Bagtzoglou, A.; Kashleva, H.; Dwivedi, P.; Diaz, P.; Vasilakos, J. Characterization of mucosal Candida albicans biofilms. PLoS ONE 2009, 4, e7967. [Google Scholar] [CrossRef] [PubMed]
  85. Dutton, L.C.; Jenkinson, H.F.; Lamont, R.J.; Nobbs, A.H. Role of Candida albicans secreted aspartyl protease Sap9 in interkingdom biofilm formation. Pathog. Dis. 2016, 74, ftw005. [Google Scholar] [CrossRef] [PubMed]
  86. Nobbs, A.H.; Vickerman, M.M.; Jenkinson, H.F. Heterologous expression of Candida albicans cell wall-associated adhesins in saccharomyces cerevisiae reveals differential specificities in adherence and biofilm formation and in binding oral Streptococcus gordonii. Eukaryot. Cell 2010, 9, 1622–1634. [Google Scholar] [CrossRef] [PubMed]
  87. Silverman, R.J.; Nobbs, A.H.; Vickerman, M.M.; Barbour, M.E.; Jenkinson, H.F. Interaction of Candida albicans cell wall Als3 protein with Streptococcus gordonii SspB adhesin promotes development of mixed-species communities. Infect. Immun. 2010, 78, 4644–4652. [Google Scholar] [CrossRef] [PubMed]
  88. Xu, H.; Sobue, T.; Thompson, A.; Xie, Z.; Poon, K.; Ricker, A.; Cervantes, J.; Diaz, P.I.; Dongari-Bagtzoglou, A. Streptococcal co-infection augments Candida pathogenicity by amplifying the mucosal inflammatory response. Cell. Microbiol. 2014, 16, 214–231. [Google Scholar] [CrossRef] [PubMed]
  89. Holmes, A.R.; van der Wielen, P.; Cannon, R.D.; Ruske, D.; Dawes, P. Candida albicans binds to saliva proteins selectively adsorbed to silicone. Oral Surg. Oral Med. Oral Pathol. Oral Radiol. Endodontol. 2006, 102, 488–494. [Google Scholar] [CrossRef] [PubMed]
  90. Zago, C.E.; Silva, S.; Sanitá, P.V.; Barbugli, P.A.; Dias, C.M.; Lordello, V.B.; Vergani, C.E. Dynamics of biofilm formation and the interaction between Candida albicans and methicillin-susceptible (MSSA) and -resistant Staphylococcus aureus (MRSA). PLoS ONE 2015, 10, e0123206. [Google Scholar] [CrossRef] [PubMed]
  91. Peters, B.M.; Jabra-Rizk, M.A.; Scheper, M.A.; Leid, J.G.; Costerton, J.W.; Shirtliff, M.E. Microbial interactions and differential protein expression in Staphylococcus aureus-Candida albicans dual-species biofilms. FEMS Immunol. Med. Microbiol. 2010, 59, 493–503. [Google Scholar] [CrossRef] [PubMed]
  92. Abe, S.; Ishihara, K.; Okuda, K. Prevalence of potential respiratory pathogens in the mouths of elderly patients and effects of professional oral care. Arch. Gerontol. Geriatr. 2001, 32, 45–55. [Google Scholar] [CrossRef]
  93. Baena-Monroy, T.; Moreno-Maldonado, V.; Franco-Martínez, F.; Aldape-Barrios, B.; Quindós, G.; Sánchez-Vargas, L.O. Candida albicans, Staphylococcus aureus and streptococcus mutans colonization in patients wearing dental prosthesis. Med. Oral Patol. Oral Cir. Bucal 2005, 10, E27–E39. [Google Scholar] [PubMed]
  94. Miyake, Y.; Iwai, T.; Sugai, M.; Miura, K.; Suginaka, H.; Nagasaka, N. Incidence and characterization of Staphylococcus aureus from the tongues of children. J. Dent. Res. 1991, 70, 1045–1047. [Google Scholar] [CrossRef] [PubMed]
  95. Smith, A.J.; Robertson, D.; Tang, M.K.; Jackson, M.S.; MacKenzie, D.; Bagg, J. Staphylococcus aureus in the oral cavity: A three-year retrospective analysis of clinical laboratory data. Br. Dent. J. 2003, 195, 701–703. [Google Scholar] [CrossRef] [PubMed]
  96. Ohara-Nemoto, Y.; Haraga, H.; Kimura, S.; Nemoto, T.K. Occurrence of staphylococci in the oral cavities of healthy adults and nasal oral trafficking of the bacteria. J. Med. Microbiol. 2008, 57, 95–99. [Google Scholar] [CrossRef] [PubMed]
  97. Coco, B.J.; Bagg, J.; Cross, L.J.; Jose, A.; Cross, J.; Ramage, G. Mixed Candida albicans and Candida glabrata populations associated with the pathogenesis of denture stomatitis. Oral Microbiol. Immunol. 2008, 23, 377–383. [Google Scholar] [CrossRef] [PubMed]
  98. Silva, S.; Henriques, M.; Hayes, A.; Oliveira, R.; Azeredo, J.; Williams, D.W. Candida glabrata and Candida albicans co-infection of an in vitro oral epithelium. J. Oral Pathol. Med. 2011, 40, 421–427. [Google Scholar] [CrossRef] [PubMed]
  99. Alves, C.T.; Wei, X.Q.; Silva, S.; Azeredo, J.; Henriques, M.; Williams, D.W. Candida albicans promotes invasion and colonisation of Candida glabrata in a reconstituted human vaginal epithelium. J. Infect. 2014, 69, 396–407. [Google Scholar] [CrossRef] [PubMed]
  100. Tay, S.T.; Tan, H.W.; Na, S.L.; Lim, S.L. Phenotypic and genotypic characterization of two closely related subgroups of Candida rugosa in clinical specimens. J. Med. Microbiol. 2011, 60, 1591–1597. [Google Scholar] [CrossRef] [PubMed]
  101. Colombo, A.L.; Melo, A.S.; Crespo Rosas, R.F.; Salomão, R.; Briones, M.; Hollis, R.J.; Messer, S.A.; Pfaller, M.A. Outbreak of Candida rugosa candidemia: An emerging pathogen that may be refractory to amphotericin B therapy. Diagn. Microbiol. Infect. Dis. 2003, 46, 253–257. [Google Scholar] [CrossRef]
