biofilm confocal de la matrix

8/18/2019 Biofilm Confocal de La Matrix

1/38

Confocal Microscopy Imaging of the Biofilm Matrix

Sebastian Schlafer, Rikke L. Meyer

PII: S0167-7012(16)30036-7

DOI: doi: 10.1016/j.mimet.2016.03.002

Reference: MIMET 4851

To appear in: Journal of Microbiological Methods

Received date: 16 October 2015

Revised date: 29 February 2016

Accepted date: 2 March 2016

Please cite this article as: Schlafer, Sebastian, Meyer, Rikke L., Confocal Mi-croscopy Imaging of the Biofilm Matrix, Journal of Microbiological Methods (2016), doi:10.1016/j.mimet.2016.03.002

This is a PDF file of an unedited manuscript that has been accepted for publication.As a service to our customers we are providing this early version of the manuscript.The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production processerrors may be discovered which could affect the content, and all legal disclaimers thatapply to the journal pertain.

http://dx.doi.org/10.1016/j.mimet.2016.03.002http://dx.doi.org/10.1016/j.mimet.2016.03.002http://dx.doi.org/10.1016/j.mimet.2016.03.002http://dx.doi.org/10.1016/j.mimet.2016.03.002


2/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Confocal Microscopy Imaging of the Biofilm Matrix

REVISED

Sebastian Schlafera*

, Rikke L. Meyerb,c

aDepartment of Dentistry, HEALTH, Aarhus University, Vennelyst Boulevard 9, 8000 Aarhus C, Denmark

bInterdisciplinary Nanoscience Center (iNANO), SCIENCE AND TECHNOLOGY, Aarhus University, Gustav

Wieds Vej 14, 8000 Aarhus C, Denmark

cDepartment of Bioscience, SCIENCE AND TECHNOLOGY, Aarhus University, Ny Munkegade 114, 8000

Aarhus C, Denmark

E-mail addresses: SS: [email protected]; RLM: [email protected]

*Corresponding author

Sponsor: Robert S. Burlage


3/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Abstract

The extracellular matrix is an integral part of microbial biofilms and an important field of research. Confocal

laser scanning microscopy is a valuable tool for the study of biofilms, and in particular of the biofilm matrix,

as it allows real-time visualization of fully hydrated, living specimens. Confocal microscopes are held by

many research groups, and a number of methods for qualitative and quantitative imaging of the matrix

have emerged in recent years. This review provides an overview and a critical discussion of techniques used

to visualize different matrix compounds, to determine the concentration of solutes and the diffusive

properties of the biofilm matrix.

Keywords: Biofilm; CLSM; confocal microscopy; extracellular matrix; EPS; fluorescent stains


4/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Abbreviations

CBM: carbohydrate-binding modules; CLSM: confocal laser scanning microscopy; DDAO: 1,3-dichloro-7-

hydroxy-9,9-dimethyl-2(9H)-acridinone; ECM: extracellular matrix; eDNA: extracellular DNA; EPS:

extracellular polymeric substances; FLIM: fluorescence lifetime imaging; FRET: Förster resonance energy

transfer; GFP: green fluorescent protein; MALDI: matrix-assisted laser desorption ionization; PI: propidium

iodide; SECM: scanning electrochemical microscopy; SIMS: secondary ion mass spectrometry; SPT: single

particle tracking; STED: stimulated emission depletion microscopy; ThT: thioflavin T; WGA: wheat germ

agglutinin


5/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

1 Introduction

The extracellular matrix of microbial biofilms is a highly complex scaffold, characterized by a multitude of

structurally and chemically heterogeneous microenvironments. Its functions are manifold: It provides

mechanical stability to the biofilm and protects the microorganisms from desiccation. It can act as a barrier

against adverse chemical and biological influences, such as osmotic stress, acid/base challenges, oxygen,

antibiotics and antiseptics, the host immune defense, and grazing protozoa. Moreover, it contributes to the

sorption and storage of nutrients and trace elements, it is the location of numerous extracellular enzymatic

reactions, and it keeps the microorganisms in tight contact to each other to facilitate genetic exchange and

bacterial communication. If the biofilm is a microbial city, then the matrix is its infrastructure.

Polysaccharides were long believed to be the main macromolecular constituent of the extracellular matrix,

and the abbreviation EPS, today used for extracellular polymeric substances, originally designated

extracellular polysaccharides. Today it is well-known that a multitude of different biopolymers, including

DNA, proteins, and lipids, i.e. in outer membrane vesicles, contribute to matrix structure and function.

For decades, the cellular components of biofilms held the center of research attention, as the

microorganisms are the driving force behind both detrimental and beneficial effects of biofilms. The past

ten years witnessed an increased focus on the matrix and its functional interplay with the microbiota. An

integrated view on both compartments is necessary to attain in-depth understanding of biofilms and to

develop target-oriented strategies for the control of biofilm-related problems.

Confocal laser scanning microscopy (CLSM) is a valuable tool for the study of biofilms, and in particular of

the biofilm matrix, as it allows real-time visualization of fully hydrated, living specimens. The past years

have brought about several new imaging technologies that improve the spatial resolution of light

microscopy, and the Nobel Prize in Chemistry was in 2014 awarded to Eric Betzig, Stefan W Hell and

William E. Moerner for their development of super-resolution optical microscopy. Confocal laser scanning


6/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

microscopes are available in many research laboratories, and consequently, methods based on CLSM have

evolved considerably in the past decade to retrieve information about the composition and the properties

of the biofilm matrix. The aim of this review is to provide an overview and to discuss the opportunities and

challenges of fluorescence labelling techniques that can be used to acquire either qualitative or

quantitative information about the biofilm matrix.

2 Qualitative confocal microscopy imaging of matrix components

The functionality of bacterial biofilms is entwined with its microscale structure, as mass transport by

diffusion and convection affects chemical gradients that dictate the limits of metabolic activity and the

conditions in the microenvironment experienced by individual cells. The physical structure also affects the

mechanical stability of the biofilm, and the protective properties of the matrix towards host immune cells

and antimicrobial agents. There is currently no fluorescence labeling method available which visualizes the

biofilm matrix in general, and this is due to the complex and highly variable composition of the matrix

produced by different bacteria and under different environmental conditions. Each matrix component must

therefore be stained individually.

2.1 Polysaccharide staining

Polysaccharides are often an important part of the biofilm matrix where they contribute to cohesion,

retention of water, sorption of organic and inorganic compounds, and protection against biocides and

grazing protozoa (for recent reviews, see Aricola et al. (Arciola et al., 2015) or Flemming and Wingender

(Flemming and Wingender, 2010)). Unfortunately a general stain for polysaccharides does not exist, as the

chemical structure of matrix polysaccharides differs among different bacteria. Calcofluor white has been

used for polysaccharide staining, but it binds only β-1,3 and β-1,4 glucans (Rasconi et al., 2009), which are

found in cellulose and chitin but not in the more common matrix polysaccharide poly-β-1,6-N-acetyl


7/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

glucosamine (Sadovskaya et al., 2005). A better approach is therefore the use of fluorescently labelled

lectins, which was pioneered by Neu et al . (Neu et al., 2001). Lectins typically recognize specific di- or tri-

saccharides. Such oligosaccharides can be present both in the matrix and as glycoconjugates on the cell

surface e.g. in the teichoic acids of Gram-positive bacteria and the lipopolysaccharides of Gram-negative

bacteria. Glycoconjugates are also highly diverse in structure (Messner et al., 2013), and lectin staining

therefore always starts with a large screening of commercial lectins to identify which are able to bind.

A new approach to carbohydrate staining was recently introduced by Nguyen et al. (Nguyen et al., 2014),

exploiting the high affinity of carbohydrate-binding modules (CBM): the non-catalytic carbohydrate-binding

domain of polysaccharide-degrading enzymes. The authors constructed a green fluorescent (GFP) fusion

protein with the carbohydrate-binding module 3 (GFP-CBM3) which has high affinity for cellulose. As a

proof of concept, they showed that this new polysaccharide label did not bind planktonic cells, but only to

E. coli biofilms and flocs induced by overexpression of the cellulose synthase BscB (Nguyen et al., 2014).

The same authors elegantly turned this concept into a quantitative and non-destructive assay for

exopolysaccharides, as GFP was cleaved off the GFP-CBM3 fusion protein by a site-specific protease and

subsequently quantified in solution (Ojima et al., 2015b). With 64 families of CBM, there is a wide scope for

using this novel approach to visualize and quantify other polysaccharides in the biofilm matrix.

