For most of the history of microbiology, microorganisms have primarily been characterized as planktonic, freely suspended cells and described on the basis of their growth characteristics in nutritionally rich culture media. The discovery of microorganisms, 1684, is usually ascribed to Antoni van Leeuwenhoek, who was the first person to publish microscopic observations of bacteria. The direct quantitative recovery techniques showed unequivocally that more than 99.9% of the bacteria grow in biofilms on a wide variety of surfaces. Although the most common mode of growth for microorganisms on earth is in surface associated communities (Stoodley et al., 2002; Sutherland, 2001), the first reported findings of microorganisms “attached in layers” were not made until the 1940s. During the 1960s and 70s the research on “microbial slimes” accelerated but the term “biofilm” was not unanimous formulated until 1984 (Bryers, 2000). Biofilm has three-dimensional (3D) structured, heterogeneous community of microbial cells enclosed in an exopolysaccharide matrix (also called glycocalyx) that are irreversibly attached to an inert or living surface. As establish, biofilm formation has a serious implications in public health and medicine. In the case of human health, a number of microbial infections are associated with surface colonization not only on live surfaces (sinusitis, pulmonary infection in cystic fibrosis patients, periodontitis, etc. (Hall-Stoodley et al., 2004) but also on medical implants (contact lenses, dental implants, intravascular catheters, urinary stents) etc. (Donlan, 2001; Hall-Stoodley et al., 2004). Biofilms affect heat exchangers, filters, etc. because they induce biocorrosion and biofouling, producing damages on metallic surfaces and the efficiency loss in industrial set-up (Dunne, 2002; Garret et al., 2008). However,biofilms have also useful applications in bioremediation (Vidali, 2001) of different environments (microorganisms degrade and convert pollutants into less toxic forms) and biolixiviation (bacteria can efficiently dissolve minerals used in industry, to obtain copper and gold).
In order that we may gain a greater insight into the ecology of the microorganisms that exist in biofilm, it is necessary not only to be able to isolate them by traditional culture methods but also to have some understanding of the way in which these individual microorganisms interact in situ in their environment. Different microscopic techniques for biofilm monitoring including Scanning Electron microscopy (SEM) have been proved to be suitable tools in order to follow the study of adhesion stage and biofilm formation. Scanning electron microscopy as a specialized field of science that employs the electron microscope as a tool and uses a beam of electrons to form an image of a specimen allowing imaging and quantification of surface topographic features.
The scope of this chapter is to illustrate the importance of scanning electron microscopy and environnemental scanning electron microscopy in biofilm examination and control. Furthermore, although we are conscious about the vast variety of biofilms in natural, clinical and industrial environments, this chapter will mainly concentrate on imaging application of SEM and ESEM biofilms.
2. Step of biofilm formation
Planktonic cells are able to attach on the surfaces and form biofilm through a process that include several steps:
The primary adhesion stage constitutes the beneficial contact between a conditioned surface and planktonic microorganisms. During the process of attachment, the organism must be brought into close proximity of the surface, propelled either randomly or in a directed fashion via chemotaxis and mobility (Prakash et al., 2003). This step is reversible and it is characterized by a number of physicochemical variables that defines the interaction between the microbial cell surface and the conditioned surface of interest (An et al., 2000; Liu et al., 2004; Singh et al., 2002).
2.2. Irreversible adhesion
The second step is the irreversible adhesion during which bacteria start to express adhesion protein such as curli or fimbriae to adhere to the surface. Microorganisms starts to produce intercellular connections (intercellular curli for example) and a polymeric matrix, usually called extracellular polymeric substances (EPS). This matrix is a complex hydrogel embedding the bacteria community and building up in three dimensions. The backbone of this gel is mainly composed of polysaccharides produced by bacteria (such as colanic acid, chitosan, alginate), other components such as enzymes, DNA, RNA, nutrients, proteins, surfactants (Flemming et al., 2007). The exact role of the matrix is not yet completely elucidated but it has been demonstrated that the matrix acts as a protective layer (Fux et al., 2005) and is microenvironment-conservative (Beech, 2004).
After the adherence of microorganism to the inert surface, the association becomes stable for micro-colonies formation (Bechmann & Eduvean, 2006; O’Toole et al., 2000). The microorganism begin to multiply while sending out chemical signals that intercommunicate among the bacterial cells. In this way, the bacteria multiply within the embedded exopolysaccharide matrix, thus giving rise to formation of a micro-colonies (Prakash et al., 2003).
