1. Introduction
Biofilms are aggregates of microbial cells embedded in a matrix of
extracellular polymeric substances (EPS) (Hans-Curt Flemming &
Wingender, 2010; Hall-Stoodley et al., 2004). They are ubiquitous in
clinical, environmental, and industrial systems and can cause human
infections, foul water filtration membranes, and promote corrosion in
pipes (Costerton et al., 1995; Hall-Stoodley et al., 2004). Biofilms
also can play beneficial roles, for example in environmental treatment
processes. Thus, biofilm removal may be sought in some cases, and its
retention in others.
Biofilm formation, persistence, deformation, and detachment are largely
determined by the biofilm mechanical properties (Kundukad et al., 2016;
Powell et al., 2013; Boudarel et al., 2018; Gloag et al., 2019). For
example, biofilms’ viscoelastic nature help them dissipate stress from
fluid flow, preventing detachment (Stoodley et al., 1999). The
characterization of biofilm mechanical properties is key to predicting
biofilm deformation and detachment (Klapper et al., 2002).
Biofilm mechanical properties can be influenced by a variety of factors,
including nutrient concentrations and microbial growth rates (Paul et
al., 2012; Van Loosdrecht et al., 2002), microbial composition (Abriat
et al., 2020; Kim et al., 2020; Yannarell et al., 2019), biofilm age
(Hwang et al., 2014; Shen et al., 2016), hydrodynamic conditions
(Dunsmore et al., 2002; Thomen et al., 2017), multivalent cation
concentration (Ahimou et al., 2007; Jones et al., 2011; Lieleg et al.,
2011), temperature (Pavlovsky et al., 2015; 2013), and pH values (Chen
& Stewart, 2000; Ho et al., 2013). For example, the microbial growth
rates and ecological stratification, which are determined by the
substrate profiles within the biofilm, can have a strong influence on
biofilm mechanical strength (Rochex et al., 2009). Higher bulk oxygen
concentrations and higher shear stresses were found to increase the
strength of biofilms (Stoodley et al., 2002; Ahimou et al., 2007a;
Pellicer-Nàcher & Smets, 2014). Also, the bulk calcium ion
(Ca2+) concentration caused biofilms to become thicker
and denser, and to significantly decrease biofilm detachment episodes
(Goode & Allen, 2011).
While biofilms are commonly thought to be mechanically homogeneous, this
may be an artifact generated by the use of bulk-scale techniques for
their mechanical characterization (Safari et al., 2015). When microscale
techniques were used, mechanical properties have been found to vary
significantly within the biofilm (Böl et al., 2012).
Biofilms usually have temporal and spatial variations of mechanical
properties. It has been widely recognized that the structure of biofilms
becomes more stable over time. For example, older biofilms were found to
be less affected by bubble disruptions, whereas younger biofilms were
easily removed by air bubbles (Jang et al., 2017). Laspidou and Rittmann
(2004b) hypothesized that biofilm increases its density over time due to
consolidation, i.e., the filling of voids within the biofilm. In a study
of the viscoelasticity of Pseudomonas aeruginosa biofilms (Gloag
et al., 2018), various temporal changes were observed in different
phenotypes. Increased stiffness or cohesive strength was observed over
biofilm depth in several studies (Ahimou et al., 2007; Derlon et al.,
2008; Olivier Galy et al., 2012). Spatial distribution of biofilm
stiffness was also found in P. aeruginosa biofilms (Hunt et al.,
2004; Karampatzakis et al., 2017).
In order to understand the spatial distribution of biofilm mechanical
properties, microscale techniques can be used. Microscale techniques
include microindentation compression (Cense et al., 2006), dedicated
microcantilever (Aggarwal et al., 2010; Poppele & Hozalski, 2003),
atomic force microscopy (AFM) indentation (Arce et al., 2009; Volle et
al., 2008), and microbead force spectroscopy (Lau et al., 2009).
However, all these methods are invasive and can compromise the biofilm
integrity.
In recent years, novel microrheological techniques have been developed
(Birjiniuk et al., 2014; Cao et al., 2016; Galy et al., 2012;
Karampatzakis et al., 2017; Thomen et al., 2017). In particular,
magnetic actuation with magnetic tweezers, coupled with magnetic
microparticles, may be ideal (Galy et al., 2014; Galy et al., 2012;
Zrelli et al., 2013). This technique can overcome the limitations of
other microscale techniques by using strong forces and in-situ
measurements. With the displacement of magnetic particles, which are
added to the media during initial growth, biofilm properties can be
mapped spatially. For example, Galy et al. (2012) used magnetic tweezers
and found that stiffness measurements at different locations in a
biofilm ranged over of two orders of magnitude, even locally. This
indicates the importance of localized spatial measurements in the study
of biofilm mechanical properties. Nevertheless, the mechanical
heterogeneity of biofilms has received little attention.
