Engineering

Synthetic Biology

Synthetic Biology represents a growing discipline which began in the late 20th century and has rapidly progressed since the early 2000s. The first international conference, Synthetic Biology 1.0 (SB 1.0), took place in 2004 at the Massachusetts Institute of Technology where researchers discussed the engineering of biology (MIT News 2004). From there, the field developed rapidly, starting with signaling circuits and genome synthesis, over metabolic engineering and minimal cells, to xenobiology (ZKBS 2018).
SynBio focuses on the design and construction of new biological parts, devices and systems or the re-design of existing, natural systems for useful applications, making SynBio an engineering science as well as an applied science. It represents an interdisciplinary field integrating concepts from genetic engineering, biochemistry, systems biology, microbiology and computer science.

Key Principles

However, SynBio is not only genetic engineering focusing on targeted DNA manipulation, but also includes an engineering perspective applied to biological systems. Key engineering principles are abstraction, modularity, standardization and characterization.
Abstraction allows biological parts to be decoupled from their natural function for use in another context, leading to completely new systems. Modularity enables the separation and recombination of system components to assemble parts to devices and complex systems. Additionally, standardization of components guarantees compatibility of parts and application of standardized tools and procedures (e.g. BioBrick standard assembly). Finally, parts are characterized, and their features are documented in the Parts Registry, serving as an information system and providing comprehensive information.

Engineering Cycle

The typical engineering cycle for synthetic biologists consists of four stages: design, build, test and learn (Fig. 2).

Design

Scientists first need to define their goal and specify their project idea. They carefully think about a certain challenge and explore possible solutions to the problem. They develop a strategy and design biological parts, devices and systems to realize their idea. This requires biological background knowledge as well as comprehensive literature research. During design, engineers make use of computational tools and ensure compatibility of parts by applying common standards like the BioBrick RFC[10] or Golden Gate Type IIS standard .

Build

In the next phase, the designed parts, devices or systems meant to solve the challenge are built in the laboratory. Common standards like the BioBrick or Golden Gate assembly are followed using construction tools like DNA synthesis and cloning. During implementation in the lab, unexpected difficulties often arise requiring detailed analysis and troubleshooting. After solving these problems, biological devices or systems are constructed and ready for testing.

Test

In the third stage of the engineering cycle, generated devices or systems are validated in the laboratory. They are tested and characterized under various conditions, providing insights towards their capability to solve the current challenge. Like the second phase, unexpected problems might occur during testing which are constantly analyzed and optimized by scientists. All information gained from the testing phase is extensively evaluated.

Learn

Ultimately, all results are analyzed and as much information as possible is considered for adjusting the design to optimize the behavior of the biological device or system. The engineering cycle is then repeated starting with a new design phase followed by implementation, testing and learning. Only when the device or system is capable of solving the challenge at hand, the cycle is finished and a novel SynBio application has been created.

Engineering in our Project

In March 2024, we decided to genetically engineer Bacillus subtilis to be used for textile waste degradation. As we divided our project into two main strategies, namely finding functional enzymes via induced expression as well as spore surface display, we visualized small gearwheels representing these strategies next to the main wheel in the middle representing our entire project (Fig. 3).

By going through our project phases, including literature research, subcloning, induced expression and spore surface display see Results page), we managed to turn the gearwheels further and further, coming closer to our final aim of textile degradation.
In the following section, we describe the design of our solution to the textile waste problem, its implementation and testing in the laboratory as well as all information we have gained from evaluating results.

Design

Literature Research

As soon as our team decided on the project topic “ReFiBa – Enzyme based Recycling of Textile Fibers using Bacillus subtilis”, we intensely reviewed literature and looked at the work of previous iGEM teams that addressed the textile waste issue.

For example, Chalmers-Gothenburg 2020 aimed to produce an enzyme cocktail produced by Escherichia coli for elastane degradation, Edinburgh 2021 immobilized cellulases on silica beads and Greatbay SCIE 2022 tested cell surface display systems for textile degradation using PETases and cellulases. Moreover, we were highly influenced by current research in the working group of Prof. Thorsten Mascher focusing on protein display on B. subtilis spores, which is based on the work of the LMU Munich 2012 iGEM team.

