Overview

The results of our project are divided into four phases: First, a comprehensive literature review in which we examined scientific papers and consulted experts with varying expertise to refine our project idea and find suitable enzymes. Second, the subcloning of enzyme candidates as an intermediate step to generate template plasmids for use in following project phases.
Third, results of our reference approach, induced expression, are explained in which enzyme candidates were cloned into inducible expression vectors and activity assays were carried out to identify enzymes functional in the host Bacillus subtilis. Fourth, and finally, promising enzymes were chosen for our final spore surface display strategy and analyzed by activity assays.

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Phase 1: Literature Research

Selection of Enzyme Candidates

As soon as our team decided on the project topic “ReFiBa – Enzyme-Based Recycling of Textile Fibers Using Bacillus subtilis”, we reviewed literature, looked at the work of previous iGEM teams and consulted local experts, all of which shaped our project (see Engineering and Human Practices page).

Our project is divided 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.

To begin, we needed to determine enzyme candidates capable of degrading cellulose and PET. All details and information, which influenced the selection of enzyme candidates, can be reviewed on the Engineering page. Due to the phylogenetic status of B. subtilis as a Firmicute, cellulase candidates from this phylum Firmicutes (Bacillota) were ultimately chosen and thought to function well in the organism. The PETase candidate had already been codon optimized for Bacillus (Xi et al., 2021). The following table (Tab. 1) displays all ten enzyme candidates, the species in which they were found and our abbreviation (which will be used from now on).

Thereby, we achieved our first milestone:

MILESTONE 1:
Decision on enzyme candidates for degradation of cellulose and PET
Endoglucanase, Exoglucanase , β-Glucosidase, PETase

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Phase 2: Subcloning

Subcloning of Enzyme Candidates into pSB1C3

After choosing ten enzyme candidates, we designed biological parts functioning as translational units containing the ribosome binding site for B. subtilis as well as the gene of interest. The parts were flanked by the BioBrick prefix and suffix sequences allowing for BioBrick standard assembly and restriction-ligation-cloning. All information on construct design can be found on the Engineering page.

In order to create a template from which these parts could be amplified, we aimed to subclone the parts into a small vector pSB1C3. For that purpose, we isolated the plasmid from E. coli DH10β provided by the laboratory collection of Prof. Thorsten Mascher, yielding a DNA concentration of 431.0 ng/µl. Afterwards, we performed a Backbone PCR with pSB1C3 (Fig. 1) and subsequently digested the amplified vector backbone and the ordered parts with EcoRI and PstI, which were purified via the HiYield^® PCR Clean-up/Gel Extraction Kit (see Experiments page for detailed protocols).

After ligation, the plasmids were transformed into chemically competent E. coli DH10β cells. Transformants were selected by chloramphenicol resistance (35 μg/ml chloramphenicol) encoded on the pSB1C3 backbone (Fig. 2). For the negative control, no DNA was added during the transformation procedure leading to no colony growth on selection plates. For the positive control, cells were transformed with the vector pSB1C3 resulting in a pink bacterial lawn due to the original RFP insert. On the selection plates of the target constructs, white colonies were tested for the presence of the correct insert by Colony PCR and agarose gel electrophoresis
(Fig. 3).

Colonies with a band the correct size of the insert were chosen for plasmid isolation according to the HiYield^® Plasmid Mini DNA Kit. Finally, ten plasmids each containing one of our ten enzyme candidates were verified via sequencing by Microsynth Seqlab GmbH. These plasmids (Tab. 2) served as templates for PCR amplification of parts required for the next phases of our project.

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Phase 3: Induced Expression

Cloning Enzyme Candidates into Inducible Expression Vectors

After subcloning our biological parts into the small vector pSB1C3, we started project phase 3 focusing on testing the functionality and activity of chosen enzyme candidates. For that purpose, the parts were cloned into xylose-inducible expression vectors to overexpress our target genes and both replicative (pBS0E-xylR-P_xylA) and integrative (pBS2E-xylR-P_xylA) vectors were used (Popp et al., 2017). Details and vector maps are documented on the Engineering page.

The vectors were isolated from E. coli DH10β provided by the laboratory collection of Prof. Thorsten Mascher, resulting in DNA concentrations of 67.5 ng/µl pBS0E-xylR-P_xylA and 165.9 ng/µl pBS2E-xylR-P_xylA. These vectors were digested with EcoRI and PstI (Fig. 4) and purified via gel extraction using the HiYield^® PCR Clean-up/Gel Extraction Kit.

Biological parts previously digested with EcoRI and PstI in the subcloning phase were ligated into the digested expression vectors. In case there was not enough restriction product left, parts were amplified via PCR (Fig. 5) using the plasmids from project phase 1 as a template (e.g. pSB1C3-BhBglA) and subsequently digested and purified via PCR clean up (see Experiments page for detailed protocols).

