Latest revision as of 08:43, 13 December 2020

Requirements of the Biofilm Displaying our Enzymes Improvement of Biofilm Formation Obviation of Sporulation Flow Chamber Atomic Force Microscopy Small Molecule Sorption

Requirements of the Biofilm

In order to implement a Bacillus subtilis biofilm to render micropollutants in wastewater less toxic, certain requirements have to be fulfilled. First of all, the enzymes we are using have to be exported into the extracellular matrix to protect our bacteria from the toxicity of the substances. We also have to avoid those enzymes from getting washed away by the water to make sure they can convert the micropollutants. Furthermore, our biofilm as a whole has to absorb the substances to enable their enzymatic degradation. Since we have to prevent genetically modified organism (GMO) release from the biofilm into the environment our biofilm needs to resist mechanical pressure to which it is exposed in wastewater treatment plants.

Biofilm Engineering

Displaying our Degradation Enzymes in the Biofilm Matrix

The Major Biofilm Matrix Component (TasA)

The Bacillus subtilis biofilm matrix consists mainly of exopolysaccharides (EPS) and proteins. One protein essential for the structure and formation of a biofilm is the major biofilm matrix component (TasA)^[1]. It polymerizes into long amyloid-like fibres which are attached to the cell wall by the TasA anchoring protein (TapA) outside of the cell^[2]. The knockout of tasA leads to a mutant that forms a weak biofilm of decreased thickness^[3]. B. subtilis secretes proteins in order to enable intercellular linkages and communication and therefore uses the Sec-dependent signal recognition particle (Sec-SRP) pathway. For proteins to be secreted via the SPR pathway, it is necessary that the proteins possess a 27 amino acids long N-terminal signal peptide, which is cleaved off during the membrane translocation. Following this scheme, TasA also possesses this N-terminal signaling peptide^[4,^5]. The Sec-SRP pathway consists of four main steps:

The signal peptide is recognized by a ribonucleoprotein complex, the signal recognition particle (SRP). 
SRP targets the Sec translocase which transports the protein through the membrane.
During membrane translocation the signal peptide is cleaved off by the SipW peptidase leading to the release of the protein.
Chaperones mediate the correct folding of the protein outside the cell^[5].

A Fusion Protein of TasA with our Degradation Enzymes

We want to immobilize our degradation enzymes in the biofilm matrix, due to the improved stability and improved enzyme activity of immobilized enzymes^[6]. Furthermore, within the biofilm matrix the degradation targets are better accessible for enzymatic degradation than within the cytoplasm. Therefore, we decided to generate fusion proteins with our degradation enzymes and the protein TasA, following previous work by Huang et al.^[7] (Figure 1).

**Figure 1:** Schematic illustration of the gene of our conceptualized fusion proteins. The 3' end of the *tasA* gene is fused to the gene of our degradation enzyme with a gene fragment encoding a glycine-serine-rich linker.

They showed that fusion proteins consisting of TasA and a second protein domain (e.g. mCherry, SpyTag) are successfully exported into the biofilm matrix (Figure 2). Even a fusion protein of TasA with the larger enzyme MHETase (63.1 kDa) was localized in the biofilm matrix of B. subtilis. Based on the research of Huang et al., we planned the fusion protein of superfolder green fluorescent protein (sfGFP) with TasA as a proof of concept. The sequence of the sfGFP gene is codon optimized for B. subtilis and fused to the 3' end of the tasA gene with a gene fragment encoding a glycine-serine-rich linker (ARGGGGSGGGGS). The display of the TasA-sfGFP fusion protein in the biofilm matrix can be verified using fluorescence microscopy. If TasA-sfGFP is successfully expressed in the biofilm matrix, we hypothesize that analogously designed TasA fusion proteins with our targeted degradation enzymes CotA, CueO and EreB will likely succeed.

**Figure 2:** The degradation enzymes are displayed in the biofilm matrix as fusion proteins with the biofilm matrix component TasA.

Integration of the TasA Fusion Proteins in the B. subtilis Genome

We want to ensure that only TasA fusion proteins, but no endogenous TasA, are secreted into the matrix in order to increase the number of immobilized degradation enzymes in the biofilm matrix. Therefore, we use a B. subtilis TasA knockout strain supplied by the group of Prof. Stülke (Georg-August-Universität Göttingen)^[8]. Then, as a proof of concept, the TasA fusion protein will be introduced in B. subtilis via plasmids (see Huang et al.). In the next step we want to integrate the TasA fusion proteins into the genome of our B. subtilis strain.

For further information on the workflow and analysis of our concept, please refer to our text on Engineering Success.