2.2 Staining of extracellular DNA (eDNA)

The significance of eDNA in the biofilm matrix was discovered when Whitchurch et al. added DNase to a

Pseudomonas aeruginosa biofilm and watched the biofilm disappear (Whitchurch et al., 2002). Since then,

it has become evident that eDNA plays an important role for bacterial attachment and the early stages of

biofilm formation in many species from across the phylogenetic tree. Despite the apparent ubiquity of

eDNA as an important matrix component, it remains unclear what it interacts with in the matrix. Co-

localization of eDNA with other elements of the matrix has led to the suggestion that eDNA interacts with a

variety of different components, such as DNA-binding integration host factor (Goodman et al., 2011),


8/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

pyocyanines (Das et al., 2013), Staphylococcus β-toxin (Huseby et al., 2010) and polysaccharides (Hu et al.,

2012; Jennings et al., 2015). These findings suggests a variety of ligands and ways for DNA to interact. The

desire to address eDNA’s role in biofilm formation and antibiotic resistance has motivated the analysis of

eDNA in the biofilm using cell-impermeant DNA-binding fluorescent stains, such as propidium iodide (PI),

1,3-dichloro-7-hydroxy-9,9-dimethyl-2(9H)-acridinone (DDAO), TOTO-1, TO-PRO 3, PicoGreen and

SYTOX stains. Most reports have used DDAO for staining eDNA in biofilms after the initial publication by

Allesen-Holm et al. (Allesen-Holm et al., 2006; Conover et al., 2011; Schooling et al., 2009). However, a

recent evaluation of eDNA stains in biofilms of Pseudomonas, Staphylococcus and Bacillus species showed

that TOTO-1, SYTOX Green and PI provide the most reliable results, whereas TO-PRO-3 and DDAO

were not completely cell impermeant (Okshevsky and Meyer, 2014). PicoGreen also becomes cell

permeant after 10-15 minutes of incubation, but by keeping incubation times short, Tang et al (Tang et al.,

2013) used this sensitive and quantitative stain to quantify eDNA in biofilms of environmental isolates. They

demonstrated a transient or a gradually increasing accumulation of eDNA over time in the biofilm matrix of

the different isolates, suggesting that the role of eDNA in the biofilm matrix is highly dynamic. Suprisingly,

eDNA strongly affected the initiation of biofilms, even when present in concentration below the detection

limit. The amount of eDNA accumulating in biofilms may therefore not necessarily reflect its importances,

as it can exert its adhesive effect at very low concentrations.

eDNA is often stained in combination with another cell-permeant DNA-binding stain, such as the

combination of PI with SYTO 9 in the LIVE/DEAD BacLightTM

kit for viability. However, using two stains

binding to the same target molecule requires optimization of the concentrations used to ensure accurate

identification of eDNA, which is exposed to both stains. One has to consider the relative concentration of

the two stains as well as stain concentration relative to the amount of DNA in the sample. Simultaneous

intercalation of the two stains can lead to Förster resonance energy transfer (FRET) if the emission

spectrum of one stain overlaps with the excitation spectrum of the other, and it turns out that this effect

works to one’s advantage in SYTO 9/ PI staining. PI cannot completely displace SYTO 9, but when used


9/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

in the right concentration, the FRET effect quenches emitted light from SYTO 9 and leads to enhanced

emission by PI. However, if not used at the right concentration, the high quantum yield of SYTO 9

compared to PI will lead to simultaneous emission from both stains (Stocks, 2004). Due to these

complications, it is recommended to stain eDNA with TOTO-1 in combination with e.g. SYTO 60, as

TOTO-1 has low intrinsic fluorescence and a high quantum yield (twice that of SYTO 60). Furthermore,

fast one-track scanning can be performed with microscopes containing two photomultipliers, as the

emission spectra of the two stains are easily separated. As an example of TOTO-1/SYTO 60 staining,

Figure 1 shows the increasing accumulation of eDNA over time in S. epidermidis biofilms. The eDNA is

tightly associated with the cell surface, and the inhomogeneous distribution demonstrates the stark cell to

cell variations in the ability to bind eDNA to the cell surface.

2.3 Protein staining

In recent years, it has become evident that proteins also can be important for the biofilm matrix, and

proteins are in some cases even more predominant than polysaccharides. For example, cell wall anchored

proteins in e.g. Staphylococcus aureus and Staphylococcus epidermidis contribute to aggregation through

homophilic interactions (Geoghegan et al., 2010; Schaeffer et al., 2015), or by interacting with matrix

components originating from the host, such as collagen, fibrin and fibronectin (Büttner et al., 2015; Foster

and Höök, 1998). The protein-component of the biofilm matrix can be visualized with non-specific stains,

such as the FilmTracerTM

SyPro

stains (Frank and Patel, 2007; Lawrence et al., 2003), but specific protein

labeling is also possible through monoclonal antibodies. So far, antibody labeling in the biofilm matrix has

mostly been used for imaging the localization of proteins by electron microscopy (Webster et al., 2006),

but the technique is easily transferrable to fluorescence microscopy by using fluorescently labeled primary

or secondary antibodies (Greiner et al., 2005). Berk et al. (2012) elegantly showed how the genetic

insertion of FLAG tags in specific matrix proteins allowed following the production and location of key

matrix proteins in real time during initiation and development of biofilms in Vibrio cholera, demonstrating


10/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

complementary architectural roles of the three proteins Bap1, RbmA and RbmC. This powerful approach

will undoubtedly reveal new insights into the specific roles of different matrix proteins, which at first glance

appear to have overlapping roles in the biofilm formation of e.g. staphylococci (Christner et al., 2010).

Proteins that fold into a cross-beta structure and polymerize into insoluble fibers are called amyloids.

Amyloids initially received attention due to their role in neurodegenerative diseases where they occur due

to a misfolding of proteins, but it turns out that bacteria purposely produce amyloids, and Larsen et al.

(2007) changed our perception of amyloids when showing that they are abundant in bacterial biofilms from

a variety of different habitats. Amyloid fibers are resistant to degradation by proteases and they contribute

to the structural integrity of biofilms by e.g. Bacillus subtilis (Romero et al., 2010) and Staphylococcus

aureus (Schwartz et al., 2012). A recent study showed that over-expression of amyloids in the matrix of

Pseudomonas fluorescens biofilms led to a 20-fold increase in the biofilm stiffness (Zeng et al., 2015). As

methods for amyloid detection improve, we will learn more about the role of amyloid production in the

ecology of biofilms.

Larsen et al. (2007) compared the specificity of thioflavin T (ThT) and Congo red staining with detection by

amyloid-specific antibodies and showed that ThT was highly sensitive and more easily penetrated biofilms

compared to the antibodies, but it was less specific, probably due to its ability to bind DNA (Ilanchelian and

Ramaraj, 2004). Antibodies are thus more appropriate for amyloid visualization in complex samples, but the

cumbersome procedure with multiple incubation steps at different temperatures to allow the primary and

secondary antibodies to bind, does not allow time-resolved imaging. However, a recent study presents a

novel fluorescent probe, CDy11, with specificity for amyloid and looks like a promising new approach that is

even suitable for in vivo detection of bacterial amyloid in e.g. Pseudomonas aeruginosa biofilm infections

(Kim et al., 2016).

2.4 Lipid staining


11/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Production of biosurfactants is an important part of the biofilm life cycle in many bacteria. Biosurfactants

include polysaccharides, proteins, lipoproteins, glycolipids and lipopeptides, and in the context of biofilms

they can have very diverse functions in the production, maturation and dispersal of biofilms (Raaijmakers et

al., 2010). Lipids in general (including membranes) can be stained with Nile red, which also binds to

hydrophobic domains of proteins. Nile red is not confined to the extracellular matrix and will therefore

stain the cell membranes as well as intracellular lipids. It has been used extensively to visualise intracellular

lipophilic storage compounds, such as polyhydroxyalkanoates (Zuriani et al., 2013). The Nile red emission

peak differs according to whether it binds to polar or nonpolar lipids (Diaz et al., 2008), and this was

exploited to show that Rhodococcus strain RC291 with a hydrophobic cell surface associated closely with

both polar and non-polar lipids in the biofilm matrix, suggesting an important role of lipids in the biofilm

architecture of this strain (Andrews et al., 2010). Alternatives to Nile Red are the hydrophobic BODIPY

dyes and carbocyanine DiD, which will stain lipids, membranes and other hydrophobic compounds (Baird et

al., 2012; Rumin et al., 2015). In contrast to Nile Red and BODIPY dyes, FM stains, which brightly stain

lipids in membranes, do not enter the cytoplasm. They are extensively used to study endocytosis and

exocytosis in plants (Bolte et al., 2004), and would be ideal to study the formation of outer membrane

vesicles, which can be a significant part of the biofilm matrix (Ojima et al., 2015a; Schooling and Beveridge,

2006) and have been suggested to interact with or contribute to release of extracellular DNA (Sahu et al.,

2012; Schooling et al., 2009).