  102. Espinel-Ingroff, A.; Pfaller, M.A.; Bustamante, B.; Canton, E.; Fothergill, A.; Fuller, J.; Gonzalez, G.M.; Lass-Flörl, C.; Lockhart, S.R.; Martin-Mazuelos, E.; et al. Multilaboratory study of epidemiological cutoff values for detection of resistance in eight Candida species to fluconazole, posaconazole, and voriconazole. Antimicrob. Agents Chemother. 2014, 58, 2006–2012. [Google Scholar] [CrossRef] [PubMed]
  103. Pfaller, M.A.; Diekema, D.J.; Gibbs, D.L.; Newell, V.A.; Ellis, D.; Tullio, V.; Rodloff, A.; Fu, W.; Ling, T.A.; Group, G.A.S. Results from the artemis disk global antifungal surveillance study, 1997 to 2007: A 10.5-year analysis of susceptibilities of Candida species to fluconazole and voriconazole as determined by CLSI standardized disk diffusion. J. Clin. Microbiol. 2010, 48, 1366–1377. [Google Scholar] [CrossRef] [PubMed]
  104. Mohammadi, R.; Badiee, P.; Badali, H.; Abastabar, M.; Safa, A.H.; Hadipour, M.; Yazdani, H.; Heshmat, F. Use of restriction fragment length polymorphism to identify Candida species, related to onychomycosis. Adv. Biomed. Res. 2015, 4, 95. [Google Scholar] [PubMed]
  105. Ghosh, A.K.; Paul, S.; Sood, P.; Rudramurthy, S.M.; Rajbanshi, A.; Jillwin, T.J.; Chakrabarti, A. Matrix-assisted laser desorption ionization time-of-flight mass spectrometry for the rapid identification of yeasts causing bloodstream infections. Clin. Microbiol. Infect. 2015, 21, 372–378. [Google Scholar] [CrossRef] [PubMed]
  106. Kirkpatrick, W.R.; Lopez-Ribot, J.L.; McAtee, R.K.; Patterson, T.F. Growth competition between Candida dubliniensis and Candida albicans under broth and biofilm growing conditions. J. Clin. Microbiol. 2000, 38, 902–904. [Google Scholar] [PubMed]
  107. Zheng, H.; Kim, J.; Liew, M.; Yan, J.K.; Herrera, O.; Bok, J.W.; Kelleher, N.L.; Keller, N.P.; Wang, Y. Redox metabolites signal polymicrobial biofilm development via the napa oxidative stress cascade in aspergillus. Curr. Biol. 2015, 25, 29–37. [Google Scholar] [CrossRef] [PubMed]
  108. Manavathu, E.K.; Vager, D.L.; Vazquez, J.A. Development and antimicrobial susceptibility studies of in vitro monomicrobial and polymicrobial biofilm models with Aspergillus fumigatus and Pseudomonas aeruginosa. BMC Microbiol. 2014, 14, 53. [Google Scholar] [CrossRef] [PubMed]
  109. Peleg, A.Y.; Hogan, D.A.; Mylonakis, E. Medically important bacterial-fungal interactions. Nat. Rev. Microbiol. 2010, 8, 340–349. [Google Scholar] [CrossRef] [PubMed]
  110. Tampakakis, E.; Peleg, A.Y.; Mylonakis, E. Interaction of Candida albicans with an intestinal pathogen, Salmonella enterica serovar typhimurium. Eukaryot. Cell 2009, 8, 732–737. [Google Scholar] [CrossRef] [PubMed]
  111. Chandra, J.; Mukherjee, P.K.; Ghannoum, M.A. In vitro growth and analysis of Candida biofilms. Nat. Protoc. 2008, 3, 1909–1924. [Google Scholar] [CrossRef] [PubMed]
  112. Doll, K.; Jongsthaphongpun, K.L.; Stumpp, N.S.; Winkel, A.; Stiesch, M. Quantifying implant-associated biofilms: Comparison of microscopic, microbiologic and biochemical methods. J. Microbiol. Methods 2016, 130, 61–68. [Google Scholar] [CrossRef] [PubMed]
  113. Krom, B.P.; Cohen, J.B.; McElhaney Feser, G.E.; Cihlar, R.L. Optimized Candidal biofilm microtiter assay. J. Microbiol. Methods 2007, 68, 421–423. [Google Scholar] [CrossRef] [PubMed]
  114. Krom, B.P.; Willems, H.M. In vitro models for Candida biofilm development. Methods Mol. Biol. 2016, 1356, 95–105. [Google Scholar] [PubMed]
  115. Trafny, E.A.; Lewandowski, R.; Zawistowska-Marciniak, I.; Stepinska, M. Use of MTT assay for determination of the biofilm formation capacity of microorganisms in metalworking fluids. World J. Microbiol. Biotechnol. 2013, 29, 1635–1643. [Google Scholar] [CrossRef] [PubMed]
  116. Kuhn, D.M.; Balkis, M.; Chandra, J.; Mukherjee, P.K.; Ghannoum, M.A. Uses and limitations of the XTT assay in studies of Candida growth and metabolism. J. Clin. Microbiol. 2003, 41, 506–508. [Google Scholar] [CrossRef] [PubMed]
  117. Martinez, L.R.; Casadevall, A. Cryptococcus neoformans cells in biofilms are less susceptible than planktonic cells to antimicrobial molecules produced by the innate immune system. Infect. Immun. 2006, 74, 6118–6123. [Google Scholar] [CrossRef] [PubMed]
  118. Pierce, C.G.; Uppuluri, P.; Tristan, A.R.; Wormley, F.L.; Mowat, E.; Ramage, G.; Lopez-Ribot, J.L. A simple and reproducible 96-well plate-based method for the formation of fungal biofilms and its application to antifungal susceptibility testing. Nat. Protoc. 2008, 3, 1494–1500. [Google Scholar] [CrossRef] [PubMed]
  119. Peeters, E.; Nelis, H.J.; Coenye, T. Comparison of multiple methods for quantification of microbial biofilms grown in microtiter plates. J. Microbiol. Methods 2008, 72, 157–165. [Google Scholar] [CrossRef] [PubMed]