2.3. Maturation of biofilm
Once bacteria have irreversibly attached to a surface, the process of biofilm maturation begins. The overall density and complexity of the biofilm increase as surface-bound organisms begin to actively replicate and extracellular components generated by attached bacteria interact with organic and inorganic molecules in the immediate environment to create the glycocalyx (Carpentier & Cerf, 1993). The maturation of biofilm generate many process already having taken place, such as quorum sensing (Nadell et al., 2008), gene transfer (Molin, 2003), persister development (Lewis, 2005) etc. All of these processes contribute to the community life of the biofilm and play an important role in biofilm survival and biofilm spreading, since they allow also detachment of biofilm parts and release of free bacteria, which is the most common way for biofilm to spread (Kaplan et al., 2003).
2.4. Detachment and dispersal of biofilm cells
As the biofilm gets older, cells detach, disperse and colonize a new niche. This detachment can be due to various factors including, fluid dynamics and shear effects of the bulk fluid (Brugnoni et al., 2007). At some point of biofilms may partially dissolve releasing cells that more away to other where a new cycle begins (Prakash et al., 2003; Singh et al., 2002).
3. Imaging application
SEM is a well-established basic method to observe the morphology of bacteria adhered on a material surfaces, the morphology of the material surface, and the relationships between them (Peters et al., 1982). SEM has been used for enumeration of adhered bacteria or tissue large number of samples. It is as a key technique that provides also information about the morphology of biofilm, presence of EPS and the nature of corrosion products (crystalline or amorphous).
3.1. SEM applied of adhesion stage
Microbial adhesion is the first step of the formation of biofilm and an extremely complicated process that is affected by many factors. In this regard, detailed investigation of microbial adhesion involved in the developmental process from single sessile bacteria to multicellular biofilm is crucial to elaborate strategies to control biofilm development. Moreover, submicrometer-scale cell surface polymers and appendages, such as curli, flagella, and exocellular polymers, have been shown to play essential roles during cell adhesion and biofilm formation (Busscher et al., 2008; Dufrêne, 2008; Rodrigues & Elimelech, 2009). A SEM image of such a curli is depicted in Figure 2.
Adhesion phenomena has been evaluated as function of substratum, liquid medium, carbone source, pH and hydrodynamics parameters including flow rate. Many of the conclusions about biofilm development, composition, distribution, and relationship to substratum have been derived from scanning electron microscopy (Bragadeewaran et al., 2010; Herald & Zottola, 1988; Pinna et al., 2000). We report here several investigations made in our laboratory used scanning electron microscopy to study adhesion phenomena. Hamadi et al., (2005) have investigated the adhesion of
The surface topography has been widely discussed as a parameter influencing microbial adhesion. In this regard, experiments made by Kouider et al., (2010) using SEM to determine the effect of stainless steel surface roughness on
3.2. SEM applied of biofilm formation
Scanning electron microscopy (SEM) is a useful technique for the investigation of surface structure of biological samples (Duckett & Ligrone, 1995; Minoura et al., 1995; Motta et al., 1994). For instance, much of the current knowledge about biofilms is due to the advances in imaging studies, especially the SEM. Early microscopic techniques used in biofilm monitoring, mainly applied during the 1980s, include scanning electron microscopy. SEM has been previously used to show a clear visualization of bacteria within a biofilm and is capable of demonstrating even a single bacterium and the relation of the biofilm to the underlying surface.
Biofilm morphology and mass are important characteristics that control the kinetics of substrate removal by biofilms. SEM is a powerful technique for revealing the fine structure of living systems and has been applied to biofilms (Eighmy et al., 1983; Richards and Turner, 1984; Weber et al., 1978). It has also been of special importance in elucidating biofilm structure for understanding the physiology and ecology of these microbial systems (Blenkinsopp & Costerton 1991). For example, electron-microscopic studies proved that the biofilm is composed of bacterial cells “wrapped” in a dense “glycocalyx”, i.e. exopolysaccharide matrix (Blenkinsopp & Costerton, 1991; Eighmy et al. 1983). In medical applications, for example, Storti et al., (2005) used scanning electron microscopy and reported that the extracellular biofilm matrix appears as an amorphous material on the catheter surface. In the same context, scanning electron microscopy (SEM) images of matrix-enclosed microbial assemblages on leaf surfaces (Surico, 1993) have led some authors to suggest that biofilms occur in the phyllosphere (Beattie and Lindow, 1995). Morris et al., (1997) have been to observe microbial biofilms directly on leaf surfaces. Bacterial aggregates in the phyllosphere have been observed previously with SEM (Surico,1993), but most have been very small (less than 20 mm long) or have lacked an obvious exopolymeric matrix (Surico,1993). Previous studies have claimed to demonstrate the presence of biofilms in situ on plant aerial surfaces using SEM (Gras et al., 1994).