After ligation, the plasmids were transformed into chemically competent E. coli DH10β and transformants were selected by ampicillin resistance (100 μg/ml ampicillin) encoded on the vector backbone. However, no growth was observed for some constructs (Fig. 6 A), which may be due to very low transformation efficiency.

For those constructs, either the entire vector or the amplified vector backbone was digested and purified by PCR clean up instead of gel extraction (DNA concentrations of digested pBS0E-xylR-P_xylA: 20.8 ng/µl, 25.9 ng/µl; digested pBS2E-xylR-P_xylA: 24.7 ng/µl, 26.0 ng/µl). This approach finally led to successful transformation (Fig. 6 B). Nevertheless, the original vector insert containing RFP was not removed. Colonies with re-ligated plasmids, however, appear pink on the plate due to the RFP insert and can therefore be distinguished from transformants with the target insert.

White colonies transformed with the expression plasmids were analyzed by Colony PCR and agarose gel electrophoresis (Fig. 7). Colonies with a band of the correct size of the insert were chosen for plasmid isolation. Finally, ten replicative and ten integrative expression plasmids containing one of our ten enzyme candidates each were verified by sequencing.

Ultimately, these expression plasmids were transformed into our target host B. subtilis. Since our reference strategy focused on the secretory expression of target enzymes (except for β-glucosidases), we chose WB800N as an expression strain. This genetically engineered variant of B. subtilis W168 features the disruption of all extracellular proteases. The eight-extracellular-protease-deficient mutant is widely used in industrial applications as it increases the stability of secreted proteins (Jeong et al. 2018).

Since we discovered background activity in initial quality activity assays (see Activity assays of secreted and intracellular enzyme candidates), we aimed to generate B. subtilis WB800N deletion strains (ΔeglS, ΔbglH) to remove the background signal caused by the native endoglucanase EglS and β-glucosidase BglH. To achieve this, we sought to replace the gene with a resistance cassette to generate knockouts.

We first amplified the spectinomycin resistance cassette and up- and downstream sequences of the gene to be knocked out by PCR before joining these fragments in an Overlap PCR (see Experiments page for detailed protocols). Here the first difficulties arose, as the ratio and concentration of the different fragments to each other are crucial for efficient amplification of the target product. Several ratios and concentrations were tested (see Experiments), with 10 ng each of up and down fragments and 30 ng of spectinomycin cassette being the best tested option (Fig. 8).

However, transformation of the constructs into B. subtilis WB800N failed several times, resulting in no growth or few false colonies. Even an attempt to use a different resistance cassette (tetracycline) resulted in problems with the efficiency of construct amplification and no transformation success (iteration details can be found on Engineering page).

After multiple failed attempts to generate the deletion strains, we decided to discard this idea and to transform expression plasmids directly into WB800N. Being aware that there is some background signal, this strain will be included as a control in all activity assays. The transformants were selected by MLS resistance (1 µg/ml erythromycin and 25 µg/ml lincomycin) encoded on the vector backbones (Fig. 9).

Since the Bacillus transformation did not work for all plasmids, the transformation was repeated with earlier addition of DNA to growing WB800N cells (at OD₆₀₀ ≈ 0.7 instead of 1.1) to not miss the timepoint of competence. Afterwards, cells were grown until OD₆₀₀ ≈ 1.1-1.3 and the same procedure was followed as in the initial protocol (see Experiments page).

Finally, all remaining plasmids could be transformed into WB800N and colonies were verified by Colony PCR. The presence of replicative plasmids was tested by two primer pairs (double check), whereas both upstream and downstream integration into the lacA locus was checked for integrative plasmids (Fig. 10). Two colonies each with the correct insert size were chosen for cryo-conservation, serving as biological duplicates.

In the end, we successfully generated all 20 expression plasmids, both integrative and replicative, as well as their corresponding E. coli and B. subtilis strains (Tab. 3, Tab. 4), the latter of which were used for subsequent activity tests.

Additionally, we noticed that our replicative vector pBS0E-xylR-P_xylA does not contain a terminator after the insertion site of our constructs. We therefore cloned all replicative plasmids with an additional terminator as a backup option in case there were any expression problems.

For this, the bidirectional terminator B0014 was amplified from a plasmid template provided by the laboratory collection, whereby the BioBrick prefix and suffix were added by oligonucleotides, followed by purification via gel extraction. All generated replicative plasmids (Tab. 3) were digested with SpeI and PstI. The terminator was digested with XbaI and PstI. Both the plasmids and the terminator were purified via PCR clean up. The terminator was then ligated into each replicative plasmid due to complementary sticky ends of XbaI/SpeI and PstI/PstI (see Experiments page for detailed protocols).

