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A Selection of Platforms to Evaluate Surface Adhesion and Biofilm Formation in Controlled Hydrodynamic Conditions

Luciana C. Gomes
Filipe J. M. Mergulhão
LEPABE—Laboratory for Process Engineering, Environment, Biotechnology and Energy, Faculty of Engineering, University of Porto, Rua Dr. Roberto Frias, 4200-465 Porto, Portugal
Author to whom correspondence should be addressed.
Microorganisms 2021, 9(9), 1993;
Submission received: 7 August 2021 / Revised: 14 September 2021 / Accepted: 17 September 2021 / Published: 21 September 2021
(This article belongs to the Special Issue Microbial Films-the Interplay of Physics and Biology)


The early colonization of surfaces and subsequent biofilm development have severe impacts in environmental, industrial, and biomedical settings since they entail high costs and health risks. To develop more effective biofilm control strategies, there is a need to obtain laboratory biofilms that resemble those found in natural or man-made settings. Since microbial adhesion and biofilm formation are strongly affected by hydrodynamics, the knowledge of flow characteristics in different marine, food processing, and medical device locations is essential. Once the hydrodynamic conditions are known, platforms for cell adhesion and biofilm formation should be selected and operated, in order to obtain reproducible biofilms that mimic those found in target scenarios. This review focuses on the most widely used platforms that enable the study of initial microbial adhesion and biofilm formation under controlled hydrodynamic conditions—modified Robbins devices, flow chambers, rotating biofilm devices, microplates, and microfluidic devices—and where numerical simulations have been used to define relevant flow characteristics, namely the shear stress and shear rate.

1. Introduction

Biofilms are surface-attached communities of microorganisms, establishing three-dimensional structures composed of bacteria surrounded by a self-made matrix [1]. This matrix consists of polysaccharides, proteins, and extracellular DNA and influences biofilm structure and morphology [2]. It is estimated that more than 90% of the bacterial cells in natural environments reside in a biofilm [3], since it gives protection against hostile conditions (pH changes, lack of nutrients, hydrodynamics, and antimicrobial compounds), encourages gene transfer, and facilitates the colonization of niches [4].
The established model for biofilm development includes five steps, starting with the (i) reversible attachment of cells to a preconditioned surface, (ii) production of extracellular polymeric substances (EPS) causing irreversible cell attachment, (iii) early development of biofilm architecture, (iv) biofilm maturation, and (v) cell dispersion from the biofilm into the surrounding environment [5,6]. An immersed substratum is rapidly covered by molecules from the liquid, forming a conditioning film that may change the properties of that surface, making it more or less suitable for bacterial adhesion [7,8]. Then, cell adsorption at the surface occurs, followed by release or reversible adhesion. The physical forces associated with conditioning film formation and reversible adhesion are electrostatic and van der Waals forces, as well as hydrophobic interactions [9]. The next step starts when the cells become irreversibly attached to the surface due to the presence of stronger attractive forces, such as covalent and hydrogen bonds, and cellular surface structures, such as fimbriae and flagella [9]. After maturation, biofilm growth and detachment/sloughing balance each other so that the biomass amount is approximately constant in time, i.e., the steady-state is attained.
Biofilm development is a problem faced by the environmental, industrial, and biomedical areas. Regardless of the setting where it takes place, it is well known that biofilm establishment and growth are affected by different factors, such as surface properties, nutrient availability, hydrodynamics, temperature, pH, and microbial cell properties [10]. Among these factors, hydrodynamics will be considered in detail in this review.
In the environment, biofilms particularly affect the efficiency of shipping, aquaculture, and coastal industries [11]. The fouling phenomenon increases the surface roughness of the ship hulls, hence increasing the friction between the fouled hull and the water [12]. This resistance increases fuel consumption and, consequently, the emissions of greenhouse gases to the atmosphere, while reducing the maneuverability of the vessel [11,12]. Additional problems related to biofouling in the environment are associated with immersed offshore structures (cages, netting, and pontoons), onshore equipment, and structures such as pumps, pipelines, and filters, due to the high drag and accelerated biocorrosion to which they are exposed [13].
Besides affecting cleaning and disinfection, biofilms formed in industrial facilities can reduce energy transfer in heat exchangers, obstruct fluid flow, and cause localized corrosion attacks [14]. It has been reported that biofilm development in industries corresponds to approximately 30% of the plant operating costs [15]. In the case of the food industry, biofilms have a detrimental effect not only on the process but also on the final product or consumer. The Center for Disease Control and Prevention (CDC) has estimated that between 1996 and 2010, 48 million illnesses, 128,000 hospitalizations, and 3,000 deaths occurred annually in the US due to the dissemination of foodborne pathogens [16].
In the biomedical field, the sessile cells are responsible for infections, as they are usually more resistant to antimicrobial therapy than their planktonic counterparts and less susceptible to host defenses [17]. The National Institutes of Health (NIH) revealed that 65% of all microbial infections are caused by biofilms [18], which can grow in indwelling medical devices and have an estimated direct cost burden of 7 billion EUR in Europe alone [19]. Biofilms formed in medical devices may contain pathogenic organisms and cause changes in surface properties and material degradation, affecting the functionality of the medical setting [20].
The ubiquitous nature of biofilms and their increasing resistance impose great challenges for the use of conventional antimicrobials and suggest the need for combined or multi-targeted approaches. In this sense, the study of strategies capable of preventing microbial adhesion through the modification of surface properties (for instance, making them antimicrobial and/or antiadhesive) may be one of the simplest and most reasonable ways to inhibit surface colonization and delay biofilm growth [21,22,23,24]. However, few studies have evaluated the effectiveness of these promising surfaces in conditions that mimic real scenarios, particularly regarding hydrodynamics. In this review, the most commonly used platforms for the in vitro assessment of microbial adhesion and biofilm formation under flow conditions—modified Robbins devices, flow chambers, rotating biofilm devices, microplates, and microfluidic devices—are introduced, and their main advantages and disadvantages are discussed. These testing platforms have been used transversally in the environmental, industrial, and medical fields, mainly with the aim of evaluating the effects of different substratum features, microbial strains, and shear forces on adhesion and biofilm formation, due to their ability to control the hydrodynamics (flow rate and/or shear stress or shear rate) and recreate in vivo flow conditions. This becomes a critical step in translating research into practical applications.

2. Effects of Hydrodynamics on Microbial Adhesion and Biofilm Formation

The flow conditions of each system where there is a surface material (natural, industrial, or biomedical) have a very strong influence on the biofilm onset. During initial adhesion, hydrodynamics dictates the rate at which macromolecules (specific for each type of fluid) and microorganisms are delivered to the surface, the time they reside close to the surface, and the shear forces at the surface-fluid interface [25]. According to Katsikogianni and Missirlis [26], there is an optimum flow rate for bacterial adhesion, reflecting the balance between the rate of cell delivery and the force acting on adhered bacteria. Furthermore, the bacteria–substratum interaction determines the shear forces that adhered bacteria will be able to withstand [26].
Besides the relevant role of hydrodynamics on the microbial adhesion step, it is also one of the most important factors in biofilm formation and structure. The fluid surrounding a biofilm is the source for nutrients and vehicle for cell by-product removal [27]. An increase in flow velocity promotes the flux of molecules (nutrients, cells, biocides, antibiotics, metabolites, etc.) by changing their concentrations in the biofilm–fluid interface. Hydrodynamics also regulates the physiological properties of the biofilm by changing the mechanical shear stresses at the interface [25]. Higher shear forces often lead to the formation of thinner, denser, and stronger biofilms [28]. Although higher flow velocities enhance molecular transport by convection, the higher density of biofilms reduces the diffusivity of the molecules inside them [29,30]. Additionally, stronger shear forces can be responsible for higher biofilm sloughing or detachment [28].
Given the importance of shear forces on initial adhesion and biofilm development, it is essential to characterize them. The vast majority of biofilm studies under flow conditions only report the tested flow rate. Nevertheless, the flow rate by itself provides little information about shear forces since it does not take into consideration the geometry of the flow system. Two main parameters should be considered to characterize shear effects: the shear rate and the shear stress. Mathematically, the shear rate is the derivative of the velocity in the perpendicular direction from the wall system [31] and quantifies the frequency at which cells contact the surface. The shear stress in Newtonian fluids is proportional to the shear rate, where fluid viscosity is the constant of proportionality [31], translating the friction from the fluid acting on the adhered cells or the biofilm. Therefore, shear stress is commonly used as a descriptor of the shear forces acting on the biofilm during maturation or detachment.
Computational fluid dynamics (CFD) are commonly used to model biofilm reactors because they enable the estimation of the fluid flow parameters of these systems, such as the shear stress and the shear rate, at relatively low cost and faster, in comparison to experimental techniques [32,33]. CFD requires that the geometry to be analyzed is divided into a finite set of volumes, called cells, forming a computational grid, called mesh. Fluid flows are described by differential equations for the conservation of mass, momentum, and energy; CFD replaces these equations with algebraic equations, which can be numerically solved for each cell, resulting in a flow field [34]. These equations describe how the single operating parameters are related. Although CFD is very useful for understanding biofilm behavior, one must bear in mind that most simulations are performed for clean surfaces. When biofilms are formed, the cross-sectional flow area is reduced, increasing the bulk flow velocity and wall shear stress. Thus, these simulations are particularly recommended for the study of initial adhesion, early stages of biofilm development (such as those usually investigated in biomedical settings), and surfaces that are frequently cleaned (as is the case with food processing equipment). In these situations, the thickness of the formed biofilms is unlikely to have a significant impact on flow dynamics and shear forces distribution [35].

3. Biofilm Platforms

In this context, biofilm reactors are platforms for the study of biofilms in laboratory conditions. One of the major obstacles to study in vitro biofilms is the choice of a suitable platform, where key variables such as flow rate and shear stress can be manipulated in order to mimic the conditions found in real scenarios. Although completely reproducible biofilms are nearly impossible to obtain, the development of in vitro platforms for biofilm studies is a foremost step towards the standardization of procedures and for better control of the environmental conditions that affect biofilm development [36,37].
Here, we describe the most commonly used platforms for microbial adhesion and biofilm formation in controlled hydrodynamic conditions, particularly those where CFD has been used to determine relevant flow characteristics. These platforms have advantages and limitations, which are summarized in Table 1.

3.1. Flow Cells: Robbins Device and Modifications, and Flow Chambers

Flow cells can be generally divided into two types: those that are based on the design of the Robbins device and those that are built for the direct inspection of biofilm development, here called flow chambers. In both types of flow cells, it is possible to test different surface materials simultaneously in similar nutritional and hydrodynamic conditions. Nevertheless, it is worth mentioning that modified Robbin devices have higher throughput and hydrodynamic range than flow chambers. The Robbins device and its modifications present a higher number of sampling ports available for analysis, allowing for multiple biofilm samples to be taken simultaneously, as well as for sampling more than a single time point during biofilm development [39]. Although both types of flow cells are useful tools for studying biofilm under controlled conditions, they need a specialized apparatus, are technically challenging, and are not suitable for rapid high throughput assays. Another weakness of these systems is that only a single microbial strain can be analysed per experiment.
The most straightforward configuration of a flow cell system is that of a bioreactor containing a batch culture of the desired microorganism so that the content of the reactor is pumped through the flow cell and the effluent drained to waste. This configuration may be interesting for adhesion studies, particularly if the flow rates to be tested are low, since the duration of the assay is limited by the cell suspension volume. Another configuration is to place the flow cell in a recycle loop so that the culture volume is no longer a limitation and assays can last longer and perform at higher flow rates [40]. However, it has the disadvantage that the composition of the batch culture is always changing. A third alternative is to have a chemostat feeding a recirculation loop so that the feed flow to the chemostat equals the drain flow from the loop. In this case, it is possible to feed the flow cell with a constant concentration of cells and nutrients, while decoupling the flow rate going through the flow cell from the dilution rate [41,42]. With this flow cell configuration, it is possible to work at very high flow rates and attain high shear stresses that are comparable to those found in the environment and industry [43].

3.1.1. Robbins Device and Modifications

The Robbins device was initially developed by Jim Robbins and Bill McCoy to monitor biofilm formation in industrial water systems [44]. Several modifications were later introduced to this design, including the use of a square-channel pipe where coupons are aligned with the inner surface without disturbing flow characteristics [45]. They are convenient for studies where a large biofilm mass amount is wanted. With the modified Robbins devices, the flow can be momentarily stopped to allow direct access to the coupon, so that time-course experiments are possible. This stop of the flow system for coupon removal involves some risk because, even if the operator is very careful that the shutdown and restart of the system are smooth, there may be some loosening of the biofilm already formed in the remaining coupons of the flow cell. For quantitative analysis of the biofilm to be carried out, destructive sampling techniques are usually required. Conventional techniques, such as total and viable cell counts, as well as protein and carbohydrate content analysis, comprise the disruption of the biofilm [42,46].
Other flow cell designs include a half-pipe geometry that more closely resembles the geometry of piping systems [43,47]. These flow cells can be operated either in laminar or turbulent regimes, but it is important to guarantee that the flow cell has an entry section that is long enough to allow for flow development before the sampling zone (thus avoiding entry effects) and that the effect of the sudden contraction on the exit zone is negligible. This will ensure that all coupons are subjected to the same hydrodynamic conditions and that biofilm samples can be directly compared [48].
In our group, a custom-made, semi-circular flow cell (identical to that shown in Figure 1) was designed to evaluate the performance of different surface coatings in preventing biofouling in the marine environment [22], food industry [24,41], and medical devices [49,50]. The hydrodynamics of this flow cell system was fully characterized by CFD [48], which allows not only the guarantee that all sampling coupons are exposed to the same shear forces but also provides knowledge of the flow rate and Reynolds number, which is necessary in order to operate this platform and simulate the shear stress and/or shear strain described for different real scenarios.