Biofilm thickness is also especially important for calculation of heat exchange or diffusion rates of antimicrobials or nutrients through a biofilm and for evaluation of the mechanical properties of a biofilm (Korstgens et al., 2001). As reported elsewhere, SEM sample (freeze-dried cross-section of Foley bladder catheter) revealed the thickness of biofilm and also the layers of embedded of slime by different strains and species of bacterial cells (Ganderton et al., 1992).
In general, other application of SEM techniques may be mentioned. Akernan et al., (1993) used scanning electron microscopy of nanobacteria - novel biofilm producing organisms in blood. Indeed, nanoscale characterization of
Scanning electron microscopy (SEM) is one of the many methods available for the visual the effect of antibacterial or antifungal on biofilm development (Camargo et al., 2005; McDowell et al., 2004; Sasidharan et al., 2010; Sevinç & Hanley, 2010; Zameer & Gopal, 2010; Zeraik & Nitschke, 2010). Sasidharan et al., (2010) used SEM for studied The effects of potential antifungal extracts from natural sources in
3.3. Advantages and disadvantages of SEM
In part, it is true that Scanning electron microscopy (SEM) present a many advantages, the more important are: (i) higher resolution of visualization microbial biofilms (Walker et al., 2001) than other imaging techniques, typically 3.5 nm, (ii) able to measure and quantify data in three dimensions. However, this technique utilizes graded solvents (alcohol, acetone, and xylene) to gradually dehydrate the specimen prior to examination, since water of hydration is not compatible with the vacuum used with the electron beam. While any pretreatment can alter specimen morphology, drying appears to significantly alter biofilms due to EPS polymers collapsing (Fassel & Edmiston, 1999; Little et al., 1991). The dehydration process results in significant sample distortion and artifacts; the extracellular polymeric substances, which are approximately 95% water and the liquid loss led them to appear more like fibers surrounding the cells than like a gelatinous matrix (Characklis & Marshall, 1990). Several ultrastructural studies have used conventional scanning electron microscopy (SEM) to investigate the glycocalyx, but these studies (Costerton et al., 1981; Fassel et al., 1991; Marshall et al., 1971) were hampered by low resolution and also by the inability to use low voltages (<5 keV), which yield increased information from small topographical features (Pawley & Erlandsen, 1989).
Typically, SEM imaging requires a high vacuum, ≤10-8 Torr (reviewed in Stewart, 1985), having ﬁrst been chemically ﬁxed, dehydrated, and coated with a conductive material (e.g. gold) to prevent charge buildup from the electron beam. Few biological specimens tolerate these conditions without rapid collapse (Heslop-Harrison, 1970) and fewer still survive (Read & Lord, 1991). Uncoated non-conductors build up local concentration of electron, referred to as-charging- that prevent the formation of usable images. Energy X-ray Spectroscopy (EDS) can be used to determine the elemental composition of surface films in the SEM, but EDS analyses must be completed prior to deposition of the thin metal coating EDS data are typically collected from an area, the specimen must be removed from the specimen chamber and coating with a conductive layer, and returned to the SEM.
To allow observations under the high vacuum conditions of SEM, many preparations of biological samples have been developed, e.g., glutaraldehyde fixation, negative staining, the Sputter–Cryo technique, and coating with gold or osmium (Allan-Wojtas et al., 2008; Hassan et al., 2003; Lamed et al., 1987). Moreover, these preparations have some positive effects on the biological sample; for instance, they enhance contrast, reduce damage, and are uncharged up by the electron beam.