2.5 Monitoring of enzyme reactions

Various enzymatic reactions are carried out in the extracellular matrix of biofilms, and a multitude of

enzyme activity assays based on the detection of fluorescent products have been developed (Barizuddin et

al., 2015; Ju et al., 2015; Ko et al., 2015; Walther et al., 2015). While diffusion of the reaction product

renders the visualization of enzymatic activities difficult, a fluorogenic phosphatase substrate that forms

precipitates at the site of reaction (ELF 97) is commercially available. Van Ommen Kloeke and Geesey


12/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

used the assay in activated sludge, combining it with fluorescence in situ hybridization, albeit with an

epifluorescence microscope (Van Ommen Kloeke, F. and Geesey, 1999). They were able to demonstrate

that the cytophaga-flavobacteria group makes an important contribution to the removal of phosphorus

from wastewater.

3 Confocal microscopy approaches to quantitative analysis of the biofilm matrix

In addition to the many qualitative descriptions of the ECM that have been performed, a number of studies

have used CLSM based approaches that investigate different properties of the biofilm matrix quantitatively.

These require the combination of confocal microscopy with digital image analysis and/or mathematical

modelling. Due to the intense development of specialized image processing software with user-friendly

interfaces and the implementation of techniques such as fluorescence lifetime imaging (FLIM) or

fluorescence recovery after photobleaching (FRAP) in commercially available confocal microscopes,

quantitative methods have seen a rise in recent years.

3.1 Geometric measurements

Geometric measurements performed on fluorescently labelled structural components of the ECM

contribute in many ways to a better understanding of matrix functionality. Area and biovolume calculations

permit to determine the predominant matrix components and their spatial distribution at different time

points during biofilm formation. Measuring of distances and colocalization patterns between different

matrix components allows drawing conclusions about the interplay between different molecules, and

analysing colocalization patterns between matrix components and microbial cells can help to identify e.g.

EPS producers in multispecies biofilms.

While these kinds of geometric measurements are routinely carried out on the cellular components of

biofilms (Bridier et al., 2010; Dige et al., 2012; Guo et al., 2013a; Lupini et al., 2011), until now most studies


13/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

investigating the ECM have been limited to qualitative or semi-quantitative descriptions. Only a few reports

performed quantitative geometric analyses of the biofilm matrix and the relationship between matrix

components and cells (Fish et al., 2015; Houari et al., 2013; Kuehn et al., 2001; Lawrence et al., 1998;

Sweity et al., 2011). The comprehensive work of Koo and collaborators who investigated the amount of

fluorescently labelled polysaccharides in the ECM of Streptococcus mutans biofilms should be noted

(Falsetta et al., 2012; Klein et al., 2009; Klein et al., 2011; Xiao and Koo, 2010). They showed that starch in

the growth medium leads to increased amounts of EPS in the presence of sucrose, and that a combination

treatment of biofilms with myricetin, farnesol and fluoride targets genes involved in matrix production and

reduces EPS production. As extracellular polysaccharides in dental biofilm play an important role for biofilm

stability and the conservation of low pH at the tooth surface, quantitative studies of the effect of different

treatments on EPS production are particularly valuable to identify new therapeutic approaches to caries

control. In general, geometric measurements of structural matrix components should be performed more

frequently to facilitate comparisons between different reports, and to reduce the bias that might arise if

phenomena are described on the basis of subjective observations that might not be representative for the

entire sample.

Digital image analysis tools for geometric analyses of confocal microscopy images are readily available:

Specially designed programs, such as Comstat, daime, CMEIAS (all freeware), Imaris, Amira, Volocity, Arivis

and the open source software environments ImageJ, Icy and BioImageXD provide a number of possibilities

for quantitative structural analysis of fluorescence images.

When geometric measurements in biofilms are performed, microscope settings must be chosen carefully,

ideally by calibration with fluorescent beads of known size (Lawrence et al., 1998). Pinhole size, detector

gain and amplifier gain/offset have been shown to influence area measurements based on confocal images

(Sekar et al., 2010). Overexposure must be avoided and microscope settings should be kept constant

throughout a series of experiments.


14/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

The most critical step in the subsequent image analysis is the segmentation process, the differentiation

between stained objects and background fluorescence. A number of different algorithms, e.g. based on

intensity thresholding, edge detection or region-growing, can be employed to identify objects. The ideal

algorithm and segmentation parameters have to be determined individually for a particular set of biofilm

samples. In any case, meticulous visual inspection of the segmented images and comparison to the original

image data are mandatory to ascertain proper calculations.

Not only the area or volume covered by different structures of the ECM, but also their spatial arrangement

can be investigated quantitatively. Colocalization analyses were first performed to determine the spatial

interaction of different bacterial populations in activated sludge flocs (Rodenacker et al., 2000). This kind of

analysis has become more widely used for bacterial cells after its incorporation into the capability of the

image analysis software daime (Augspurger et al., 2010; Kara et al., 2007; Maixner et al., 2006; Schillinger

et al., 2012). The program employs a linear dipole algorithm (Reed and Howard, 1999) to determine if two

different populations of fluorescently labelled objects co-localize (Daims et al., 2006). Images subjected to

colocalization analysis must be cleared carefully for artifacts (i.e. by using algorithms that remove objects

up to a certain pixel size), as the presence of fluorescent objects other than the investigated populations,

irrespective of their size, will lead to erroneous results. Moreover, the physical properties of the samples

must be taken into account. Areas without biofilm formation, including carrier materials, need to be

excluded from the image analysis. So far, colocalization analyses of ECM components have been employed

in laboratory biofilms to determine the spatial relationship between dental bacteria and fluorescently

labelled dextrans (Xiao et al., 2012), and to study the colocalization of eDNA and wheat germ agglutinin

(WGA) targeted exopolysaccharides in Myxococcus xanthus biofilms (Hu et al., 2012). The software Duostat

(www.imageanalysis.dk) and the ImageJ plugin JACoP (Schneider et al., 2012) were used for calculations in

the respective studies. An increased focus on quantitative analyses of spatial arrangements in the biofilm

matrix is highly desirable, as it can expand our knowledge on the production and function of different ECM

components, especially in multispecies biofilms.


15/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

3.2 Measuring concentrations of diffusing molecules

Many dissolved molecules are important for biofilm development, maintenance and virulence. Oxygen and

carbon dioxide concentrations, pH, and metal ions such as Ca2+

, Mg2+

, Zn2+

and Fe3+

have dramatic effects

on metabolic processes carried out in biofilms. For all of these molecules, different fluorescent probes are

commercially available, and within a certain range, the fluorescence intensity correlates with the molecule

concentration. At first glance, it might seem straightforward to use fluorescence intensity measurements to

determine local concentrations of small solutes. However, a number of factors other than bulk

concentration of the dye affect the fluorescence. Microscope parameters such as laser power, detector

gain and amplifier offset/gain are crucial and difficult to standardize, and fluorescent probes are bleached

to a different extent during measurements, depending on the sample processing. Moreover, the local

probe concentration in a biofilm is unknown. All molecules face differences in penetration, reaction-

diffusion limitations and compartmentalization in biofilms, which makes it impossible to use calibration

data obtained from homogenous buffer solutions. Finally, interactions with biopolymers or other,

simultaneously employed dyes might alter the fluorescence properties of a probe. Consequently, only

semiquantitative comparisons can be made, even if specimens are handled in identical ways and imaged

with identical settings (Epstein et al., 2011; Guo et al., 2013b).

3.2.1 Immobilization of fluorescent dyes in particles

Several strategies can be employed to circumvent these problems. A fluorescent probe for a particular

solute might be applied along with a reference dye that emits light in a different part of the spectrum and

does not show a spectral response to the solute (Barker et al., 1998; de los Rios, Asuncion et al., 2003).

Calculating the ratio between the fluorescence from both dyes would then allow determination of the

solute concentration, but only if the concentration ratio of the dyes is identical in every location of the

biofilm. As the diffusive properties and the binding patterns of two different stains in a biofilm might differ,


16/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

both stains have to be immobilized, tied together with a fixed concentration ratio to enable reliable

measurements.

To date, only two studies have taken this approach, immobilizing a sensitive fluorescent dye and a

reference dye on particles. Hidalgo et al. (2009) investigated pH microenvironments in E. coli and mixed

species wastewater biofilms, using core shell silica nanoparticles containing covalently bound pH sensitive

fluorescein isothiocyanate, and Cy5 as the reference stain (Hidalgo et al., 2009). Acosta et al. monitored

oxygen gradients in S. aureus biofilms using silica microparticles containing Ru(Ph2phen3)Cl2, which is

quenched in an oxygen dependent fashion, and Nile blue chloride as reference (Acosta et al., 2012). Dyes

immobilized on a particle offer good photostability, and due to the embedding in the silica matrix,

interactions with biopolymers and their potential influence on the emission spectra are reduced. Still, the

effect of biofilm components on fluorescence should be tested, as done by Hidalgo et al. (2009). Moreover,

calibration should be performed at the same temperature as the actual pH measurements, as the

fluorescence intensity might be temperature dependent. The simultaneous use of two dyes brings about

some difficulties. FRET between the two dyes must be excluded, and the emission spectra of the dyes must

not overlap. Visualization of the bacterial biomass with a third stain is complicated, as any overlap between

the dyes would affect the calculated fluorescence ratios. Acosta et al. (2012) used DyLightTM

488-labelled

antibodies, the emission of which partly overlaps with the one of Ru(Ph2phen3)Cl2. Hidalgo et al. (2009)

refrained from employing a third stain and used bright-field images to visualize bacterial cells, at the cost of

losing three-dimensional information.

