Understanding the mechanism of biofilm distribution and detachment is very important to effectively improve water treatment and prevent blockage in porous media. The existing research is more related to the local biofilm evolving around one or few microposts and the lack of the integral biofilm evolution in a micropost array for a longer growth period. This study combines microfluidic experiments and mathematical simulations to study the distribution and detachment of biofilm in porous media. Microfluidic chips with an array of microposts with different sizes are designed to simulate the physical pore structure of soil. The research shows that the initial formation and distribution of biofilm are influenced by bacterial transport velocity gradients within the pore space. Bacteria prefer to aggregate areas with smaller microposts, leading to the development of biofilm in those regions. Consequently, impermeable blockage structures form in this area. By analyzing experimental images of biofilm structures at the later stages, as well as coupling fluid flow and porous medium, and the finite element simulation, we find that the biofilm detachment is correlated with the morphology and permeability (kb) (from 10−15 to 10−9 m2) of the biofilm. The simulations show that there are two modes of biofilm detachment, such as internal detachment and external erosion.

  • The distribution of biofilms is related to the structure of porous medium.

  • This study combines the microfluidic experiment and mathematical simulation to study the distribution and detachment of biofilm in porous media.

  • The biofilm detaching is related to the morphology and permeability.

  • Two modes of biofilm detachment are internal detachment and external erosion.

In nature, biofilm is a community of bacteria that can be either superficially attached or free-floating. The bacteria are held together by extracellular polymeric substances secreted by the bacteria (Nadell et al. 2008; Flemming et al. 2016). Bacteria can survive almost anywhere (Conrad & Poling-Skutvik 2018), and soil is the most complex component of the earth's ecosystem and is suitable for microbial growth. Therefore, it is necessary to understand the formation process and characteristics of biofilm communities in soil.

The porous structure of soil affects the spatial distribution and colony behavior of bacteria (Raynaud & Nunan 2014; Coyte et al. 2016). The formation of biofilms in porous media has been studied by many researchers. Scheidweiler et al. (2019) found that biofilms differentiate into an annular base biofilm coating the microcolumns and into streamers protruding from the microcolumns into the pore space. The shape and distribution of streamers are influenced by multiple factors, such as flow velocity (Valiei et al. 2012), channel size, and flow direction (Marty et al. 2012). In addition, Hassanpourfard et al. (2015) show that bacterial flocs (i.e. bacterial aggregates encapsulated in extracellular polymeric substances) can lead to the formation of streamers through large deformation processes. The secondary flow formed by tortuous channels promotes the development of streamers (Marty et al. 2014). Under certain flow conditions, the biofilm attached to the porous medium forms a network of streamers that capture the separated cells, resulting in uneven distribution and even clogging of the biofilm (Drescher et al. 2013). This heterogeneity causes biofilm separation by increasing local fluid shear forces (Karimifard et al. 2021). Hassanpourfard et al. (2016) observed that the clogged bacterial biomass exhibited an instability phenomenon marked by localized streamer breakage and failure leading to the formation of extended water channels, but the origin of this mode of failure is not yet fully understood.

The finite element method is widely used to provide better understanding and explanation of biofilm growth in fluid environments. Traditionally, biofilms have been studied as impermeable domains. In such models, no water can enter biofilms, and contaminants can only enter via molecular diffusion (Bottero et al. 2013; Peszynska et al. 2016). Benioug et al. (2017) studied the biofilm flow in porous media using the Lattice–Boltzmann method and simulated biofilm growth. However, other researchers believe that this assumption is incorrect, and experimental work has shown that biofilms have uneven morphology, including voids and channels, and there are both flowing and stagnant water in biofilms (Lewandowski 2000), so biofilms are permeable. Some researchers have considered the effect of biofilm permeability on the flow of porous media. Deng et al. (2013) studied the effect of permeable biofilm on the flow of porous media and developed a model to predict the overall permeability based on the biofilm permeability and biofilm volume ratio (i.e. the ratio of biofilm volume to preinoculation pore-space volume). The model proves that biofilm permeability affects the shear stress distribution. Even though much work has been done on biofilms in porous media, most of them only focus on the structure of biofilms. Microfluidic platforms have not been used to study how different pore sizes affect the spatial distribution and detachment of biofilms.