4. Biofilm formation: Environmental Scanning Electron Microscopy (ESEM)
A new SEM technique is now available which allows overcoming these obstacles. a modified, low-vacuum scanning electron microscopy technique for biofilm monitoring that enables imaging of hydrated specimens, termed environmental scanning electron microscopy (ESEM) also called variable pressure SEM (VP-SEM),was introduced in the mid-1990s (Little et al., 1991). The environmental SEM (reviewed in Stokes & Donald, 2000) uses a series of pressure limiting apertures (Muscariello et al., 2005) while preventing gas leakage from the specimen chamber, which can be maintained at 1–20 Torr. The ESEM is based upon the gaseous detection device (GDD). The main feature distinguishing ESEM from conventional SEM is the presence of a gas in the specimen chamber. Gases may include nitrous oxide, helium, argon and other, but water vapour is the most efficient amplifying gas found and the most common gas used in ESEM. The ionization GDD uses the ionization of the gas for the detection of secondary electrons from the specimen surface. It is a conical electrode about 1 cm in diameter that is positioned with the apex downward and concentric with the beam at the bottom of the pole piece. Secondary electrons emitted from the sample collide with water molecules in the chamber producing additional electrons and positive ions. The positive ions are attracted to the sample surface and eliminate the charging artifacts. A proportional cascade amplification of the original secondary electron signal results. With the GDD both secondary and backscattered electron images can be produced. Detailed technical explanations about this device can be found elsewhere (Danilatos, 1990).
The balance of gas flows into and out of the ESEM sample chamber determines its pressure.
The multiple apertures are situated below the objective lens and separate the sample chamber from the column. This feature allows the column to remain at high vacuum while the specimen chamber may sustain pressures as high as 50 Torr. The temperature and humidity of the sample can also be manually controlled to provide a suitable environment for maintaining the biological samples in their natural state.
The relative humidity in an ESEM specimen chamber can be controlled (Stokes & Donald, 2000), so ESEM is particularly useful for hydrated materials (Muscariello et al., 2005; Stokes & Donald, 2000; Stokes, 2001). A gaseous secondary electron detector (GSED) exploits the gas in the specimen chamber for signal amplification. BSED operation produces positive ions that have the added benefit of limiting charging of non-conductive specimens (Stokes & Donald, 2000). It does not require prior fixing and staining of the biofilm, minimizes biofilm dehydration and thus preserves native morphologies including surface structures (Walker et al., 2001) and native morphologies of bacteria and biofilms (e.g. Priester et al., 2007) and is able to achieve high magnifications, comparable with SEM. Shrinkage is prevented and artefact formation is reduced.
Additional advantages of ESEM include minimal processing of samples. It results in shorter time scales and lower costs while reducing the possibility of introducing artefacts. Samples can be preserved in saline in a common refrigerator (in fresh) if examination is to be deferred a few hours (Ramírez-Camacho et al., 2008). ESEM provides spatial resolutions of 10 nm or less. Compared to SEM, ESEM produces different, perhaps complementary, information for biological specimens (Doucet et al., 2005; Surman et al., 1996). Cell structures are visible with SEM, but external polymers around cells are more apparent in ESEM (Callow et al., 2003; Doucet et al., 2005; S. Douglas & D.D. Douglas, 2001).
4.1. ESEM applied of biofilm formation
Sutton et al., (1994) used this technique to study the structure of a
Scanning electron microscopes are frequently equipped with an energy dispersive x-ray analyser. This equipment permits elemental analysis with a high horizontal resolution of the inspected specimens. In this same context, mineral structures formed by bacterial and microalgal biofilms growing on the archaeological surface in Maltese hypogea were studied using Energy Dispersive X-Ray Spectroscopy (EDS) coupled to Environmental Scanning Electron Microscopy (ESEM), are reported by Zammit et al., (2011). These techniques have shown that mineral structures having different morphologies and chemical composition were associated with the microorganisms in the subaerophytic biofilm (Figure.5).
Interestingly, Shen et al., (2011) have been proposed a novel method for measuring an adhesion force of single yeast cell based on a nanorobotic manipulation system inside an environmental scanning electron microscope (ESEM) and Dubey & Ben-Yehuda (2011) report the identiﬁcation of analogous nanotubular channels formed among bacterial cells grown on solid surface. They demonstrate that nanotubes connect bacteria of the same and different species, thereby providing an effective conduit for exchange of intracellular content.
Scanning electron microscopy is a key tool to study the effect of physicochemical properties on adhesion phenomena (pH, roughness, topography, temperature, etc). SEM plays also a paramount role for assessing the microbial populations, three-dimensional structure, physiology, thickness, etc.
SEM proved to be an invaluable method for ultra-structural investigation, allowing imaging of the overall appearance and/or specific features of biofilms formed in different environments, e.g. microbial colonies and individual cells, the glycocalyx, and the presence of inorganic products within the biofilm.