The replicative plasmids containing the terminator were transformed into chemically competent E. coli DH10β and transformants were selected by ampicillin (100 μg/ml ampicillin). White colonies were checked for the presence of the terminator by colony PCR and agarose gel electrophoresis (Fig. 11). Colonies with a band of the correct size were selected for plasmid isolation using the HiYield^® Plasmid Mini DNA Kit. The insertion of the terminator into all replicative plasmids was verified via sequencing by Microsynth Seqlab GmbH.

However, these generated plasmids (Tab. 5) were not used further in our project, since no expression problems occurred during testing of expression plasmids without a terminator (see Activity assays of secreted and intracellular enzyme candidates).

Activity Assays of Secreted and Intracellular Enzyme Candidates

For the expression and testing of heterologously expressed enzymes in B. subtilis, we induced protein production by adding 0.5 % xylose when cultures reached an OD₆₀₀ of 0.5–0.6. After 24 hours, supernatants were collected to test secreted enzymes (endoglucanases, exoglucanases, PETase), while intracellular enzymes (β-glucosidases) were tested using liquid cultures and cell lysate. The enzyme activity of lysates and supernatants was assessed, with additional samples analyzed by SDS-PAGE to confirm protein expression. Detailed procedures are available on the Experiments page.

From now on, we will refer to B. subtilis WB800N transformed with the replicative vector pBS0E-xylR-P_xylA containing one of our enzymes, such as BhBglA, as "pBS0EX-BhBglA". Similarly, B. subtilis WB800N transformed with the integrative plasmid containing the same enzyme will be referred to as "pBS2EX-BhBglA".

Endoglucanases
The functionality of the endoglucanases was investigated with a qualitative plate test using 1 % carboxymethyl cellulose (CMC) as substrate. First, the protocol was tested with a commercially available cellulase mixture Accellerase 1500 (Genencor International, United States). After staining the plates with congo red and subsequently destaining them with 1 M NaCl solution, a yellow halo was visible (as shown in Fig. 12).

Next, the background activity of different B. subtilis strains was tested. Overnight cultures of all studied strains were prepared, adjusted to OD₆₀₀ of 0.5, and 10 µl of each culture was pipetted onto the LB-Agar-CMC plates. As positive control 10 μl of diluted 1:100 cellulase mix Accellerase 1500 was added to the well made on the plate as it is described on the Experiments page. All plates were incubated for 24 hours at 37 °C. Then, they were stained with congo red solution and destained with 1 M NaCl to visualize halos (see Experiments page). The results are shown in Fig. 13.

Only with B. subtilis W168 trpC2 ΔeglS::erm, mls^r in which eglS, gene encoding an endoglucanase, was deleted, no background activity was detected. As earlier it was decided to produce enzymes of interest in B. subtilis WB800N, an attempt was made to delete eglS in particularly this strain. However, it failed, and the production of enzymes was performed in B. subtilis WB800N without further deletions.

To determine CMCase activity of heterologous expressed endoglucanases (AtCelA, AtCelG, BsEglS, BpEglA) in B. subtilis, we first conducted a qualitative assay on CMC-Agar plates. For that, we first choose to test the strains containing the replicative pBS0E-xylR-P_xylA vector assuming that the activity will be higher than with the integrative pBS2E-xylR-P_xylA one.

After the expression of the proteins, we applied 15 µl of the supernatant to the wells in the center of the CMC-Agar plates. After 24 hours incubation at 50 °C, we stained plates with congo red method to visualize enzyme activity. The results are shown in Fig. 14. The control strain WB800N and cultures without addition of inducer were used as controls. After induction with 0.5 % xylose, the activity was determined based on halo formation, which indicates CMC degradation. Negligible halos were observed on the plate of WB800N, indicating basal endoglucanase activity in the control strain and confirming the presence of the eglS gene in B. subtilis. Similarly, negligible halo formation was observed in the uninduced cultures, verifying that endoglucanase expression did not occur without induction, with only the basal activity of EglS being present.

Clear halos were observed for BsEglS, BpEglA and AtCelA, indicating successful expression and activity of these endoglucanases. The halo formation suggests that these enzymes are capable of degrading CMC. Since the activity of BsEglS appeared to be comparable to that of BpEglA and higher than that of AtCelA, the latter was not considered for further experiments. Unfortunately, no halo formation was observed on plates with the supernatant of AtCelG, indicating failed production in B. subtilis or very low endoglucanase activity under the conditions tested.

To decide whether BsEglS or BpEglA should be used for further immobilization on the spore surface, a discontinuous DNS assay was performed which is typically applied to estimate the amount of the reducing ends produced during the CMC degradation. At first, for this purpose a calibration curve was generated using calibration standards with glucose concentrations ranging from 200 to 2000 μg/ml (as shown in Fig. 15).