In this paper, we used microfluidic platforms for research. Microfluidic platforms enabled us to have better control over the flow of the bacterial solution. We used the microscope to observe the solution for a long time. These devices were disposable and prevented cross-contamination during the experiment. Most importantly, researchers can easily change the section shape and various sizes of the channel to meet the experimental requirements (Rusconi et al. 2011). More and more researchers used this method to explore the dynamic changes of biofilm. In this work, we designed and manufactured microfluidic chip embedded with a micropost array, and the pores between the microposts simulate the interval among soil particles. Our devices made us observe the movement of bacteria in a porous structure that mimics the soil topography. Bacillus subtilis was cultured in these microfluidic channels at different volume flow rates. We described the evolution of the initial spatial distribution of biofilms in porous media and related it to hydrodynamic factors. We found the bacterial transport velocity gradients across pore space have influence on the initial biofilm formation. However, as the biofilm grew, the channels were gradually blocked, and the biofilm was locally unstable and detached. Considering permeability and porosity, we conducted computational fluid dynamics simulations by coupling the Navier–Stokes equations with Brinkman equation and proposed a mode of biofilm detachment.

Biofilm growth

Bacillus subtilis can be isolated from various environments (soil, marine habitat, etc.) (Yan et al. 2016; Yahya et al. 2021) and has become a model bacterium for studying biofilm formation (Lemon et al. 2008). So, we used Bacillus subtilis 3610 in our experiments, as it is one of the most studied (Gram-positive bacteria) and most widely used to study biofilm growth (Dervaux et al. 2014; Douarche et al. 2015; Gingichashvili et al. 2022). We cultured biofilms by feeding the minimal-salts-glutamate-glycerol (MSgg) solution at OD600 = 0.4 with a flow rate of Q = 10 and 25 μL h−1 in microfluidic channels. We cultured at a flow rate of Q = 10 μL h−1 for 25 h; on this basis, culture was carried out at a flow rate of Q = 25 μL h−1. MSgg is composed of 5 mM potassium phosphate (pH 7), 100 mM 4-morpholinepropanesulfonic acid (MOPS) (pH 7), 2 mM MgCl2, 700 μm CaCl2, 50 μm MnCl2, 50 μm FeCl2, 1 μm ZnCl2, 2 μm thiamine, 0.5% glycerol, 0.5% glutamate, 50 μg/ml tryptophan, 50 μg/ml phenylalanine, and 50 μg/ml threonine. The MSgg used in the experiment had been sterilized. All experiments were repeated five times.

Microfluidic experiments

The structure of the porous media in the actual soil was very complex, with non-uniformity and uniformity, which made it difficult to carry out accurate simulation. Therefore, we set the disordered porous media as microposts with certain regular arrangement. In this study, we fabricated a polydimethylsiloxane microfluidic device using standard soft lithography technology and designed two types of channels. Two types of micromodel pore network structures are used in this study: microposts of non-uniform distribution and microposts of uniform distribution. Polydimethylsiloxane (PDMS) was cast (on a reticle master obtained by photolithography of a SU8 photoresist) over a silicon wafer and then bonded to a glass slide. The microfluidic device was disinfected using 70% ethanol for 5 min, followed by rinsing with MSgg solution to remove any ethanol left behind. The final microfluidic channel had dimensions of width = 600 μm, height = 20 μm (Figure 1). For this study, the flow direction, width and height correspond to the X, Y and Z axes, respectively. We tracked the biofilm formation process in channels using a wide-field fluorescent stereoscopic microscope.
Figure 1

(a) A schematic of experiment set-up under pressure-driven flow with constant volume flow rate (Q). (b) Layout of staggered pattern porous media. The channel width (W) is 600 μm. The distance between the center of posts (L2) is 42 μm and two rows of consecutive posts (L1) is 50 μm. The diameter of the pillars (d) is 16–36 μm, and the height of the device is 20 μm. (c) The diameter of the posts (d) is 32 μm, and the other dimensions are consistent with Figure 1( image (b).