  120. Jiang, L.M.; Hoogenkamp, M.A.; van der Sluis, L.W.; Wesselink, P.R.; Crielaard, W.; Deng, D.M. Resazurin metabolism assay for root canal disinfectant evaluation on dual-species biofilms. J. Endod. 2011, 37, 31–35. [Google Scholar] [CrossRef] [PubMed]
  121. Shopova, I.; Bruns, S.; Thywissen, A.; Kniemeyer, O.; Brakhage, A.A.; Hillmann, F. Extrinsic extracellular dna leads to biofilm formation and colocalizes with matrix polysaccharides in the human pathogenic fungus Aspergillus fumigatus. Front. Microbiol. 2013, 4, 141. [Google Scholar] [CrossRef] [PubMed]
  122. Fessia, S.L.; Griffin, M.J. A method for assaying biofilm capacity on polyurethane-coated slides. Perit. Dial. Int. 1991, 11, 144–146. [Google Scholar] [PubMed]
  123. Di Bonaventura, G.; Pompilio, A.; Picciani, C.; Iezzi, M.; D’Antonio, D.; Piccolomini, R. Biofilm formation by the emerging fungal pathogen Trichosporon asahii: Development, architecture, and antifungal resistance. Antimicrob. Agents Chemother. 2006, 50, 3269–3276. [Google Scholar] [CrossRef] [PubMed]
  124. Sohm, J.A.; Edwards, B.R.; Wilson, B.G.; Webb, E.A. Constitutive extracellular polysaccharide (EPS) production by specific isolates of Crocosphaera watsonii. Front. Microbiol. 2011, 2, 229. [Google Scholar] [CrossRef] [PubMed]
  125. Tote, K.; Vanden Berghe, D.; Maes, L.; Cos, P. A new colorimetric microtitre model for the detection of Staphylococcus aureus biofilms. Lett. Appl. Microbiol. 2008, 46, 249–254. [Google Scholar] [CrossRef] [PubMed]
  126. Hawser, S.P.; Baillie, G.S.; Douglas, L.J. Production of extracellular matrix by Candida albicans biofilms. J. Med. Microbiol. 1998, 47, 253–256. [Google Scholar] [CrossRef] [PubMed]
  127. Lawrence, J.R.; Swerhone, G.D.; Leppard, G.G.; Araki, T.; Zhang, X.; West, M.M.; Hitchcock, A.P. Scanning transmission X-ray, laser scanning, and transmission electron microscopy mapping of the exopolymeric matrix of microbial biofilms. Appl. Environ. Microbiol. 2003, 69, 5543–5554. [Google Scholar] [CrossRef] [PubMed]
  128. Priester, J.H.; Horst, A.M.; Van de Werfhorst, L.C.; Saleta, J.L.; Mertes, L.A.; Holden, P.A. Enhanced visualization of microbial biofilms by staining and environmental scanning electron microscopy. J. Microbiol. Methods 2007, 68, 577–587. [Google Scholar] [CrossRef] [PubMed]
  129. Bridier, A.; Meylheuc, T.; Briandet, R. Realistic representation of Bacillus subtilis biofilms architecture using combined microscopy (CLSM, ESEM and FESEM). Micron 2013, 48, 65–69. [Google Scholar] [CrossRef] [PubMed]
  130. Dige, I.; Nilsson, H.; Kilian, M.; Nyvad, B. In situ identification of streptococci and other bacteria in initial dental biofilm by confocal laser scanning microscopy and fluorescence in situ hybridization. Eur. J. Oral Sci. 2007, 115, 459–467. [Google Scholar] [CrossRef] [PubMed]
  131. 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. 2009, 43, 63–76. [Google Scholar] [CrossRef] [PubMed]
  132. Surman, S.B.; Walker, J.T.; Goddard, D.T.; Morton, L.H.G.; Keevil, C.W.; Weaver, W.; Skinner, A.; Hanson, K.; Caldwell, D.; Kurtz, J. Comparison of microscope techniques for the examination of biofilms. J. Microbiol. Methods 1996, 25, 57–70. [Google Scholar] [CrossRef]
  133. Repp, K.K.; Menor, S.A.; Pettit, R.K. Microplate alamar blue assay for susceptibility testing of Candida albicans biofilms. Med. Mycol. 2007, 45, 603–607. [Google Scholar] [CrossRef] [PubMed]
  134. Pettit, R.K.; Weber, C.A.; Kean, M.J.; Hoffmann, H.; Pettit, G.R.; Tan, R.; Franks, K.S.; Horton, M.L. Microplate alamar blue assay for Staphylococcus epidermidis biofilm susceptibility testing. Antimicrob. Agents Chemother. 2005, 49, 2612–2617. [Google Scholar] [CrossRef] [PubMed]
  135. Muszkieta, L.; Beauvais, A.; Pahtz, V.; Gibbons, J.G.; Anton Leberre, V.; Beau, R.; Shibuya, K.; Rokas, A.; Francois, J.M.; Kniemeyer, O.; et al. Investigation of Aspergillus fumigatus biofilm formation by various “Omics” approaches. Front. Microbiol. 2013, 4, 13. [Google Scholar] [CrossRef] [PubMed]
  136. De Hoog, C.L.; Mann, M. Proteomics. Annu. Rev. Genom. Hum. Genet. 2004, 5, 267–293. [Google Scholar] [CrossRef] [PubMed]
  137. Azevedo, N.F.; Lopes, S.P.; Keevil, C.W.; Pereira, M.O.; Vieira, M.J. Time to “Go large” On biofilm research: Advantages of an omics approach. Biotechnol. Lett. 2009, 31, 477–485. [Google Scholar] [CrossRef] [PubMed]
  138. Rajendran, R.; May, A.; Sherry, L.; Kean, R.; Williams, C.; Jones, B.L.; Burgess, K.V.; Heringa, J.; Abeln, S.; Brandt, B.W.; et al. Integrating Candida albicans metabolism with biofilm heterogeneity by transcriptome mapping. Sci. Rep. 2016, 6, 35436. [Google Scholar] [CrossRef] [PubMed]
  139. Otto, A.; Bernhardt, J.; Hecker, M.; Becher, D. Global relative and absolute quantitation in microbial proteomics. Curr. Opin. Microbiol. 2012, 15, 364–372. [Google Scholar] [CrossRef] [PubMed]
  140. Thomas, D.P.; Bachmann, S.P.; Lopez-Ribot, J.L. Proteomics for the analysis of the Candida albicans biofilm lifestyle. Proteomics 2006, 6, 5795–5804. [Google Scholar] [CrossRef] [PubMed]