Improvement of Biofilm Formation

The matrix protein TasA is encoded in the tapA-sipW-tasA operon which is regulated by the repressing transcription factor SinR^[1]. SinR also regulates exopolysaccharide synthesis by controlling the expression of the epsA-O operon^[9].
Consequently, the genomic deletion of sinR improves the biofilm formation of B. subtilis, because involved genes are expressed without repressing effects by SinR (Figure 3)^[10]. In addition, sinR knockout mutants have been found to show nonmotile phenotypes and thus are not able to disperse from the biofilm^[11]. We are therefore planning the knockout of the sinR gene in our B. subtilis strain.

**Figure 3:** The repressing transcription factor *sinR* regulates the expression of the matrix protein TasA. The absence of *sinR* leads to the expression of TasA and therefore the improvment of biofilm formation.

Additionally, we hypothesize that the overexpression of tasA or respectively of the tapA-sipW-tasA operon could improve biofilm formation. To our knowledge no direct attempts are published. However, Lei et al. overexpressed the small protein Veg which led to a highly increased expression of tasA^[12]. Although the authors are strongly suggesting that Veg negatively regulates the expression of sinR, we cannot assume that the overexpression of Veg will additionally improve stability after sinR knockout as Veg appears to mainly affect sinR expression and does not affect the biofilm stability via another mechanism. For investigating the mechanism of action, a comparison of a veg overexpressing strain with a veg overexpressing strain and simultaneous sinR knockout is necessary.
Overall, the approach of overexpressing veg sounds promising to gain a strain with high stability.
For comparison of the various strains we would use our flow chamber specifically designed for testing mechanical stability of biofilms.

Obviation of Sporulation

Bacillus subtilis is able to form endospores. During B. subtilis biofilm maturation, cells can sporulate and leave the biofilm which could cause an escape of our genetically modified organisms into the environment^[13]. Since endospores are physiologically inactive, they do not express enzymes and thus do not contribute to micropollutant degradation. For these reasons, we aim to prevent any sporulation in the biofilm population.
The sigma factor F (σ^F) plays a critical role in the sporulation of B. subtilis by controlling several required genes^[14]. The absence of σ^F renders B. subtilis unable to sporulate (Figure 4), which is why we want to knockout the sigF gene in the genome of our B. subtilis^[15].

**Figure 4:** The knockout of *sinR* improves biofilm formation while the knockout of *sigmaF* prevents sporulation.

Testing

Flow Chamber

It is essential that our biofilm is stable enough to withstand the rough conditions that prevail in a wastewater treatment plant (WWTP), otherwise GMOs are likely to escape into the environment. Therefore, we sought for a method to measure the stability of biofilms. In order to simulate the most extreme conditions prevailing in WWTPs, we decided to design a custom-made flow chamber consisting of three parts which are stacked on top of each other.
We printed the 3D models using a Creality Cr10V2 3D printer. The upper part contains two holes the for the influx and efflux water hoses. The bottom part is used to grow a biofilm on. The middle part is designed to direct the liquid to flow over the biofilm in a thin layer, simulating the most extreme conditions in a WWTP. Two clamps are used to compress the stacked parts and prevent leakage. We estimate that the roughness of the used printing material (ecoPLA filament by 3DJAKE) enables to form a biofilm on the carrier surface. The flow chamber is assembled as you can see in our video and our manual.