Surely, Scanning Electron Microscope (SEM) is a powerful research tool, but since it requires high vacuum conditions, the wet materials and biological samples must undergo a complex preparation that limits the application of SEM on this kind of specimen and often causes the introduction of artifacts. The introduction of Environmental Scanning Electron Microscope (ESEM), working in gaseous atmosphere, represented a new perspective in biofilm monitoring with high resolution without prior fixing and staining.
ESEM could be useful as a complementary technique to help in the characterization of the structure and architecture of biofilms. In fact, ESEM could reveal the exact topography of intact, live and fully hydrated biofilms, with a higher magnification than the other microscopy techniques. In general, a combination of several techniques is to be recommended when investigating biofilms as the different techniques offer distinctly valuable information about different aspects of biofilm development.
Akernan K. K. Kuronen Ilpo. Olavi Kajander. E. 1993Scanning electron microscopy of nanobacteria- Novel biofilm producing organisms in blood. 15Supplement III.
Allan-Wojtas P. Hansen L. T. Paulson A. T. 2008Microstructural studies of probiotic bacteria loaded alginate microcapsules using standard electron microscopy techniques and anhydrous fixation. 41 1(January 2008), 101 108.
An Y. H. Dickinson R. B. Doyle R. J. 2000Mechanisms of bacterial adhesion and pathogenesis of implant and tissue infections. 1 27. In An, Y. H. & Friedman, R. J. (ed.), Handbook of bacterial adhesion: principles, methods, and applications. Humana Press, Totowa, N.J.
Bacteria to Polystyrene Surfaces: Effect of Temperature and hydrophobicity. 61(December 2010 554 559.
Beattie G. A. Lindow S. E. 1995The secret life of foliar bacterial pathogens on leaves. 33(September 1995), 145 117.
Bechmann R. T. Eduvean R. G. C. 2006AFM Study of the colonization of stainless steel by Aquabecterium commune. , 58 3-4, (October-December 2006), 112 118.
Beech I. 2004Biocorrosion: towards understanding interactions between biofilms and metals. , 15 3(Jun 2004), 181 186.
AS Blenkinsopp Costerton J. W. 1991Understanding bacterial biofilms. 9 1(January 1991), 138 143.
Bragadeeswaran S. Balasubramanian S. T. Raffi S. M. Rani Sophia. S. 2010Scanning electron microscopy elemental studies of primary film. . 10 2 169 172.
Brugnoni-I L. Lozano-E J. Cubitto-A M. 2007Potential of yeast isolated from apple juice to adhere to stainless steel surfaces in the apple juice processing industry. 40 3(April 2007), 332 340.
Bryers J.D. 2000Biofilms: an introduction, in Biofilms II: process analysis and applications, In: Bryers, JD, (Ed.), 3 11, Wiley-Liss, New York.
Busscher H. J. van de Belt-Gritter B. Dijkstra R. J. B. Norde W. van der Mei H. C. 2008Streptococcus mutans and Streptococcus intermedius adhesion to fibronectin films are oppositely influenced by ionic strength. , 24N.19, (August 2008), 10968 10973.
Callow J. A. Osborne M. P. Callow M. E. Baker F. Donald A. M. 2003Use of environmental scanning electron microscopy to image the spore adhesive of the marine alga Enteromorpha in its natural hydrated state. 27 4(Jun 2003), 315 321.
Camargo G. M. P. A. Pizzolitto A. C. Pizzolitto E. L. 2005Biofilm formation on catheters used after cesarean section as observed by scanning electron microscopy. , 90(August 2005), 148 149.
Carpentier B. Cerf O. 1993Biofilms and their consequences, with particular reference to hygiene in the food industry. , 75 6(March 1993), 499 511.
Characklis W. G. Marshall K. C. 1990Bioﬁlms: a basis for an interdisciplinary approach, 3 15. In: Characklis, W.G. & Marshall, K.C. (ed.), Bioﬁlms. John Wiley & Sons, New York, N.Y.
Costerton J. W. Irvin R. T. Cheng-J K. 1981The bacterial glycocalyx in nature and disease. , 35(October 1981), 299 324
Costerton J. W. Stewart P. S. Greenberg E. P. 1999Bacterial biofilms: a common cause of persistent infections. , 284 5418(May 1999), 1318 1322.
Danilatos G. D. 1990Theory of the gaseous detector device in the environmental scanning electron microscope. , 78 1 102.
Darkin M. G. Gilpin C. Williams J. B. Sangha C. M. 2001Direct wet surface imaging of an anaerobic biofilm by environmental scanning electron microscopy: application to landfill clay liner barriers, , 23 5 346 350.