Inspired by their work, we were able to meet several experts (see Human Practices page) who significantly shaped our project. We divided the project into two main strategies: 1) Induced expression as a reference approach, in which we aimed to find active cellulases and PETases for working with B. subtilis as an expression host, and 2) Spore surface display as a final strategy, in which chosen enzymes are immobilized on the surface of B. subtilis spores.
As previously mentioned, former iGEM teams have already worked on engineering devices and systems for textile degradation, but none so far have attempted to immobilize textile-degrading enzymes on Bacillus spores as a display platform. Thus, our project represents a unique strategy to tackle the textile waste problem.

We first searched for suitable enzymes for our aim to degrade cellulose and PET using B. subtilis. Even though fungal cellulases are most efficient in breaking down cellulose, we decided to only focus on bacterial cellulases due to difficulties associated with eukaryotic gene expression in bacteria, e.g. lacking glycosylation which may affect enzyme folding and activity.
We chose cellulase candidates out of the phylum Firmicutes (Bacillota), as B. subtilis belongs to this phylum as well. Hence, these enzymes provide the highest chance of functioning well in Bacillus without applying codon harmonization. Consequently, we excluded all fungal candidates as well as genes from other phyla (e.g. Actinomycetota) and focused on cellulases from Bacillota species. For PETase, we chose a candidate that was already codon optimized for Bacillus combined with the signal peptide SP_aprE from B. subtilis (Xi et al., 2021).

We defined following criteria:

• biosafety level: S1 Bacillota (for cellulases)
• gene size: < 2000 bp (max. 2500 bp)
• active state: monomer
• localization: extracellular (if possible)
• pH range: ≈ 5 - 8
• temperature: ≈ 30 - 60 °C

We decided on ten enzyme candidates (Tab. 1).:

Subcloning

After choosing ten enzyme candidates, we started to design the biological parts in the cloud-based design tool Benchling. Based on the literature, we designed translational units containing the optimal ribosome binding site (RBS) “TAAGGAGG” for expression in B. subtilis, a 7 bp spacer “AAAAAAA” (Vellanoweth & Rabinowitz 1992) and the coding sequence (CDS) of each enzyme candidate (Fig. 4). Design of all parts is documented on respective Parts Registry pages (links on Contribution page).

For cloning via the BioBrick assembly standard, these parts were flanked with the BioBrick prefix and suffix sequences and 4 bp “GATC” as an overhang to facilitate binding of restriction enzymes. The restriction sites EcoRI, XbaI, SpeI, PstI and NotI were therefore removed from the CDS. To make our parts compatible with the Type IIS standard, BsaI and SapI sites were removed as well. Additionally, HindIII sites were taken out to enable cloning with pET expression vectors in E. coli if needed. This was achieved by codon exchange using the codon usage table of Bacillus subtilis (Codon Usage Database Kazusa). These adjusted biological parts were ordered via gene synthesis from IDT.
Ultimately, these parts were to be cloned into the small vector pSB1C3 (Fig. 5), which is commonly used for subcloning, as it comprises a small vector backbone without promoters or other parts.

Induced Expression

In this project phase, we focused on testing the functionality and activity of our chosen enzyme candidates. For that purpose, parts were cloned into xylose-inducible expression vectors to overexpress the target genes. We chose both replicative and integrative vectors, pBS0E-xylR-P_xylA and pBS2E-xylR-P_xylA, respectively (Fig. 6, Fig. 7), with a xylose-inducible promoter for induced expression and a xylose repressor to decrease basal promoter activity (Popp et al., 2017).
Whereas replicative plasmids provide a high copy number and result in high concentrations of target proteins, genomic integration (in this case into the lacA locus) ensures high stability but results in lower protein concentrations. For comparison, we planned to clone parts in both vectors.