3.1.2. Flow Chambers

In spite of the many advantages of modified Robbins devices, they are neither adequate for monitoring the initial cell adhesion to a surface nor for the direct analysis of biofilm development. For these purposes, several models of flow chambers that can be mounted on a microscope stage and used with video capture systems have been developed, enabling real-time observation of microbial adhesion, particularly when used with transparent surfaces. The employment of fluorescent probes coupled with confocal laser scanning microscopy (CLSM) makes flow chambers especially appreciated for in situ gene expression studies [51].
The most well-known flow system to study cell adhesion is the parallel-plate flow chamber (PPFC) developed by Bos et al. [52]. Adhesion can be studied in the PPFC system under controlled hydrodynamics that mimics, for instance, physiologically relevant conditions [40,53] using a wide range of microorganisms and surfaces with different properties. This system requires low volumes and, consequently, has a reduced cost when compared to modified Robbins devices; additionally, it presents one or more glass viewing ports that permit non-destructive, real-time adhesion (single-cell visualization) and biofilm observation. Despite their versatility, one must bear in mind that PPFCs have a much lower throughput than microplates and larger flow cells based on the Robbins device. Additionally, when real-time monitoring of adhesion is performed, a decrease in the initial adhesion rates is often observed along the experimental time, which is related to a phenomenon called hydrodynamic blocking [54,55]. Hydrodynamic blocking can reduce the adhesion of cells since the area behind each adhered cell is screened from incoming cells. Adhesion rates obtained in such conditions are not truly representative of the interaction between a single cell and the surface. Thus, initial adhesion assays in these setups should be conducted so that low surface coverage is attained, and the absence of blocking should be confirmed so that consistent results can be obtained [54].
Flow chamber systems have been designed to analyse cell adhesion [23,56,57] and biofilm formation [58,59], including a PPFC coupled to a jacketed tank and connected to centrifugal pumps and a valve via a silicone tubing system (Figure 2). The valve allows the bacterial suspension to circulate through the system at a controlled flow rate [40], and the recirculating water bath is connected to the tank jacket to enable temperature control.

3.2. Rotating Biofilm Devices

Two types of rotating biofilm reactors are commonly used in the assessment of material and fluid flow effects on biofilm development: the rotating disk reactor and the rotating cylinder reactor. These reactors have different designs. The rotating disk reactor consists of a 1-L vessel with a magnetically driven rotor at the bottom, which holds removable coupons for biofilm formation (Figure 3) [60]. The hydrodynamic conditions under which the biofilm is formed are controlled by adjusting the disk rotation speed [60], and the shear stress on the coupons’ surface can be estimated from the Navier–Stokes equations. The rotating cylinder reactor is often composed of four cylindrical sections that can be rotated at variable speeds within four concentric chambers [61]. Unlike the rotating disk reactor, this platform can be used to test different cell suspensions, since each chamber of the cylinder reactor has independent feeding and sampling ports [61].

3.3. Microfluidic Devices

Microfluidic platforms have demonstrated high potential and versatility for the study of microbial adhesion and biofilm formation under different growth conditions. Compared with traditional flow cell systems, microfluidics enables greater control over flow conditions, can be used to explore a much wider range of shear rates with high flexibility in designing different flow geometries, and facilitate the parallelization of experiments [62,63]. Although microfluidic devices can be fabricated by different techniques and from a diversity of materials, the flexible elastomer polydimethylsiloxane (PDMS) has been the material of choice for the construction of these devices. Several other surfaces can be studied using xerographic construction techniques that enable different polymers to be incorporated into microfluidic flow cells [53]. Concerning the analysis methods, although off-chip detection with conventional methods is feasible, on-chip detection by optical and/or fluorescence microscopy is preferred, in order to visualize in situ and real-time effects (Figure 4) [36].
Although there is a tendency to develop biofilm models in miniaturized devices, microfluidic-based devices also have their limitations: the small liquid volumes used in microfluidics may further impede molecular analysis, and the spatial confinement may generate different biofilms from those formed in more open systems [64]. Additionally, this platform requires specialized technical abilities for device fabrication and experimental setup, and system clogging can occur due to the small dimensions [36]. Air bubbles are another recurring issue in microfluidics [65]. Because of the micrometric dimensions of the tubes and channels, air bubbles can be very difficult to remove, leading to fluid flow instability and most likely to the detachment of adhered cells or biofilm portions.

3.4. Microplates

Microplates are currently the most widely used platform for biofilm development studies. They consist of plates with multiple wells arranged in a rectangular array with a 2:3 aspect ratio, resulting in 6, 12, 24, 48, 96, and 384 wells. The volume of each well can range from tens of microliters to few milliliters, depending on the number of wells [66]. Although most researchers use microplates in static conditions, they can be placed in orbital incubators and used for dynamic biofilm studies under controlled fluid conditions [67,68]. These devices are easy to handle, which allows for studying the adhesion of different microbial strains and consequent biofilm formation in rapid and inexpensive assays, due to their reduced volume [69]. Depending on the format used, they enable high throughput at an affordable cost and sometimes non-invasive imaging through optical coherence tomography (OCT) [70,71] and confocal laser scanning microscopy (CLSM) [72]. Particularly for larger well dimensions, it is possible to place coupons at the bottom of the wells so that different surface materials can be tested [70,73,74]. The main limitations of microplates are that loosely attached cells may not be measured correctly due to detachment during washing and that biofilms formed in this platform are affected by sedimentation.

3.4.1. 96-Well Microplates

This is the most intensively used microplate format, mainly for screening purposes. Biofilm formation in this platform is severely affected by sedimentation, and the direct inspection of the biofilm is possible but technically difficult [75,76]. They are particularly suited for short-term experiments, as they operate in batch mode with the intrinsic exhaustion of nutrients and accumulation of toxic metabolites. Results obtained in this platform often lack reproducibility, possibly due to the washing steps that are researcher-dependent and the existence of several protocol versions for biofilm analysis [36]. These plates are generally not compatible with the use of coupons, as the bottom surface is relatively small; so, only a limited number of surfaces can be assayed (limited to the construction materials of these plates).

3.4.2. 12- and 6-Well Microplates

These microplates are very attractive formats. Although theoretically their throughput is lower than the 96-well plates, the results obtained with these platforms are more reproducible due to the higher liquid volume, decreasing the need for a large number of replicate wells. These two types of plates also sustain microbial growth for longer periods, but medium replacement can be necessary. Large coupons can be used for biofilm formation (square surfaces of up to 1.5 cm can be placed on the bottom of the 12-well plates), and uniform shear forces can be obtained. Even though the shear stress in the coupon varies with the radial distance to the center, each coupon has identical average shear stress values [71].
The hydrodynamics inside the wells of 12-well microplates have been simulated to assess the effect of orbital shaking frequency on shear stress. Numerical simulations were performed at 25 °C, with an orbital diameter of 25 mm, a liquid volume of 3 mL, and shaking frequencies of 40 and 180 rpm (Figure 5). As expected, higher shear stresses at the bottom of the wells can be attained at higher shaking frequencies; values up to 0.07 Pa and shear rates of 42 s−1 were achieved. These values are much higher than those obtained with 96-well microplates [8,77].

4. Adhesion and Biofilm Studies Performed under Controlled Hydrodynamics

In this section, illustrative examples of the application of the described in vitro platforms are given, when appropriate, for the investigation of initial microbial adhesion, biofilm formation and its treatment under controlled shear conditions in different fields: environment, industry, and medicine.

4.1. Environmental Applications

Table 2 presents typical shear values that can be found in the environmental field. In a natural environment, a shear rate range between 4 and 125,000 s−1 can be obtained.
Most of the research in this area has been devoted to the impact of shear and surface characteristics on biofilm formation, giving less relevance to microbial cell adhesion (Table 3). It was also observed that flow systems, namely modified Robbins devices and rotating biofilm devices, are the main choice to emulate the turbulent flows and high wall shear stresses found in water systems [79,80,81]. However, in the last few years, efforts have been made to predict flow conditions in easy-to-handle biofilm platforms like microplates [68,71]. A detailed hydrodynamic analysis of the 12-well microplates [71] allows us to define the operational conditions that should be used in the laboratory bench to further assess the biofilm formation capacity of marine bacteria [70,71] and the antibiofilm activity of novel surface coatings [22,82] under hydrodynamic conditions prevailing in natural aquatic environments.
Table 2. Environmental scenarios and their typical shear ranges.
Table 2. Environmental scenarios and their typical shear ranges.
Environmental ScenarioShear Stress (Pa)Shear Strain (s−1) References
Drinking-water distribution systems0.13–9.10n.d.[79,83]
Ship in harborn.d.50[84]
Ship navigation (turbulent flow)n.d.125,000[84]
Marine environmentsn.d.4 and 40[71]
Tumbling and pouringn.d.10–100[84]
Channels within a biofilmn.d.60–300[84]
n.d.—not disclosed.

4.2. Industrial Applications

Similar to what was observed in environmental systems (Table 3), in the industrial field, the modified Robbins devices and rotating devices were the most reported reactors for biofilm formation and treatment studies. Different groups have used these flow systems in shear stress intervals of great amplitude [94,95,96], covering a huge range of shear values that can be found in the industry (Table 4). Our research group, in particular, has operated a semi-circular flow cell system (Figure 1) in different conditions and was able to attain shear stress values up to 0.6 Pa during biofilm formation [42,48], confirming the versatility of this platform and its capacity to mimic the hydrodynamic conditions that can be found, for instance, in the food industry (Table 4).
When the aim was to study microbial adhesion in an industrial environment, biofilm researchers preferred to use flow chambers [97,98] or microplates [24,41], since they are faster to operate and may allow for direct inspection by microscopic techniques (Table 5).
Table 4. Examples of industrial processes and their associated shear stress ranges.
Table 4. Examples of industrial processes and their associated shear stress ranges.
Industrial Equipment or PhenomenonShear Stress (Pa)References
Pipeline elbows0.009[99]
Dead ends0.05–18.9[100]
Removal of deposits from stainless steel tubes0.09[101]
Corners of a washing tank<0.1[102]
Angles of a washing tank0.1–0.4[102]
Mix proof valve0–0.25[103]
Three-way valve0.4–1.7[104]
Half-open butterfly valve0–190[100]
Product fill valve0–1180[105]
Milk spray dryer0–0.4[106]
Cleaning-in-place pilot plant0–5[107]
Plate heat exchanger for yoghurt processing6.7 and 20–46[108,109]
Plate heat exchanger of an ice slurry system50–100[110]
Pilot-scale plate heat exchanger for milk treatment150–450[108]