Donlan R. M. 2001Biofilms and Device-Associated Infections. , 7 2(March-April 2001), 277 281.
Doucet F. J. Lead J. R. Maguire L. Achterberg E. P. Millward G. E. 2005Visualisation of natural aquatic colloids and particles- a comparison of conventional high vacuum and environmental scanning electron microscopy., 7 2(January 2005), 115 121.
Douglas S. Douglas D. D. 2001Structural and geomicrobiological characteristics of a microbial community from a cold sulfide spring. 18 4(November 2001), 401 422.
Dubey G. P. Ben-Yehuda S. 2011Intercellular nanotubes mediate bacterial communication. 144 4(February 2011), 590 600.
Duckett J. G. Ligrone R. 1995The formation of catenate foliar gemmae and the origin of oil bodies in the liverwort Odontoschisma denudatum (Mart.) dum (Jungermanniales): a light and electron microscope study. , 76(October 1995), 405 419.
(2008).Towards nanomicrobiology using atomic force microscopy. Dufrêne Y. F. (2008 6 6N.9, (September 2008), 674 680.
Dunne W. M. 2002Bacterial Adhesion: Seen Any Good Biofilms Lately? , 15 2(April 2002), 155 166.
Eighmy T. T. Maratea D. Bishop P. L. 1983Electron microscopic examination of wastewater biofilm formation and structural components. , 45 6 1921 1931.
Fassel T. A. Edmiston C. E. 1999Bacterial biofilms: strategies for preparing glycocalyx for electron microscopy. , 310 194 203.
Fassel T. A. Van Over J. E. Hauser C. C. Edmiston C. E. Sanger J. R. 1991Adhesion of staphylococci to breast prosthesis biomaterials: an electron microscopic evaluation. 1 199 208.
Filloux A. Vallet I. 2003Biofilm: set-up and organization of a bacterial community. , 19 1(January 2003), 77 83.
Flemming H. Neu T. R. Wozniak D. J. 2007The EPS Matrix: The "House of Biofilm Cells". , 189 22(November 2007), 7945 7947.
Fux C. Costerton J. Stewart P. Stoodley P. 2005Survival strategies of infectious biofilms. , 13 1(January 2005), 34 40.
Ganderton L. Chawla J. Winters C. Wimpenny J. Stickler D. 1992Scanning electron microscopy of bacterial biofilms on indwelling bladder catheters. , 11 9(September 1992), 789 796.
Garrett T. R. Bhakoo M. Zhang Z. 2008Bacterial adhesion and biofilms on surfaces. , 18 9(September 2008), 1049 1056.
Gilpin C. J. Sigee D. C. 1995X-ray microanalysis of wet biological specimens in the environmental scanning electron microscope. 1. Reduction of specimen distance under different atmospheric conditions. 1(July 1995), 22 28.
Gras M. H. Druetmichaud C. Cerf O. 1994La ﬂore bactérienne des feuilles de salade fraiche. 14 2 173 188.
Hall-Stoodley L. Costerton J. W. Stoodley P. 2004Bacterial Biofilms: from the natural environment to infectious diseases. , 2 2(February 2004), 95 108.
Hamadi F. Latrache H. Mabrrouki M. Elghmari A. Outzourhit A. Ellouali M. Chtaini A. 2005 Effect of pH on distribution and adhesion of Staphylococcus aureus to glass. , 19 1(November 2004), 73 85.
Hassan A. N. Frank J. F. . Elsoda M. 2003 Observation of bacterial exopolysaccharide in dairy products using cryo-scanning electron microscopy. , 13 9(July 2003), 755 762.
Herald P. J. Zottola E. A. 1988Scanning electron microscopic examination of attached to stainless steel at selected temperatures and pH values. Journal of Food Protection, 51 6(Jun 1988), 445 448
Heslop-Harrison Y. 1970Scanning electron microscopy of fresh leaves of . Science, 167 3815(January 1970), 172 174.
Kaplan J. B. Meyenhofer M. F. Fine D. H. 2003 Biofilm Growth and Detachment of Actinobacillus actinomycetemcomitans., 185 4(February 2003), 1399 1404.
Korstgens V. Flemming H. C. Wingender J. Borchard W. 2001 Influence of calcium ions on the mechanical properties of a model biofilm of mucoid Pseudomonas aeruginosa., 13 6 49 57.