Spore Surface Display

After choosing enzymes for spore surface display, we finally entered the last project phase including completely new construct design. In contrast to our reference approach (induced expression), in which the enzyme candidates were secreted due to a signal peptide (except β-glucosidases), these signal peptides are not required for protein immobilization on B. subtilis spores.
Therefore, each signal peptide was removed from the CDS of the chosen enzymes BsEglS, AtCelO, AtCelS and BhrPET. As the chosen β-glucosidases (BhBglA, PpBglB) do not contain natural signal peptides, the original CDS could be used for construct design. Design of all parts is documented on the respective Parts Registry pages (links on Contribution page).

To express target genes only under sporulation, the sporulation-dependent promoter P_cotYZ of B. subtilis was chosen. In previous studies, this promoter has so far provided the highest activity for spore surface display (Bartels et al., 2018, unpublished data of Elif Öztel). Downstream from the promoter, the same RBS with a 7 bp spacer followed as in the construct design of our reference approach.
To anchor the target enzymes on the spore surface, these were fused to the N-terminus of the anchor protein CotY. This anchor is located in the crust, the outermost spore layer, and has been shown to be well suited for protein immobilization (McKenney et al., 2013; Bartels et al., 2018; Lin et al., 2020). We chose to start testing N-terminal fusions because they provided higher reporter signals when working with sfGFP (Bartels et al., 2018). Due to time constraints during the iGEM competition, we were not able to test C-terminal fusions.

Moreover, different linkers between the fused target enzyme and anchor protein were analyzed, as these proteins may affect the folding and stability of each other and, eventually, lead to misfolding and reduced activity. Whereas flexible linkers promote the movement of joined proteins and are usually composed of small amino acids (e.g. Gly, Ser, Thr), rigid linkers are usually applied to maintain a fixed distance between the domains (Chen et al., 2013).
We chose to test three linkers: 1) A short flexible GA linker (L1) encoding the small amino acids Gly and Ala, 2) A long flexible linker (GGGGS)₄ (L2) which is one of the most common flexible linkers consisting of Gly and Ser residues and 3) A rigid linker GGGEAAAKGGG (L3) in which the EAAAK motif results in the formation of an alpha helix providing high stability (Chen et al., 2013).

Following the CDS of cotY, we inserted a spacer consisting of 10 bp of the natural genome sequence downstream from the cotYZ operon. This created space before the terminator and ensures that the ribosome is able to read the full length of the CDS. Secondary mRNA structures were predicted by RNAfold WebServer and showed no differences in stem loop formation (Fig. 8). Each construct ends with the terminator B0014, a bidirectional terminator consisting of B0012 and B0011.

The entire construct was flanked with the BioBrick prefix and suffix, allowing for cloning via BioBrick assembly standard (Fig. 9). The vector pBS1C (Fig. 10) from the Bacillus BioBrickBox was used as an integrative plasmid backbone (Radeck et al., 2013), thereby enabling genomic integration into the amyE locus of B. subtilis.

Build

Subcloning

As soon as the experimental plan was finalized, we started with the subcloning of ordered biological parts into the small vector pSB1C3, which was provided by the laboratory collection. After restriction digest and ligation, the plasmids were transformed into chemically competent Escherichia coli DH10β cells. Successful transformations were verified by Colony PCR and sequencing. Respective cloning results can be found on the Results page and respective Parts Registry pages (links on Contribution page).

Induced Expression

In our reference approach, genes of interest were cloned into inducible expression vectors to verify enzyme functionality in B. subtilis as a host organism. Parts were amplified and cloned into xylose inducible vectors pBS0E-xylR-P_xylA (replicative) and pBS2E-xylR-P_xylA (integrative). After restriction digest and ligation, expression plasmids were transformed into chemically competent E. coli DH10β cells and transformants were verified by Colony PCR and sequencing.
Due to the limited time available within the framework of the iGEM competition, our reference approach was essential to test the functionality of the enzyme candidates in B. subtilis before subjecting immobilized proteins to testing.

Engineering cycles during cloning of expression plasmids

However, building our expression plasmids came with unforeseen challenges that led to us going through rounds of the design, build, test and learn stages. Select the iteration to find out how we dealt with difficulties during cloning.