The goal of this experiment is analysing the amount of living cells which are washed off the biofilm at different flow velocities. We chose phosphate-buffered saline (PBS) as liquid since it has physiological osmolarity and ion concentrations. The test is conducted using a PBS reservoir of around 50 mL which is pumped circularly through the flow chamber by a peristaltic pump (Cole-Parmer Masterflex L/S Model 7523-60 Digital Peristaltic Pump with Easy-Load II Head).
After flowing PBS with a defined flow rate over the biofilm for 3 hours, the flow chamber is deconstructed, and the used PBS buffer is collected in a falcon tube and centrifuged at 6000 xg for at least 6 min. Subsequently, the supernatant is discarded and the pellet is resuspended in 2 mL lysogeny broth (LB) media. These first samples are later used to quantify the number of cells that have been washed out the biofilm. In addition, multiple areas of defined size are scrapped of the biofilm, each area separately resuspended in 2 mL LB media. These second samples are the remaining cells per mm² to which we can compare the amount of washed-out living cells. As controls are essential, we simultaneously conduct the assay without a biofilm and with a tasA deletion strain of Bacillus subtilis which misses the major protein component of biofilm matrices and is thus assumed instable biofilm ^[3].
Quantifying the number of cells and cells within a biofilm matrix in our samples is not trivial. Although we found a method based on flow cytometry ^[16], we sought for an alternative as many laboratories worldwide do not have access to flow cytometry. As biofilms fluctuate in their density and thickness and thus in their cell number, quantifying the ratio and not the total numbers of washed-out living cells appears to be a reasonable approach. We therefore theoretically developed a microplate reader assay to roughly quantify the amount of washed-out living cells which yet has to be tested in practice. Both samples, of the remaining biofilm and the washed-out cells (see previous paragraph), are transferred into a microtiter plate and the samples are serial diluted in LB media. After a few hours of initial growth, a growth curve of the cells in the plate is recorded by measuring the optical density at 600 nm (OD₆₀₀) of each well. Comparing optical densities of the serial diluted samples of the biofilm and samples, we are able to estimate the ratio of cells that have been washed out compared to all cells in the biofilm.
We had a short time frame of two weeks in the lab to establish biofilm growth and the assay, but were not able to grow a B. subtilis biofilm following a method based on MSgg media ^[17]. However, for illustration of the assay setup, we used red dye to prepare serial dilutions on microtiter plates. The plate represents samples after a few hours of growth under suitable conditions. The blue rectangle marks the resuspended cells of the biofilm in 2 mL LB media. Each dilution step in our example was performed 1:1.5 for the sake of a visible serial dilution over multiple steps. Assuming that the X^th (e.g. 7^th) dilution of the biofilm sample corresponds to the optical density of the flow sample, we can calculate the ratio of cells washed out by using the following formulas.
Formula 1.

Formula 2.

In case of our example with a dilution factor of 1.5 and 7 dilution steps, 5.53 % of all living cells were washed out. The corresponding dilutions are outlined with a yellow rectangle.

**Figure 7:** Exemplary layout of the microtiter plate assay for estimating the amount of washed-out living cells using serial dilution of red dye instead of *Bacillus subtilis* cultures. In practice, rows A-D would contain samples of the resuspended biofilm in LB media. Rows E-H contain flow samples. Serial dilutions in LB media are horizontally prepared from the samples in column 1. The blue rectangle outlines undiluted cells of a certain area of the biofilm. Yellow rectangles show samples of equal optical intensity and thus roughly similar numbers of cells. Measurements can be conducted in common plate readers with absorbance function.

With our designed flow chamber, we are able to compare different B. subtilis strains regarding their biofilm stability. These information are very important for our project as we display our degradation enzymes in the biofilm matrix fused to the major protein biofilm component TasA. For every new enzyme we want to display in the biofilm, we have to prove that TasA remains able to connect B. subtilis cells to provide a stable biofilm. This assay also allows to compare strains with different genotypes (e.g. strains harboring gene knockouts). We argue that knocking out the gene sinR, encoding the transcription factor which represses tasA expression, will stabilize B. subtilis biofilms – here we present a microplate reader assay to validate our hypothesis ^[10]. Hopefully, we can also contribute to future iGEM projects which may want to use our flow chamber or want to build up on this concept. Therefore, we have created a manual and provide the files for 3D printing the three flow chamber parts. Find the manual, the files and more information on our Contribution page!

Atomic Force Microscopy

An Atomic Force Microscopy (AFM) is a probe microscope with an up to 1000-fold higher resolution than common microscopes^[18,^19]. By measuring the surface, it will give precise information about the topology of our biofilm. A typical AFM setup is shown in Figure 8. The imaging process works with the offset of the laser pointing at the cantilver’s back, as well as the mechanical force bending the cantilever back.
AFM is capable of different measuring modes, that are favourable in varying conditions. For a biofilm’s soft surface, the non-contact mode is preferable since it reduces the friction of the cantilever’s tip on the surface^[20].
Using our 3D printed flow chamber experiments in combination with AFM data analysis, we can determine the stability of our biofilm. For more details see Experiments.

AFM Picture — **Figure 8:** Setup of an AFM that can be used to measure the surface of a biofilm.

Assay Small Molecule Sorption into the Biofilm

We want to produce our pollutant-degrading enzymes fused to one of the B. subtilis biofilm-forming proteins, the major protein component (TasA). This way it will be displayed in the matrix of the biofilm. We need to make sure that the substances are able to enter the biofilm to be converted by our displayed enzymes. Here we focused on the sorption of diclofenac because it poses the biggest issue in wastewater treatment plants. Torresi et al. recently established an assay to measure the uptake of small molecules into biofilms of various thickness on which our assay is based on^[21]. The following assay has been developed theoretically and could not be tested in the lab.