4.3. Biomedical Applications

Several studies were found in the literature where biofilm assays were performed under characterized hydrodynamic conditions similar to those of medical settings. Depending on the biomedical scenario, the shear stress range can vary between 0.02 and 88.3 Pa, and the shear strain between 0.1 and 80,000 s−1 (Table 6). Flow chambers have particularly been used in the medical field to evaluate the antiadhesive activity of novel surface materials for biomedical devices, including urinary tract and implanted devices (Table 7), since they are adequate for low fluid shear stresses and laminar flow applications, as well as for real-time insight into the dynamic process of microbial cell adhesion [21,40,57]. Furthermore, the dimensions of the flow cell or the flow rate can be adjusted to attain the required shear stress/shear rate, in order to resemble in vivo flow conditions.
Microfluidic platforms have also demonstrated high potential and flexibility for the study of microbial adhesion [113,114] and biofilm formation [115,116] under different hydrodynamic conditions.
Table 6. Characteristic shear conditions found in biomedical scenarios.
Table 6. Characteristic shear conditions found in biomedical scenarios.
Human Body or Biomedical DeviceShear Stress (Pa)Shear Strain (s−1)References
Blood flow in veins0.076–3.420–800[117,118]
Blood flow in arteries0.2–150–650[117,118]
Fluid in the oral cavityn.d.0.1–50[84]
Kidney collecting duct cells0.02–2n.d.[119]
Cerebral circulationn.d.>100[121]
Urinary cathetern.d.15[84,122]
Hemodialysis catheter52.6–88.320,000–80,000[123]
Catheter sheath introducer0.03n.d.[124]
Endovascular stent0.22–6.72n.d.[125]
Prosthetic valve0.06–27.84n.d.[126]
Contact lens motionn.d.1000[127]
n.d.—not disclosed.
Table 7. Biomedical studies performed on different biofilm platforms to evaluate the initial adhesion, as well as the biofilm formation and treatment under the defined shear conditions.
Table 7. Biomedical studies performed on different biofilm platforms to evaluate the initial adhesion, as well as the biofilm formation and treatment under the defined shear conditions.
PlatformFieldBiofilm StageStudy AimHydrodynamicsAssay TimeSurface MaterialOrganismsConcluding RemarksReferences
Modified Robbins deviceGeneral medical devicesBiofilm formationEffect of flow rate variation on mass transfer and biofilm developmentFlow rates of 374 and 242 L h−1, corresponding to shear stresses between 0.183 and 0.511 Pa9 daysPolyvinyl chlorideEscherichia coliBiofilm formation was favored at the lowest flow rate because shear stress effects were more important than mass transfer limitations.
This flow cell system generates wall shear stresses that are similar to those found in some biomedical settings.
Urinary devicesBiofilm formationEvaluation of the potential of antiadhesive coatings when immobilized onto medical-grade polyurethaneFlow rate of 53 mL s−1, corresponding to 15 s−148 hPolyurethane
Polyurethane coated with CyanoCoating through a polydopamine layer application, or O2- plasma, N2-plasma, and O3 activation
Escherichia coliWhen the coating was produced via O3 activation, CyanoCoating was able to decrease the biofilm biovolume by 88% and the surface coverage by 95%, compared to the uncoated surface. [50]
Investigation of the role of uncommon bacteria on the Escherichia coli microbial consortiumFlow rate of 300 mL min−1, corresponding to 15 s−172 hSilicone rubberEscherichia coli
Delftia tsuruhatensis
E. coli and D. tsuruhatensis were able to form single- and dual-species biofilms.
Both bacteria tend to co-aggregate and cooperate over time, persisting in a stable microbial community.
Development of new functional coatings using magnetron co-sputtering to deposit triple TiO2/SiO2/Ag nanocomposite thin filmsFlow rate of 53 mL s−1, corresponding to 15 s−148 hGlass
TiO2/SiO2 coated glass with different Ag contents (0 to 19.8 at %)
Escherichia coliBiofilm formation was reduced down to 92% compared to a control glass surface.
The coatings are promising candidates for antimicrobial protection of urinary tract devices for at least 48 h, suggesting benefits over longer periods.
Flow chamberGeneral medical devicesAdhesionAssessment of interactions of bacteria with specific biomaterial surface chemistries under flow conditions50, 500, 1000, and 2000 s−12 hGlass
Glass with alkyl silane monolayers
Staphylococcus epidermidisThe increase in the ionic strength enhanced adhesion to the different surfaces, in accordance with the Derjaguin–Landau–Verwey–Overbeek (DLVO) theory, under low shear rates.
The increase in the shear rate restricted the predictability of the theory.
Effect of shear stress on bacterial adhesion to biomedical materialsFlow rates of 2 and 4 mL s−1, corresponding to shear stresses of 0.01 and 0.022 Pa0.5 hGlass
Poly(L-lactic acid)
Escherichia coliSimilar adhesion rates were obtained on glass and polydimethylsiloxane.
The highest adhesion rates were obtained on glass and polydimethylsiloxane, and the lowest on poly(L-lactic acid).
Effect of fluid composition and shear conditions on bacterial adhesion to an antifouling peptide-coated surfaceFlow rates of 2 and 4 mL s−1, corresponding to 15 and 30 s−10.5 hGlass
Peptide-coated glass
Poly(L-lactic acid)
Escherichia coliAdhesion reductions of 40–50% were attained at a shear rate of 15 s−1 on the peptide-coated surfaces compared with glass.
The performance of the peptide-based antifouling coating was superior to poly(L-lactic acid).
Effect of shear stress on bacterial adhesion to antifouling polymer brushesFlow rates of 2 and 4 mL s−1, corresponding to 0.010 and 0.024 Pa0.5 hGlass
Poly[N-(2-hydroxypropyl) methacrylamide] brush
Poly[oligo(ethyleneglycol) methyl ether methacrylate] brush
Escherichia coliBoth polymer brushes reduced the initial adhesion up to 90% when compared to glass.[56]
Evaluate the antiadhesive activity of carbon nanotube compositesFlow rate of 2 mL s−1, corresponding to 15 s−10.5 hPolydimethylsiloxane
Carbon nanotube/polydimethylsiloxane composites
Escherichia coliThe introduction of carbon nanotubes composites in the polydimethylsiloxane matrix yielded less bacterial adhesion than the polydimethylsiloxane alone.
Less adhesion was obtained on the composites with pristine rather than functionalized carbon nanotubes.
Incorporation of higher amounts of carbon nanotubes in polymer composites can affect bacterial adhesion by more than 40%.
Composites enabling a 60% reduction in cell adhesion were obtained by carbon nanotube treatment by ball-milling.
Devices and implantsAdhesionPrevention of microbial adhesion to silicone rubber using polyacrylamide brush coatingsFlow rate of 0.025 mL s−1, corresponding to 10 s−14 hSilicone wafers
Silicone rubber
Polyacrylamide brushes
Staphylococcus aureus
Streptococcus salivarius
Escherichia coli
Candida albicans
A high reduction (52–92%) in microbial adhesion to the polyacrylamide brushes was observed compared to untreated silicon surfaces.
The polymer brush did not cause surface deterioration and discouraged microbial adhesion, even after 1-month of exposure to physiological fluids.
Implanted medical devicesAdhesionStudy of adhesion of bacterial and yeast strains to a poly(ethylene oxide) brush covalently attached to the glassFlow rate of 0.025 mL s−1, corresponding to 10 s−14 hGlass
Poly(ethylene oxide) brushes on glass
Staphylococcus epidermidis
Staphylococcus aureus
Streptococcus salivarius
Escherichia coli
Pseudomonas aeruginosa
Candida albicans
Candida tropicalis
The poly(ethylene oxide) brush yielded more than 98% reduction in bacterial adhesion, although for the more hydrophobic P. aeruginosa a smaller reduction was observed.
For yeast species, adhesion suppression was less effective than for the bacteria.
Evaluation of the role of surface free energy on bacterial adhesion to plasma-modified films50 and 200 s−12.5 hPolyethylene terephthalate
Plasma treated polyethylene terephthalate
Staphylococcus epidermidisPlasma treatments reduced bacterial adhesion, in comparison to the untreated polymer.
The ageing effect and the subsequent decrease in the surface free energy seemed to favor bacterial adhesion and aggregation.
The increase in the shear rate restricted the predictability of the thermodynamic models.
Adhesion and biofilm formationEvaluation of the effectiveness of different formulations of a biomedical-grade polyetherurethane at inhibiting bacterial colonization under flow conditions2.03 PaAdhesion: 2, 4 and 6 h
Biofilm: 8, 20 and 24 h
Polyetherurethane with triglyme
Polyetherurethane with poly(butyl methyacrylate) barrier membrane releasing ciprofloxacin
Pseudomonas aeruginosaThe rate of adherent cell accumulation was zero for the polyetherurethane with a poly(butyl methyacrylate) barrier membrane releasing ciprofloxacin.[135]
Surgical, catheters, and haemodialysis devicesAdhesionEvaluation of the adhesion behavior of bacterial strains to hydrophilic and hydrophobic surfaces using theoretical predictionsFlow rate of 0.025 mL s−1, corresponding to 6 s−12 hGlass
Indium tin oxide-coated glass
Pseudomonas stutzeri
Staphylococcus epidermidis
P. stutzeri has a much better adhesion rate than S. epidermidis for both material surfaces.
Both bacterial strains adhered better to the hydrophobic indium tin oxide-coated glass than to the hydrophilic glass.
Orthopedic implantsAdhesionStudy the bacterial adhesion to polymers that show promise as orthopedic materialsFlow rate of 1 mL min−1, corresponding to a shear rate of 1.9 s−11 hPoly(orthoester)
Poly(L-lactic acid)
Poly(ether ether ketone)
Staphylococcus epidermidis
Pseudomonas aeruginosa
Escherichia coli
Tryptic soy broth decreased adhesion to polymers, when compared to phosphate-buffered saline.
The estimated values of the free energy of adhesion correlated with the amount of adherent P. aeruginosa.
There was 50% more adhesion of E. coli and P. aeruginosa on poly(orthoester) and poly(L-lactic acid) pre-exposed to hyaluronic acid.
P. aeruginosa was the most adherent strain, while S. epidermidis was the least adherent strain.
Urinary devicesAdhesionExamination of the ability of probiotic strains to displace adhering cells from hydrophobic and hydrophilic substrata15 s−14.5 hGlass
Fluorinated ethylene propylene
Enterococcus faecalisEnt. faecalis was displaced by lactobacilli (31%) and streptococci (74%) from fluorinated ethylene propylene in buffer, and that displacement by lactobacilli was even more effective on glass in urine (54%).
The passage of an air–liquid interface impacted adhesion, especially when the surface had been challenged with lactobacilli (up to 100%) or streptococci (up to 94%).
Potential of biosurfactant layer to inhibit adhesion of uropathogensFlow rate of 0.034 mL s−1, corresponding to 15 s−14 hGlass
Silicone rubber coated with different concentrations of a biosurfactant
Enterococcus faecalisBiosurfactant layers inhibited the initial deposition rates (> 30%) and adhesion numbers (≈ 70–100%) in a dose-related way.For urine experiments, biosurfactant coatings caused higher adhesion reductions.[122]
Effect of supplementation on human urine and uropathogen adhesionFlow rate of 0.034 mL s−1, corresponding to 15 s−14 hSilicone rubberEscherichia coli
Enterococcus faecalis
Staphylococcus epidermidis
Pseudomonas aeruginosa
Candida albicans
Cranberry and ascorbic acid supplementation can provide a degree of protection against adhesion and colonization of biomaterials by some uropathogens.[139]
Effect of combined surface chemistry and topography on bacterial adhesionFlow rates of 2 and 4 mL s−1, corresponding to 0.010 and 0.024 Pa0.5 hSmooth polydimethylsiloxane
Smooth polydimethylsiloxane with peptide coating
Micropatterned polydimethylsiloxane
Micropatterned polydimethylsiloxane with peptide coating
Escherichia coliThe highest adhesion was obtained on the smooth polydimethylsiloxane, whereas the micropatterned polydimethylsiloxane coated with peptide totally inhibited adhesion.
The peptide addition to the smooth surface reduced the adhesion by 43–58%, while the micropatterned surface reduced the adhesion by 99%.
Biofilm formationImpact of temperature and surface on the biofilm-forming capacity of uropathogensFlow rate of 4 × 10−3 mL s−1, corresponding to 33 s−120–24 hSilicone
Silicone coated with plasma polymerized vinylpyrrolidone
Escherichia coliTemperature had a considerable influence upon the adhesion and biofilm-forming capacity of some of the isolates, and the influence of surface chemistry also depended on the temperature.[140]
Effect of applying different current densities to platinum electrodes as a possible catheter coating materialFlow rate of 3333 mL s−1, corresponding to 200 s−16 daysPlatinum electrodesProteus mirabilisBy applying alternating microcurrent densities, a self-regenerative surface is produced, which removed the conditioning film and reduced bacterial adherence, growth, and survival.[141]
Biofilm formation and treatmentPotential of using a polymer brush on the prevention of biofilm formation and susceptibilityFlow rate of 2 mL s−1, corresponding to 15 s−1Biofilm: 24 h
Treatment: 8 h
Poly[oligo(ethyleneglycol) methyl ether methacrylate] brush
Escherichia coliThe polymer brush reduced the surface area and the number of total adhered cells by 57%.
The antibiotic treatment potentiated cell death and removal (88%).
The polymer brush has the potential to prevent biofilm growth and in eradicating biofilms developed in urinary devices.
Effect of using a polymer brush on biofilm cell composition and architectureFlow rate of 2 mL s−1, corresponding to 15 s−1Biofilm: 24 h
Treatment: 8 h
Poly[N-(2-hydroxypropyl) methacrylamide] brush
Escherichia coliInitial adhesion and surface coverage decreased on the polymer brush.
Viable but nonculturable cells were completely removed from the brush.
The polymer brush may reduce biofilm growth and antibiotic resistance in urinary catheters.
96-well microplatesBiomedical scenariosBiofilm formationEvaluation of the combined effects of shear forces and nutrient levels on biofilm formation and definition of the operational conditions to be used to simulate relevant biomedical scenariosOrbital shaking with 25 and 50 mm diameter incubators at 150 rpm (average shear rate of 23 and 46 s−1)60 hPolystyreneEscherichia coliHigher glucose concentrations enhanced E. coli adhesion in the first 24 h, but variations in peptone and yeast extract concentrations had no significant impact on biofilm formation.
Numerical simulations indicate that 96-well microplates can be used to simulate a variety of biomedical scenarios if the operating conditions are carefully set.
Microfluidic deviceGeneral medical devicesAdhesionDevelopment of a fabrication method to produce a microfluidic device to test cell adhesion0.01–1 Pa0.5 hPolyamide
Polyethylene oxide
Poly(L-lactic acid)
Escherichia coliBacterial adhesion increased linearly over time.
The evaluation performed with polydimethylsiloxane for shear stresses between 0.02 and 1 Pa showed that the lowered surface (inherent weakness of the fabrication method) did not influence adhesion.
Study the initial cell adhesion dependence on local wall shear stress in a microchannel with intercalate zones of constrictions and expansions0.2–10 Pa0.5 hGlassEscherichia coliBacterial adhesion increased in locations with a sudden increase in shear stress.[142]
Examination of the role of surface properties on bacterial adhesion0.002–0.042 Pan.d.Smooth silicone
Patterned silicone
Escherichia coliCell attachment was observed to be strongly dependent upon the topographical features.
The highest attachment density was observed on smooth surfaces.
Biofilm formationComparison of the biofilm-forming capacities of various Methicillin-resistant Staphylococcus aureus clones0.05 Pa18 hGlassMethicillin-resistant Staphylococcus aureusFrom tested isolates, 51% successfully formed biofilms under shear flow.
Differences in biofilm formation might also be due to the different adherent surfaces.
Study of biofilm formation and host–pathogen interactions0.05–1 Pa24 hGlass
Eukaryotic cells (HRT-18)
Escherichia coliBiofilm formation on glass was observed for most strains in M9 medium at 30 °C.
HRT-18 cell monolayers enhanced E. coli binding and biofilm formation.
Implanted medical deviceBiofilm formationInvestigation of how environmental factors, such as surface geometry and chemistry, as well as fluid flow, affect biofilm development0.02–1 Pa16 h Uncoated and human blood plasma-coated channelsStaphylococcus aureusThe flow was the major contributor to the shape of biofilm structures, whereas bacterial motility was less significant.[115]
Mammary environmentBiofilm formationEvaluation of the effect of coagulase-negative staphylococci isolates with a weak- biofilm phenotype0.05 Pa24 hGlassCoagulase-negative staphylococciCoagulase-negative staphylococci with a weak biofilm phenotype did not inhibit the growth of isolates with a strong-biofilm phenotype.[145]
Intravascular catheterBiofilm formationInvestigation of flow as an environmental signal for biofilm formationFlow rates of 1–10 mL h−1, corresponding to 0.065–1.14 Pa24 hChannels treated with octyl(tri-ethoxy)silaneStaphylococcus epidermidisFluid shear alone induced the formation of polysaccharide intracellular adhesin-positive biofilms and influenced the biofilm structure.[116]
Urinary devicesAdhesionDevelopment of microfluidic-based devices replicating the urodynamic field within different configurations of an occluded and stented ureterUp to 0.175 Pa1 hPolydimethylsiloxane Pseudomonas fluorescensThe unobstructed device showed no bacterial attachment, including in regions of low shear stress (<0.04 Pa).
For the obstructed devices, the cavity region, and the nearby proximal side-hole (shear stresses of 0.131–0.175 Pa) exhibited greater levels of bacterial attachment (18%) compared to other regions of the model.
n.d.—not disclosed; D. tsuruhatensis—Delftia tsuruhatensis, Ent. faecalis—Enterococcus faecalis, E. coli—Escherichia coli, P. aeruginosa—Pseudomonas aeruginosa, P. stutzeri—Pseudomonas stutzeri, S. epidermidis—Staphylococcus epidermidis.