The use of particle sensors for solute visualization has some inherent disadvantages. Production of the

sensors requires knowledge foreign to the field of microbiology, and unless the relative concentrations of

both dyes on the particles are stable, calibration must be performed for every new batch of particles.

Furthermore, biofilm penetration and the spatial distribution of the particles pose problems. The

microparticles employed by Acosta et al. did not penetrate established S. aureus biofilms and had to be


17/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

applied before and during biofilm growth. This precludes their use in in situ grown medical biofilms. Hidalgo

et al. tested different sizes of nanoparticles and found that only the smallest, 10 nm particles penetrated

the biofilms sufficiently. Incubation had to be performed for several hours, which makes the rapid

examination of in situ grown biofilms impossible. The 10 nm particles employed were not distributed

evenly across the biofilms, but left certain cell-free areas unstained, precluding calculation of pH in the

entire extracellular matrix. While bigger particles settle in the extracellular matrix, it cannot be excluded

that small particles are internalized by microbial cells. Interactions with intracellular macromolecules might

change their fluorescence properties, and moreover, the concentration of the investigated solute might

differ considerably between intracellular and extracellular compartments, due to bacterial homeostasis. In

the case of pH, averaging fluorescence ratios deriving from both intra- and extracellular areas is of little

relevance.

3.2.2 Fluorescence lifetime imaging (FLIM) and pH ratiometry

Some of the problems encountered with two dyes immobilized in a particle can be avoided when solutes

are quantified by FLIM of a single probe or when intrinsically ratiometric dyes are employed. For FLIM, dye

molecules are excited by a short light pulse, and the resulting fluorescence is recorded in a time-resolved

manner. Fluorescence decay over time changes with the surrounding environmental conditions and follows

an exponential function that is independent of the initial fluorescence intensity and thus probe

concentration. Vroom et al. made use of the pH dependent fluorescence lifetime of carboxyfluorescein to

determine pH in a 10-species laboratory dental biofilm (Vroom et al., 1999). The ratio of fluorescence

intensities recorded in two different time gates was calculated and translated into pH values.

FLIM uses the inherent fluorescence properties of a single dye in a concentration independent way to

quantify solutes. Immobilization is thus unnecessary, penetration of the dye into the biofilm is

unproblematic and interactions between different stains can be avoided. The same advantages also apply

to ratiometric dyes, and in addition, their use requires less advanced microscopy equipment. Ratiometric


18/38


19/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

They could show that C-SNARF-4 is upconcentrated in bacterial cells at acidic pH, and they used a digital

image analysis procedure based on intensity thresholding to remove the bacterial biomass from the

confocal microscopy images (Figure 2) (Schlafer et al., 2015). As this procedure offers a simple solution to

the problems described above, the authors recommend digital image post-processing to ascertain

adequate monitoring of extracellular pH. In dental biofilms, microscale landscaping of extracellular pH

contributes to our understanding of the caries process. It allows studying how pH profiles and mineral loss

of the underlying dental tissues correlate and it is a valuable tool to investigate the effect of caries

controlling agents on pH (Schlafer et al., 2016).

For all confocal microscopy approaches to solute quantification in biofilms the importance of the

calibration procedure needs to be stressed. A variety of environmental factors, such as temperature, the

presence of biopolymers and other fluorescent stains, and varying concentrations of other solutes than the

one in question might affect the fluorescence intensity of the chosen fluoroprobe. Ratiometric calcium-

sensitive dyes might serve as an example. The emission spectra of probes such as Fura-2 and Indo-1 are

dependent on calcium-binding, but also on temperature and pH (Larsson et al., 1999; Oliver et al., 2000). If

they were to be applied in an acid-producing biofilm, local variations in pH would render an appropriate

interpretation of the effect of calcium concentration on the observed fluorescence ratios impossible. While

a large number of fluorescent probes for different solutes are commercially available, increased attention

needs to be paid to their specific target-oriented calibration prior to experimental use. Ideally, calibration

of a dye should be performed in the presence of biofilms and all other stains employed concomitantly to

exclude adverse effects on the performed measurements. If the described technical difficulties can be

overcome, confocal microscopy measurements of solute concentrations can make an important

contribution to our understanding of the metabolic processes carried out in different microenvironments of

biofilms.


20/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Given the cell permeability of some pH-sensitive dyes, it is tempting to employ these fluoroprobes to

monitor intracellular changes of pH at the individual cell level (Shabala et al., 2006). For calibration, trans-

membrane gradients of bacterial cells can be collapsed using permeant acids and bases, but for reliable

intracellular pH measurements, it must be ascertained that the probe is exclusively located inside the

bacterial cells and not bound on the outside of the cell wall. It is impossible to determine this based on

confocal images alone. Martinez et al . elegantly circumvented this problem using genetically engineered

strains of E. coli and Bacillus subtilis that expressed the intracellular ratiometric protein pHluorin (Martinez

et al., 2012). Their approach, however, cannot be extended to in situ grown biofilms.

3.2.3 Förster resonance energy transfer (FRET)

FRET-based biosensing is another technique that might be exploited for concentration measurements. FRET

is widely used in cell biology for in situ, real-time monitoring of macromolecule interactions and

concentrations of intracellular metabolites and signaling molecules through FRET-based biosensors (Mohsin

et al., 2015; Shrestha et al., 2015). FRET-based biosensing involves a fluorescent donor (typically a

fluorescent protein) and an acceptor, which quenches the fluorescence of the donor upon interaction,

resulting in excitation and emission of light from the acceptor fluorophore. Although applications in

microbiology are less common, FRET-based biosensing is making its way into biofilm research, and a

biosensor was recently developed to detect the intracellular c-di-GMP concentration in Caulobacter

crescentus (Christen et al., 2010) and Salmonella typhimurium (Mills et al., 2015) through conformational

changes in the c-di-GMP binding protein YcgR fused with a donor and acceptor fluorescent protein at either

end. Despite the great potential for characterizing inter-molecular interactions and extracellular

concentrations of e.g. signaling molecules in biofilm, this approach is yet to see its first application in

studies of the biofilm matrix. The main requirement for extracellular detection is of course that the

genetically encoded biosensor must be transported outside of the cell, and secondly that the sensing

molecule then remains in the biofilm. One way to achieve this could be to initially focus the effort on


21/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

detecting interactions of cell wall- or cell membrane anchored proteins, for which fluorescent fusion

proteins can be generated. This was recently done for the membrane bound nuclease Nuc2 in S. aureus,

and a FRET-based assay was used – albeit not in combination with microscopy – to study the activity of this

extracellular nuclease in vitro and in vivo (Kiedrowski et al., 2014). Hence FRET-based biosensing in the

extracellular environment of a biofilm might not be so far into the future.

3.3 Measuring diffusion properties

3.3.1 Time-lapse imaging of fluorescent solutes

A number of different confocal microscopy techniques can be used to quantitatively investigate diffusion

properties in biofilms. Time-lapse imaging of fluorescent or fluorescently labelled molecules is a

straightforward approach: A biofilm is exposed to a fluorescent molecule under static or dynamic

conditions, and repeated confocal microscopy images are taken in a particular location. The fluorescence

intensity is recorded in a semiquantitative way, using the intensity in the bulk fluid or the intensity of a

stain that already has penetrated as a reference, until equilibrium is reached. The publications of Stewart

and coworkers should be noted, who found that antibiotic-sized tracers, daptomycin and even a variety of

macromolecular solutes penetrated bacterial clusters (Ø>100 µm) in laboratory biofilms within a few

minutes (Rani et al., 2005; Stewart et al., 2009; Takenaka et al., 2009). Their work and a similar study

conducted by Stone et al. (2002) proved the long standing theory wrong that the biofilm matrix constitutes

a significant penetration barrier against antibiotics.

3.3.2 Single particle tracking (SPT) microscopy

Time-lapse imaging can also be employed to track the fate of single nano- or microscale particles in

biofilms. Determining particle trajectories under flow conditions, as performed by Stoodley et al . (1994)

and Kuehn et al . (2001), allows collecting detailed information on the hydrodynamics in different parts of

the biofilm matrix, e.g. in bioreactors for wastewater treatment. Moreover, studies of particle diffusion


22/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

properties are useful to develop suitable carrier particles for controlled drug delivery to biofilms. Two

recent reports investigated the diffusional behavior of particles with different size and surface chemistry in

biofilms under static conditions (Birjiniuk et al., 2014; Forier et al., 2013). Both studies found that neutral

(PEGylated) particles diffuse more rapidly through biofilms than charged particles and that diffusion is size-

dependent. Moreover, Birjiniuk et al. (2014) showed that matrix density of E. coli biofilms increased with

age, leading to reduced diffusion, and that charge density increased in deeper layers of the biofilm. By

comparing the motion paths of particles present during biofilm growth and particles added after biofilm

growth they evidenced the presence of channels permitting rapid diffusion.