Kouider N. Hamadi F. Mallouki B. Bengoram J. Mabrouki M. Zekraoui M. Ellouali M. Latrache H. 2010Effect of stainless steel surface roughness on Staphylococcus aureus adhesion. , 4 1(August 2009), 1 7.
Lamed R. Naimark J. Morgenstern E. Bayer E. A. 1987 Scanning electron microscopic delineation of bacterial surface topology using cationized ferritin. 7 4-5, (December 1987), 233 240.
Lewis K. 2005 Persister cells and the riddle of biofilm survival. (Moscow), 70 2(February 2005), 267 274.
Lim,J. 2008 Nanoscale characterization of Escherichia coli biofilm formed under laminar flow using atomic force microscopy (AFM) and scanning electron microscopy. , 11 2114 2118.
Little B. Wagner P. Ray R. Pope R. Scheetz R. 1991 Biofilms: an ESEM evaluation of artifacts introduced during SEM preparation. , 8 4 213 222.
Liu-Q Y. Liu Y. Tay-H J. 2004 The effects of extracellular polymeric substances on the formation and stability of biogranules 65 2(Jun 2004), 143 148.
Mallouki B. Latrache H. Mabrouki M. Outzourhit A. Hamadi F. Muller D. Ellouali M. 2007. The inhibitory effect of fucans on adhesion and production of slime of . Microbiologie Hygiène Alimentaire, 19 19 55(July 2007), 64 71.
Marshall K. C. Stout R. Mitchell R. 1971 Mechanism of the initial events in the sorption of marine bacteria to surfaces. Journal of Genetic of , 68 3(November 1971), 337 348
Mc Dowell J. W. Daryl B. S. Paulson S. Mitchell J. A. 2004 A simulated-use evaluation of a strategy for preventing biofilm formation in dental unit waterlines. 135 6(Jun 2004), 799 805.
Minoura N. Aiba S. I. Higuchi M. Gotoh Y. Tsukada M. Imai Y. 1995 Attachment and growth of fibroblast cells on silk fibroin. , 208 2(March 1995), 511 516.
Molin S. 2003 Gene transfer occurs with enhanced efficiency in biofilms and induces enhanced stabilisation of the biofilm structure., 14 3(Jun 2003), 255 261.
Morris C. E. Monier J. M. Jacques M. A. 1997 Methods for observing microbial biofilms directly on leaf surfaces and recovering them for isolation of culturable microorganism. 63 4(April 1997), 1570 1576.
Motta P. M. Makabe S. Naguro T. Correr S. 1994 Oocyte follicle cells association during development of human ovarian follicle. A study by high resolution scanning and transmission electron microscopy., 57 4(October 1994), 369 394.
Muscariello L. Rosso F. Marino G. Giordano A. Barbarisi M. Cafiero G. Barbarisi A. 2005 A critical overview of ESEM applications in the biological field., 205(Jun 2005), 328 334.
Nadell C. D. Xavier J. B. Levin S. A. Foster K. R. 2008 The Evolution of Quorum Sensing in Bacterial Biofilms,, 6 1(January 2008), e14 EOF.
O’Toole G. Kaplan H. B. Kolter R. 2000Biofilm formation as microbial development. 54 49 79.
Pawley J. B. Erlandsen S. L. 1989 The case for low voltage high resolution scanning electron microscopy of biological samples., 3(suppl), 16 173.
Peters G. Locci R. Pulverer G. 1982 Adherence and growth of coagulase-negative staphylococci on surfaces of intravenous catheters, 146 4 479 482.
Pinna A. Sechi L. A. Zanetti S. Delogu D. Carta F. 2000 Adherence of Ocular Isolates of Staphylococcus Epidermidis to ACRYSOF Intraocular Lenses. , 107 12(October 1982), 2162 2166.
Prakash B. Veeregowda B. M. Krishnappa G. 2003Biofilms: A survival strategy of bacteria. , 85 9(November 2003), 9 10.
Priester J. H. Horst A. M. Van De Werfhorst L. C. Saleta J. L. Mertes L. A. K. Holden P. A. 2007Enhanced visualization of microbial biofilms by staining and environmental scanning electron microscopy. 68 2(March 2007), 577 587.
Ramírez-Camacho R. González-Tallón A. I. Gómez D. Trinidad A. Ibáñez A. García-Berrocal J. R. Verdaguer J. M. González-García J. A. San Román. J. 2008Environmental scanning electron microscopy for biofilm detection in tonsils. 59 1(January 2008), 16 20.