  141. Seneviratne, C.J.; Wang, Y.; Jin, L.; Abiko, Y.; Samaranayake, L.P. Candida albicans biofilm formation is associated with increased anti-oxidative capacities. Proteomics 2008, 8, 2936–2947. [Google Scholar] [CrossRef] [PubMed]
  142. Martinez-Gomariz, M.; Perumal, P.; Mekala, S.; Nombela, C.; Chaffin, W.L.; Gil, C. Proteomic analysis of cytoplasmic and surface proteins from yeast cells, hyphae, and biofilms of Candida albicans. Proteomics 2009, 9, 2230–2252. [Google Scholar] [CrossRef] [PubMed]
  143. Santi, L.; Beys-da-Silva, W.O.; Berger, M.; Calzolari, D.; Guimaraes, J.A.; Moresco, J.J.; Yates, J.R., 3rd. Proteomic profile of Cryptococcus neoformans biofilm reveals changes in metabolic processes. J. Proteome Res. 2014, 13, 1545–1559. [Google Scholar] [CrossRef] [PubMed]
  144. Buescher, J.M.; Driggers, E.M. Integration of omics: More than the sum of its parts. Cancer Metab. 2016, 4, 4. [Google Scholar] [CrossRef] [PubMed]
  145. Kojic, E.M.; Darouiche, R.O. Candida infections of medical devices. Clin. Microbiol. Rev. 2004, 17, 255–267. [Google Scholar] [CrossRef] [PubMed]
  146. Uppuluri, P.; Chaturvedi, A.K.; Lopez-Ribot, J.L. Design of a simple model of Candida albicans biofilms formed under conditions of flow: Development, architecture, and drug resistance. Mycopathologia 2009, 168, 101–109. [Google Scholar] [CrossRef] [PubMed]
  147. Nett, J.E.; Andes, D.R. Fungal biofilms: In vivo models for discovery of anti-biofilm drugs. Microbiol. Spectr. 2015, 3, E30. [Google Scholar] [CrossRef] [PubMed]
  148. Yanagisawa, N.; Li, D.Q.; Ljungh, A. Protein adsorption on ex vivo catheters and polymers exposed to peritoneal dialysis effluent. Perit. Dial. Int. 2004, 24, 264–273. [Google Scholar] [PubMed]
  149. Dongari-Bagtzoglou, A. Mucosal biofilms: Challenges and future directions. Expert Rev. Anti Infect. Ther. 2008, 6, 141–144. [Google Scholar] [CrossRef] [PubMed]
  150. Martinez, L.R.; Casadevall, A. Cryptococcus neoformans biofilm formation depends on surface support and carbon source and reduces fungal cell susceptibility to heat, cold, and UV light. Appl. Environ. Microbiol. 2007, 73, 4592–4601. [Google Scholar] [CrossRef] [PubMed]
  151. Ronsani, M.M.; Mores Rymovicz, A.U.; Meira, T.M.; Trindade Grégio, A.M.; Guariza Filho, O.; Tanaka, O.M.; Ribeiro Rosa, E.A. Virulence modulation of Candida albicans biofilms by metal ions commonly released from orthodontic devices. Microb. Pathog. 2011, 51, 421–425. [Google Scholar] [CrossRef] [PubMed]
  152. Chandra, J.; McCormick, T.S.; Imamura, Y.; Mukherjee, P.K.; Ghannoum, M.A. Interaction of Candida albicans with adherent human peripheral blood mononuclear cells increases C. Albicans biofilm formation and results in differential expression of pro- and anti-inflammatory cytokines. Infect. Immun. 2007, 75, 2612–2620. [Google Scholar] [CrossRef] [PubMed]
  153. Martinez, L.R.; Casadevall, A. Specific antibody can prevent fungal biofilm formation and this effect correlates with protective efficacy. Infect. Immun. 2005, 73, 6350–6362. [Google Scholar] [CrossRef] [PubMed]
  154. Donlan, R.M. Biofilms and device-associated infections. Emerg. Infect. Dis. 2001, 7, 277–281. [Google Scholar] [CrossRef] [PubMed]
  155. Harriott, M.M.; Lilly, E.A.; Rodriguez, T.E.; Fidel, P.L.; Noverr, M.C. Candida albicans forms biofilms on the vaginal mucosa. Microbiology 2010, 156, 3635–3644. [Google Scholar] [CrossRef] [PubMed]
  156. Andes, D.R. In Vivo Candida Device Biofilm Models. In Candida albicans: Cellular and Molecular Biology; Prasad, R., Ed.; Springer International Publishing: Cham, Switzerland, 2017; pp. 93–113. [Google Scholar]
  157. Ghannoum, M.; Roilides, E.; Katragkou, A.; Petraitis, V.; Walsh, T.J. The role of echinocandins in Candida biofilm-related vascular catheter infections: In vitro and in vivo model systems. Clin. Infect. Dis. 2015, 61, S618–S621. [Google Scholar] [CrossRef] [PubMed]
  158. Andes, D.; Nett, J.; Oschel, P.; Albrecht, R.; Marchillo, K.; Pitula, A. Development and characterization of an in vivo central venous catheter Candida albicans biofilm model. Infect. Immun. 2004, 72, 6023–6031. [Google Scholar] [CrossRef] [PubMed]
  159. Martinez, L.R.; Mihu, M.R.; Tar, M.; Cordero, R.J.; Han, G.; Friedman, A.J.; Friedman, J.M.; Nosanchuk, J.D. Demonstration of antibiofilm and antifungal efficacy of chitosan against Candidal biofilms, using an in vivo central venous catheter model. J. Infect. Dis. 2010, 201, 1436–1440. [Google Scholar] [CrossRef] [PubMed]
  160. Schinabeck, M.K.; Long, L.A.; Hossain, M.A.; Chandra, J.; Mukherjee, P.K.; Mohamed, S.; Ghannoum, M.A. Rabbit model of Candida albicans biofilm infection: Liposomal amphotericin B antifungal lock therapy. Antimicrob. Agents Chemother. 2004, 48, 1727–1732. [Google Scholar] [CrossRef] [PubMed]