3.3.3 Fluorescence recovery after photobleaching (FRAP) and fluorescence correlation spectroscopy (FCS)

While time-lapse imaging is a valuable tool for diffusion measurements, it has a limited spatial and

temporal resolution. Diffusion is quantified over a rather large volume of the biofilm, i.e. cell clusters with a

radial dimension of 100 – 200 µm (Stewart et al., 2009), and local differences in diffusivity cannot be

resolved. Acquisition times for microscopic images are in the order of seconds, and the diffusion and

reactivity of the molecules in question cannot be tracked any longer, once equilibrium is reached. FRAP and

FCS can overcome these problems, and today both techniques can be implemented in commercially

available confocal microscopes. FRAP measures the increase of fluorescence intensity in a defined volume

of the matrix after irreversible photobleaching of the fluorophore. The resulting fluorescence recovery

curve is dependent on the inward diffusion of the fluorescent molecule into the bleached area. For FCS,

fluorescent dyes or fluorescently labelled molecules are applied in nanomolar concentrations, and the

durations and amplitudes of fluctuations in the fluorescence intensity are recorded in a defined volume of

the matrix. Both techniques allow determining diffusion coefficients in small volumes of the biofilm with a

temporal resolution in the order of milliseconds. Early FRAP approaches date back to the 1990’s (Bryers and

Drummond, 1998; Lawrence et al., 1994), but in particular, the effort of Briandet and coworkers should be

mentioned, who have improved both techniques considerably and applied them to study diffusion of


23/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

bacteriophages, dextrans and antibiotics in different biofilms (Briandet et al., 2008; Daddi Oubekka et al.,

2012; Guiot et al., 2002; Lacroix-Gueu et al., 2005; Waharte et al., 2010). For a review, see Bridier et al.

(2011). As biofilms are the causative agents of many diseases and responsible for biofouling in different

industrial fields, a detailed understanding of the diffusional behavior of macromolecules in biofilms is of the

utmost importance and contributes to the rational design of anti-biofilm agents.

3.3.4 Rheology measurements

As outlined above (3.3.2) the behavior of nano- or microscale particles can be used to describe diffusion

processes in biofilms, but it can also be employed to determine the rheological properties of the matrix.

Galy et al . (2012) embedded 2.8 µm sized magnetic particles in E. coli biofilms during growth and applied

defined forces to the particles via magnetic tweezers. Tracking the movements of the particles allowed

plotting creep curves from which the viscoelastic behavior of the matrix in defined locations could be

extracted. A similar approach was taken by Rogers et al. (2008) who tracked the trajectories of bacteria in

S. aureus and P. aeruginosa biofilms exposed to different flow regimes, albeit with bright field microscopy.

Mathias and Stoodley (2009) used digital image correlation to quantify the strains deriving from different

flow rates, equally based on bright field microscopy. The latter two approaches might be combined with

confocal imaging to quantitatively assess the mechanical properties of biofilms.


24/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

4 Conclusions

In recent years, there has been an increased research focus on the extracellular matrix in biofilms, which

has led to a better understanding of matrix complexity, its structural, chemical and physical organization.

Confocal microscopy techniques allow preserving the three-dimensional structure of biofilms, investigating

the matrix in fully hydrated state. Different structural matrix components and their spatial arrangement can

be studied, the concentrations of solutes and their role in biofilm physiology and virulence determined, and

the mechanical and diffusive properties of different microenvironments in the matrix probed. Being a

standard analysis tool in many research laboratories, the use of confocal microscopy evolves constantly and

will continuously provide deeper insight into the structure and functioning of the extracellular matrix.


25/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Figure captions

Figure 1. eDNA in Staphylococcus epidermidis biofilm becomes more abundant over time. The biofilm

was grown in Tryptic Soy Broth for 24 (A) or 48 hours (B) and stained with 2 µm TOTO-1 for eDNA (green),

and 20 µM SYTO 60 for intracellular DNA (red). Bar = 5 µm.

Figure 2. Ratiometric imaging of pH in in vivo grown dental biofilm exposed to glucose. The ratiometric

pH-sensitive dye C-SNARF-4 is employed in combination with digital image post-processing to visualize

extracellular pH in real-time. A) C-SNARF-4 is taken up by all bacterial cells in the biofilm, yielding

different levels of fluorescent intensity in extracellular and intracellular compartments. An overlay of

fluorescent emission in the green and red spectra is shown. B) The bacterial biomass has been removed

from the image in panel A) using digital image analysis. Extracellular pH has been calculated by dividing the

green by the red fluorescent intensities pixel-wise. A color lookup table was used to visualize pH. Acid

production in the observed field of view is moderate. With an average extracellular pH of 6.28, 5 min after

exposure to glucose, levels critical for enamel dissolution (ca. 5.5) have not yet been reached. Bars = 20 µm


26/38


27/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Büttner, H., Mack, D., Rohde, H., 2015. Structural basis of Staphylococcus epidermidis biofilm formation:

mechanisms and molecular interactions. Frontiers in cellular and infection microbiology 5, 14.

Christen, M., Kulasekara, H.D., Christen, B., Kulasekara, B.R., Hoffman, L.R., Miller, S.I., 2010. Asymmetrical

distribution of the second messenger c-di-GMP upon bacterial cell division. Science (New York, N.Y.)

328 (5983), 1295 –1297.

Christner, M., Franke, G.C., Schommer, N.N., Wendt, U., Wegert, K., Pehle, P., Kroll, G., Schulze, C., Buck, F.,

Mack, D., Aepfelbacher, M., Rohde, H., 2010. The giant extracellular matrix-binding protein of

Staphylococcus epidermidis mediates biofilm accumulation and attachment to fibronectin. Molecular

microbiology 75 (1), 187 –207.

Conover, M.S., Mishra, M., Deora, R., 2011. Extracellular DNA is essential for maintaining Bordetella biofilm

integrity on abiotic surfaces and in the upper respiratory tract of mice. PloS one 6 (2), e16861.

Daddi Oubekka, S., Briandet, R., Fontaine-Aupart, M.-P., Steenkeste, K., 2012. Correlative time-resolved

fluorescence microscopy to assess antibiotic diffusion-reaction in biofilms. Antimicrobial agents and

chemotherapy 56 (6), 3349 –3358.Daims, H., Lücker, S., Wagner, M., 2006. daime, a novel image analysis program for microbial ecology and

biofilm research. Environmental microbiology 8 (2), 200 –213.

Das, T., Kutty, S.K., Kumar, N., Manefield, M., 2013. Pyocyanin facilitates extracellular DNA binding to

Pseudomonas aeruginosa influencing cell surface properties and aggregation. PloS one 8 (3), e58299.

de los Rios, Asuncion, Wierzchos, J., Sancho, L.G., Ascaso, C., 2003. Acid microenvironments in microbial

biofilms of antarctic endolithic microecosystems. Environmental microbiology 5 (4), 231 –237.

Diaz, G., Melis, M., Batetta, B., Angius, F., Falchi, A.M., 2008. Hydrophobic characterization of intracellular

lipids in situ by Nile Red red/yellow emission ratio. Micron (Oxford, England : 1993) 39 (7), 819 –824.

Dige, I., Schlafer, S., Nyvad, B., 2012. Difference in initial dental biofilm accumulation between night and

day. Acta odontologica Scandinavica 70 (6), 441 –447.Epstein, A.K., Pokroy, B., Seminara, A., Aizenberg, J., 2011. Bacterial biofilm shows persistent resistance to

liquid wetting and gas penetration. Proceedings of the National Academy of Sciences of the United

States of America 108 (3), 995 –1000.

Falsetta, M.L., Klein, M.I., Lemos, J.A., Silva, B.B., Agidi, S., Scott-Anne, K.K., Koo, H., 2012. Novel antibiofilm

chemotherapy targets exopolysaccharide synthesis and stress tolerance in Streptococcus mutans to

modulate virulence expression in vivo. Antimicrobial agents and chemotherapy 56 (12), 6201 –6211.

Fish, K.E., Collins, R., Green, N.H., Sharpe, R.L., Douterelo, I., Osborn, A.M., Boxall, J.B., 2015.

Characterisation of the physical composition and microbial community structure of biofilms within a

model full-scale drinking water distribution system. PloS one 10 (2), e0115824.