Subsequently, these generated expression plasmids were transformed into B. subtilis WB800N, a genetically engineered variant of W168 in which eight extracellular proteases are deleted (Jeong et al., 2018). As we seek to secrete most of our enzyme candidates, this strain is perfectly suited for our reference approach. Successful transformants were verified by Colony PCR.
All results related to cloning as well as transformations of E. coli and B. subtilis can be found on the Results page and the respective Parts Registry pages (links on Contribution page).

Engineering cycles during transformation of B. subtilis WB800N

To test the activity of our enzyme candidates, the expression plasmids needed to be transformed into B. subtilis WB800N. Here, we had to go through the engineering stages as well. Select the iteration to see how we dealt with difficulties during the transformation of B. subtilis.

Spore Surface Display

In our final spore display strategy, we aimed to fuse genes of interest to an anchor gene out of the spore crust of B. subtilis. In sporulation-dependent expression, enzymes will be immobilized on the spore surface. All biological parts, including enzyme candidates with varying linkers, the promoter P_cotYZ, the terminator B0014 and the anchor gene cotY were amplified, assembled and ligated into pBS1C.
After restriction digest and ligation, plasmids were transformed into chemically competent E. coli DH10β cells and transformants were verified by Colony PCR and sequencing. The generated spore display plasmids were linearized and transformed into B. subtilis W168. Successful transformants were verified by starch assay, as the vector backbone provides genomic integration into the amyE locus encoding an amylase.
All results related to cloning, transformation of E. coli and B. subtilis can be found on the Results page and the respective Parts Registry pages (links on Contribution page).

Engineering cycles during cloning of spore display plasmids

Building our spore display plasmids also included some challenges. We therefore went through the design, build, test and learn stages once again. Select the iteration to find out how we dealt with difficulties during cloning.

Test

During the development of enzyme activity assays for both exoglucanase and β-glucosidase, we employed an iterative engineering cycle to address challenges and optimize our methods. For exoglucanase, our goal was to develop a reliable and quantitative activity assay, which required the evaluation of various substrates, staining techniques, and incubation conditions. We successfully established a straightforward visualization technique using phosphoric acid swollen cellulose (PACS) and avicel (PASA).

Our work on β-glucosidase activity assays faced challenges, particularly with background interference and substrate suitability. Initial attempts using LB-Agar-Esculin plates and cellobiose degradation assays highlighted the complexities associated with substrate specificity and lysate interference, respectively. By exploring alternative substrates like pNPG and optimizing assay conditions, we improved the reliability and accuracy of our measurements.

Engineering cycles during the development of a quantitative exoglucanase activity assay

During tests with exoglucanases we noticed that our assays did not work as planned. Select the iteration to find out how we dealt with difficulties during the development of a qualitative assay for exoglucanase activity determination.

1

Iteration 1

2

Iteration 2

3

Iteration 3

4

Iteration 4

Iteration 1

Design
Initially, the protocol for exoglucanase activity examination was adapted, with multiple corrections, from the work of Singh et al. (2022). Since endoglucanases primarily target amorphous regions of cellulose while exoglucanases act on its dense, crystalline regions, avicel, a substrate with high crystallinity, was selected for exoglucanase activity determination.
Build
LB-Agar-Avicel media containing 1% avicel was prepared as described in the Experiments section. Wells were created in the medium, and 10 µl of the commercially available cellulase mix, Accellerase 1500 (containing exoglucanases as indicated by the manufacturer), was added into the wells. Water was used as a negative control. The plates were then incubated at 50 °C for 24 hours. After incubation, the plates were stained with congo red and subsequently destained using 1 M NaCl solution (see Experiments).
Test
No exoglucanase activity could be determined using the applied method. Instead, red stains of higher intensity were observed around the wells, as the congo red solution infiltrated the wells and reached the bottom of the plate.

Learn
Possibly, the staining with congo red did not work out or the applied enzyme could not effectively degrade substrate under the applied conditions.