5. Current Challenges and Future Directions of Biofilm Platforms Research

Although biofilms are a recognized problem for the environment, industry, and medicine, and act as a possible reservoir of pathogens, there is a lack of reliable standard procedures to evaluate the efficacy of methods for biofilm prevention and removal. Consequently, it is very difficult to compare data obtained in different laboratories. As discussed before, laboratory reactors are available for growing biofilms that are more representative of a clinical situation [37,146] and industrial environment [147]. Although commercially available reactors with standardized protocols exist (e.g., ASTM Method E2871-13 and 2562-12 for the CDC biofilm reactor [148]), they are usually expensive and, thus, not accessible to all biofilm researchers, besides that the operation of these reactors has specific limitations. For instance, some of them cannot be used to test different surface materials, have reduced sampling areas, require specialized labor for operation, and the fluid dynamics are rarely well-characterized. While factors such as the temperature, microbial composition, and carbon source may be similar across different protocols and biofilm platforms, the fluid dynamics, namely the shear stress and shear rate, are a defining feature of a particular reactor operation. Whether the researchers are using a commercial or custom-made biofilm setup, computational simulations of hydrodynamics are extremely valuable, as they enable a more informed decision about whether the flow behavior in that specific biofilm reactor is suitable for their research.
Nevertheless, not all interactions between early adhered cells or established biofilms and fluid flow phases (gas and/or liquid) are considered when using the CFD technique. Almost all the flows in the described biofilm reactors deal with multiphase (gas–liquid, solid–liquid, and gas–liquid–solid), but some simplifications are introduced to reduce the model complexity [78,149]. For example, the aeration of flow cell systems is often not taken into account in the CFD study [43]. Furthermore, one must bear in mind that numerical simulations are mostly performed for clean and perfectly smooth surfaces. However, as biofilms grow or different coupon materials are used, the surface properties (such as roughness and hydrophobicity) should be considered for their impact on the wall shear stress. Therefore, there is still a great challenge in the integration of physical and biological processes in biofilm reactors.
Small flow chambers and microfluidic platforms are promising for screening new possible antibiofilm approaches. They need smaller volumes of media and reagents to run continuous biofilm experiments, when compared to the Robbins device and rotating biofilm reactors, enabling high-throughput assays. Additionally, the Bioflux [150] and other microfluidic devices [151,152] are dynamic systems with significant potential for monitoring heterogeneity in the biofilm microenvironment [153]. This can be achieved with specific stains and examination by confocal microscopy. However, direct biofilm observation might not be feasible, specific stains/probes may not be available (for nutrients or metabolites), or the time scale may be too slow. Introducing sensing techniques, such as microsensors or electrochemical probes in microfluidic chips, is an important development for online biofilm detection and microenvironment analysis [153].

6. Conclusions

Studying microbial adhesion and biofilm growth is crucial for understanding the physiology of sessile organisms and forming the basis for the development of novel antimicrobial materials. Fluid hydrodynamics is one of the most important factors affecting cell adhesion, as well as biofilm structure and behavior. Therefore, to simulate the relevant biofilms of different fields (environment, industry, and medicine) in the laboratory, it is of utmost importance to select an adequate biofilm platform and be able to operate it at hydrodynamic conditions that are as close as possible to those encountered in a real scenario.

Author Contributions

Conceptualization, L.C.G. and F.J.M.M.; investigation, L.C.G.; writing—original draft preparation, L.C.G.; writing—review and editing, L.C.G. and F.J.M.M.; supervision, F.J.M.M. All authors have read and agreed to the published version of the manuscript.


This work was financially supported by: Project PTDC/CTM-COM/4844/2020 funded by the Portuguese Foundation for Science and Technology (FCT); Project PTDC/BII-BIO/29589/2017-POCI-01-0145-FEDER-029589 funded by FEDER funds through COMPETE2020-Programa Operacional Competitividade e Internacionalização (POCI) and by national funds (PIDDAC) through FCT/MCTES; Base Funding—UIDB/00511/2020 of the Laboratory for Process Engineering, Environment, Biotechnology and Energy -LEPABE- funded by national funds through the FCT/MCTES (PIDDAC); Project HealthyWaters (NORTE-01-0145- FEDER-000069) and 2SMART (NORTE-01-0145-FEDER-000054) supported by Norte Portugal Regional Operational Programme (NORTE 2020) under the PORTUGAL 2020 Partnership Agreement and through the European Regional Development Fund (ERDF). L.C.G. thanks FCT for the financial support of her work contract through the Scientific Employment Stimulus -Individual Call-[CEECIND/01700/2017].