Figure 1

(a) A schematic of experiment set-up under pressure-driven flow with constant volume flow rate (Q). (b) Layout of staggered pattern porous media. The channel width (W) is 600 μm. The distance between the center of posts (L2) is 42 μm and two rows of consecutive posts (L1) is 50 μm. The diameter of the pillars (d) is 16–36 μm, and the height of the device is 20 μm. (c) The diameter of the posts (d) is 32 μm, and the other dimensions are consistent with Figure 1( image (b).

Close modal

COMSOL simulation

We used the COMSOL software to carry out related simulation in fluid flow and used CAD software to draw the two-dimensional shape of porous media. Fluid space was set to have the material properties of water, with the inlet having a velocity boundary condition of fully developed flow and the outlet having a pressure boundary condition of zero Pa. The mesh used was a free triangular mesh.

In the simulation, we employed the incompressible form of the Navier–Stokes and continuity equations. The Navier–Stokes equation is a mathematical equation used to describe fluid motion and can be regarded as Newton's second law of fluid motion. For compressible Newtonian fluids, one can obtain:
(1)
where u is the velocity, p is the pressure, ρ is the density, and μ is the viscosity. The terms in the formula correspond to the inertial force, pressure, viscous force, and the external force acting on the fluid.
In our experiments, fluid flows from left to right through the entire geometry with the velocity less than 10−3 m/s, and then the maximum Reynolds number is less than 0.01. Because there is no external force (gravity is ignored), the force term is also equal to zero. From this, the Navier–Stokes equation can be simplified as:
(2)
At the inlet, we consider several parameters including the average flow velocity Um, the constant volume flow rate Q, the cross-sectional area A, the liquid density , the liquid viscosity , the Reynolds number Re, and the channel hydraulic diameter Dh, the width is W, the height is H, the perimeter is L (Wang et al. 2016):
(3)
In our study, since (low Reynolds number) Stokes flow prevails and the fluid is Newtonian, shear stress is linearly related to shear rate (Wang et al. 2016):
(4)
In this study, bacteria in the suspension were represented by suspended particles. Therefore, the density of particles was selected as 1,080 kg/m3 and the diameter of particles was set as 1 μm, which was close to the density of the fluid to achieve the effect of suspension. In the initial process, because bacteria were relatively less adhered to the micropost, the particles did not have enough inertia to significantly interfere with the mobile phase, so the influence of particles on the continuous phase could be ignored. There was no need to consider the reaction force of particles on the fluid and the interaction between particles. In this simulation, it was unidirectional coupling. Therefore, only the effect of fluid drag on the particles needs to be considered. Drag force was the main force of particle motion. When particles were moved in a viscous fluid, there was a velocity difference between the particles and the fluid medium, and the resistance of the fluid acting on the particles was called fluid drag force. The drag force model selected was Schiller–Naumann model, and the fluid drag force of this model can be expressed as:
(5)
When Reynolds number Re << 0.1, CD can be expressed as:
(6)

FD represents the fluid drag force on the particle, CD represents the drag coefficient of the particle, and dp represents the diameter of the particle.

The particles enter at the same time as the water flows, and the particles were released at the inlet boundary. The release time was set at 0.05 s, and 200 particles were released each time. In the experiment, the number of bacteria is not easy to be quantified. The number of simulated particles does not correspond to the number of bacteria but only qualitatively studies the movement law of bacteria, and we simulated a different number of particles and found that their motion laws were consistent.