Iteration 2

Design
The assay design remained largely unchanged, still utilizing avicel as the substrate. However, adjustments were made to the avicel concentration in the media, the incubation time, and the sample volume, which was doubled. These changes aimed to verify whether the issues encountered were due to the assay setup itself rather than the enzymes applied.
Build
The assay was performed as previously described, with the following adjustments: LB-Agar-Avicel media were prepared with either 1% or 0.5% avicel, 20 µl of Accellerase 1500 solution was used, and plates were incubated for 48 hours at 37 °C.
Test
Again, no exoglucanase activity was detected with the assay.

Learn
The results indicated that the congo red solution might not penetrate the avicel medium effectively, suggesting that the staining solution was not suitable for the assay.

Iteration 3

Design
To improve staining efficiency, a new assay design with a cellulose overlay was developed, hoping that this setup would allow the congo red staining method to work properly.
Build
10 μl of Accellerase 1500 mix was added to LB agar plates, dried out and then 0.2 % cellulose overlay was poured onto the medium (see Experiments section). The plates were incubated at 37 °C.
Test
No enzyme activity was detected, even though the staining appeared to work better.

Learn
Since none of the methods applied thus far yielded successful results, it was decided to change both the substrate and the assay setup.

Iteration 4

Design
Through literature research, it was discovered that phosphoric acid swollen avicel (PASA) is sometimes used as a substrate for exoglucanase tests due to its more accessible structure for enzymes. Based on this finding, a new assay was designed using PASA, as well as testing phosphoric acid swollen cellulose (PASC).
Build
PASC and PASA agar media were prepared with approximately 0.2% PASC and PASA, respectively, as described in the Experiments section. 20 µl of commercially available cellobiohydrolase I was added into wells made in the medium. The plates were incubated at 50 °C for 24 hours.
Test
After incubation, visible zones of clearance appeared without the need for staining, indicating the degradation of PASC and PASA.

Learn
An assay for determining exoglucanase activity was successfully developed. To the best of our knowledge, this type of assay had not been previously reported. The fact that no staining solution is needed to visualize the zone of substrate degradation is highly advantageous, as it allows for easier observation of the degradation process at different time points. Moreover, congo red is a carcinogenic reagent, and avoiding its use is an additional benefit.

Engineering cycles during the development of a β-glucosidase activity assay

During the evaluation of β-glucosidase enzymatic activity, we discovered that our initial assay did not perform as intended. Select each iteration below to learn how we addressed the challenges encountered during the development of both qualitative and quantitative assays for β-glucosidase activity determination.

1

Iteration 1

2

Iteration 2

3

Iteration 3

4

Iteration 4

5

Iteration 5

6

Iteration 6

7

Iteration 7

Iteration 1

Design
Initially, we decided to test the activity of the intracellularly expressed β-glucosidases (AtBglA, BhBglA, PpBglB) on LB-Agar-Esculin plates, aiming to observe black halo formation around the colonies as an indicator of enzyme activity.
Build
LB-Agar-Esculin media was prepared as described in the Experiments section. Overnight cultures of our strains were prepared, and the next day, 10 µl of OD₆₀₀ = 0.5 culture was spotted onto plates (with and without 0.5 % xylose inducer). E. coli and WB800N strains were used as controls. The plates were incubated overnight, and halo formation was monitored and documented every hour for up to 6 hours, and again at 24 hours.
Test
No halo formation was observed before 6 hours, and after 24 hours, black halos were formed around every colony, regardless of induction (Fig. 15). This suggested that even if β-glucosidase activity existed, it was obscured by high background activity.

Learn
We learned that either the background activity was too high, preventing detection of β-glucosidase activity, or that our selected enzymes were not capable of degrading esculin effectively. The high background potentially resulted from aryl-β-glucosidases, which are natively produced in B. subtilis. These enzymes can specifically degrade aryl-glucosides, but are not able to cleave cellobiose (Ouyang B. et al., 2023).