For the DNS assay, supernatants diluted 1:14 were used, obtained from induced B. subtilis WB800N strains containing the respective replicative pBS0E-xylR-P_xylA plasmids. The reaction was conducted with CMC diluted in 50 mM phosphate buffer (pH 7) at 50 °C for different time intervals, ranging from 1 to 13 hours. DNS stop solution was then applied as described on the Experiments page and absorbance was measured at 540 nm (see Fig. 16). Controls included supernatants from B. subtilis WB800N, and supernatants from cultures without the addition of a protein expression inducer.

Since B. subtilis naturally produces BsEglS, a notable background activity was detected after 13 hours of reaction, as previously shown with the qualitative CMC assay. However, this background activity did not interfere with the assessment of recombinant protein activity. The amount of reducing ends produced in samples from uninduced cultures containing plasmids with BsEglS and BpEglA genes was comparable to the WB800N control, reaching 123.33 μg/ml and 113.33 μg/ml, respectively, after 13 hours. Supernatants containing BpEglA led to the formation of 360.00 μg/ml reducing ends, while BsEglS-containing supernatants showed the highest activity, correlating with 500.00 μg/ml of reducing ends produced in 13 hours. Therefore, BsEglS was selected for further spore immobilization experiments.

Noteworthy, since the experiments were carried out with unpurified endoglucanases, the activity values were not standardized to the amount of enzyme responsible for the reaction, which led to limited comparability of the results. It is possible that BsEglS appears more active than BpEglA due to being produced in larger quantities by B. subtilis WB800N under the test conditions rather than having inherently higher activity. Additionally, the chosen discontinuous assay does not reflect the initial velocity of enzyme catalysis and could be influenced by factors such as thermostability. Despite these considerations, BsEglS remained the candidate for spore immobilization experiments, as it is naturally produced by B. subtilis, which may enhance its chances for successful production on the spore surface of this organism.

Exoglucanases
The activity of exoglucanases was tested using a qualitative phosphoric acid swollen cellulose (PASC) assay and the phosphoric acid swollen avicel (PASA) assay, which was developed as described on the Engineering page. The samples used included supernatants from untransformed B. subtilis WB800N and B. subtilis WB800N transformed with replicative pBS0E-xylR-P_xylA vectors containing genes for AtCelS and AtCelO, respectively. Unfortunately, no enzyme activity was detected in any of the samples, as illustrated by the example of AtCelS (shown in Fig. 17). The experiment was repeated with supernatants from cultures in which protein expression was induced, not only at 37 °C as before, but also at 28 °C and 42 °C, however, still without success (data not shown). Because of this, we decided to proceed with both exoglucanase genes for protein immobilization on the spore surface.

β-Glucosidases
To examine the functioning of β-glucosidases, initially LB-Agar-Esculin plates were intended to be used. For that purpose, 10 μl of cellulase mix Accellerase 1500 was added to the well made on the plate as it is described on the Experiments page. Additionally, different strains were tested to study the presence of background activity. Overnight cultures of all generated strains were prepared, adjusted to OD₆₀₀ of 0.5, and 10 µl of each culture was pipetted onto the plates, which were incubated for 24 hours at 37 °C (as shown in Fig. 18). High background activity was observed in all studied strains. Nevertheless, it was decided to test this qualitative assay for the recombinant proteins investigated in our project.

To evaluate β-glucosidase activity, LB-Agar plates containing esculin were prepared, as described on the Experiments page. Additionally, LB-Agar-Esculin plates, with and without 0.5 % xylose, were used to test whether our generated intracellular β-glucosidases could degrade esculin as an alternative substrate for cellobiose. Overnight cultures of all generated strains were grown, adjusted to OD₆₀₀ of 0.5, and 10 µl of each culture was added to the plates, which were incubated for 24 hours at 37 °C (as shown in Fig. 19). We tested both B. subtilis clones that were generated in the cloning procedure; however, data for strains containing the integrative vector (pBS2E-xylR-P_xylA) is not shown, as no other results were obtained.

The assay revealed the formation of a black halo around all cultures, indicating esculin degradation. We hoped for increased activity on plates containing xylose, which was expected to induce expression of our enzymes, but no increase was observed. This may have been due to the background activity being too high to detect any differences. Unfortunately, even the negative control (WB800N) showed similar halo formation, suggesting high background activity from enzymes in B. subtilis that are capable of degrading esculin. Due to this background activity, the assay was discontinued.

Due to the high background activity observed in the previous assay (LB-Agar-Esculin plates), we decided to quantify the amount of glucose produced from the degradation of cellobiose because of the induced expression of our glucosidases. To access the intracellular enzymes to perform the reaction with cellobiose, we subsequently had to lyse the cells.