We grow the biofilm directly on carriers used in wastewater treatment to make the experiment as realistic as possible. After the biofilm is formed on the carriers, we test the diclofenac uptake (Figure 9). Therefore, we incubate the carriers with different concentrations of diclofenac and take samples of both the solution and the biofilm at certain time points. The biofilm sample is resuspended in water, centrifuged and washed repeatedly. After that, the cells are lysed via sonification and the suspension is centrifuged again to clear the lysate. The supernatants of this step and the samples of the diclofenac solutions are quantified via UV after HPLC separation. If diclofenac is absorbed by the biofilm at the assayed concentrations, we will do the same with concentrations that can be found in wastewater in Germany and then analyze the taken samples via LC-MS because it is more sensitive than HPLC with UV detection^[22]. For more details see Experiments.

**Figure 9:** The biofilm is grown on plastics carrier. This takes approximately 5 days. Afterwards, the biofilm is put in diclofenac solution and samples are taken at certain time points to analyze via HPLC.

Importantly, plastics has shown adsorption of hydrophobic substances^[23]. On that account, we perform the same assay with an empty carrier in diclofenac solution to see potential adsorption to the carrier itself.

Evaluation

We are confident that the biofilm will grow on these carriers, because the material is the same as used in the flow chamber where it grew. Furthermore, the carrier material was recommended by Prof. Dr. Lackner and these carriers are a common method in moving bed biofilm reactors^[24]. If the biofilm does not grow, we will try and use a different more porous carrier such as pumice stone or ceramics. Biofilm growth on the carrier is detectable by a mucous layer on the floating body which is visible by eye.
Diclofenac is a hydrophobic molecule which is positively charged at pH 7.4^[25]. The B. subtilis biofilm matrix is negatively charged and hydrophobic as well so diclofenac should be able to be absorbed into the biofilm^[9]. This test will be successful if the HPLC analysis shows a decrease of diclofenac even at low concentrations over time so we can test at real-life concentrations. The paper we based our assay on also tested diclofenac biofilm sorption but could not see a significant decrease of diclofenac in their bioreactor^[21]. As they used activated sludge and not a B. subtilis biofilm, we cannot directly compare results. Other reasons why this assay might not work could be wrong charge of the matrix or the size of the pores in the matrix. If this situation would occur, we could try to change the pH of the solution to consequently change the charge of the molecule of the matrix, e.g. the extracellular polymer poly-γ-glutamate. However, this approach would likely be not applicable in WWTP due to the high volume and later drain into the environment. Rather, we could change the conditions in which the biofilm is formed because that can affect the biofilm matrix and it also could be done in our proposed implementation^[26]. Furthermore, we would search for other solutions to make the biofilm matrix more receptive to diclofenac. That could be tried by overexpressing genes responsible for resorptive extracellular polymer substance synthesis^[9].
Here we exemplary focused on diclofenac, but there are further substances whose uptake into the biofilm we could test since they are relevant for this project .

References

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[20] Sean A. James, Lydia C. Powell and Chris J. Wright. Atomic Force Microscopy of Biofilms—Imaging, Interactions, and Mechanics, DOI 10.5772/63312. [21] Torresi, E.; Polesel, F.; Bester, K. Diffusion and Sorption of Organic Micropollutants in Biofilms with Varying Thicknesses. Water Res. 2017, 123, 388–400, DOI 10.1016/j.watres.2017.06.027. [22] Abdel-Hamid, M. E. Comparative LC-MS and HPLC Analyses of Selected Antiepileptics and Beta-Blocking Drugs. Farmaco 2000, 55 (2), 136–145, DOI 10.1016/S0014-827X(00)00006-9. [23] Zhang, H.; Pap, S.; Taggart, M. A. A Review of the Potential Utilisation of Plastic Waste as Adsorbent for Removal of Hazardous Priority Contaminants from Aqueous Environments. Environmental Pollution. Elsevier Ltd March 1, 2020, p 113698 [24] Andersson, S., Nilsson, M., Dalhammar, G. (2008). Assessment of carrier materials for biofilm formation and denitrification. Vatten, 64, 201–207. http://urn.kb.se/resolve?urn=urn:nbn:se:kth:diva-10154 (accessed on Oct 3, 2020) [25] National Center for Biotechnology Information (2020). PubChem Compound Summary for CID 3033, Diclofenac. https://pubchem.ncbi.nlm.nih.gov/compound/Diclofenac (accessed on October 4, 2020) [26] Shukla, A.; Mehta, K.; Parmar, J. Depicting the Exemplary Knowledge of Microbial Exopolysaccharides in a Nutshell. European Polymer Journal. Elsevier Ltd October 1, 2019, pp 298–310, DOI 10.1016/j.eurpolymj.2019.07.044.

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