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Costerton, J.W.; Stewart, P.S.; Greenberg, E.P. Bacterial biofilms: A common cause of persistent infections. Science 1999, 284, 1318–1322. [Google Scholar] [CrossRef] [Green Version]
  2. Flemming, H.-C.; Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 2010, 8, 623–633. [Google Scholar] [CrossRef]
  3. Petrova, O.E.; Sauer, K. Sticky Situations: Key Components That Control Bacterial Surface Attachment. J. Bacteriol. 2012, 194, 2413–2425. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Nikolaev, Y.A.; Plakunov, V.K. Biofilm—“City of microbes” or an analogue of multicellular organisms? Microbiology 2007, 76, 125–138. [Google Scholar] [CrossRef]
  5. Monroe, D. Looking for Chinks in the Armor of Bacterial Biofilms. PLoS Biol. 2007, 5, e307. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  6. Stoodley, P.; Sauer, K.; Davies, D.G.; Costerton, J.W. Biofilms as Complex Differentiated Communities. Annu. Rev. Microbiol. 2002, 56, 187–209. [Google Scholar] [PubMed] [Green Version]
  7. Slate, A.J.; Wickens, D.; Wilson-Nieuwenhuis, J.; Dempsey-Hibbert, N.; West, G.; Kelly, P.; Verran, J.; Banks, C.E.; Whitehead, K.A. The effects of blood conditioning films on the antimicrobial and retention properties of zirconium-nitride silver surfaces. Colloids Surf. B Biointerfaces 2019, 173, 303–311. [Google Scholar] [CrossRef]
  8. Moreira, J.M.R.; Gomes, L.C.; Whitehead, K.A.; Lynch, S.; Tetlow, L.A.; Mergulhão, F.J. Effect of surface conditioning with cellular extracts on Escherichia coli adhesion and initial biofilm formation. Food Bioprod. Process. 2017, 104, 1–12. [Google Scholar] [CrossRef]
  9. Renner, L.D.; Weibel, D.B. Physicochemical regulation of biofilm formation. MRS Bull. 2011, 36, 347–355. [Google Scholar] [CrossRef] [Green Version]
  10. Donlan, R.M. Biofilms: Microbial life on surfaces. Emerg. Infect. Dis. 2002, 8, 881–890. [Google Scholar] [CrossRef] [PubMed]
  11. de Carvalho, C.C.C.R. Marine Biofilms: A Successful Microbial Strategy with Economic Implications. Front. Mar. Sci. 2018, 5. [Google Scholar] [CrossRef] [Green Version]
  12. Demirel, Y.K.; Turan, O.; Incecik, A. Predicting the effect of biofouling on ship resistance using CFD. Appl. Ocean. Res. 2017, 62, 100–118. [Google Scholar] [CrossRef] [Green Version]
  13. Bannister, J.; Sievers, M.; Bush, F.; Bloecher, N. Biofouling in marine aquaculture: A review of recent research and developments. Biofouling 2019, 35, 631–648. [Google Scholar] [CrossRef] [Green Version]
  14. Bott, T.R. Industrial Biofouling. In Industrial Biofouling; Bott, T.R., Ed.; Elsevier: Amsterdam, The Netherlands, 2011; pp. 1–5. [Google Scholar]
  15. Flemming, H.-C. Microbial Biofouling: Unsolved Problems, Insufficient Approaches, and Possible Solutions. In Biofilm Highlights; Flemming, H.-C., Wingender, J., Szewzyk, U., Eds.; Springer: Berlin/Heidelberg, Germany, 2011; pp. 81–109. [Google Scholar]
  16. Srey, S.; Jahid, I.K.; Ha, S.-D. Biofilm formation in food industries: A food safety concern. Food Control 2013, 31, 572–585. [Google Scholar] [CrossRef]
  17. Shunmugaperumal, T. Introduction and overview of biofilm. In Biofilm Eradication and Prevention; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2010; pp. 36–72. [Google Scholar]
  18. Jamal, M.; Ahmad, W.; Andleeb, S.; Jalil, F.; Imran, M.; Nawaz, M.A.; Hussain, T.; Ali, M.; Rafiq, M.; Kamil, M.A. Bacterial biofilm and associated infections. J. Chin. Med. Assoc. JCMA 2018, 81, 7–11. [Google Scholar] [CrossRef]
  19. Marschang, S.; Bernardo, G. Prevention and control of healthcare-associated infection in Europe: A review of patients’ perspectives and existing differences. J. Hosp. Infect. 2015, 89, 357–362. [Google Scholar] [CrossRef] [PubMed]
  20. Kaali, P.; Strömberg, E.; Karlsson, S. Prevention of biofilm associated infections and degradation of polymeric materials used in biomedical applications. In Biomedical Engineering, Trends in Materials Science; Citeseer: Princeton, NJ, USA, 2011. [Google Scholar]
  21. Dolid, A.; Gomes, L.C.; Mergulhão, F.J.; Reches, M. Combining chemistry and topography to fight biofilm formation: Fabrication of micropatterned surfaces with a peptide-based coating. Colloids Surf. B Biointerfaces 2020, 196, 111365. [Google Scholar] [CrossRef] [PubMed]
  22. Silva, E.R.; Tulcidas, A.V.; Ferreira, O.; Bayón, R.; Igartua, A.; Mendoza, G.; Mergulhão, F.J.M.; Faria, S.I.; Gomes, L.C.; Carvalho, S.; et al. Assessment of the environmental compatibility and antifouling performance of an innovative biocidal and foul-release multifunctional marine coating. Environ. Res. 2021, 198, 111219. [Google Scholar] [CrossRef]
  23. Vagos, M.R.; Gomes, M.; Moreira, J.M.R.; Soares, O.S.G.P.; Pereira, M.F.R.; Mergulhão, F.J. Carbon Nanotube/Poly(dimethylsiloxane) Composite Materials to Reduce Bacterial Adhesion. Antibiotics 2020, 9, 434. [Google Scholar]
  24. Moreira, J.M.R.; Fulgêncio, R.; Alves, P.; Machado, I.; Bialuch, I.; Melo, L.F.; Simões, M.; Mergulhão, F.J. Evaluation of SICAN performance for biofouling mitigation in the food industry. Food Control 2016, 62, 201–207. [Google Scholar] [CrossRef] [Green Version]
  25. Martinuzzi, R.J.; Salek, M.M. Numerical Simulation of Fluid Flow and Hydrodynamic Analysis in Commonly Used Biomedical Devices in Biofilm Studies. In Numerical Simulations—Examples and Applications in Computational Fluid Dynamics; Angermann, L., Ed.; InTech: London, UK, 2010; pp. 193–212. [Google Scholar]
  26. Katsikogianni, M.; Missirlis, Y.F. Concise review of mechanisms of bacterial adhesion to biomaterials and of techniques used in estimating bacteria-material interactions. Eur. Cells Mater. 2004, 8, 37–57. [Google Scholar] [CrossRef]
  27. Gjersing, E.L.; Codd, S.L.; Seymour, J.D.; Stewart, P.S. Magnetic resonance microscopy analysis of advective transport in a biofilm reactor. Biotechnol. Bioeng. 2005, 89, 822–834. [Google Scholar] [CrossRef]
  28. Liu, Y.; Tay, J.-H. The essential role of hydrodynamic shear force in the formation of biofilm and granular sludge. Water Res. 2002, 36, 1653–1665. [Google Scholar] [CrossRef]
  29. Stewart, P.S. A review of experimental measurements of effective diffusive permeabilities and effective diffusion coefficients in biofilms. Biotechnol. Bioeng. 1998, 59, 261–272. [Google Scholar] [CrossRef]
  30. Stewart, P.S. Diffusion in Biofilms. J. Bacteriol. 2003, 185, 1485–1491. [Google Scholar] [CrossRef] [Green Version]
  31. Munson, B.R.; Young, D.F.; Okiishi, T.H. Fundamentals of Fluid Mechanics, 4th ed.; John Wiley & Sons, Inc.: Chicago, IL, USA, 2002. [Google Scholar]
  32. Werner, S.; Kaiser, S.C.; Kraume, M.; Eibl, D. Computational fluid dynamics as a modern tool for engineering characterization of bioreactors. Pharm. Bioprocess. 2014, 2, 85–89. [Google Scholar] [CrossRef]
  33. Ramírez-Muñoz, J.; Guadarrama-Pérez, R.; Alvarado-Lassman, A.; Valencia-López, J.J.; Márquez-Baños, V.E. CFD study of the hydrodynamics and biofilm growth effect of an anaerobic inverse fluidized bed reactor operating in the laminar regime. J. Environ. Chem. Eng. 2021, 9, 104674. [Google Scholar] [CrossRef]
  34. Sharma, C.; Malhotra, D.; Rathore, A.S. Review of Computational fluid dynamics applications in biotechnology processes. Biotechnol. Prog. 2011, 27, 1497–1510. [Google Scholar] [CrossRef] [PubMed]
  35. Salek, M.M.; Jones, S.M.; Martinuzzi, R.J. The influence of flow cell geometry related shear stresses on the distribution, structure and susceptibility of Pseudomonas aeruginosa 01 biofilms. Biofouling 2009, 25, 711–725. [Google Scholar] [CrossRef]
  36. Azeredo, J.; Azevedo, N.F.; Briandet, R.; Cerca, N.; Coenye, T.; Costa, A.R.; Desvaux, M.; Di Bonaventura, G.; Hébraud, M.; Jaglic, Z.; et al. Critical review on biofilm methods. Crit. Rev. Microbiol. 2017, 43, 313–351. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  37. Ramstedt, M.; Ribeiro, I.A.C.; Bujdakova, H.; Mergulhao, F.J.M.; Jordao, L.; Thomsen, P.; Alm, M.; Burmolle, M.; Vladkova, T.; Can, F.; et al. Evaluating Efficacy of Antimicrobial and Antifouling Materials for Urinary Tract Medical Devices: Challenges and Recommendations. Macromol. Biosci. 2019, 19, e1800384. [Google Scholar] [CrossRef] [Green Version]
  38. Gomes, I.B.; Simões, M.; Simões, L.C. An overview on the reactors to study drinking water biofilms. Water Res. 2014, 62, 63–87. [Google Scholar] [CrossRef] [Green Version]
  39. Hall-Stoodley, L.; Rayner, J.C.; Stoodley, P.; Lappin-Scott, H.M. Establishment of Experimental Biofilms Using the Modified Robbins Device and Flow Cells. In Environmental Monitoring of Bacteria; Edwards, C., Ed.; Humana Press: Totowa, NJ, USA, 1999; pp. 307–319. [Google Scholar]
  40. Moreira, J.M.R.; Araújo, J.D.P.; Miranda, J.M.; Simões, M.; Melo, L.F.; Mergulhão, F.J. The effects of surface properties on Escherichia coli adhesion are modulated by shear stress. Colloids Surf. B Biointerfaces 2014, 123, 1–7. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  41. Moreira, J.M.R.; Fulgêncio, R.; Oliveira, F.; Machado, I.; Bialuch, I.; Melo, L.F.; Simões, M.; Mergulhão, F.J. Evaluation of SICON® surfaces for biofouling mitigation in critical process areas. Food Bioprod. Process. 2016, 98, 173–180. [Google Scholar] [CrossRef] [Green Version]
  42. Teodósio, J.S.; Simões, M.; Melo, L.F.; Mergulhão, F.J. Flow cell hydrodynamics and their effects on E. coli biofilm formation under different nutrient conditions and turbulent flow. Biofouling 2011, 27, 1–11. [Google Scholar] [CrossRef]
  43. Teodósio, J.S.; Silva, F.C.; Moreira, J.M.R.; Simões, M.; Melo, L.F.; Alves, M.A.; Mergulhão, F.J. Flow cells as quasi-ideal systems for biofouling simulation of industrial piping systems. Biofouling 2013, 29, 953–966. [Google Scholar] [CrossRef] [PubMed]
  44. McCoy, W.F.; Bryers, J.D.; Robbins, J.; Costerton, J.W. Observations of fouling biofilm formation. Can. J. Microbiol. 1981, 27, 910–917. [Google Scholar] [CrossRef] [PubMed]
  45. Stoodley, P.; Warwood, B.K. Use of flow cells an annular reactors to study biofilms. In Biofilms in Medicine, Industry and Environmental Biotechnology: Characteristics, Analysis and Control, 1st ed.; Lens, P., O’Flaherty, V., Moran, A.P., Stoodley, P., Mahony, T., Eds.; IWA Publishing: Cornwall, UK, 2003; pp. 197–213. [Google Scholar]
  46. Gomes, L.C.; Mergulhão, F.J. Heterologous protein production in Escherichia coli biofilms: A non-conventional form of high cell density cultivation. Process. Biochem. 2017, 57, 1–8. [Google Scholar] [CrossRef]
  47. Pereira, M.O.; Kuehn, M.; Wuertz, S.; Neu, T.; Melo, L.F. Effect of flow regime on the architecture of a Pseudomonas fluorescens biofilm. Biotechnol. Bioeng. 2002, 78, 164–171. [Google Scholar] [CrossRef] [PubMed]
  48. Teodósio, J.S.; Simões, M.; Alves, M.A.; Melo, L.F.; Mergulhão, F.J. Setup and Validation of Flow Cell Systems for Biofouling Simulation in Industrial Settings. Sci. World J. 2012, 2012, 361496. [Google Scholar] [CrossRef]
  49. Vladkova, T.; Angelov, O.; Stoyanova, D.; Gospodinova, D.; Gomes, L.C.; Soares, A.; Mergulhao, F.J.; Ivanova, I. Magnetron co-sputtered TiO2/SiO2/Ag nanocomposite thin coatings inhibiting bacterial adhesion and biofilm formation. Surf. Coat. Technol. 2020, 384, 125322. [Google Scholar] [CrossRef]
  50. Matinha-Cardoso, J.; Mota, R.; Gomes, L.C.; Gomes, M.; Mergulhão, F.J.; Tamagnini, P.; Martins, M.C.L.; Costa, F. Surface activation of medical grade polyurethane for the covalent immobilization of an anti-adhesive biopolymeric coating. J. Mater. Chem. B 2021, 9, 3705–3715. [Google Scholar] [CrossRef] [PubMed]
  51. Zou, F.; Bai, L. Using time-lapse fluorescence microscopy to study gene regulation. Methods 2019, 159–160, 138–145. [Google Scholar] [CrossRef] [PubMed]
  52. Bos, R.; van der Mei, H.C.; Busscher, H.J. Physico-chemistry of initial microbial adhesive interactions—Its mechanisms and methods for study. FEMS Microbiol. Rev. 1999, 23, 179–230. [Google Scholar] [CrossRef]
  53. Moreira, J.M.R.; Ponmozhi, J.; Campos, J.B.L.M.; Miranda, J.M.; Mergulhão, F.J. Micro- and macro-flow systems to study Escherichia coli adhesion to biomedical materials. Chem. Eng. Sci. 2015, 126, 440–445. [Google Scholar] [CrossRef] [Green Version]
  54. Alves, P.; Moreira, J.M.; Miranda, J.M.; Mergulhão, F.J. Analysing the Initial Bacterial Adhesion to Evaluate the Performance of Antifouling Surfaces. Antibiotics 2020, 9, 421. [Google Scholar] [CrossRef]
  55. Busscher, H.J.; van der Mei, H.C. Microbial adhesion in flow displacement systems. Clin. Microbiol. Rev. 2006, 19, 127–141. [Google Scholar] [CrossRef] [Green Version]
  56. Lopez-Mila, B.; Alves, P.; Riedel, T.; Dittrich, B.; Mergulhão, F.; Rodriguez-Emmenegger, C. Effect of shear stress on the reduction of bacterial adhesion to antifouling polymers. Bioinspir. Biomim. 2018, 13, 065001. [Google Scholar] [CrossRef]
  57. Alves, P.; Nir, S.; Reches, M.; Mergulhão, F. The effects of fluid composition and shear conditions on bacterial adhesion to an antifouling peptide-coated surface. MRS Commun. 2018, 8, 938–946. [Google Scholar] [CrossRef]
  58. Alves, P.; Gomes, L.C.; Vorobii, M.; Rodriguez-Emmenegger, C.; Mergulhão, F.J. The potential advantages of using a poly(HPMA) brush in urinary catheters: Effects on biofilm cells and architecture. Colloids Surf. B Biointerfaces 2020, 191, 110976. [Google Scholar] [CrossRef]
  59. Alves, P.; Gomes, L.C.; Rodríguez-Emmenegger, C.; Mergulhão, F.J. Efficacy of A Poly(MeOEGMA) Brush on the Prevention of Escherichia coli Biofilm Formation and Susceptibility. Antibiotics 2020, 9, 216. [Google Scholar] [CrossRef]
  60. Cotter, J.J.; O’Gara, J.P.; Stewart, P.S.; Pitts, B.; Casey, E. Characterization of a modified rotating disk reactor for the cultivation of Staphylococcus epidermidis biofilm. J. Appl. Microbiol. 2010, 109, 2105–2117. [Google Scholar] [CrossRef] [Green Version]
  61. Willcock, L.; Gilbert, P.; Holah, J.; Wirtanen, G.; Allison, D.G. A new technique for the performance evaluation of clean-in-place disinfection of biofilms. J. Ind. Microbiol. Biotechnol. 2000, 25, 235–241. [Google Scholar] [CrossRef]