The glucose concentration in the reactions was determined using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit. This assay involved preparing a working solution containing Amplex^® Red reagent, horseradish peroxidase (HRP), and glucose oxidase, which was then mixed with glucose standards and samples (a more detailed protocol can be found on the Experiments page). A glucose calibration curve was generated using standard concentrations ranging from 0 µM to 150 µM, allowing for accurate quantification of glucose in subsequent reactions (as shown in Fig. 20).

To test the enzymatic activity of our glucosidases, we performed the expression protocol followed by the cell lysis to obtain cell lysate, that was tested for β-glucosidases activity.
The lysates of pBS0EX-AtBglA, pBS0EX-BhBglA, and pBS0EX-PpBglB were incubated with 50 mM cellobiose at 50 °C for up to 24 hours, followed by termination of the reaction. The glucose produced from cellobiose degradation was measured using absorbance at 560 nm with the glucose assay to evaluate enzyme performance, as shown in Fig. 21.

The results indicate that the induced expression (0.5 % xylose was used as an inducer) of pBS0EX-BhBglA produced more glucose compared to its control and other enzymes, suggesting high enzymatic activity. pBS0EX-AtBglA showed moderate activity, with slightly increased glucose production in the induced state compared to the control. Interestingly, pBS0EX-PpBglB exhibited higher absorbance in the control sample compared to the induced state, suggesting unexpected glucose production without induction.

The green bar, representing the cellobiose standard absorbance at 560 nm, suggests the presence of impurities in the cellobiose, resulting in high background signal of the substrate. Furthermore, the lower absorbance values of the enzyme samples compared to the substrate indicate potential interference from the lysate with the glucose assay's coupled reaction, making the assay unsuitable for accurately testing lysate samples. The difference in absorbance between cellobiose and pBS0EX-BhBglA corresponds to approx. 10 µM of glucose. Since the assay involves a 1:2 dilution of the sample, due to addition of working solution, the calculated glucose concentration should be doubled. Therefore, the actual glucose concentration is approx. 20 µM. However, it should be noted that the results might be inaccurate due to potential interference between the lysate and the assay.

Due to these unexpected results and potential interference from the lysate, we decided to use commercially available p-nitrophenyl-β-D-glucopyranoside (pNPG) as an alternative substrate for analyzing the β-glucosidase activity of our intracellular expressed enzymes.

The activity of β-glucosidase was determined by measuring the hydrolysis of pNPG, using the initial rate of accumulation of the colored product, p-nitrophenol (pNP), following the method of Korotkova et al. (2009) (see in Experiments). For activity assessment, 20 µL of β-glucosidase lysate (pBS0EX-BhBglA, pBS0EX-PpBglB, pBS0EX-AtBglA, pBS2EX-BhBglA, pBS2EX-PpBglB, pBS2EX-AtBglA) was mixed with 180 µL of 5 mM pNPG substrate, dissolved in 50 mM sodium phosphate buffer (pH 7.0) and incubated at 50 °C for 10 minutes. The reaction was stopped by adding 100 µL of ice-cold 0.5 M Na₂CO₃, allowing for subsequent measurement of pNP formation at 405 nm. The results are shown in Fig. 22. A negative control, without lysate, was included to determine the background signal of the substrate, and the control values were subtracted from the obtained results to reflect actual enzyme activity.

The induced sample of pBS0EX-BhBglA shows a higher absorbance (approx. A₄₀₅ = 2.5) compared to the control, indicating β-glucosidase activity. pBS0EX-AtBglA, pBS2EX-AtBglA, pBS2EX-BhBglA, pBS0EX-PpBglB, and pBS2EX-PpBglB, show comparable absorbance between induced and control samples, indicating minimal or no increase in activity upon induction. This could be due to insufficient enzyme yield during cell lysis, leading to reduced detectable activity. The negative control, WB800N, shows similar low absorbance for both conditions, confirming the absence of enzyme activity.

The results suggest that pBS0EX-BhBglA in the induced state exhibits higher enzyme activity, while other samples display almost no or minimal increase, suggesting lower or absent induction effects or absent enzyme activity towards the substrate.

Based on the results shown in Fig. 21 and in Fig. 22, BhBglA demonstrated the highest activity, particularly among the replicate vector (pBS0EX strains), in both the glucose and pNPG assays, establishing it as the most promising candidate out of three. Therefore, BhBglA was selected for enzyme display on spores. Additionally, due to challenges associated with testing lysate, PpBglB was also chosen for spore display, supported by favorable findings from literature, despite its comparatively lower activity in the assays.