Read N. D. Lord K. M. 1991Examination of living fungal spores by scanning electron microscopy. Experimental mycology, 15 2 132 139.
Richards S. R. Turner R. J. 1984 A comparative study of techniques for the examination of biofilms by scanning electron microscopy. , 18 6 767 773.
Rodrigues D. F. Elimelech M. 2009Role of Type 1 Fimbriae and Mannose in the Development of K12 Biofilm: From Initial Cell Adhesion to Biofilm Formation. Biofouling, 25 5(July 2009), 401 411.
Sasidharan S. Yoga Latha. L. Angeline T. 2010Imaging In vitro Anti-biofilm Activity to Visualize the Ultrastructural Changes. Microscopy: Science, Technology, Applications and Education A. Méndez-Vilas & J. Díaz (Eds.) Formatex, 2010, 622 626.
Schwartz T. Jungfer C. Heißler S. Friedrich F. Faubel W. Obst U. 2009Combined use of molecular biology taxonomy, Raman spectrometry, and ESEM imaging to study natural biofilms grown on filter materials at waterworks. 77 2(September 2009), 249 257.
Sevinç B. A. Hanley L. 2010 Antibacterial activity of dental composites containing zinc oxide nanoparticles. Part B: Applied Biomaterials, 94 1(July 2010), 22 31.
Shen Y. Nakajima M. Ahmad M. R. Kojima S. Hommac M. Fukuda T. 2011Effect of ambient humidity on the strength of the adhesion force of single yeast cell inside environmental-SEM. 8 1176 1183.
Singh P. K. Parsek M. R. Greenberg E. P. Welsh M. J. 2002 A component of innate immunity prevents bacterial biofilm development., 417 6888(May 2002), 552 555.
Stewart A. D. G. 1985The origins and development of scanning electron microscopy. 139 2(August 1985), 121 127.
Stokes D. J. Donald A. M. 2000In situ mechanical testing of dry and hydrated breadcrumb in the environmental scanning electron microscope (ESEM). 35 3(December 2000), 599 607.
Stokes D. J. Donald A. M. 2000In situ mechanical testing of dry and hydrated breadcrumb in the environmental scanning electron microscope (ESEM). 35 3(December 2000), 599 607.
Stokes D. J. 2001Characterization if soft condensed matter and delicate materials using environmental scanning electron microscopy (ESEM). 3 3 126 130.
Stoodley P. Sauer K. Davies D. G. Costerton J. W. 2002 Biofilms as complex differentiated communities., 56(January 2002), 187 209.
Storti A. CA Pizzolitto Pizzolitto L. E. 2005 Detection of mixed microbial biofilms on central venous catheters removed from intensive care unit patients. , 36 275 280.
Surico G. 1993 Scanning electron microscopy of olive and oleander leaves colonized by Pseudomonas syringae subsp. savastanoi. 138 1(May 1993), 31 40.
Surman S. B. Walker J. T. Goddard D. T. Morton L. H. G. Keevil C. W. Weaver W. Skinner A. Hanson K. Caldwell D. 1996 Comparison of microscope techniques for the examination of biofilms. , 25 1(March 1996) 57 70.
Sutherland I. W. 2001The biofilm matrix- an immobilized but dynamic microbial environment. , 9 5(May 2001), 222 227.
Sutton N. A. Hughes N. Handley P. S. 1994A comparison of conventional SEM techniques, low temperature SEM and the electroscan wet scanning electron microscope to study the structure of a biofilm of Streptococcus crista CR3. , 76 5(May 1994), 448 454.
Vidali M. 2001Bioremediation. A overview. , 73 7 1163 1172.
Walker J. T. Verran J. Boyd R. D. Percival S. 2001 Microscopy methods to investigate structure of potable water biofilms., 337 2001(July 2004), 243 255.
Weber W. J. J. Pirbazari M. Melson G. L. 1978 Biological growth on activated carbon: an investigation by scanning electron microscopy. , 12 7(July 1978), 817 819.
Zameer F. Gopal S. 2010Evaluation of antibiotic susceptibility in mixed culture biofilms. 6 1 93 99.
Zammit G. Sánchez-Moral S. Albertano P. 2011 Bacterially mediated mineralisation processes lead to biodeterioration of artworks in Maltese catacombs. , 409 14(Jun 2011), 2773 2783.
Zeraik A. E. Nitschke M. 2010Biosurfactants as agents to reduce adhesion of pathogenic