  62. Kim, J.; Park, H.D.; Chung, S. Microfluidic approaches to bacterial biofilm formation. Molecules 2012, 17, 9818–9834. [Google Scholar] [CrossRef] [Green Version]
  63. De Grazia, A.; LuTheryn, G.; Meghdadi, A.; Mosayyebi, A.; Espinosa-Ortiz, E.J.; Gerlach, R.; Carugo, D. A Microfluidic-Based Investigation of Bacterial Attachment in Ureteral Stents. Micromachines 2020, 11, 408. [Google Scholar] [CrossRef]
  64. Yawata, Y.; Nguyen, J.; Stocker, R.; Rusconi, R. Microfluidic Studies of Biofilm Formation in Dynamic Environments. J. Bacteriol. 2016, 198, 2589–2595. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Huang, C.; Wippold, J.A.; Stratis-Cullum, D.; Han, A. Eliminating air bubble in microfluidic systems utilizing integrated in-line sloped microstructures. Biomed. Microdevices 2020, 22, 76. [Google Scholar] [CrossRef] [PubMed]
  66. Kumar, S.; Wittmann, C.; Heinzle, E. Minibioreactors. Biotechnol. Lett. 2004, 26, 1–10. [Google Scholar] [CrossRef]
  67. Moreira, J.M.R.; Gomes, L.C.; Araújo, J.D.P.; Miranda, J.M.; Simões, M.; Melo, L.F.; Mergulhão, F.J. The effect of glucose concentration and shaking conditions on Escherichia coli biofilm formation in microtiter plates. Chem. Eng. Sci. 2013, 94, 192–199. [Google Scholar] [CrossRef]
  68. Gomes, L.C.; Moreira, J.M.R.; Teodósio, J.S.; Araújo, J.D.P.; Miranda, J.M.; Simões, M.; Melo, L.F.; Mergulhão, F.J. 96-well microtiter plates for biofouling simulation in biomedical settings. Biofouling 2014, 30, 535–546. [Google Scholar] [CrossRef] [PubMed]
  69. Stepanovic, S.; Vukovic, D.; Hola, V.; Di Bonaventura, G.; Djukic, S.; Cirkovic, I.; Ruzicka, F. Quantification of biofilm in microtiter plates: Overview of testing conditions and practical recommendations for assessment of biofilm production by staphylococci. APMIS Acta Pathol. Microbiol. Immunol. Scand. 2007, 115, 891–899. [Google Scholar] [CrossRef]
  70. Faria, S.I.; Teixeira-Santos, R.; Romeu, M.J.; Morais, J.; Jong, E.D.; Sjollema, J.; Vasconcelos, V.; Mergulhão, F.J. Unveiling the Antifouling Performance of Different Marine Surfaces and Their Effect on the Development and Structure of Cyanobacterial Biofilms. Microorganisms 2021, 9, 1102. [Google Scholar] [CrossRef]
  71. Romeu, M.J.; Alves, P.; Morais, J.; Miranda, J.M.; de Jong, E.D.; Sjollema, J.; Ramos, V.; Vasconcelos, V.; Mergulhão, F.J.M. Biofilm formation behaviour of marine filamentous cyanobacterial strains in controlled hydrodynamic conditions. Environ. Microbiol. 2019, 21, 4411–4424. [Google Scholar] [CrossRef] [PubMed]
  72. Bridier, A.; Dubois-Brissonnet, F.; Boubetra, A.; Thomas, V.; Briandet, R. The biofilm architecture of sixty opportunistic pathogens deciphered using a high throughput CLSM method. J. Microbiol. Methods 2010, 82, 64–70. [Google Scholar] [CrossRef] [PubMed]
  73. Moreira, J.M.R.; Gomes, L.C.; Simões, M.; Melo, L.F.; Mergulhão, F.J. The impact of material properties, nutrient load and shear stress on biofouling in food industries. Food Bioprod. Process. 2015, 95, 228–236. [Google Scholar] [CrossRef] [Green Version]
  74. Gomes, M.; Gomes, L.C.; Teixeira-Santos, R.; Pereira, M.F.R.; Soares, O.S.G.P.; Mergulhão, F.J. Optimizing CNT Loading in Antimicrobial Composites for Urinary Tract Application. Appl. Sci. 2021, 11, 4038. [Google Scholar] [CrossRef]
  75. Gomes, L.C.; Moreira, J.M.; Miranda, J.M.; Simões, M.; Melo, L.F.; Mergulhão, F.J. Macroscale versus microscale methods for physiological analysis of biofilms formed in 96-well microtiter plates. J. Microbiol. Methods 2013, 95, 342–349. [Google Scholar] [CrossRef]
  76. Gomes, L.C.; Moreira, J.M.R.; Simões, M.; Melo, L.F.; Mergulhão, F.J. Biofilm Localization in the Vertical Wall of Shaking 96-Well Plates. Scientifica 2014, 2014, 6. [Google Scholar] [CrossRef]
  77. Gomes, L.C.; Moreira, J.M.R.; Araújo, J.D.P.; Mergulhão, F.J. Surface conditioning with Escherichia coli cell wall components can reduce biofilm formation by decreasing initial adhesion. AIMS Microbiol. 2017, 3, 613–628. [Google Scholar] [CrossRef]
  78. Gomes, L.C.; Miranda, J.; Mergulhão, F.J. Operation of Biofilm Reactors for the Food Industry Using CFD. In Computational Fluid Dynamics in Food Processing, 2nd ed.; Sun, D.-W., Ed.; CRC Press: Boca Raton, FL, USA, 2019. [Google Scholar]
  79. Cowle, M.W.; Webster, G.; Babatunde, A.O.; Bockelmann-Evans, B.N.; Weightman, A.J. Impact of flow hydrodynamics and pipe material properties on biofilm development within drinking water systems. Environ. Technol. 2020, 41, 3732–3744. [Google Scholar] [CrossRef] [Green Version]
  80. Gomes, I.B.; Lemos, M.; Mathieu, L.; Simões, M.; Simões, L.C. The action of chemical and mechanical stresses on single and dual species biofilm removal of drinking water bacteria. Sci. Total Environ. 2018, 631–632, 987–993. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  81. Ferreira, O.; Rijo, P.; Gomes, J.; Santos, R.; Monteiro, S.; Guedes, R.; Serralheiro, M.L.; Gomes, M.; Gomes, L.C.; Mergulhão, F.J.; et al. Antimicrobial Ceramic Filters for Water Bio-Decontamination. Coatings 2021, 11, 323. [Google Scholar] [CrossRef]
  82. Faria, S.I.; Teixeira-Santos, R.; Gomes, L.C.; Silva, E.R.; Morais, J.; Vasconcelos, V.; Mergulhão, F.J.M. Experimental Assessment of the Performance of Two Marine Coatings to Curb Biofilm Formation of Microfoulers. Coatings 2020, 10, 893. [Google Scholar] [CrossRef]
  83. Mathieu, L.; Bertrand, I.; Abe, Y.; Angel, E.; Block, J.C.; Skali-Lami, S.; Francius, G. Drinking water biofilm cohesiveness changes under chlorination or hydrodynamic stress. Water Res. 2014, 55, 175–184. [Google Scholar] [CrossRef]
  84. Bakker, D.P.; Plaats, A.v.d.; Verkerke, G.J.; Busscher, H.J.; Mei, H.C.v.d. Comparison of Velocity Profiles for Different Flow Chamber Designs Used in Studies of Microbial Adhesion to Surfaces. Appl. Environ. Microbiol. 2003, 69, 6280–6287. [Google Scholar] [CrossRef] [Green Version]
  85. Liu, C.; Zhao, Q. The CQ ratio of surface energy components influences adhesion and removal of fouling bacteria. Biofouling 2011, 27, 275–285. [Google Scholar] [CrossRef]
  86. Liu, C.; Zhao, Q. Influence of Surface-Energy Components of Ni–P–TiO2–PTFE Nanocomposite Coatings on Bacterial Adhesion. Langmuir 2011, 27, 9512–9519. [Google Scholar] [CrossRef]
  87. Gomes, I.B.; Simões, L.C.; Simões, M. Influence of surface copper content on Stenotrophomonas maltophilia biofilm control using chlorine and mechanical stress. Biofouling 2020, 36, 1–13. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  88. Azevedo, N.F.; Pinto, A.R.; Reis, N.M.; Vieira, M.J.; Keevil, C.W. Shear stress, temperature, and inoculation concentration influence the adhesion of water-stressed Helicobacter pylori to stainless steel 304 and polypropylene. Appl. Environ. Microbiol. 2006, 72, 2936–2941. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  89. Romeu, M.J.; Alves, P.; Morais, J.; Ramos, V.; Vasconcelos, V.; Mergulhão, F.J. Biofilm formation by a cyanobacterial strain belonging to a distinct Leptolyngbya phylotype: Surface effect. In Biofilms: Characterization, Applications and Recent Advances; Henderson, J., Ed.; Nova Science Publishers: Hauppauge, NY, USA, 2016; pp. 153–174. [Google Scholar]
  90. Faria, S.I.; Teixeira-Santos, R.; Romeu, M.J.; Morais, J.; Vasconcelos, V.; Mergulhão, F.J. The Relative Importance of Shear Forces and Surface Hydrophobicity on Biofilm Formation by Coccoid Cyanobacteria. Polymers 2020, 12, 653. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  91. Romeu, M.J.L.; Domínguez-Pérez, D.; Almeida, D.; Morais, J.; Campos, A.; Vasconcelos, V.; Mergulhão, F.J.M. Characterization of planktonic and biofilm cells from two filamentous cyanobacteria using a shotgun proteomic approach. Biofouling 2020, 36, 631–645. [Google Scholar] [CrossRef] [PubMed]
  92. Faria, S.I.; Teixeira-Santos, R.; Morais, J.; Vasconcelos, V.; Mergulhão, F.J. The association between initial adhesion and cyanobacterial biofilm development. FEMS Microbiol. Ecol. 2021, 97, fiab052. [Google Scholar] [CrossRef] [PubMed]
  93. Romeu, M.J.; Domínguez-Pérez, D.; Almeida, D.; Morais, J.; Araújo, M.J.; Osório, H.; Campos, A.; Vasconcelos, V.; Mergulhão, F.J. Quantitative proteomic analysis of marine biofilms formed by filamentous cyanobacterium. Environ. Res. 2021, 201, 111566. [Google Scholar] [CrossRef]
  94. Perrin, A.; Herbelin, P.; Jorand, F.P.A.; Skali-Lami, S.; Mathieu, L. Design of a rotating disk reactor to assess the colonization of biofilms by free-living amoebae under high shear rates. Biofouling 2018, 34, 368–377. [Google Scholar] [CrossRef]
  95. Lemos, M.; Mergulhão, F.; Melo, L.; Simões, M. The effect of shear stress on the formation and removal of Bacillus cereus biofilms. Food Bioprod. Process. 2015, 93, 242–248. [Google Scholar] [CrossRef] [Green Version]
  96. Brugnoni, L.I.; Cubitto, M.A.; Lozano, J.E. Role of shear stress on biofilm formation of Candida krusei in a rotating disk system. J. Food Eng. 2011, 102, 266–271. [Google Scholar] [CrossRef]
  97. Szlavik, J.; Paiva, D.S.; Mørk, N.; van den Berg, F.; Verran, J.; Whitehead, K.; Knøchel, S.; Nielsen, D.S. Initial adhesion of Listeria monocytogenes to solid surfaces under liquid flow. Int. J. Food Microbiol. 2012, 152, 181–188. [Google Scholar] [CrossRef] [Green Version]
  98. Skovager, A.; Whitehead, K.; Siegumfeldt, H.; Ingmer, H.; Verran, J.; Arneborg, N. Influence of flow direction and flow rate on the initial adhesion of seven Listeria monocytogenes strains to fine polished stainless steel. Int. J. Food Microbiol. 2012, 157, 174–181. [Google Scholar] [CrossRef]
  99. Piepiórka-Stepuk, J.; Mierzejewska, S. Simulation tests of liquid flow in the pipeline elements. Agric. Eng. 2014, 1, 175–184. [Google Scholar]
  100. Jensen, B.B.B.; Friis, A. CFD Prediction of Hygiene in Food Processing Equipment. In Computational Fluid Dynamics in Food Processing; Sun, D.-W., Ed.; CRC Press: Boca Raton, FL, USA, 2007. [Google Scholar]
  101. Bergman, B.-O.; Tragardh, C. An approach to study and model the hydrodynamic cleaning effect. J. Food Process. Eng. 1990, 13, 135–154. [Google Scholar] [CrossRef]
  102. Cunault, C.; Faille, C.; Bouvier, L.; Föste, H.; Augustin, W.; Scholl, S.; Debreyne, P.; Benezech, T. A novel set-up and a CFD approach to study the biofilm dynamics as a function of local flow conditions encountered in fresh-cut food processing equipments. Food Bioprod. Process. 2015, 93, 217–223. [Google Scholar] [CrossRef]
  103. Jensen, B.B.B.; Stenby, M.; Nielsen, D.F. Improving the cleaning effect by changing average velocity. Trends Food Sci. Technol. 2007, 18 (Suppl. 1), S58–S63. [Google Scholar] [CrossRef]
  104. Lelièvre, C.; Legentilhomme, P.; Legrand, J.; Faille, C.; Bénézech, T. Hygienic Design: Influence of the Local Wall Shear Stress Variations on the Cleanability of a Three-Way Valve. Chem. Eng. Res. Des. 2003, 81, 1071–1076. [Google Scholar] [CrossRef]
  105. Rahaman, M.F.; Bari, S.; Veale, D. Flow investigation of the product fill valve of filling machine for packaging liquid products. J. Food Eng. 2008, 85, 252–258. [Google Scholar] [CrossRef]
  106. Jin, Y.; Chen, X.D. A fundamental model of particle deposition incorporated in CFD simulations of an industrial milk spray dryer. Dry. Technol. 2010, 28, 960–971. [Google Scholar] [CrossRef]
  107. Lelièvre, C.; Legentilhomme, P.; Gaucher, C.; Legrand, J.; Faille, C.; Bénézech, T. Cleaning in place: Effect of local wall shear stress variation on bacterial removal from stainless steel equipment. Chem. Eng. Sci. 2002, 57, 1287–1297. [Google Scholar] [CrossRef]
  108. Afonso, I.M.; Hes, L.; Maia, J.M.; Melo, L.F. Heat transfer and rheology of stirred yoghurt during cooling in plate heat exchangers. J. Food Eng. 2003, 57, 179–187. [Google Scholar] [CrossRef] [Green Version]
  109. Fernandes, C.S.; Dias, R.; Nóbrega, J.M.; Afonso, I.M.; Melo, L.F.; Maia, J.M. Simulation of stirred yoghurt processing in plate heat exchangers. J. Food Eng. 2005, 69, 281–290. [Google Scholar] [CrossRef] [Green Version]
  110. Nørgaard, E.; Sørensen, T.A.; Hansen, T.M.; Kauffeld, M. Performance of components of ice slurry systems: Pumps, plate heat exchangers, and fittings. Int. J. Refrig. 2005, 28, 83–91. [Google Scholar] [CrossRef]
  111. Moreira, J.M.R.; Teodósio, J.S.; Silva, F.C.; Simões, M.; Melo, L.F.; Mergulhão, F.J. Influence of flow rate variation on the development of Escherichia coli biofilms. Bioprocess. Biosyst. Eng. 2013, 36, 1787–1796. [Google Scholar] [CrossRef]
  112. Moreira, J.M.R.; Simões, M.; Melo, L.F.; Mergulhão, F.J. The combined effects of shear stress and mass transfer on the balance between biofilm and suspended cell dynamics. Desalination Water Treat. 2015, 53, 3348–3354. [Google Scholar] [CrossRef] [Green Version]
  113. Ponmozhi, J.; Moreira, J.M.R.; Mergulhão, F.J.; Campos, J.B.L.M.; Miranda, J.M. Fabrication and Hydrodynamic Characterization of a Microfluidic Device for Cell Adhesion Tests in Polymeric Surfaces. Micromachines 2019, 10, 303. [Google Scholar] [CrossRef] [Green Version]
  114. Graham, M.V.; Mosier, A.P.; Kiehl, T.R.; Kaloyeros, A.E.; Cady, N.C. Development of antifouling surfaces to reduce bacterial attachment. Soft Matter 2013, 9, 6235–6244. [Google Scholar] [CrossRef]
  115. Kim, M.K.; Drescher, K.; Pak, O.S.; Bassler, B.L.; Stone, H.A. Filaments in curved streamlines: Rapid formation of Staphylococcus aureus biofilm streamers. New J. Phys. 2014, 16, 065024. [Google Scholar]
  116. Weaver, W.M.; Milisavljevic, V.; Miller, J.F.; Di Carlo, D. Fluid flow induces biofilm formation in Staphylococcus epidermidis polysaccharide intracellular adhesin-positive clinical isolates. Appl. Environ. Microbiol. 2012, 78, 5890–5896. [Google Scholar] [CrossRef] [Green Version]
  117. Inauen, W.; Baumgartner, H.R.; Bombeli, T.; Haeberli, A.; Straub, P.W. Dose- and shear rate-dependent effects of heparin on thrombogenesis induced by rabbit aorta subendothelium exposed to flowing human blood. Arterioscler. Off. J. Am. Heart Assoc. Inc. 1990, 10, 607–615. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  118. Michelson, A. Platelets, 2nd ed.; Academic Press: New York, NY, USA, 2002. [Google Scholar]