Mathematical models describing biofilm mechanical behavior can improve
our understanding of biofilm structures and properties (Böl et al.,
2012). Such models include the simulation of biofilm deformation under
applied stress (Li et al., 2020; Picioreanu et al., 2018; Picioreanu et
al., 2001; Towler et al., 2007). However, the mechanical properties are
typically assumed to be homogeneous and are based on large-scale
measurements. It is not clear whether mechanical heterogeneity is
significant, and whether it should be considered for accurate
predictions of biofilm deformation.
As discussed above, very few studies have addressed the spatial
distribution and variability of mechanical properties of biofilms in a
non-destructive fashion (Cao et al., 2016; Galy et al., 2012;
Karampatzakis et al., 2017). Microscale heterogeneities could have
important impacts on biofilm formation and behavior (Böl et al., 2012),
yet the impact of this variability has not been explored.
In this study, we used magnetic tweezers to determine the spatial
distribution of biofilm mechanical properties for different flow
conditions, dissolved oxygen and Ca2+ concentrations
in P. aeruginosa biofilms. P. aeruginosa is
commonly used as a model species for biofilm research, as it is an
important human pathogen and also widespread in the environment. We then
used a fluid-structure interaction mathematical model to evaluate the
mechanical behavior of a biofilm (i.e., biofilm deformation) considering
average mechanical properties versus the heterogeneity observed in the
experiments.
2.Methods
Biofilm growth
conditions
P. aeruginosa ATCC strain 15692 (PAO1), tagged with green
fluorescent protein (GFP), was used for biofilm growth (Shrout et al.,
2006). The bacterial strain was grown overnight to an optical density of
0.4-0.5 (OD600) in Luria-Bertani (LB) broth at 37°C on
an orbital shaker. A 1 mL aliquot of the culture was mixed with red
fluorescent magnetic beads (Dynabeads M-270 Amine, Invitrogen, Carlsbad,
CA), with a 2.8 μm diameter, at a final concentration of
2.5\(\times\)106 particles/mL. These beads are
superparamagnetic, hydrophilic, and contain surface amino groups that
form covalent bonds with cell and EPS components. The hydrophilic
surface ensures non-specific binding to the biofilm. The
superparamagnetic behavior allows for high levels of magnetization and
the absence of magnetization without an external magnetic field. As a
control, fluorescent non-magnetic particles of 1 μm diameter were used
(Fluoresbrite 18660, Polysciences, Germany).
Biofilms were grown in borosilicate glass capillaries with a 1 mm
internal side and 150 μm wall thickness (Friedrich and Dimmock Inc.,
Millville, NJ). The capillaries were inoculated with the mixed
suspension and kept under static conditions for 2 h before starting a
continuous flow of 10% LB for the growth period. Flow was provided
using a syringe pump (Kd Scientific, KDS-220, Holliston, MA). Flow rates
were kept between 0.1 and 5 mL/h to maintain Reynolds numbers (Re)
between 0.28 and 13.9 (laminar flow). Biofilms were grown with dissolved
oxygen (DO) concentrations of either 8 or 1 mg/L, and with or without a
Ca2+ addition of 100 mg/L as CaCl2. Anin-situ magnetic actuation technique was then applied to
non-destructively estimate the mechanical behavior of the biofilm after
5 days of growth. The general procedures were followed according to Galy
et al. (2012). The detailed information of the magnetic tweezers and
force calibration procedure is provided in Supporting Information
Material (SI).
Microscopy and particle tracking
A confocal laser scanning microscopy (CLSM) (Nikon 90i A1R, Melville,
NY) equipped with a 40x Plan Fluor WD 0.2 objective was used to image
the biofilm and particles large-spectrum fluorescence signals.
Typically, the first examined plane was located between 5 to 10 μm above
the capillary bottom. A z-stack with a depth step size of 0.5 μm was
collected to verify biofilm thickness and abundance of magnetic beads.
For a given plane, particle motion upon magnetic force application was
recorded at a scanning speed of 30 frames per second (fps) over a period
of 20 s, and further processed using a particle tracking software
(ImageJ, NIH, Bethesda, MD). About 3 to 4 fields of 250 μm \(\times\)250 μm (with each field) were collected along the capillary width before
changing the plane (z-direction) (Figure 1). After each field was
obtained, the field view was changed in the x-direction. Vertical planes
were analyzed every 10 μm across the height of the biofilm. About 10-15
particles were analyzed per field. A more visual description of the
procedure can also be found in Galy et al. (2014).
Creep curves and mechanical behavior
analysis