Flemming, H.-C., Wingender, J., 2010. The biofilm matrix. Nature reviews. Microbiology 8 (9), 623 –633.

Forier, K., Messiaen, A.-S., Raemdonck, K., Deschout, H., Rejman, J., Baets, F. de, Nelis, H., De Smedt,

Stefaan C, Demeester, J., Coenye, T., Braeckmans, K., 2013. Transport of nanoparticles in cystic fibrosis

sputum and bacterial biofilms by single-particle tracking microscopy. Nanomedicine (London, England)

8 (6), 935 –949.

Foster, T.J., Höök, M., 1998. Surface protein adhesins of Staphylococcus aureus. Trends in microbiology 6

(12), 484 –488.

Frank, K.L., Patel, R., 2007. Poly-N-Acetylglucosamine Is Not a Major Component of the Extracellular Matrix

in Biofilms Formed by icaADBC-Positive Staphylococcus lugdunensis Isolates. Infection and immunity 75

(10), 4728 –4742.


28/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Franks, A.E., Nevin, K.P., Jia, H., Izallalen, M., Woodard, T.L., Lovley, D.R., 2009. Novel strategy for three-

dimensional real-time imaging of microbial fuel cell communities: monitoring the inhibitory effects of

proton accumulation within the anode biofilm. Energy Environ. Sci. 2 (1), 113 –119.

Galy, O., Latour-Lambert, P., Zrelli, K., Ghigo, J.-M., Beloin, C., Henry, N., 2012. Mapping of bacterial biofilm

local mechanics by magnetic microparticle actuation. Biophysical journal 103 (6), 1400 –1408.

Geoghegan, J.A., Corrigan, R.M., Gruszka, D.T., Speziale, P., O'Gara, J.P., Potts, J.R., Foster, T.J., 2010. Role

of surface protein SasG in biofilm formation by Staphylococcus aureus. Journal of bacteriology 192 (21),

5663 –5673.

Greiner, L.L., Edwards, J.L., Shao, J., Rabinak, C., Entz, D., Apicella, M.A., 2005. Biofilm Formation by

Neisseria gonorrhoeae. Infection and immunity 73 (4), 1964 –1970.

Guiot, E., Georges, P., Brun, A., Fontaine-Aupart, M.P., Bellon-Fontaine, M.N., Briandet, R., 2002.

Heterogeneity of Diffusion Inside Microbial Biofilms Determined by Fluorescence Correlation

Spectroscopy Under Two-photon Excitation¶. Photochemistry and Photobiology 75 (6), 570 –578.

Guo, K., Freguia, S., Dennis, P.G., Chen, X., Donose, B.C., Keller, J., Gooding, J.J., Rabaey, K., 2013a. Effects ofsurface charge and hydrophobicity on anodic biofilm formation, community composition, and current

generation in bioelectrochemical systems. Environmental science & technology 47 (13), 7563 –7570.

Guo, L., Hu, W., He, X., Lux, R., McLean, J., Shi, W., 2013b. investigating acid production by Streptococcus

mutans with a surface-displayed pH-sensitive green fluorescent protein. PloS one 8 (2), e57182.

Hassan, A.N., Frank, J.F., Farmer, M.A., Schmidt, K.A., Shalabi, S.I., 1995a. Formation of Yogurt

Microstructure and Three-Dimensional Visualization as Determined by Confocal Scanning Laser

Microscopy. Journal of Dairy Science 78 (12), 2629 –2636.

Hassan, A.N., Frank, J.F., Farmer, M.A., Schmidt, K.A., Shalabi, S.I., 1995b. Observation of Encapsulated

Lactic Acid Bacteria Using Confocal Scanning Laser Microscopy. Journal of Dairy Science 78 (12), 2624 –

2628.Hidalgo, G., Burns, A., Herz, E., Hay, A.G., Houston, P.L., Wiesner, U., Lion, L.W., 2009. Functional

tomographic fluorescence imaging of pH microenvironments in microbial biofilms by use of silica

nanoparticle sensors. Applied and environmental microbiology 75 (23), 7426 –7435.

Houari, A., Seyer, D., Kecili, K., Heim, V., Di Martino, P., 2013. Kinetic development of biofilm on NF

membranes at the Méry-sur-Oise plant, France. Biofouling 29 (2), 109 –118.

Hu, W., Li, L., Sharma, S., Wang, J., McHardy, I., Lux, R., Yang, Z., He, X., Gimzewski, J.K., Li, Y., Shi, W., 2012.

DNA builds and strengthens the extracellular matrix in Myxococcus xanthus biofilms by interacting with

exopolysaccharides. PloS one 7 (12), e51905.

Hunter, R.C., Beveridge, T.J., 2005. Application of a pH-sensitive fluoroprobe (C-SNARF-4) for pH

microenvironment analysis in Pseudomonas aeruginosa biofilms. Applied and environmental

microbiology 71 (5), 2501 –2510.

Huseby, M.J., Kruse, A.C., Digre, J., Kohler, P.L., Vocke, J.A., Mann, E.E., Bayles, K.W., Bohach, G.A.,

Schlievert, P.M., Ohlendorf, D.H., Earhart, C.A., 2010. Beta toxin catalyzes formation of nucleoprotein

matrix in staphylococcal biofilms. Proceedings of the National Academy of Sciences of the United States

of America 107 (32), 14407 –14412.

Ilanchelian, M., Ramaraj, R., 2004. Emission of thioflavin T and its control in the presence of DNA. Journal of

Photochemistry and Photobiology A: Chemistry 162 (1), 129 –137.


29/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Ju, H., Ryu, B.H., Doohun Kim, T., 2015. Identification, characterization, immobilization of a novel type

hydrolase (LmH) from Listeria monocytogenes. International journal of biological macromolecules 72,

63 –70.

Kara, D., Luppens, Suzanne B I, van Marle, J., Ozok, R., ten Cate, Jacob M, 2007. Microstructural differences

between single-species and dual-species biofilms of Streptococcus mutans and Veillonella parvula,

before and after exposure to chlorhexidine. FEMS microbiology letters 271 (1), 90 –97.

Kiedrowski, M.R., Crosby, H.A., Hernandez, F.J., Malone, C.L., McNamara, J.O., Horswill, A.R., 2014.

Staphylococcus aureus Nuc2 is a functional, surface-attached extracellular nuclease. PloS one 9 (4),

e95574.

Kim, J.-Y., Sahu, S., Yau, Y.-H., Wang, X., Shochat, S.G., Nielsen, P.H., Dueholm, M.S., Otzen, D.E., Lee, J.,

Delos Santos, May Margarette Salido, Yam, J.K.H., Kang, N.-Y., Park, S.-J., Kwon, H., Seviour, T., Yang, L.,

Givskov, M., Chang, Y.-T., 2016. Detection of Pathogenic Biofilms with Bacterial Amyloid Targeting

Fluorescent Probe, CDy11. J. Am. Chem. Soc. 138 (1), 402 –407.

Klein, M.I., Duarte, S., Xiao, J., Mitra, S., Foster, T.H., Koo, H., 2009. Structural and molecular basis of therole of starch and sucrose in Streptococcus mutans biofilm development. Applied and environmental


Klein, M.I., Xiao, J., Heydorn, A., Koo, H., 2011. An analytical tool-box for comprehensive biochemical,

structural and transcriptome evaluation of oral biofilms mediated by mutans streptococci. Journal of

visualized experiments : JoVE (47).

Ko, K.-C., Lee, B., Cheong, D.-E., Han, Y., Choi, J.H., Song, J.J., 2015. Bacterial cell surface display of a

multifunctional cellulolytic enzyme screened from a bovine rumen metagenomic resource. Journal of

microbiology and biotechnology.

Kuehn, M., Mehl, M., Hausner, M., Bungartz, H.J., Wuertz, S., 2001. Time-resolved study of biofilm

architecture and transport processes using experimental and simulation techniques: the role of EPS.Water Science and Technology 43 (6), 143 –151.

Lacroix-Gueu, P., Briandet, R., Lévêque-Fort, S., Bellon-Fontaine, M.-N., Fontaine-Aupart, M.-P., 2005. In

situ measurements of viral particles diffusion inside mucoid biofilms. Comptes rendus biologies 328

(12), 1065 –1072.

Larsen, P., Nielsen, J.L., Dueholm, M.S., Wetzel, R., Otzen, D., Nielsen, P.H., 2007. Amyloid adhesins are

abundant in natural biofilms. Environmental microbiology 9 (12), 3077 –3090.

Larsson, D., Larsson, B., Lundgren, T., Sundell, K., 1999. The effect of pH and temperature on the

dissociation constant for fura-2 and their effects on [Ca(2+)](i) in enterocytes from a poikilothermic

animal, Atlantic cod (Gadus morhua). Analytical Biochemistry 273 (1), 60 –65.