In addition, we did a different flow simulation to study the detaching of biofilms, and the flow in pore space was coupled with the flow within biofilm (green in Figure 7) using the Brinkman equations:
(7)
(8)
where is the biofilm porosity, is the biofilm permeability (m2), is the Darcy flux vector, and is the Forchheimer coefficient that is defined as follows:
(9)
where is the dimensionless friction coefficient and is calculated as follows:
(10)

Distribution of biofilms

Our microfluidic device consisted of a sequence of PDMS microposts in a periodic staggered grid pattern (Figure 1(b) and 1(c)). Using the syringe pump, the bacterial suspension was injected into the device with a constant volume flow rate of the fluid (Q) for each experiment. A few minutes after the injection, biofilms could be observed adhering to the microposts in the inlet, regardless of whether the microposts were non-uniformly or uniformly distributed (Figure 2(a) and 2(c)).
Figure 2

Time-lapse microscopic images of microfluidic channels with the Bacillus subtilis MSgg solution flow in two-type devices. (a) 5 min in the non-uniform device. (b) 2 h in the non-uniform device. (c) 5 min in the uniform device. (d) 2 h in the uniform device (scale bars:100 μm).

Figure 2

Time-lapse microscopic images of microfluidic channels with the Bacillus subtilis MSgg solution flow in two-type devices. (a) 5 min in the non-uniform device. (b) 2 h in the non-uniform device. (c) 5 min in the uniform device. (d) 2 h in the uniform device (scale bars:100 μm).

Close modal

As the experiment progressed, the biofilm continued to grow and formed large clusters with compact and dense structures that blocked the pores in the inlet (Figure 2(b) and 2(d)) (Xiao et al. 2020; Ke et al. 2021). Although the biofilm adhesion in both devices occurred for 5 min, the distribution was different. The initial formation of biofilm in the non-uniformly distributed micropost was mainly located in the region with small microposts (Supplementary Movie S1). On the contrary, only a few biofilms formed in the region with bigger microposts, showing an asymmetric distribution. However, the initial formation of biofilm in uniformly distributed microposts was more uniform in space (Supplementary Movie S2).

To explore the biofilm distribution in two types of devices in our experiment, we obtained the fluid flow velocity distributions in two kinds of devices through the finite element simulation (Figure 3). Before the biofilm blockage, the micropost distribution determined the flow velocity field. The velocity field in the uniform micropost structure is uniform, while the velocity field in the inhomogeneous micropost structure was not uniform. The inhomogeneity was represented by the higher flow velocity in the local area, which formed the dominant channels with higher nutrient supply, thereby facilitating the biofilm formation (Ke et al. 2021).
Figure 3

Simulation of velocity field in different devices. (a) Microposts of non-uniform distribution. (b) Microposts of uniform distribution. The color bar represents the flow field velocity.

Figure 3

Simulation of velocity field in different devices. (a) Microposts of non-uniform distribution. (b) Microposts of uniform distribution. The color bar represents the flow field velocity.

Close modal
To further explore the influence of velocity field on the biofilm distribution, we randomly released solid particles of the same size at the entrance of the flow field and found that similar results were produced (Figure 4), and the color represented the velocity of the particle. In the non-uniform micropost device, due to the uneven diameter of the micropost, most of the particles moved to the area with small diameter of the micropost at beginning and then gradually a small number of particles appeared in the area with bigger microposts. In contrast, in the device of uniform microposts, the particles were distributed symmetrically and diffused toward the outlet. The simulation results were consistent with experimental observation.
Figure 4

Simulated particle velocity distribution in the two-type devices. The color bar represents the particle velocity. (a) t = 0.25 s in the non-uniform device. (b) t = 0.55 s in the non-uniform device. (c) t = 0.15 s in the uniform device. (d) t = 0.5 s in the uniform device.

Figure 4

Simulated particle velocity distribution in the two-type devices. The color bar represents the particle velocity. (a) t = 0.25 s in the non-uniform device. (b) t = 0.55 s in the non-uniform device. (c) t = 0.15 s in the uniform device. (d) t = 0.5 s in the uniform device.