  119. Cai, Z.; Xin, J.; Pollock, D.M.; Pollock, J.S. Shear stress-mediated NO production in inner medullary collecting duct cells. Am. J. Physiology. Ren. Physiol. 2000, 279, F270–F274. [Google Scholar] [CrossRef]
  120. Nauman, E.A.; Ott, C.M.; Sander, E.; Tucker, D.L.; Pierson, D.; Wilson, J.W.; Nickerson, C.A. Novel Quantitative Biosystem for Modeling Physiological Fluid Shear Stress on Cells. Appl. Environ. Microbiol. 2007, 73, 699–705. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  121. Singh, P.K.; Marzo, A.; Howard, B.; Rufenacht, D.A.; Bijlenga, P.; Frangi, A.F.; Lawford, P.V.; Coley, S.C.; Hose, D.R.; Patel, U.J. Effects of smoking and hypertension on wall shear stress and oscillatory shear index at the site of intracranial aneurysm formation. Clin. Neurol. Neurosurg. 2010, 112, 306–313. [Google Scholar] [CrossRef]
  122. Velraeds, M.M.C.; Van Der Mei, H.C.; Reid, G.; Busscher, H.J. Inhibition of initial adhesion of uropathogenic Enterococcus faecalis to solid substrata by an adsorbed biosurfactant layer from Lactobacillus acidophilus. Urology 1997, 49, 790–794. [Google Scholar] [CrossRef]
  123. Mareels, G.; De Wachter, D.S.; Verdonck, P.R. Computational fluid dynamics-analysis of the Niagara hemodialysis catheter in a right heart model. Artif. Organs 2004, 28, 639–648. [Google Scholar] [CrossRef]
  124. Frumento, R.J.; Hirsh, A.L.; Parides, M.K.; Bennett-Guerrero, E. Differences in arterial and venous thromboelastography parameters: Potential roles of shear stress and oxygen content. J. Cardiothorac. Vasc. Anesth. 2002, 16, 551–554. [Google Scholar] [CrossRef]
  125. Nicoud, F.; Vernhet, H.; Dauzat, M. A numerical assessment of wall shear stress changes after endovascular stenting. J. Biomech. 2005, 38, 2019–2027. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  126. Morsi, Y.; Kogure, M.; Umezu, M. Relative blood damage index of the jellyfish valve and the Bjork-Shiley tilting-disk valve. J. Artif. Organs 1999, 2, 163–169. [Google Scholar] [CrossRef]
  127. Tran, V.B.; Fleiszig, S.M.J.; Evans, D.J.; Radke, C.J. Dynamics of flagellum- and pilus-mediated association of Pseudomonas aeruginosa with contact lens surfaces. Appl. Environ. Microbiol. 2011, 77, 3644–3652. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  128. Azevedo, A.S.; Almeida, C.; Gomes, L.C.; Ferreira, C.; Mergulhão, F.J.; Melo, L.F.; Azevedo, N.F. An in vitro model of catheter-associated urinary tract infections to investigate the role of uncommon bacteria on the Escherichia coli microbial consortium. Biochem. Eng. J. 2017, 118, 64–69. [Google Scholar] [CrossRef] [Green Version]
  129. Katsikogianni, M.G.; Missirlis, Y.F. Interactions of bacteria with specific biomaterial surface chemistries under flow conditions. Acta Biomater. 2010, 6, 1107–1118. [Google Scholar] [CrossRef]
  130. Vagos, M.R.; Moreira, J.M.R.; Soares, O.S.G.P.; Pereira, M.F.R.; Mergulhão, F.J. Incorporation of carbon nanotubes in polydimethylsiloxane to control Escherichia coli adhesion. Polym. Compos. 2019, 40, E1697–E1704. [Google Scholar] [CrossRef]
  131. Cringus-Fundeanu, I.; Luijten, J.; van der Mei, H.C.; Busscher, H.J.; Schouten, A.J. Synthesis and Characterization of Surface-Grafted Polyacrylamide Brushes and Their Inhibition of Microbial Adhesion. Langmuir 2007, 23, 5120–5126. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  132. Fundeanu, I.; van der Mei, H.C.; Schouten, A.J.; Busscher, H.J. Polyacrylamide brush coatings preventing microbial adhesion to silicone rubber. Colloids Surf. B Biointerfaces 2008, 64, 297–301. [Google Scholar] [CrossRef]
  133. Roosjen, A.; Kaper, H.J.; van der Mei, H.C.; Norde, W.; Busscher, H.J. Inhibition of adhesion of yeasts and bacteria by poly(ethylene oxide)-brushes on glass in a parallel plate flow chamber. Microbiology 2003, 149, 3239–3246. [Google Scholar] [CrossRef] [Green Version]
  134. Katsikogianni, M.; Amanatides, E.; Mataras, D.; Missirlis, Y.F. Staphylococcus epidermidis adhesion to He, He/O2 plasma treated PET films and aged materials: Contributions of surface free energy and shear rate. Colloids Surf. B Biointerfaces 2008, 65, 257–268. [Google Scholar] [CrossRef]
  135. Hendricks, S.K.; Kwok, C.; Shen, M.; Horbett, T.A.; Ratner, B.D.; Bryers, J.D. Plasma-deposited membranes for controlled release of antibiotic to prevent bacterial adhesion and biofilm formation. J. Biomed. Mater. Res. 2000, 50, 160–170. [Google Scholar] [CrossRef]
  136. Bayoudh, S.; Othmane, A.; Mora, L.; Ben Ouada, H. Assessing bacterial adhesion using DLVO and XDLVO theories and the jet impingement technique. Colloids Surf. B Biointerfaces 2009, 73, 1–9. [Google Scholar] [CrossRef]
  137. Barton, A.J.; Sagers, R.D.; Pitt, W.G. Bacterial adhesion to orthopedic implant polymers. J. Biomed. Mater. Res. 1996, 30, 403–410. [Google Scholar] [CrossRef]
  138. Millsap, K.; Reid, G.; van der Mei, H.C.; Busscher, H.J. Displacement of Enterococcus faecalis from hydrophobic and hydrophilic substrata by Lactobacillus and Streptococcus spp. as studied in a parallel plate flow chamber. Appl. Environ. Microbiol. 1994, 60, 1867–1874. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  139. Habash, M.B.; Mei, H.C.V.d.; Busscher, H.J.; Reid, G. The effect of water, ascorbic acid, and cranberry derived supplementation on human urine and uropathogen adhesion to silicone rubber. Can. J. Microbiol. 1999, 45, 691–694. [Google Scholar] [CrossRef]
  140. Andersen, T.E.; Kingshott, P.; Palarasah, Y.; Benter, M.; Alei, M.; Kolmos, H.J. A flow chamber assay for quantitative evaluation of bacterial surface colonization used to investigate the influence of temperature and surface hydrophilicity on the biofilm forming capacity of uropathogenic Escherichia coli. J. Microbiol. Methods 2010, 81, 135–140. [Google Scholar] [CrossRef] [PubMed]
  141. Gabi, M.; Hefermehl, L.; Lukic, D.; Zahn, R.; Vörös, J.; Eberli, D. Electrical microcurrent to prevent conditioning film and bacterial adhesion to urological stents. Urol. Res. 2011, 39, 81–88. [Google Scholar] [CrossRef]
  142. Neves, S.F.; Ponmozhi, J.; Mergulhão, F.J.; Campos, J.B.L.M.; Miranda, J.M. Cell adhesion in microchannel multiple constrictions—Evidence of mass transport limitations. Colloids Surf. B Biointerfaces 2020, 198, 111490. [Google Scholar] [CrossRef]
  143. Vanhommerig, E.; Moons, P.; Pirici, D.; Lammens, C.; Hernalsteens, J.-P.; De Greve, H.; Kumar-Singh, S.; Goossens, H.; Malhotra-Kumar, S. Comparison of Biofilm Formation between Major Clonal Lineages of Methicillin Resistant Staphylococcus aureus. PLoS ONE 2014, 9, e104561. [Google Scholar] [CrossRef] [PubMed]
  144. Tremblay, Y.D.; Vogeleer, P.; Jacques, M.; Harel, J. High-throughput microfluidic method to study biofilm formation and host-pathogen interactions in pathogenic Escherichia coli. Appl. Environ. 2015, 81, 2827–2840. [Google Scholar] [CrossRef] [Green Version]
  145. Goetz, C.; Tremblay, Y.D.N.; Lamarche, D.; Blondeau, A.; Gaudreau, A.M.; Labrie, J.; Malouin, F.; Jacques, M. Coagulase-negative staphylococci species affect biofilm formation of other coagulase-negative and coagulase-positive staphylococci. J. Dairy Sci. 2017, 100, 6454–6464. [Google Scholar] [CrossRef] [PubMed]
  146. Brackman, G.; Coenye, T. In Vitro and In Vivo Biofilm Wound Models and Their Application. Adv. Exp. Med. Biol. 2016, 897, 15–32. [Google Scholar]
  147. Goeres, D.M.; Walker, D.K.; Buckingham-Meyer, K.; Lorenz, L.; Summers, J.; Fritz, B.; Goveia, D.; Dickerman, G.; Schultz, J.; Parker, A.E. Development, standardization, and validation of a biofilm efficacy test: The single tube method. J. Microbiol. Methods 2019, 165, 105694. [Google Scholar] [CrossRef]
  148. Johnson, E.; Petersen, T.; Goeres, D.M. Characterizing the Shearing Stresses within the CDC Biofilm Reactor Using Computational Fluid Dynamics. Microorganisms 2021, 9, 1709. [Google Scholar] [CrossRef]
  149. Wu, B. Advances in the use of CFD to characterize, design and optimize bioenergy systems. Comput. Electron. Agric. 2013, 93, 195–208. [Google Scholar] [CrossRef]
  150. Benoit, M.R.; Conant, C.G.; Ionescu-Zanetti, C.; Schwartz, M.; Matin, A. New device for high-throughput viability screening of flow biofilms. Appl. Environ. Microbiol. 2010, 76, 4136–4142. [Google Scholar] [CrossRef] [Green Version]
  151. Straub, H.; Eberl, L.; Zinn, M.; Rossi, R.M.; Maniura-Weber, K.; Ren, Q. A microfluidic platform for in situ investigation of biofilm formation and its treatment under controlled conditions. J. Nanobiotechnol. 2020, 18, 166. [Google Scholar] [CrossRef]
  152. Zhang, X.Y.; Sun, K.; Abulimiti, A.; Xu, P.P.; Li, Z.Y. Microfluidic System for Observation of Bacterial Culture and Effects on Biofilm Formation at Microscale. Micromachines 2019, 10, 606. [Google Scholar] [CrossRef] [Green Version]
  153. Blanco-Cabra, N.; López-Martínez, M.J.; Arévalo-Jaimes, B.V.; Martin-Gómez, M.T.; Samitier, J.; Torrents, E. A new BiofilmChip device for testing biofilm formation and antibiotic susceptibility. NPJ Biofilms Microbiomes 2021, 7, 62. [Google Scholar] [CrossRef]
Figure 1. Modified Robbins device with the fluid behavior fully characterized by CFD: (a) schematic representation and (b) photograph. The system is mainly composed of a recirculating tank and one vertical semi-circular flow cell (about a meter high) with removable coupons, as well as peristaltic and centrifugal pumps.
Figure 1. Modified Robbins device with the fluid behavior fully characterized by CFD: (a) schematic representation and (b) photograph. The system is mainly composed of a recirculating tank and one vertical semi-circular flow cell (about a meter high) with removable coupons, as well as peristaltic and centrifugal pumps.
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Figure 2. Flow chamber system setup: (a) schematic representation and (b) photograph. The PPFC is coupled to a glass tank connected to four centrifugal pumps and a tubing system to conduct adhesion or biofilm formation assays.
Figure 2. Flow chamber system setup: (a) schematic representation and (b) photograph. The PPFC is coupled to a glass tank connected to four centrifugal pumps and a tubing system to conduct adhesion or biofilm formation assays.
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Figure 3. Scheme of a rotating disk reactor.
Figure 3. Scheme of a rotating disk reactor.
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Figure 4. Schematic diagram of a microfluidic setup.
Figure 4. Schematic diagram of a microfluidic setup.
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Figure 5. Shear stress magnitudes (Pa) on the bottom of 12-well microplates shaken at (a) 40 and (b) 180 rpm (orbital diameter of 25 mm, liquid volume of 3 mL, and temperature of 25 °C). Adapted from Gomes et al. [78].
Figure 5. Shear stress magnitudes (Pa) on the bottom of 12-well microplates shaken at (a) 40 and (b) 180 rpm (orbital diameter of 25 mm, liquid volume of 3 mL, and temperature of 25 °C). Adapted from Gomes et al. [78].
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Table 1. Main advantages and limitations of standard platforms for in vitro biofilm studies [36,38].
Table 1. Main advantages and limitations of standard platforms for in vitro biofilm studies [36,38].
Modified Robbins deviceLarge amount of biomass is producedComplex setup
High/moderate throughputEntry effects
Allows periodical samplingSampling can affect the biofilm
Can be run for very long periods without interventionLimited in situ biofilm visualization
Large dynamic rangeBiofilm destruction for most quantitative analysis
Flow chamberOptimized for online in situ microscopyLow throughput
Allows direct and nondestructive observation of biofilm developmentInability to study adhesion to nontransparent surfaces
Complex setup
Rotating biofilm devicesPossibility to study different materials in the same runThe flow pattern changes in the boundaries of the coupons
Shear stress and feed flow rate can be set independentlyLack of sampling surface area
Easy to control the operational conditionsComplex setup
MicroplatesHigh-throughput analysisDirect observation under the microscope can be difficult
Simple to runBatch system
Needs small spaceLoosely attached biofilm may not be correctly quantified
InexpensiveOperator dependent
Microfluidic devicesNoninvasive techniqueRequires special equipment for manufacturing and running systems
Allows real-time visualization of biofilm developmentClogging can occur due to small dimensions
Requires small volumes Laborious operation
Can be custom made for specific purposesAir bubbles may be an issue
Rapid and precise analysisViscosity effects may arise
Compatible with single-cell analysis
Table 3. Environmental studies performed on different biofilm platforms to evaluate the initial adhesion, biofilm formation, and treatment under defined shear conditions.
Table 3. Environmental studies performed on different biofilm platforms to evaluate the initial adhesion, biofilm formation, and treatment under defined shear conditions.
PlatformFieldBiofilm StageStudy AimHydrodynamicsAssay TimeSurface
OrganismsConcluding RemarksReferences
Modified Robbins deviceDrinking- water distribution systemsBiofilm formationInvestigate the combined impact of flow hydrodynamics and pipe material0.13 and 0.24 Pa100 daysPolyvinyl chloride
Structured wall high-density polyethylene
Solid wall high-density polyethylene
Natural flora present in drinking waterThe biomass amount was greater for the biofilms formed at lower shear stress.
The opportunistic pathogens have limited ability to propagate within biofilms under high shear conditions without protection (surface roughness).
Water treatmentBiofilm formationEvaluate the application of non-biocide release coatings as coated filters for biofouling preventionFlow rate of 300 L h−1, corresponding to an average shear stress of 0.25 Pa2 daysPolyurethane coating
Polyurethane coating with incorporated Econea
Polyurethane coating with grafted Econea