  161. Kucharikova, S.; Neirinck, B.; Sharma, N.; Vleugels, J.; Lagrou, K.; van Dijck, P. In vivo Candida glabrata biofilm development on foreign bodies in a rat subcutaneous model. J. Antimicrob. Chemother. 2015, 70, 846–856. [Google Scholar] [CrossRef] [PubMed]
  162. Sangeorzan, J.A.; Bradley, S.F.; He, X.; Zarins, L.T.; Ridenour, G.L.; Tiballi, R.N.; Kauffman, C.A. Epidemiology of oral candidiasis in HIV-infected patients: Colonization, infection, treatment, and emergence of fluconazole resistance. Am. J. Med. 1994, 97, 339–346. [Google Scholar] [CrossRef]
  163. Solis, N.V.; Filler, S.G. Mouse model of oropharyngeal candidiasis. Nat. Protoc. 2012, 7, 637–642. [Google Scholar] [CrossRef] [PubMed]
  164. Nett, J.E.; Marchillo, K.; Spiegel, C.A.; Andes, D.R. Development and validation of an in vivo Candida albicans biofilm denture model. Infect. Immun. 2010, 78, 3650–3659. [Google Scholar] [CrossRef] [PubMed]
  165. Freire, F.; Ferraresi, C.; Jorge, A.O.; Hamblin, M.R. Photodynamic therapy of oral Candida infection in a mouse model. J. Photochem. Photobiol. B 2016, 159, 161–168. [Google Scholar] [CrossRef] [PubMed]
  166. Seleem, D.; Benso, B.; Noguti, J.; Pardi, V.; Murata, R.M. In vitro and in vivo antifungal activity of lichochalcone-a against Candida albicans biofilms. PLoS ONE 2016, 11, e0157188. [Google Scholar] [CrossRef] [PubMed]
  167. Vila, T.V.; Chaturvedi, A.K.; Rozental, S.; Lopez-Ribot, J.L. In vitro activity of miltefosine against Candida albicans under planktonic and biofilm growth conditions and in vivo efficacy in a murine model of oral candidiasis. Antimicrob. Agents Chemother. 2015, 59, 7611–7620. [Google Scholar] [CrossRef] [PubMed]
  168. Wang, X.; Fries, B.C. A murine model for catheter-associated candiduria. J. Med. Microbiol. 2011, 60, 1523–1529. [Google Scholar] [CrossRef] [PubMed]
  169. Nett, J.E.; Brooks, E.G.; Cabezas-Olcoz, J.; Sanchez, H.; Zarnowski, R.; Marchillo, K.; Andes, D.R. Rat indwelling urinary catheter model of Candida albicans biofilm infection. Infect. Immun. 2014, 82, 4931–4940. [Google Scholar] [CrossRef] [PubMed]
  170. Sun, Y.; Chandra, J.; Mukherjee, P.; Szczotka-Flynn, L.; Ghannoum, M.A.; Pearlman, E. A murine model of contact lens-associated Fusarium keratitis. Investig. Ophthalmol. Vis. Sci. 2010, 51, 1511–1516. [Google Scholar] [CrossRef] [PubMed]
  171. Pinnock, A.; Shivshetty, N.; Roy, S.; Rimmer, S.; Douglas, I.; MacNeil, S.; Garg, P. Ex vivo rabbit and human corneas as models for bacterial and fungal keratitis. Graefes Arch. Clin. Exp. Ophthalmol. 2017, 255, 333–342. [Google Scholar] [CrossRef] [PubMed]
  172. Zhang, H.; Wang, L.; Li, Z.; Liu, S.; Xie, Y.; He, S.; Deng, X.; Yang, B.; Liu, H.; Chen, G.; et al. A novel murine model of Fusarium solani keratitis utilizing fluorescent labeled fungi. Exp. Eye Res. 2013, 110, 107–112. [Google Scholar] [CrossRef] [PubMed]
  173. Benaducci, T.; Sardi Jde, C.; Lourencetti, N.M.; Scorzoni, L.; Gullo, F.P.; Rossi, S.A.; Derissi, J.B.; de Azevedo Prata, M.C.; Fusco-Almeida, A.M.; Mendes-Giannini, M.J. Virulence of Cryptococcus sp. Biofilms in vitro and in vivo using Galleria mellonella as an alternative model. Front. Microbiol. 2016, 7, 290. [Google Scholar] [CrossRef] [PubMed]
  174. Rajendran, R.; Borghi, E.; Falleni, M.; Perdoni, F.; Tosi, D.; Lappin, D.F.; O'Donnell, L.; Greetham, D.; Ramage, G.; Nile, C. Acetylcholine protects against Candida albicans infection by inhibiting biofilm formation and promoting hemocyte function in a Galleria mellonella infection model. Eukaryot. Cell 2015, 14, 834–844. [Google Scholar] [CrossRef] [PubMed]
  175. Cirasola, D.; Sciota, R.; Vizzini, L.; Ricucci, V.; Morace, G.; Borghi, E. Experimental biofilm-related Candida infections. Future Microbiol. 2013, 8, 799–805. [Google Scholar] [CrossRef] [PubMed]
  176. Flemming, H.C.; Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 2010, 8, 623–633. [Google Scholar] [CrossRef] [PubMed]
  177. Mitchell, K.F.; Zarnowski, R.; Andes, D.R. The extracellular matrix of fungal biofilms. Adv. Exp. Med. Biol. 2016, 931, 21–35. [Google Scholar] [PubMed]
  178. Mathe, L.; van Dijck, P. Recent insights into Candida albicans biofilm resistance mechanisms. Curr. Genet. 2013, 59, 251–264. [Google Scholar] [CrossRef] [PubMed]
  179. Sardi, J.; Pitangui, N.; Gullo, F.; Fusco-Almeida, A.; Mendes-Giannini, A. Fungal biofilms: Formation, resistance and pathogenicity. In Medical Mycology: Current Trends and Future Prospects; Razzaghi-Abyaneh, M., Shams-Ghahfarokhi, M., Rai, M., Eds.; Taylor & Francis Group: Oxford, UK, 2015; Volume 1, pp. 291–314. [Google Scholar]
  180. Perumal, P.; Mekala, S.; Chaffin, W.L. Role for cell density in antifungal drug resistance in Candida albicans biofilms. Antimicrob. Agents Chemother. 2007, 51, 2454–2463. [Google Scholar] [CrossRef] [PubMed]
  181. Seneviratne, C.J.; Jin, L.; Samaranayake, L.P. Biofilm lifestyle of candida: A mini review. Oral Dis. 2008, 14, 582–590. [Google Scholar] [CrossRef] [PubMed]