Close modal

Detachment of biofilms

With the culture time, we found the biofilm blocked the channel, similar to the findings of Drescher et al. (2013), which can make disastrous consequences. Meanwhile, we also observed that the blockage elimination induces by the localized failure of biofilms. To find the reason of biofilm detachment, taking the biofilm in uniform micropost device as an example, two typical biofilm morphologies were selected (Figures 5 and 6). The front micropost column of the channel was completely covered by biofilm (Figure 5), and the selected section including the biofilm-covered micropost column was named morphology 1 and showed the morphology 1 evolution containing the biofilm detachment (Figure 5(a)–5(d)).
Figure 5

Time-lapse microscopic image of morphology 1. The red arrow shows the inlet direction. Blue areas are biofilm detaching areas. Here t0 is 60 min after the imposed flow velocity of 25 μL/h.

Figure 5

Time-lapse microscopic image of morphology 1. The red arrow shows the inlet direction. Blue areas are biofilm detaching areas. Here t0 is 60 min after the imposed flow velocity of 25 μL/h.

Close modal
Figure 6

Time-lapse microscopic image of morphology 2. The red arrow shows the inlet direction. Blue areas are biofilm detaching areas. The channel referred to by the white arrow is the area within the device that is not covered by biofilms. Here t0 is 11 h after the imposed flow velocity of 25 μL/h.

Figure 6

Time-lapse microscopic image of morphology 2. The red arrow shows the inlet direction. Blue areas are biofilm detaching areas. The channel referred to by the white arrow is the area within the device that is not covered by biofilms. Here t0 is 11 h after the imposed flow velocity of 25 μL/h.

Close modal
Figure 7

Experiment and schematic of two biofilm morphologies. The red area represents the nutrient solution flowing in the pores. Green areas represent biofilms. The gray area represents the microposts set-up in the experiment (scale bars: 50 μm).

Figure 7

Experiment and schematic of two biofilm morphologies. The red area represents the nutrient solution flowing in the pores. Green areas represent biofilms. The gray area represents the microposts set-up in the experiment (scale bars: 50 μm).

Close modal

The front micropost column of the channel was not completely covered by biofilm (Figure 6), and the selected section including the biofilm not completely covered micropost column was named as morphology 2 and showed the morphology 2 evolution containing the biofilm detachment (Figure 6(a)–6(d)).

To further analyze the mechanism of biofilm detachment, we used finite element simulation to obtain the flow velocity field and shear stress distribution in morphology 1 and morphology 2, which had the same size of 380 μm × 600 μm (Figure 7). The channel width was 600 μm, and the segment along the channel length was 380 μm, as the formation of biofilm was a dynamic process and the growth of biofilm followed the same evolutionary trend. The selected section with a length of 380 μm represents the biofilm dynamics along the whole channel.

This paper assumed that the thickness of biofilm was uniform and did not consider the effect of thickness. In the simulation, the porosity of biofilm was set to be 0.4, which was proven to have no effect on the distribution of flow field and shear stress field (Deng et al. 2013). The biofilm morphology and permeability were considered, and biofilm permeability ranged from 1 × 10−15 to 1 × 10−9 m2 and showed the flow velocity field under different biofilm morphologies and different permeabilities (Figure 8).
Figure 8

Images show the normalized velocity by the maximum flow velocity.

Figure 8

Images show the normalized velocity by the maximum flow velocity.

Close modal

As for biofilms in morphology 1, at lower biofilm permeability, Kb = 1 × 10−15 m2, the fluid flows through the biofilm very slowly, while the flow velocity was large in the area without the biofilm, that is because the lower biofilm permeability induces high pressure differences in the interface between the biofilm and the fluid (Figure 8(d)).

When Kb = 1 × 10−12 m2, the flow becomes easy to permeate through the biofilm; however, the flow velocity of the fluid in the rear was smaller and the flow path did not change significantly, compared to the situation with Kb = 1 × 10−15 m2 (Figure 8(c)).