Iteration 2

Design
Due to the high background activity observed in the previous assay (LB-Agar-Esculin plates), we decided to quantify the glucose produced from cellobiose degradation due to the induced expression of our β-glucosidases.
Build
To access the intracellular enzymes, we lysed the cells (see Experiments) prior to the reaction with cellobiose instead of using liquid cultures. The glucose concentration in the reactions was determined using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit (a detailed protocol is provided on the Experiments page).
Test
The lysate of pBS0EX-AtBglA, pBS0EX-BhBglA, and pBS0EX-PpBglB was incubated with 50 mM cellobiose at 50 °C for up to 24 hours, after which the reaction was terminated. Glucose production from cellobiose degradation was measured at 560 nm using the glucose assay to evaluate enzyme performance, as shown in Fig. 16.
The results indicated that induced expression (0.5 % xylose) of pBS0EX-BhBglA produced more glucose compared to its control and the other enzymes, indicating high enzymatic activity. pBS0EX-AtBglA showed moderate activity, with a slight increase in glucose production upon induction. Interestingly, pBS0EX-PpBglB exhibited higher absorbance in the control sample compared to the induced state, suggesting unexpected glucose production without induction. The high absorbance of the cellobiose standard (green bar) suggested the presence of impurities (e.g., glucose), resulting in a high background signal.

Learn
The lower absorbance of enzyme samples compared to the substrate suggested potential lysate interference, rendering the assay unsuitable for accurately quantifying the glucose in the applied samples. We learned that interference from the lysate with the glucose assay's coupled reaction made this approach unsuitable for accurate testing.

Iteration 3

Design
In this iteration, our goal was to determine the maximum lysate concentration that could be used without interfering with the glucose assay's coupled reaction. We aimed to optimize the lysate concentration in order to minimize interference while still retaining sufficient β-glucosidase activity for accurate measurement.
Build
The assay was set up using varying amounts of lysate from WB800N, serving two purposes: the culture contained 0.5 % xylose during the expression phase to evaluate any impact on glucose levels, and to ensure no β-glucosidase activity from heterologously expressed enzymes, thereby providing a baseline for comparison with our enzymes.
Test
The lysate (at varying percentages) was mixed with 50 mM cellobiose, 200 µM glucose, and 1X reaction buffer from the Amplex™ Red Glucose/Glucose Oxidase Assay Kit. Glucose concentration was measured using the kit, and absorbance was recorded at 560 nm. The results showed that reducing the lysate volume decreased background interference, as shown in in Fig. 17, with the aim of consistently detecting 200 µM glucose.

Learn
We learned that optimizing lysate concentration is crucial for minimizing interference from the lysate. After diluting the reaction with the glucose working solution, we determined that the effective lysate concentration should be a maximum of 0.5 % in reactions involving the glucose assay. However, even at 0.5 % lysate, a decrease in glucose detection was observed, indicating some residual interference. (Note: This value effectively doubles due to the dilution factor introduced by the glucose assay working solution.) Therefore, in reactions with cellobiose, we decided to use up to 1% lysate in the reaction mixture to achieve reliable results, balancing enzyme activity and interference.

Iteration 4

Design
In this iteration, we aimed to evaluate the β-glucosidase activity of our enzymes over a prolonged incubation period. The goal was to determine glucose production from cellobiose degradation using an optimized lysate concentration that minimized interference.
Build
Reactions were set up with our β-glucosidases (pBS0EX-AtBglA, pBS0EX-BhBglA, pBS0EX-PpBglB and pBS2EX-AtBglA, pBS2EX-BhBglA, pBS2EX-PpBglB) for 24 hours at 50 °C, using a maximum lysate concentration of 1 % with 50 mM cellobiose. After incubation, the glucose concentration was determined using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit.
Test
Reducing the lysate concentration led to decreased enzyme activity, resulting in lower glucose production, as shown in Fig. 18. No difference was observed between the induced and control samples, and in some cases, the control even showed higher absorbance than the induced sample. However, both were still higher than the cellobiose-only control.

Learn
We observed a high background signal from cellobiose, possibly indicating impurities or inhibition of glucose detection. Additionally, reducing the lysate concentration may have led to some loss of enzyme activity, impacting overall glucose production.