Lawrence, J., Neu, T., Swerhone, G., 1998. Application of multiple parameter imaging for the quantification

of algal, bacterial and exopolymer components of microbial biofilms. Journal of microbiological

methods 32 (3), 253 –261.

Lawrence, J.R., Swerhone, G. D. W., Leppard, G.G., Araki, T., Zhang, X., West, M.M., Hitchcock, A.P., 2003.

Scanning Transmission X-Ray, Laser Scanning, and Transmission Electron Microscopy Mapping of the

Exopolymeric Matrix of Microbial Biofilms. Applied and environmental microbiology 69 (9), 5543 –5554.

Lawrence, J.R., Wolfaardt, G.M., Korber, 1994. Determination of Diffusion Coefficients in Biofilms by

Confocal Laser Microscopy. Applied and environmental microbiology 60 (4), 1166 –1173.


30/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Lupini, G., Proia, L., Di Maio, M., Amalfitano, S., Fazi, S., 2011. CARD-FISH and confocal laser scanner

microscopy to assess successional changes of the bacterial community in freshwater biofilms. Journal of

microbiological methods 86 (2), 248 –251.

Maixner, F., Noguera, D.R., Anneser, B., Stoecker, K., Wegl, G., Wagner, M., Daims, H., 2006. Nitrite

concentration influences the population structure of Nitrospira-like bacteria. Environmental

microbiology 8 (8), 1487 –1495.

Martinez, K.A., Kitko, R.D., Mershon, J.P., Adcox, H.E., Malek, K.A., Berkmen, M.B., Slonczewski, J.L., 2012.

Cytoplasmic pH response to acid stress in individual cells of Escherichia coli and Bacillus subtilis

observed by fluorescence ratio imaging microscopy. Applied and environmental microbiology 78 (10),

3706 –3714.

Mathias, J.D., Stoodley, P., 2009. Applying the digital image correlation method to estimate the mechanical

properties of bacterial biofilms subjected to a wall shear stress. Biofouling 25 (8), 695 –703.

Messner, P., Schäffer, C., Kosma, P., 2013. Bacterial cell-envelope glycoconjugates. Advances in

carbohydrate chemistry and biochemistry 69, 209 –272.Mills, E., Petersen, E., Kulasekara, B.R., Miller, S.I., 2015. A direct screen for c-di-GMP modulators reveals a

Salmonella Typhimurium periplasmic ʟ-arginine-sensing pathway. Science signaling 8 (380), ra57.

Mohsin, M., Ahmad, A., Iqbal, M., 2015. FRET-based genetically-encoded sensors for quantitative

monitoring of metabolites. Biotechnology letters 37 (10), 1919 –1928.

Neu, T., Swerhone, G.D., Lawrence, J.R., 2001. Assessment of lectin-binding analysis for in situ detection of

glycoconjugates in biofilm systems. Microbiology (Reading, England) 147 (Pt 2), 299 –313.

Nguyen, M.H., Ojima, Y., Sakka, M., Sakka, K., Taya, M., 2014. Probing of exopolysaccharides with green

fluorescence protein-labeled carbohydrate-binding module in Escherichia coli biofilms and flocs induced

by bcsB overexpression. Journal of bioscience and bioengineering 118 (4), 400 –405.

Ojima, Y., Nguyen, M.H., Yajima, R., Taya, M., 2015a. Flocculation of Escherichia coli Cells in Associationwith Enhanced Production of Outer Membrane Vesicles. Applied and environmental microbiology 81

(17), 5900 –5906.

Ojima, Y., Suparman, A., Nguyen, M.H., Sakka, M., Sakka, K., Taya, M., 2015b. Exopolysaccharide assay in

Escherichia coli microcolonies using a cleavable fusion protein of GFP-labeled carbohydrate-binding

module. Journal of microbiological methods 114, 75 –77.

Okshevsky, M., Meyer, R.L., 2014. Evaluation of fluorescent stains for visualizing extracellular DNA in

biofilms. Journal of microbiological methods 105, 102 –104.

Oliver, A.E., Baker, G.A., Fugate, R.D., Tablin, F., Crowe, J.H., 2000. Effects of Temperature on Calcium-

Sensitive Fluorescent Probes. Biophysical journal 78 (4), 2116 –2126.

Rani, S.A., Pitts, B., Stewart, P.S., 2005. Rapid diffusion of fluorescent tracers into Staphylococcus

epidermidis biofilms visualized by time lapse microscopy. Antimicrobial agents and chemotherapy 49

(2), 728 –732.

Rasconi, S., Jobard, M., Jouve, L., Sime-Ngando, T., 2009. Use of calcofluor white for detection,

identification, and quantification of phytoplanktonic fungal parasites. Applied and environmental

microbiology 75 (8), 2545 –2553.

Reed, Howard, 1999. Stereological estimation of covariance using linear dipole probes. Journal of

Microscopy 195 (2), 96 –103.


31/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Rodenacker, K., Brühl, A., Hausner, M., Kühn, M., Liebscher, V., Wagner, M., Wuertz, S., 2000.

Quantification of biofilms in multi-spectral digital volumes from confocal laser scanning microscopes.

Image Anal. Stereol. 19, 151 –156.

Rogers, S.S., van der Walle, C, Waigh, T.A., 2008. Microrheology of bacterial biofilms in vitro:

Staphylococcus aureus and Pseudomonas aeruginosa. Langmuir : the ACS journal of surfaces and

colloids 24 (23), 13549 –13555.

Romero, D., Aguilar, C., Losick, R., Kolter, R., 2010. Amyloid fibers provide structural integrity to Bacillus

subtilis biofilms. Proceedings of the National Academy of Sciences of the United States of America 107

(5), 2230 –2234.

Rumin, J., Bonnefond, H., Saint-Jean, B., Rouxel, C., Sciandra, A., Bernard, O., Cadoret, J.-P., Bougaran, G.,

2015. The use of fluorescent Nile red and BODIPY for lipid measurement in microalgae. Biotechnology

for biofuels 8, 42.

Raaijmakers, J.M., Bruijn, I. de, Nybroe, O., Ongena, M., 2010. Natural functions of lipopeptides from

Bacillus and Pseudomonas: more than surfactants and antibiotics. FEMS microbiology reviews 34 (6),1037 –1062.

Sadovskaya, I., Vinogradov, E., Flahaut, S., Kogan, G., Jabbouri, S., 2005. Extracellular carbohydrate-

containing polymers of a model biofilm-producing strain, Staphylococcus epidermidis RP62A. Infection

and immunity 73 (5), 3007 –3017.

Sahu, P.K., Iyer, P.S., Oak, A.M., Pardesi, K.R., Chopade, B.A., 2012. Characterization of eDNA from the

clinical strain Acinetobacter baumannii AIIMS 7 and its role in biofilm formation.

TheScientificWorldJournal 2012, 973436.

Schaeffer, C.R., Woods, K.M., Longo, G.M., Kiedrowski, M.R., Paharik, A.E., Büttner, H., Christner, M.,

Boissy, R.J., Horswill, A.R., Rohde, H., Fey, P.D., 2015. Accumulation-associated protein enhances

Staphylococcus epidermidis biofilm formation under dynamic conditions and is required for infection ina rat catheter model. Infection and immunity 83 (1), 214 –226.

Schillinger, C., Petrich, A., Lux, R., Riep, B., Kikhney, J., Friedmann, A., Wolinsky, L.E., Göbel, U.B., Daims, H.,

Moter, A., 2012. Co-localized or randomly distributed? Pair cross correlation of in vivo grown

subgingival biofilm bacteria quantified by digital image analysis. PloS one 7 (5), e37583.

Schlafer, S., Birkedal, H., Olsen, J., Skovgaard, J., Sutherland, D.S., Wejse, P.L., Nyvad, B., Meyer, R.L., 2016.

Calcium-Phosphate-Osteopontin Particles for Caries Control. Biofouling (In press).

Schlafer, S., Dige, I., 2015. Ratiometric imaging of extracellular pH in dental biofilms. Journal of visualized

experiments : JoVE, Accepted.

Schlafer, S., Garcia, J.E., Greve, M., Raarup, M.K., Nyvad, B., Dige, I., 2015. Ratiometric imaging of

extracellular pH in bacterial biofilms with C-SNARF-4. Applied and environmental microbiology 81 (4),

1267 –1273.

Schlafer, S., Raarup, M.K., Meyer, R.L., Sutherland, D.S., Dige, I., Nyengaard, J.R., Nyvad, B., 2011. pH

landscapes in a novel five-species model of early dental biofilm. PloS one 6 (9), e25299.

Schneider, C.A., Rasband, W.S., Eliceiri, K.W., 2012. NIH Image to ImageJ: 25 years of image analysis. Nat

Meth 9 (7), 671 –675.

Schooling, S.R., Beveridge, T.J., 2006. Membrane vesicles: an overlooked component of the matrices of

biofilms. Journal of bacteriology 188 (16), 5945 –5957.