  182. Nailis, H.; Vandenbosch, D.; Deforce, D.; Nelis, H.J.; Coenye, T. Transcriptional response to fluconazole and amphotericin b in Candida albicans biofilms. Res. Microbiol. 2010, 161, 284–292. [Google Scholar] [CrossRef] [PubMed]
  183. Nett, J.E. Future directions for anti-biofilm therapeutics targeting candida. Expert Rev. Anti Infect. Ther. 2014, 12, 375–382. [Google Scholar] [CrossRef] [PubMed]
  184. LaFleur, M.D.; Kumamoto, C.A.; Lewis, K. Candida albicans biofilms produce antifungal-tolerant persister cells. Antimicrob. Agents Chemother. 2006, 50, 3839–3846. [Google Scholar] [CrossRef] [PubMed]
  185. Al-Fattani, M.A.; Douglas, L.J. Biofilm matrix of Candida albicans and Candida tropicalis: Chemical composition and role in drug resistance. J. Med. Microbiol. 2006, 55, 999–1008. [Google Scholar] [CrossRef] [PubMed]
  186. Diez-Orejas, R.; Molero, G.; Navarro-Garcia, F.; Pla, J.; Nombela, C.; Sanchez-Perez, M. Reduced virulence of Candida albicans mkc1 mutants: A role for mitogen-activated protein kinase in pathogenesis. Infect. Immun. 1997, 65, 833–837. [Google Scholar] [PubMed]
  187. Mowat, E.; Lang, S.; Williams, C.; McCulloch, E.; Jones, B.; Ramage, G. Phase-dependent antifungal activity against Aspergillus fumigatus developing multicellular filamentous biofilms. J. Antimicrob. Chemother. 2008, 62, 1281–1284. [Google Scholar] [CrossRef] [PubMed]
  188. Niimi, M.; Firth, N.A.; Cannon, R.D. Antifungal drug resistance of oral fungi. Odontology 2010, 98, 15–25. [Google Scholar] [CrossRef] [PubMed]
  189. Ramage, G.; Rajendran, R.; Sherry, L.; Williams, C. Fungal biofilm resistance. Int. J. Microbiol. 2012, 2012, 528521. [Google Scholar] [CrossRef] [PubMed]
  190. Reichhardt, C.; Stevens, D.A.; Cegelski, L. Fungal biofilm composition and opportunities in drug discovery. Future Med. Chem. 2016, 8, 1455–1468. [Google Scholar] [CrossRef] [PubMed]
  191. Jennings, L.K.; Storek, K.M.; Ledvina, H.E.; Coulon, C.; Marmont, L.S.; Sadovskaya, I.; Secor, P.R.; Tseng, B.S.; Scian, M.; Filloux, A.; et al. Pel is a cationic exopolysaccharide that cross-links extracellular DNA in the Pseudomonas aeruginosa biofilm matrix. Proc. Natl. Acad. Sci. USA 2015, 112, 11353–11358. [Google Scholar] [CrossRef] [PubMed]
  192. Taff, H.T.; Mitchell, K.F.; Edward, J.A.; Andes, D.R. Mechanisms of Candida biofilm drug resistance. Future Microbiol. 2013, 8, 1325–1337. [Google Scholar] [CrossRef] [PubMed]
  193. Ramage, G.; Mowat, E.; Jones, B.; Williams, C.; Lopez-Ribot, J. Our current understanding of fungal biofilms. Crit. Rev. Microbiol. 2009, 35, 340–355. [Google Scholar] [CrossRef] [PubMed]
  194. Wongsuk, T.; Pumeesat, P.; Luplertlop, N. Fungal quorum sensing molecules: Role in fungal morphogenesis and pathogenicity. J. Basic Microbiol. 2016, 56, 440–447. [Google Scholar] [CrossRef] [PubMed]
  195. Albuquerque, P.; Casadevall, A. Quorum sensing in fungi—A review. Med. Mycol. 2012, 50, 337–345. [Google Scholar] [CrossRef] [PubMed]
  196. Ramage, G.; Saville, S.P.; Thomas, D.P.; Lopez-Ribot, J.L. Candida biofilms: An update. Eukaryot. Cell 2005, 4, 633–638. [Google Scholar] [CrossRef] [PubMed]
  197. Hornby, J.M.; Jensen, E.C.; Lisec, A.D.; Tasto, J.J.; Jahnke, B.; Shoemaker, R.; Dussault, P.; Nickerson, K.W. Quorum sensing in the dimorphic fungus Candida albicans is mediated by farnesol. Appl. Environ. Microbiol. 2001, 67, 2982–2992. [Google Scholar] [CrossRef] [PubMed]
  198. Enjalbert, B.; Whiteway, M. Release from quorum-sensing molecules triggers hyphal formation during Candida albicans resumption of growth. Eukaryot. Cell 2005, 4, 1203–1210. [Google Scholar] [CrossRef] [PubMed]
  199. Sharma, M.; Prasad, R. The quorum-sensing molecule farnesol is a modulator of drug efflux mediated by ABC multidrug transporters and synergizes with drugs in Candida albicans. Antimicrob. Agents Chemother. 2011, 55, 4834–4843. [Google Scholar] [CrossRef] [PubMed]
  200. Ramage, G.; Saville, S.P.; Wickes, B.L.; Lopez-Ribot, J.L. Inhibition of Candida albicans biofilm formation by farnesol, a quorum-sensing molecule. Appl. Environ. Microbiol. 2002, 68, 5459–5463. [Google Scholar] [CrossRef] [PubMed]
  201. Semighini, C.P.; Hornby, J.M.; Dumitru, R.; Nickerson, K.W.; Harris, S.D. Farnesol-induced apoptosis in Aspergillus nidulans reveals a possible mechanism for antagonistic interactions between fungi. Mol. Microbiol. 2006, 59, 753–764. [Google Scholar] [CrossRef] [PubMed]
  202. Jabra-Rizk, M.A.; Shirtliff, M.; James, C.; Meiller, T. Effect of farnesol on Candida dubliniensis biofilm formation and fluconazole resistance. FEMS Yeast Res. 2006, 6, 1063–1073. [Google Scholar] [CrossRef] [PubMed]
  203. Lewis, K. Persister cells. Annu. Rev. Microbiol. 2010, 64, 357–372. [Google Scholar] [CrossRef] [PubMed]
  204. Lewis, K. Persister cells: Molecular mechanisms related to antibiotic tolerance. Handb. Exp. Pharmacol. 2012, 121–133. [Google Scholar] [CrossRef]