Iteration 5

Design
To address the high interference observed in the previous iteration, we aimed to purify the cellobiose to reduce interference in glucose detection.
Build
The cellobiose substrate was purified by treating it with glucose oxidase at 37 °C for 3 hours. The reaction was then stopped by heating to 85 °C, and impurities were removed by filtration before using the substrate in a follow-up reaction with lysate.
Test
The glucose assay with the purified cellobiose showed no improvement, as the background signal remained. Additionally, the H₂O₂ produced from the reaction between glucose and glucose oxidase interfered with resorufin, the color-changing substrate in the glucose assay kit.
Learn
Purification of the cellobiose substrate did not reduce the background signal. We concluded that using an alternative coupled assay to determine the concentration of glucose might be useful. Another option would be to use a commercial chromogenic substrate, such as p-nitrophenyl-β-D-glucopyranoside (pNPG).

Iteration 6

Design
To overcome the interference issues observed with cellobiose, we decided to use pNPG as the substrate for the β-glucosidase activity assay, as it provides a direct colorimetric measurement.
Build
The reaction was set up with 5 mM pNPG instead of cellobiose (see Experiments), and the absorbance was measured at 405 nm to quantify β-glucosidase activity.
Test
The β-glucosidase activity of pBS0EX-AtBglA, pBS0EX-BhBglA, pBS0EX-PpBglB, pBS2EX-AtBglA, pBS2EX-BhBglA, and pBS2EX-PpBglB was determined using pNPG as the substrate. Absorbance at 405 nm was measured, indicating the formation of p-nitrophenol (pNP) from pNPG degradation by the different β-glucosidases, as shown in Fig. 19.

Learn
Using pNPG as a substrate effectively eliminated the background interference issue, providing a more reliable and straightforward measurement of β-glucosidase activity. This approach proved to be a suitable alternative for assessing enzyme performance, especially when using unpurified lysate.

Iteration 7

Design
In this iteration, we aimed to evaluate the thermostability of β-glucosidase enzymes displayed on spores using pNPG as the substrate. The goal was to understand how enzyme activity changes over time under different conditions.
Build
Spores displaying β-glucosidase (BhBglA-L1, BhBglA-L2, BhBglA-L3) were pre-incubated for 2 hours at temperatures ranging from 40 °C to 90 °C. After incubation, the spores were tested with pNPG to assess residual enzyme activity.
Test
The results indicated that enzyme activity decreased after 2 hours of incubation, suggesting a loss of activity at elevated temperatures over extended periods, as shown in Fig. 20.

Learn
This result highlighted the need to revisit the cellobiose degradation experiments and reduce the incubation time to a maximum of 2 hours instead of 24 hours.

Learn

Cloning Strategy

In both of our strategies, induced expression and spore surface display, we faced difficulties during cloning. Especially, the ppurification of the digested vector backbone appears to be crucial for subsequent ligation. We would therefore directly purify digested vectors by PCR clean up instead of gel extraction in future experiments.
Using this approach, a re-ligation control during E. coli transformation must be included, as the original insert containing RFP is not removed. Colonies with re-ligated plasmids, however, appear pink on the plate due to the RFP insert and can therefore be distinguished from correct transformants via pink-white selection.

During the construction process of our spore display constructs, cloning issues occurred as well, particularly concerning Overlap PCR. Parts were most likely misassembled due to complementary sequences, as bands of wrong sizes were detected after agarose gel electrophoresis. In the future, we would adjust the design regarding overhangs added by oligonucleotides to avoid unspecific binding and misassembly.

Transformation of B. subtilis

After having discovered background activity of the WB800N strain in initial quality assays regarding endoglucanase and β-glucosidase activity, we planned to generate WB800N deletion strains (ΔeglS, ΔbglH) to remove the background signal for our reference approach.
This idea was ultimately discarded, though, as multiple attempts to transform WB800N with the knockout constructs including antibiotic resistance cassettes failed. Consequently, we always included WB800N as a control in activity assays. We learned that deletion strains are not necessary for testing enzyme activity, as a visible difference between some expression strains (e.g. endoglucanase BsEglS) and the control could be detected.