Schooling, S.R., Hubley, A., Beveridge, T.J., 2009. Interactions of DNA with biofilm-derived membrane

vesicles. Journal of bacteriology 191 (13), 4097 –4102.


32/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Schwartz, K., Syed, A.K., Stephenson, R.E., Rickard, A.H., Boles, B.R., 2012. Functional amyloids composed of

phenol soluble modulins stabilize Staphylococcus aureus biofilms. PLoS pathogens 8 (6), e1002744.

Sekar, R., Griebe, T., Flemming, H.-C., 2010. Influence of Image Acquisition Parameters on Quantitative

Measurements of Biofilms using Confocal Laser Scanning Microscopy. Biofouling 18 (1), 47 –56.

Shabala, L., McMeekin, T., Budde, B.B., Siegumfeldt, H., 2006. Listeria innocua and Lactobacillus delbrueckii

subsp. bulgaricus employ different strategies to cope with acid stress. International journal of food


Shrestha, D., Jenei, A., Nagy, P., Vereb, G., Szöllősi, J., 2015. Understanding FRET as a research tool for

cellular studies. International journal of molecular sciences 16 (4), 6718 –6756.

Stewart, P.S., Davison, W.M., Steenbergen, J.N., 2009. Daptomycin rapidly penetrates a Staphylococcus

epidermidis biofilm. Antimicrobial agents and chemotherapy 53 (8), 3505 –3507.

Stocks, S.M., 2004. Mechanism and use of the commercially available viability stain, BacLight. Cytometry.

Part A : the journal of the International Society for Analytical Cytology 61 (2), 189 –195.

Stone, G., Wood, P., Dixon, L., Keyhan, M., Matin, A., 2002. Tetracycline Rapidly Reaches All the ConstituentCells of Uropathogenic Escherichia coli Biofilms. Antimicrobial agents and chemotherapy 46 (8), 2458 –

2461.

Stoodley, P., Beer, D. de, Lewandowski, Z., 1994. Liquid flow in biofilm systems. Applied and environmental

microbiology 60 (8), 2711 –2716.

Sweity, A., Ying, W., Ali-Shtayeh, M.S., Yang, F., Bick, A., Oron, G., Herzberg, M., 2011. Relation between EPS

adherence, viscoelastic properties, and MBR operation: Biofouling study with QCM-D. Water research

45 (19), 6430 –6440.

Takenaka, S., Pitts, B., Trivedi, H.M., Stewart, P.S., 2009. Diffusion of macromolecules in model oral

biofilms. Applied and environmental microbiology 75 (6), 1750 –1753.

Tang, L., Schramm, A., Neu, T.R., Revsbech, N.P., Meyer, R.L., 2013. Extracellular DNA in adhesion andbiofilm formation of four environmental isolates: a quantitative study. FEMS microbiology ecology 86

(3), 394 –403.

Van Ommen Kloeke, F., Geesey, G.G., 1999. Localization and Identification of Populations of Phosphatase-

Active Bacterial Cells Associated with Activated Sludge Flocs. Microbial Ecology 38 (3), 201 –214.

Vroom, J.M., Grauw, K.J. de, Gerritsen, H.C., Bradshaw, D.J., Marsh, P.D., Watson, G.K., Birmingham, J.J.,

Allison, C., 1999. Depth Penetration and Detection of pH Gradients in Biofilms by Two-Photon Excitation

Microscopy. Applied and environmental microbiology 65, 3502 –3511.

Waharte, F., Steenkeste, K., Briandet, R., Fontaine-Aupart, M.-P., 2010. Diffusion measurements inside

biofilms by image-based fluorescence recovery after photobleaching (FRAP) analysis with a commercial

confocal laser scanning microscope. Applied and environmental microbiology 76 (17), 5860 –5869.

Walther, E., Richter, M., Xu, Z., Kramer, C., Grafenstein, S. von, Kirchmair, J., Grienke, U., Rollinger, J.M.,

Liedl, K.R., Slevogt, H., Sauerbrei, A., Saluz, H.P., Pfister, W., Schmidtke, M., 2015. Antipneumococcal

activity of neuraminidase inhibiting artocarpin. International journal of medical microbiology : IJMM

305 (3), 289 –297.

Webster, P., Wu, S., Gomez, G., Apicella, M., Plaut, A.G., St Geme, Joseph W, 2006. Distribution of bacterial

proteins in biofilms formed by non-typeable Haemophilus influenzae. The journal of histochemistry and

cytochemistry : official journal of the Histochemistry Society 54 (7), 829 –842.

Whitchurch, C.B., Tolker-Nielsen, T., Ragas, P.C., Mattick, J.S., 2002. Extracellular DNA required for bacterial

biofilm formation. Science (New York, N.Y.) 295 (5559), 1487.


33/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Xiao, J., Klein, M.I., Falsetta, M.L., Lu, B., Delahunty, C.M., Yates, J.R., Heydorn, A., Koo, H., 2012. The

exopolysaccharide matrix modulates the interaction between 3D architecture and virulence of a mixed-

species oral biofilm. PLoS pathogens 8 (4), e1002623.

Xiao, J., Koo, H., 2010. Structural organization and dynamics of exopolysaccharide matrix and microcolonies

formation by Streptococcus mutans in biofilms. Journal of applied microbiology 108 (6), 2103 –2113.

Zeng, G., Vad, B.S., Dueholm, M.S., Christiansen, G., Nilsson, M., Tolker-Nielsen, T., Nielsen, P.H., Meyer,

R.L., Otzen, D.E., 2015. Functional bacterial amyloid increases Pseudomonas biofilm hydrophobicity and

stiffness. Frontiers in microbiology 6, 1099.

Zuriani, R., Vigneswari, S., Azizan, M. N. M., Majid, M. I. A., Amirul, A.A., 2013. A high throughput Nile red

fluorescence method for rapid quantification of intracellular bacterial polyhydroxyalkanoates.

Biotechnol Bioproc E 18 (3), 472 –478.


34/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Figure 1


35/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Figure 2


36/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Table 1. Overview of methods for visualizing the components of the biofilm matrix by confocal microscopy

Target molecule Staining principle References

Polysaccharide

adhesin

Calcofluor (for β-1,3 and β-1,4

glucans)Fluorescently labeled lectins (for

various di- or tri-saccharides)

GFP fusion protein with

carbohydrate-binding modules

from polysaccharide-

degrading enzymes

Rasconi et al. 2009

Reviewed by Neu et al. 2001

Nguyen et al. 2014

Extracellular DNA Cell impermeant DNA-binding

fluorescent stains: TOTO-1,

TO-PRO 3, PicoGreen,

DDAO, propidium iodide andSYTOX stains

Specificity of stains evaluated by

Okshevsky and Meyer 2014

Extracellular proteins Fluorescent stains that bind to all

proteins: FilmTracerTM

SyPro

Fluorescently labeled antibodies

with specificity for individual

proteins

Dye with specificity for amyloid

proteins

Lawrence et al. 2003, Frank et al.

2007

Berk et al. 2012

Kim et al. 2016

Extracellular amyloid Thioflavin T stainingFluorescently labeled antibodies

with specificity for amyloid

proteins

Ilanchelian and Ramaraj 2004Larsen et al. 2007, Kim et al.

2016


37/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Table 2. Overview of methods for quantitative analyses of the biofilm matrix by confocal microscopy

Employed technique References

Geometric measurements Area/volume quantification

of fluorescently stainedmatrix components using

digital image analysis

Lawrence et al. 1998, Xiao et al.

2010

Colocalization analyses of

bacteria and matrix

components using digital

image analysis

Hu et al. 2012

Measurements of solute

concentrations

Immobilization of pH/O2-

sensitive fluorescent dyes

in particles

Hidalgo et al. 2009, Acosta et al.

2012

Fluorescence lifetimeimaging (FLIM) of pH-

sensitive fluorescent dyes

Vroom et al. 1999

pH ratiometry Xiao et al. 2012, Schlafer et al.

2015

Measurements of diffusion

properties

Time-lapse imaging Takenaka et al. 2009

Single particle tracking (SPT) Stoodley et al. 1994, Birjiniuk et

al. 2014

Fluorescence recovery after

photobleaching (FRAP)

Waharte et al. 2010, Daddi

Oubekka et al. 2012

Fluorescence correlationspectroscopy (FCS) Briandet et al. 2008; DaddiOubekka et al. 2012

Rheology measurements Magnetic microparticle

actuation

Galy et al. 2012


38/38

A C C E P

T E D

M A N U S C R I P T

ACCEPTED MANUSCRIPT

Highlights

● Confocal laser scanning microscopy is a valuable tool to study the biofilm matrix

● Various extracellular compounds can be labelled fluorescently and visualized in 3D

● Concentrations of diffusing molecules in the matrix can be monitored in real-time

● Diffusion of solutes through the matrix can be quantified

biofilm confocal de la matrix

Documents