  205. Mukherjee, P.K.; Chandra, J.; Kuhn, D.M.; Ghannoum, M.A. Mechanism of fluconazole resistance in Candida albicans biofilms: Phase-specific role of efflux pumps and membrane sterols. Infect. Immun. 2003, 71, 4333–4340. [Google Scholar] [CrossRef] [PubMed]
  206. Soto, S.M. Role of efflux pumps in the antibiotic resistance of bacteria embedded in a biofilm. Virulence 2013, 4, 223–229. [Google Scholar] [CrossRef] [PubMed]
  207. Nickerson, K.W.; Atkin, A.L.; Hornby, J.M. Quorum sensing in dimorphic fungi: Farnesol and beyond. Appl. Environ. Microbiol. 2006, 72, 3805–3813. [Google Scholar] [CrossRef] [PubMed]
  208. Martins, M.; Henriques, M.; Azeredo, J.; Rocha, S.M.; Coimbra, M.A.; Oliveira, R. Morphogenesis control in Candida albicans and Candida dubliniensis through signaling molecules produced by planktonic and biofilm cells. Eukaryot. Cell 2007, 6, 2429–2436. [Google Scholar] [CrossRef] [PubMed]
  209. Shirtliff, M.E.; Krom, B.P.; Meijering, R.A.; Peters, B.M.; Zhu, J.; Scheper, M.A.; Harris, M.L.; Jabra-Rizk, M.A. Farnesol-induced apoptosis in Candida albicans. Antimicrob. Agents Chemother. 2009, 53, 2392–2401. [Google Scholar] [CrossRef] [PubMed]
  210. Alem, M.A.; Oteef, M.D.; Flowers, T.H.; Douglas, L.J. Production of tyrosol by Candida albicans biofilms and its role in quorum sensing and biofilm development. Eukaryot. Cell 2006, 5, 1770–1779. [Google Scholar] [CrossRef] [PubMed]
  211. Chen, H.; Fink, G.R. Feedback control of morphogenesis in fungi by aromatic alcohols. Genes Dev. 2006, 20, 1150–1161. [Google Scholar] [CrossRef] [PubMed]
  212. Cordeiro Rde, A.; Teixeira, C.E.; Brilhante, R.S.; Castelo-Branco, D.S.; Alencar, L.P.; de Oliveira, J.S.; Monteiro, A.J.; Bandeira, T.J.; Sidrim, J.J.; Moreira, J.L.; et al. Exogenous tyrosol inhibits planktonic cells and biofilms of Candida species and enhances their susceptibility to antifungals. FEMS Yeast Res. 2015, 15, fov012. [Google Scholar] [CrossRef] [PubMed]
  213. Lorek, J.; Poggeler, S.; Weide, M.R.; Breves, R.; Bockmuhl, D.P. Influence of farnesol on the morphogenesis of Aspergillus niger. J. Basic Microbiol. 2008, 48, 99–103. [Google Scholar] [CrossRef] [PubMed]
  214. Dichtl, K.; Ebel, F.; Dirr, F.; Routier, F.H.; Heesemann, J.; Wagener, J. Farnesol misplaces tip-localized ρ proteins and inhibits cell wall integrity signalling in Aspergillus fumigatus. Mol. Microbiol. 2010, 76, 1191–1204. [Google Scholar] [CrossRef] [PubMed]
  215. Brilhante, R.S.; de Lima, R.A.; Marques, F.J.; Silva, N.F.; Caetano, E.P.; Castelo-Branco Dde, S.; Bandeira Tde, J.; Moreira, J.L.; Cordeiro Rde, A.; Monteiro, A.J.; et al. Histoplasma capsulatum in planktonic and biofilm forms: In vitro susceptibility to amphotericin B, itraconazole and farnesol. J. Med. Microbiol. 2015, 64, 394–399. [Google Scholar] [CrossRef] [PubMed]
  216. Derengowski, L.S.; De-Souza-Silva, C.; Braz, S.V.; Mello-De-Sousa, T.M.; Bao, S.N.; Kyaw, C.M.; Silva-Pereira, I. Antimicrobial effect of farnesol, a Candida albicans quorum sensing molecule, on Paracoccidioides brasiliensis growth and morphogenesis. Ann. Clin. Microbiol. Antimicrob. 2009, 8, 13. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Models of biofilm development in filamentous fungi (a) and C. albicans (b). The stages of development are similar, although the morphology and number of stages are different. In the first model (a), six stages were proposed by Harding et al. [12]: (I) adsorption, (II) active attachment, (III) first formation of microcolony through germination and/or monolayer development, (IV) mycelial development, (V) biofilm maturation, and (VI) dispersion of conidia and/or arthroconidia. The second model corresponds to classical C. albicans biofilm development (b) which includes five stages, such as in bacteria: (i) adsorption, (ii) adhesion, (iii) microcolony formation, (iv) mature biofilm, and (v) dispersion. Modified from Harding et al. [12]. T. rubrum mature biofilm Costa-Orlandi et al. [7]; Pires et al. [18].
Figure 1. Models of biofilm development in filamentous fungi (a) and C. albicans (b). The stages of development are similar, although the morphology and number of stages are different. In the first model (a), six stages were proposed by Harding et al. [12]: (I) adsorption, (II) active attachment, (III) first formation of microcolony through germination and/or monolayer development, (IV) mycelial development, (V) biofilm maturation, and (VI) dispersion of conidia and/or arthroconidia. The second model corresponds to classical C. albicans biofilm development (b) which includes five stages, such as in bacteria: (i) adsorption, (ii) adhesion, (iii) microcolony formation, (iv) mature biofilm, and (v) dispersion. Modified from Harding et al. [12]. T. rubrum mature biofilm Costa-Orlandi et al. [7]; Pires et al. [18].