PETase
To demonstrate the functionality of the produced BhrPET, a continuous p-nitrophenyl-acetate (pNPAc) assay was conducted using supernatants containing this enzyme. In this assay, pNPAc is degraded to acetate and pNP, which imparts a yellow color to the solution. Notably, the reaction also occurs in the absence of enzymes, which is referred to as autohydrolysis. To rule out the influence of autohydrolysis on the measured activity values, a negative control without BhrPET was included. Additionally, supernatants obtained from an untransformed B. subtilis WB800N were used as controls to check the ability of B. subtilis to degrade pNPAc natively. The reaction was performed in 50 mM phosphate buffer with pH 7 at 40 °C for 10 minutes (see Experiments page). The results are shown in Fig. 23.

The volume activity was calculated with extinction coefficient ε = 6.52 mM^-1cm^-1 (Kademi et al., 2000). Notably, the influence of DMSO on the pNP absorbance was neglected. In the future, to obtain more precise values for volume activity, the extinction coefficient should be determined experimentally for the exact conditions used. However, since the focus of our project was on the enzyme immobilization on spore surface of B. subtilis, no further assay optimizations were performed.

With pNPAc assay no background activity could be detected with supernatants obtained from B. subtilis WB800N, while the activity of the BhrPET-containing supernatant was successfully demonstrated, with an activity of 0.375 U/ml. Interestingly, supernatants from the uninduced culture showed slightly higher activity compared to the WB800N controls, indicating basal expression of the vector.

SDS-PAGE
To determine the molecular weight of the expressed proteins, sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed using gels of varying concentrations (8 % for exoglucanases, 10 % for endoglucanases and β-glucosidases, and 12 % for PETases). Samples were prepared by mixing with SDS loading buffer and heating for protein denaturation. After loading the samples and molecular weight marker (PageRuler™ Plus Prestained Protein Ladder, 10 bis 250 kDa), gels were run at 100 V for approximately 1.5 hours. The gels were stained with Coomassie Blue and subsequently destained to visualize the proteins. Further details of the procedure can be found on the Experiments page.

To determine the molecular weight of heterologously expressed β-glucosidases, we expressed the strains containing the pBS0E-xylR-P_xylA vector (shown in Fig. 24). As enzyme purification was not prioritized, and we applied cell lysate directly onto the gel, the presence of multiple protein bands was expected. These bands represent both the target enzymes and other cellular proteins. Consequently, the gel displays several protein bands, which makes it challenging to identify the specific band corresponding to the expressed β-glucosidase gene.

We expected to observe a band for AtBglA, BhBglA and for PpBglB between 51 – 53 kDa. However, due to the overlapping bands from other cellular proteins, it remains difficult to definitively pinpoint the specific bands for these expressed β-glucosidases.

We also analyzed the debris from β-glucosidase expression (as shown in Fig. 25). Like the results obtained from the cell lysate, multiple bands were observed on the gel, making it difficult to accurately identify the target protein bands based on molecular weight. The presence of these overlapping bands further complicates the identification of the specific β-glucosidase proteins.

We also performed SDS-PAGE analysis for other heterologous expressed proteins, including endoglucanases (BpEglA and BsEglS), exoglucanases (AtCelO and AtCelS), and PETase (BhrPET), using the supernatant fraction (see Experiments page).

Since we did not detect any exoglucanase activity, we further analyzed the insoluble fraction (pellet) alongside the supernatant, as shown in Fig. 26. Due to the lack of purification steps, it was difficult to identify AtCelO and AtCelS. The expected molecular weights for the target proteins are 75 kDa for AtCelO and 86 kDa for AtCelS. Additionally, we attempted to express both enzymes at different temperatures (28 °C, 37 °C, and 42 °C), but no bands or activity were detected in any of the conditions (data not shown).

For endoglucanases, since activity was detected in our initial expression, we focused on BsEglS and BpEglA in the SDS-PAGE analysis (shown in Fig. 27). Distinct bands were observed in the induced culture lanes, corresponding to 55 kDa for BsEglS and 72 kDa for BpEglA, verifying the expression of these proteins. However, since BsEglS is originally a B. subtilis gene, slight bands were observed in all samples, confirming some basal expression of this enzyme even without induction.

For our BhrPET we used both generated strains (pBS0EX-BhrPET and pBS2EX-BhrPET). The expected molecular weight for the target protein, BhrPET, is 25 kDa. Since no purification step was performed, multiple protein bands are visible, indicating the presence of both target and non-target proteins in the supernatant, as shown in Fig. 28. A very faint band is observed at the expected molecular weight. However, to confirm the presence of the target protein, further purification is necessary.