When Kb = 1 × 10−9 m2, the flow becomes much easier permeating the biofilm, and the flow path was significantly altered and the fluid velocity is largely reduced, resulting in almost no velocity gradient (Figure 8(b)).

As for biofilms in morphology 2, at lower biofilm permeability, Kb = 1 × 10−15 m2, a clear water fluid flows through the preferential path emerged, there was an obvious velocity gradient, and the water fluid flow barely conducted in the biofilm because the liquid could hardly permeate through the thick biofilm (Figure 8(h)).

When Kb = 1 × 10−12 m2, the flow path was not significantly changed, but the velocity was relatively reduced (Figure 8(g)).

When Kb = 1 × 10−9 m2, the flow path was relatively unchanged, the flow velocity decreased further, and the flow also proceeded in the biofilm region (Figure 8(f)).

Shear stress was very important for biofilm detachment, which was induced when the shear stress is larger than the strength of the biofilm and showed the shear stress under different biofilm morphologies and different permeabilities (Figure 9).
Figure 9

Images show the normalized shear stress by the maximum shear stress.

Figure 9

Images show the normalized shear stress by the maximum shear stress.

Close modal

As for biofilms in morphology 1, at lower biofilm permeability, Kb = 1 × 10−15 m2, biofilms were virtually impermeable but cause higher shear stress (maximum = 5.88 Pa) inside biofilms relative to that at the liquid biofilm interface, detaching was more likely to occur as an internal detach than as an external erosion (Figure 9(d)).

When Kb = 1 × 10−12 m2, fluid flow was still not easily permeable to biofilms, at which point the shear stress inside the biofilm was not obviously different from that at the liquid biofilm interface, which can proceed either as an internal detach or as an external erosion (Figure 9(c)).

When Kb = 1 × 10−9 m2, the fluid flow was easily permeable to the biofilm, and the shear stress inside the biofilm was similar to that at the liquid biofilm interface, resulting in both the internal detach and interfacial erosion (Figure 9(b)).

As for biofilms in morphology 2, at lower biofilm permeability, Kb = 1 × 10−15 m2, a clear shear stress gradient was found, with a large shear stress at the liquid biofilm interface (maximum = 4.14 Pa) than that at the interior of the biofilm, at which point the biofilm was more likely to detach in the form of an external erosion (Figure 9(b)).

When the permeability increased to 1 × 10−12 and 1 × 10−9 m2, a shear stress gradient was also found, shear stress at the liquid biofilm interface was still greater than that at the interior of the biofilm (Figure 9(f)–9(g)), so it was more likely that biofilms undergo detaching in the form of external erosion.

In this work, we studied the biofilm formation and detachment dynamics in porous structed microfluidic channels. Combined with experimental observation and mathematical modeling, we obtained the following conclusions:

  • It showed that the bacterial transport velocity gradients across pore space influenced the initial biofilm growth, and the biofilm clogging preferentially occurred in the area with small microposts. Previous studies have also shown that rapid bacterial aggregation occurs at the pore throat (i.e. the region of channel contraction and expansion), partly due to the large velocity gradient (Lee et al. 2023).

  • The study showed that the detaching of biofilms was related to its morphology and permeability (Karimifard et al. 2021; Wang et al. 2022), which cause biofilm detaching through shear stress with a value of about 5 Pa. In our previous study, about the biofilm growth in the microfluidic channel with a single micropost inside (Liu et al. 2022), we found that the shear stress threshold that is suitable for the biofilm adhesion was 0.3 Pa and proposed that there are two detaching modes of biofilm: internal detachment and external erosion.

The authors would like to thank Professor David A. Weitz and Professor Shmuel Rubinstein from Harvard University for their experimental support. The authors would like to thank the the National Natural Science Foundation of China for funding support (12372321, 11972074, 11772047 and 11620101001).

Y.T., C.T., Z.Z., S.L., F.D., D.Z., J.Z. and X.W. contributed to the study conception and design.

All relevant data are included in the paper or its Supplementary Information.

The authors declare there is no conflict.

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