Enterococcus faecalisBiocidal polyurethane-based surfaces were less prone to biofilm formation, with an average reduction of 60%, compared to pristine polyurethane.[81]
Flow chamberMan-made equipment (heat exchangers, ship hulls, and pipelines) Biofilm formationStudy the influence of surface energy components on the adhesion and removal of fouling9.8 × 10−4, 4.6 × 10−4, and 2.1 × 10−4 Pa10 days316 L Stainless steel
Ni–P–TiO2–polytetrafluoroethylene nanocomposite coatings
Pseudomonas fluorescences
Cobetia marina
Vibrio alginolyticus
Coatings with the lowest ratio between the Lifshitz van der Waals apolar component and the electron donor component had the lowest bacterial adhesion or the highest bacterial removal.[85,86]
Rotating cylinder reactorDrinking- water distribution systemsBiofilm formation and treatmentEffect of chemical and mechanical stresses on single and dual- species biofilm removalBiofilm formation: 1 Pa
Treatment: 1–23 Pa
7 daysPolyvinyl chlorideAcinectobacter
Stenotrophomonas maltophilia
Dual species biofilms were the most susceptible to chemical and mechanical removal.
Stenotrophomonas maltophilia biofilms demonstrated high tolerance to chemical and mechanical stress.
Biofilm formationAction of copper materials on biofilm formation and control by chemical and mechanical stress0.1 Pa7 daysStainless steel
Copper alloys (100, 96, and 57%)
Stenotrophomonas maltophiliaChemical, mechanical, and combined shocks were not effective in biofilm control.
Copper surfaces were found to reduce the number of non-damaged cells.
6-well microplatesDrinking- water distribution systemsAdhesion and biofilm formationInfluence of shear stress, temperature, and inoculation concentration on water-stressed Helicobacter pylori0, 60, and 120 rpm corresponding to 0, 0.138, and 0.317 Pa2, 6, 12, 24, 48, 96, and 192 h304 stainless steel
Helicobacter pyloriHigh shear stresses negatively influenced the adhesion to the substrata.
However, the temperature and inoculation concentration appeared to not affect adhesion.
12-well microplatesMarine environmentBiofilm formationEffect of surface hydrophobicity on biofilm development by a filamentous cyanobacteriumOrbital shaking with a 25 mm diameter incubator at 185 rpm (average shear stress of 0.07 Pa) 3 weeksGlass
Leptolyngbya mycoidea LEGE 06118Higher biofilm growth was observed on perspex, the most hydrophobic surface.
Effect of different marine coatings on biofilm formation by microfoulersOrbital shaking with a 25 mm diameter incubator at 185 rpm (average shear rate of 40 s−1)7 weeks
Epoxy-coated glass
Silicone hydrogel coating
Cyanobium sp. LEGE 10375
Pseudoalteromonas tunicata (marine bacterium)
Epoxy-coated surface was effective in inhibiting biofilm formation at the initial stages, while silicone coating showed high antibiofilm efficacy during maturation.
Silicone coating was less prone to biofilm formation.
The efficacy of silicone may be dependent on the organism, while the performance of epoxy-coated surface was strongly influenced by a combined effect of surface and microorganism.
Effect of different materials on biofilm structure 7 weeksGlass
Epoxy-coated glass
Silicone hydrogel coating
Synechocystis salina LEGE 00041
Cyanobium sp. LEGE 06098
Cyanobium sp. LEGE 10375
Silicone coating was effective in inhibiting cyanobacterial biofilm formation.
Cyanobacterial biofilms formed on silicone coating showed a lower percentage and size of empty spaces among all surfaces.
Study the environmental compatibility of an innovative biocidal foul-release multifunctional coating 7 weeksPolydimethylsiloxane
Polydimethylsiloxane coating with grafted Econea
Pseudoalteromonas tunicataPolydimethylsiloxane coating with grafted Econea was more effective in inhibiting biofilm formation than the bare polydimethylsiloxane (reductions of 77%, 60%, and 73% on biovolume, thickness, and substratum coverage, respectively).
Long-lasting antifouling performances were observed in simulated and real scenarios.
Effect of shear forces on biofilm development by filamentous cyanobacteriaOrbital shaking with a 25 mm diameter incubator at 40 rpm (average shear rate of 4 s−1) and 185 rpm (average shear rate of 40 s−1)7 weeksGlass
Nodosilinea sp. LEGE 06020
Nodosilinea sp. LEGE 06022
Unidentified filamentous Synechococcales LEGE 07185
Biofilm formation was higher under low shear conditions.
The hydrodynamics was more effective on biofilm maturation than during initial cell adhesion.
Different shear rates affected biofilm architecture.
Effect of shear forces and surface hydrophobicity on biofilm development by coccoid cyanobacteria with different biofilm formation capacities 6 weeksGlass
Epoxy-coated glass
Synechocystis salina LEGE 00041
Cyanobium sp. LEGE 06097
Biofilms developed in both surfaces at lower shear conditions had a higher number of cells, wet weight, thickness, and chlorophyll a content.
The impact of hydrodynamics was generally stronger than the impact of surface hydrophobicity.
The antibiofilm performance of the polymeric coating was confirmed.
Qualitative proteomic analyses of filamentous cyanobacterial biofilms formed under different shear rates 7 weeks
Nodosilinea sp. LEGE 06145
Nodosilinea sp. LEGE 0611
Biofilm formation was higher under low shear conditions.
Biofilm development of Nodosilinea sp. LEGE 06145 was higher than LEGE 06119, but no significant differences were found between surfaces.
Adhesion and biofilm formationPotential of adhesion assays on the estimation of biofilm development behavior at different hydrodynamic conditions Adhesion:
7.5 h
Biofilm: 6 weeks
Epoxy-coated glass
Synechocystis salina LEGE 00041
Synechocystis salina LEGE 06155
Cyanobium sp. LEGE 06097
For both adhesion and biofilm assays, the number of adhered cells was higher under low shear conditions.
Higher biofilm development at 4 s−1 was confirmed by biofilm wet weight, thickness, and chlorophyll a content. Initial adhesion assays can be used to estimate marine biofilm development.
Quantitative proteomic analyses of biofilms formed on different surfaces 7 weeksGlass
Unidentified filamentous cyanobacterium LEGE 06007After 7 weeks, high biofilm thickness was observed in biofilms formed at 4 s−1 on glass when compared to perspex.
Differences in protein expression were more noticeable in biofilms formed under low shear conditions.
Proteomic analysis revealed differentially expressed proteins between surfaces.
Table 5. Industrial studies performed on different biofilm platforms to evaluate the initial adhesion, as well as biofilm formation and treatment under defined shear conditions.
Table 5. Industrial studies performed on different biofilm platforms to evaluate the initial adhesion, as well as biofilm formation and treatment under defined shear conditions.
PlatformBiofilm StageStudy AimHydrodynamicsAssay TimeSurface
OrganismsConcluding RemarksReferences
Modified Robbins deviceBiofilm formationEffect of flow rate/shear stress variation on mass transfer and biofilm development in a flow cell that mimics industrial pipingFlow rates of 374 and 242 L h−1, corresponding to wall shear stresses between 0.183 and 0.511 Pa9 daysPolyvinyl chlorideEscherichia coliBiofilm formation was favored at the lowest flow rate.
Shear stress effects were more important than mass transfer limitations.
This flow cell system generates wall shear stresses that are similar to those found in some industrial settings.
Biofilm formation and treatmentEvaluation of a modified diamond-like carbon surface for biofouling mitigation in critical process areasFlow rate of 300 L h−1, corresponding to an average shear stress of 0.25 PaBiofilm formation: 5 days
Treatment: 6, 18, and 24 h
316 L Stainless steel
Escherichia coli
Natural flora present in the water from an industrial salad washing line
Biofilm formation was reduced on SICON® (1–2 Log).
Biofilm cleaning with chlorine was more efficient when SICON® was used (3.5-Log reduction and 15% removal).
Industries with cleaning frequencies up to 6 h may benefit from the use of SICON®.
Biofilm formation and treatmentEvaluation of SICAN for biofouling mitigation in the food industry Biofilm formation: 5 days
Treatment: 6, 18 and 24 h
316 L Stainless Steel
Escherichia coli
Natural flora present in the water from an industrial salad washing line
Biofilm formation on SICAN and stainless steel were similar.
Processes with cleaning intervals of about 6 h could potentially use SICAN surfaces on critical areas.
AdhesionEffect of strain, shear stress, surface soiling, and growth conditions on Listeria monocytogenes adhesionFlow rates of 0.76 and 10.9 mL min−1, corresponding to wall shear stresses of 0.0505 and 0.7620 Pa30 minGlass
Polyvinyl chloride
Glass coated with beef extract, casein, and milk
Listeria monocytogenesStrain differences influenced the initial adhesion rate to all the surfaces at both low and high shear stress.
There was a significant effect of the surfaces on the adhesion ability of almost all strains.
The initial adhesion rate decreased at high shear stress for most strains.
Effect of flow direction and flow rate on the initial adhesion of Listeria monocytogenes strainsFlow rates of 0.75 and 8.40 mL min−1, corresponding to wall shear stresses of 0.10 and 1.20 Pa15 minFine polished stainless steelListeria monocytogenesInitial adhesion rates were influenced by flow rate and strain specificity.
The flow direction, in relation to the orientation of surface features, could be disregarded.
Biofilm formationEffect of surface conditioning on adhesion and biofilm formation under conditions that are prevalent in the food industryFlow rate of 11 mL s−1, corresponding to an average shear stress of 0.07 Pa24 hPolystyrene
Polystyrene conditioned with cell extracts and cell wall components
Escherichia coliUnder flow conditions, all conditioning films reduced biofilm formation, except mannose.
Surface conditioning affected the amount and clustering of bacteria on surfaces.
Rotating cylinder reactorBiofilm formation and treatmentEffect of shear stress on the formation and removal of biofilms0.02, 0.12, and 0.17 PaBiofilm formation: 7 days
Treatment: 0.5 h
AISI 316 Stainless steelBacillus cereusBiofilm density increased with the shear stress, while the thickness decreased.
The biocide treatment promoted the higher removal of biofilms formed under higher shear stress.
Biofilms formed under higher shear stress were more resistant to the mechanical and combined biocide and mechanical treatments.
Rotating disk reactorBiofilm formationEffect of shear stress on biofilm formationRotational speeds of 350 and 800 rpm, corresponding to shear stresses between 0 and 91 Pa4 daysAISI 304 2B food grade stainless steelCandida kruseiThe early development of a biofilm (24 h) was unaffected by shear stress.
In a mature biofilm, shear stress determined the disposition of biofilm cells onto the surface.
Biofilms formed under higher shear stress differ in their arrangement, as compared with those formed under lower shear conditions.
Assessment of the colonization of biofilms by free-living amoebaeShear rates between 31,000 and 85,000 s−1, representative of cooling circuits10 daysStainless steelFreshwater containing free-living amoebae and bacteriaFree-living amoebae were able to establish in biofilms under shear rate as high as 85,000 s−1.
The developed reactor seems to be ideal for studying the effects of high shear stress on surface colonization by microorganisms.
96-well microplatesAdhesionEffect of surface conditioning on adhesion and biofilm formation under conditions that are prevalent in the food industryOrbital shaking with a 50 mm diameter incubator at 150 rpm (average shear stress of 0.07 Pa) 1 hPolystyrene Polystyrene conditioned with cell extracts and cell wall componentsEscherichia coliTotal cell extract, cytoplasm with cellular debris, myristic, and palmitic acid decreased initial adhesion. Adhesion increased when periplasmic extract was used.
Adhesion was dependent on the conditioning film concentration.
6-well microplatesAdhesionEvaluation of the antiadhesive activity of SICON®Orbital shaking with a 25 mm diameter incubator (average shear stress of 0.25 Pa)0.5, 2, and 6 h316 L Stainless steel
Escherichia coli
Natural flora present in the water from an industrial salad washing line
Bacterial adhesion on SICON® and stainless steel were similar. [41]
AdhesionEvaluation of the antiadhesive activity of SICANOrbital shaking with a 25 mm diameter incubator (average shear stress of 0.25 Pa)0.5, 2, and 6 h316 L Stainless steel
Escherichia coli
Natural flora present in the water from an industrial salad washing line
Adhesion on SICAN and stainless steel were similar.
Escherichia coli and the flora from industrial water had similar adhesion behaviour.
Adhesion and biofilm formationAssessment of the impact of material properties, nutrient load, and shear stress on biofouling in food industriesStatic and orbital shaking with a 25 mm diameter incubator at 115 rpm (average shear stress of 0.27 Pa)Adhesion: 0.5 h
Biofilm: 6 h
Stainless steel
Escherichia coliSurface material was the most important factor in adhesion and biofilm formation.
Adhesion and biofilm formation were correlated with surface hydrophobicity.
The effect of surface properties was dependent on the nutrient load and shear stress.
Initial adhesion performance may be a good predictor for biofilm formation.
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Gomes, L.C.; Mergulhão, F.J.M. A Selection of Platforms to Evaluate Surface Adhesion and Biofilm Formation in Controlled Hydrodynamic Conditions. Microorganisms 2021, 9, 1993.

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Gomes LC, Mergulhão FJM. A Selection of Platforms to Evaluate Surface Adhesion and Biofilm Formation in Controlled Hydrodynamic Conditions. Microorganisms. 2021; 9(9):1993.

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Gomes, Luciana C., and Filipe J. M. Mergulhão. 2021. "A Selection of Platforms to Evaluate Surface Adhesion and Biofilm Formation in Controlled Hydrodynamic Conditions" Microorganisms 9, no. 9: 1993.

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