The SDS-PAGE analysis of β-glucosidases, endoglucanases, exoglucanases, and PETase consistently revealed multiple protein bands, indicating the presence of both target and non-target proteins. The absence of a purification step contributed to the complexity of the results, making it difficult to definitively identify the target protein bands, particularly when multiple overlapping bands were present. For some proteins, such as BhrPET, only faint bands were observed at the expected molecular weight, whereas clear bands at the correct size were only detected for BsEglS and BhEglA.

To address these challenges, purification using methods such as immobilized metal affinity chromatography (IMAC) with the His-tag of the proteins before SDS-PAGE could help reduce background proteins, allowing for clearer identification of the target bands. Additionally, Western blotting using specific antibodies could confirm the presence of the target proteins, even when they are present at low concentrations. Achieving higher protein expression levels could also improve the detection of target proteins in the gel. Alternatively, using a more sensitive staining method, such as silver staining instead of Coomassie Blue, could enhance the visibility of faint bands. Another possible solution would be to concentrate on the protein samples. However, this would be most effective after a purification step to avoid concentrating of non-target proteins as well.

In conclusion, our SDS-PAGE results indicate a need for further optimization of sample preparation and analysis techniques to clearly identify and confirm the presence of the target proteins. However, since the induced expression was primarily intended to identify candidates with desired enzymatic activity for spore display, we did not pursue additional solutions to improve the purity of the samples. Instead, we concluded that the observed enzyme activity was sufficient for our protein selection, and therefore focused on activity assays for further analysis.

MILESTONE 2:
Decision on final enzymes for spore surface display
Endoglucanase: BsEglS
Exoglucanase: AtCelO, AtCelS
β-Glucosidase: BhBglA, PpBglB
PETase: BhrPET

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Phase 4: Spore Surface Display

Cloning Enzyme Candidates Fused to Anchor Protein Into pBS1C

After choosing enzymes for the spore surface display, we finally entered project phase 4 in which we aimed to immobilize enzymes on B. subtilis spores. To express target genes only under sporulation, a sporulation-dependent promoter P_cotYZ was chosen. The target enzymes were fused to the N-terminus of the anchor protein CotY to anchor them on the spore surface.

We first tested N-terminal fusions, and due to limited time capacities, we were not able to test C-terminal fusions as well. Additionally, different linkers were analyzed: L1 – a short flexible linker, L2 – a long flexible linker and L3 – a rigid linker. All details about construct design are documented on the Engineering page.

First, all biological parts including enzyme candidates with varying linkers (Fig. 29) as well as the promoter P_cotYZ, the terminator B0014 and the anchor gene cotY (Fig. 30) were amplified by PCR and purified using the HiYield^® PCR Clean-up/Gel Extraction Kit (see Experiments page for detailed protocols). Plasmids generated in the subcloning phase (Tab. 2) served as templates for PCR of enzyme candidates with the addition of linkers by oligonucleotides. Promoter and anchor genes were amplified from genomic DNA of B. subtilis W168 and the terminator from a plasmid provided by the laboratory collection.

Parts were subsequently assembled via Overlap PCR by complementary overhangs which were designed and added by oligonucleotides (Fig. 31). All Overlap PCR products of BsEglS, BhBglA, PpBglB and BhrPET could be constructed. However, difficulties occurred during cloning of larger enzyme candidates AtCelO and AtCelS, whose parts could not be assembled in time. This was most likely due to faulty assembly caused by similarities between the overhangs and sequences in the parts. Of the exoglucanases, only one Overlap PCR product was successfully constructed: P_cotYZ-AtCelO-L2-cotY-B0014.

Afterwards, Overlap PCR products were cloned into the vector backbone pBS1C enabling genome integration into the amyE locus in
B. subtilis (Radeck et al., 2013). For that purpose, inserts as well as pBS1C were digested with EcoRI and PstI, whereas inserts were purified by PCR clean up and the vector by gel extraction (Fig. 32) using the HiYield^® PCR Clean-up/Gel Extraction Kit (see Experiments page for detailed protocols).

After ligation, the plasmids were transformed into chemically competent E. coli DH10β cells and transformants were selected by ampicillin resistance (100 μg/ml ampicillin) encoded on the vector backbone. However, no colonies grew on selection plates, most likely due to very low transformation efficiency.
As we already encountered these problems during phase 3 of our project, we repeated the restriction of pBS1C with purification by PCR clean up instead of gel extraction (DNA concentration: 25.0 ng/µl, 29.2 ng/µl). Using this approach, we did not get rid of the original insert containing RFP, but colonies with re-ligated plasmids appear pink on the plate due to the RFP insert and can therefore be distinguished from correct transformants. Transformation plates of spore display plasmids based on vector purification by PCR clean up are shown below (Fig. 33).