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Figure 2. Scheme of the mechanisms and factors that promote fungal biofilm resistance, which are common to several fungi. Adapted from Ramage et al. [189].
Figure 2. Scheme of the mechanisms and factors that promote fungal biofilm resistance, which are common to several fungi. Adapted from Ramage et al. [189].
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Table 1. Microtiter plates assay for susceptibility testing and biofilm characterization.
Table 1. Microtiter plates assay for susceptibility testing and biofilm characterization.
Microtiter Plates AssaysCharacteristics
MTTMTT is a yellow soluble salt, which in the presence of metabolic activity, is reduced to an insoluble purple formazan crystal. This method is used to determine the metabolic activity of some microorganisms in planktonic and biofilm forms. Moreover, this method shows excellent correlation with biomass determination by dry weight. Fast and convenient [24].
XTTTetrazolium salt (yellow) is reduced by the activity of fungal mitochondrial dehydrogenase to formazan salt (orange), which is correlated with cell viability. It is also used to determine metabolic activity in the developmental stages of biofilms and in antifungal susceptibility tests [46,116,118]. The method is simple and, reproducible, but some disadvantages were reported by Khun et al. [116].
Alamar Blue and ResazurinReduction is dependent on metabolic activity. The methods are fast and simple and measurement can be conducted spectrofluorometrically or spectrophotometrically. Resazurin is the active principle of Alamar Blue. The reagents are nontoxic to humans and fungi and the method is reproducible. Good correlation with XTT assay and CFU/mL [133,134]. Used for biofilm quantification. Blue dye resazurin is converted to pink resorufin in the presence of metabolic activity. Nontoxic and soluble in water [119,120,121].
SafraninDye easy to use for ECM quantification Difficult interpretation; low-cost [7,42,122].
Crystal Violet (CV)Used for biomass quantification. CV stains living and dead cells, and thus it is not indicated to verify antifungal activity in biofilms [119]. Low cost and easy [31].
Alcian BlueMeasures mass quantity of biofilm ECM [123,124].
1,9-Dimethyl Methylene Blue (DMMB)Quantification of biofilm matrix [119,125].
Table 2. Resistance mechanisms associated with biofilm formation. Adapted from Mathé and Dijick [178] and Sardi et al. [179].
Table 2. Resistance mechanisms associated with biofilm formation. Adapted from Mathé and Dijick [178] and Sardi et al. [179].
Resistance MechanismsEffectReferences
Cellular densityQuorum sensingPerumal et al. [180]; Seneviratne et al. [181].
Differential regulation drug targetAlteration in target levels; Associated with changes in target structure that make the drug unable to bind to the target.Nailis et al. [182].
Upregulation drug efflux pumpsAntifungal is pumped out of cells and thus cannot perform its intracellular function.Nett et al. [183]
Persister cellsBecause of the dormant state of the persisters, antifungal targets are inactive.LaFleur et al. [184]
Presence of a matrixSpecific binding of antifungals to β-1,3-glucans, a major component of the matrix, prevents antifungal agents from reaching their targets.Al-Fattani and Douglas [185]; Mitchell et al. [177].
Diverse stress responsesPossible indirect effects through the regulation of other resistance mechanisms.Diez-Orejas et al. [186]
Table 3. Role of QSMs (quorum-sensing molecules) in yeasts and dimorphic fungi. Adapted from Wongsuk et al. [194].
Table 3. Role of QSMs (quorum-sensing molecules) in yeasts and dimorphic fungi. Adapted from Wongsuk et al. [194].
OrganismQSMsRole of QSMs in Molds and Dimorphic FungiReferences
C. albicansFarnesol Inhibited hyphal development
Involved in morphogenesis
Inhibited biofilm formation
Induced apoptosis
Antifungal activity
Modulated drug extrusion
Nickerson et al. [207]
Martins et al. [208]
Ramage et al. [200]
Shirtliff et al. [209]
Sardi et al. [10]
Sharma et al. [199]
Tyrosol Promoted germ tube formation
Stimulated hypha production during the early stages of biofilm development
Antifungal activity
Alem et al. [210]
Chen et al. [211]
Cordeiro et al. [212]
A. nigerFarnesol Inhibited conidiation
Reduced intracellular cAMP levels
Lorek et al. [213]
A. fumigatusFarnesol Altered growth phenotype
Perturbed cell wall
Dichtl et al. [214]
H. capsulatumFarnesol Inhibited biofilm formation
Antifungal activity
Brilhante et al. [215]
P. brasiliensisFarnesol Inhibited growth
Delayed the dimorphic transition
Antifungal activity
Derengowski et al. [216]

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Costa-Orlandi, C.B.; Sardi, J.C.O.; Pitangui, N.S.; De Oliveira, H.C.; Scorzoni, L.; Galeane, M.C.; Medina-Alarcón, K.P.; Melo, W.C.M.A.; Marcelino, M.Y.; Braz, J.D.; Fusco-Almeida, A.M.; Mendes-Giannini, M.J.S. Fungal Biofilms and Polymicrobial Diseases. J. Fungi 2017, 3, 22.

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Costa-Orlandi CB, Sardi JCO, Pitangui NS, De Oliveira HC, Scorzoni L, Galeane MC, Medina-Alarcón KP, Melo WCMA, Marcelino MY, Braz JD, Fusco-Almeida AM, Mendes-Giannini MJS. Fungal Biofilms and Polymicrobial Diseases. Journal of Fungi. 2017; 3(2):22.

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Costa-Orlandi, Caroline B., Janaina C. O. Sardi, Nayla S. Pitangui, Haroldo C. De Oliveira, Liliana Scorzoni, Mariana C. Galeane, Kaila P. Medina-Alarcón, Wanessa C. M. A. Melo, Mônica Y. Marcelino, Jaqueline D. Braz, Ana Marisa Fusco-Almeida, and Maria José S. Mendes-Giannini. 2017. "Fungal Biofilms and Polymicrobial Diseases" Journal of Fungi 3, no. 2: 22.

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