Regarding the B. subtilis transformation itself, we discovered that the protocol has to be adjusted when using the strain WB800N. To test the secretory expression of additional enzyme candidates, we would recommend earlier addition of DNA (OD₆₀₀ ≈ 0.7) to not miss the timepoint of competence. Cells should then be grown until OD₆₀₀ ≈ 1.1-1.3, following the same procedure as in the initial protocol (see Experiments).
As the earlier addition of DNA has no negative effect on either DNA or cells, this approach could also be applied to the transformation of W168. In transformation experiments for the spore display strategy, we implemented this adjustment and obtained successful transformants.

Quantitative exoglucanase activity assay

Throughout the development of a quantitative exoglucanase activity assay, variations in incubation conditions, such as temperature, incubation time, and enzyme concentration, were tested across different iterations. It was found that longer incubation times and higher temperatures did not always improve activity detection when using avicel as the substrate with congo red staining.

Using substrates like PASA and PASC, which do not require staining, made it easier to visualize enzyme activity while avoiding the use of harmful chemicals like congo red. A method that allows for direct visualization of activity, such as observing zones of clearance, proved to be more reliable and safer compared to using staining reagents. Eliminating the staining step improved the clarity and simplicity of the assay, allowing for a more straightforward evaluation of exoglucanase activity.

Although we could not detect any exoglucanase activity with our strains, we successfully demonstrated the functionality of the assay using a commercial exoglucanase. The next steps would involve optimizing the expression of AtCelO or AtCelS, or generating spores displaying exoglucanases, to further explore enzyme activity determination.

β-glucosidase activity assay

During the evaluation of β-glucosidase activity, the initial assay using LB-Agar-Esculin plates encountered high background activity, making it impossible to accurately detect enzyme activity. This may have been due to endogenous activity exhibited by the B. subtilis strain or the inability of the chosen enzymes to effectively degrade esculin.

When using cell lysates, interference in the glucose assay prevented the accurate quantification of glucose produced from cellobiose degradation. Moreover, the glucose standards needed careful control for impurities to avoid misleading results. Reducing the lysate concentration decreased background interference, which improved the reliability of β-glucosidase activity detection. However, using too little lysate reduced enzyme activity, leading to lower glucose production. Additionally, longer incubation times resulted in a decline in enzyme activity and increased background signal from cellobiose, potentially due to substrate degradation or assay interference.

Attempts to purify the cellobiose substrate did not reduce the background interference, and the H₂O₂ produced during the glucose oxidase reaction interfered with resorufin detection in the glucose assay kit, rendering purification ineffective. Switching to a more direct colorimetric assay or using alternative substrates, such as pNPG was found to mitigate these complications.

Using pNPG as a substrate provided a direct and reliable measurement of β-glucosidase activity, effectively eliminating the background interference observed with cellobiose. The use of pNPG offered a more straightforward and interference-free method for assessing β-glucosidase activity, especially when working with unpurified lysates. However, since pNPG is not the substrate, which we aim to degrade, substrate specificity influence on the enzyme performance was neglected during these experiments. In the future, this parameter must be considered in order to select enzymes with the best characteristics.

Conclusion

The goal of our project was to develop a sustainable approach for textile waste degradation using genetically engineered B. subtilis spores. By leveraging the inherent stability and robustness of bacterial spores, we aimed to create an efficient and reusable biocatalyst for breaking down cellulose and PET, which are prevalent in textile waste.

The use of spore display presents several key advantages: it eliminates the need for enzyme purification, provides a natural platform for enzyme attachment, and allows immobilized enzymes to remain functional. Additionally, the spores could be reused, offering a potential route to cost-effective enzyme applications. However, we observed that enzyme activity decreased after repeated use, likely due to enzyme loss during washing steps or inactivation over time due to heat exposure.

Despite challenges during the development of enzyme activity assays, particularly for exoglucanase and β-glucosidase activities, we successfully demonstrated the promise of our approach. Testing enzymes on spore surface display provided valuable insights into enzyme performance.

In the future, enhancing the stability of enzymes displayed on spores could be achieved by exploring different linker strategies or alternative spore anchoring proteins. We also plan to examine the pH stability of the enzymes and determine their optimal pH for activity. Expanding the selection of enzymes, including optimizing candidates like AtCelO and AtCelS, will be crucial for improving the overall efficiency of textile degradation.

References