White colonies transformed with the spore display plasmids were analyzed by Colony PCR and agarose gel electrophoresis (Fig. 34). Colonies with a band of the correct size of the insert were chosen for plasmid isolation and verification by sequencing. All plasmids of BsEglS, BhBglA and BhrPET with three varying linkers were successfully generated.
For PpBglB, only the constructs with linkers L1 and L3 could be built. The PpBglB-L2 assembly product resulted in white transformants with correct insert size after Colony PCR, but sequencing revealed that none of the inserts had the correct sequence. After repeating ligation, E. coli transformation and Colony PCR, no correct colonies could be found and, unfortunately, we could not manage to generate this construct in time. Of the exoglucanases, only the pBS1C-P_cotYZ-AtCelO-L2-cotY-B0014 plasmid could be constructed due to the cloning difficulties explained above.

Ultimately, these plasmids had to be transformed into the wildtype strain W168 of our target host B. subtilis. The protease-deficient strain WB800N was not required for the spore display approach since proteins are not secreted in this case. Transformants were selected by chloramphenicol resistance (5 μg/ml) encoded on the pBS1C vector backbone (Fig. 35).

The successful genomic integration was verified via starch assay (Fig. 36). Four colonies per construct were transferred onto replica and starch plates. If integration into the amyE locus was successful, the native amylase of Bacillus is not produced correctly, resulting in the organism's inability to degrade starch. Correct transformants displayed no halo around the cells. The wildtype W168 with a functional amylase was used as a control and produces a clear halo. Two biological duplicates were chosen for cryo-conservation.

In the end, we successfully generated 12 of 18 planned spore display plasmids (six enzyme candidates: BsEglS, AtCelO, AtCelS, BhBglA, PpBglB, BhrPET, three linkers each: L1, L2, L3) as well as their corresponding E. coli and B. subtilis strains (Tab. 6), with the latter being used for subsequent activity tests.

Activity Assays of Enzymes Immobilized on B.subtilis Spores

Spores were prepared by culturing cells in LB medium with chloramphenicol until reaching the exponential growth phase (OD₆₀₀ of 0.4 – 0.6). After washing and resuspension in DSM, the culture was incubated for 24 hours at 37 °C to induce sporulation. The cells were lysed using lysozyme and washed with dH₂O and SDS to remove vegetative cell residues. The spore suspension was adjusted to an OD₆₀₀ of 2 for the glucose assay, the pNPG assay, and the pNPAc assay, and to 2.8 for the DNS assay. For qualitative plate assays, an OD₆₀₀ of 0.2 was used. Further details are available on the Experiments page.

Endoglucanases
To determine the CMCase activity of BsEglS displaying spores, we first performed a qualitative assay using CMC-Agar plates following the protocol described on the Experiments page. The OD₆₀₀ of the spore solution was adjusted to 0.2, and 15 µL of the suspension was applied onto the CMC-Agar plates. After an incubation period of 24 hours at 50 °C, the plates were stained with congo red solution and subsequently destained with 1 M NaCl to observe halo formation. The enzyme activity was visualized by the appearance of clear halos on the congo red stained agar, indicating areas of CMC degradation. The results are shown in Fig. 37.

In the control plate containing W168 spores, there was no visible activity observed, as expected. This confirmed that the control spores, which do not display the BsEglS enzyme, exhibit no enzyme activity. In contrast, the plates with spores displaying BsEglS linked via different linkers (L1, L2, L3) showed clear halos, demonstrating that enzyme activity was present.

To quantitatively compare the enzyme activities associated with the different linkers, we subsequently applied the DNS assay, which allowed us to measure the reducing sugars produced during the enzymatic degradation of carboxymethylated cellulose, providing a more detailed comparison of enzyme activity across the different linker systems.

Initially, DNS assay with BsEglS displaying spores was performed with CMC as substrate for 24 hours at 50 °C to quantitively compare the influence of different linkers on BsEglS activity. Spores from W168 showed no activity as expected since endoglucanases are not present natively on the surface of B. subtilis spores. Small differences between the corresponding activity values could be detected, as shown in Fig. 38. The resulted concentrations of reducing ends were 585.02 μg/ml, 616.81 μg/ml and 626.35 μg/ml for BsEglS-L1, BsEglS-L2 and BsEglS-L3 respectively.

The assay was repeated with the reaction conducted for 30 minutes, as shown in Fig. 39. This time BsEglS immobilized via linker L2 showed the lowest activity which resulted in 101.67 μg/ml of produced reducing ends. BsEglS immobilized via linker L1 and L3 performed similarly, with BsEglS-L1 showing slightly higher activity. In samples with BsEglS-L1 293.33 μg/ml of reducing ends were formed and with BsEglS-L3 275.00 μg/ml.

There are different possible explanations as to why no differences were detected after 24 hours. It is possible that, after a certain period, enzyme activity was inhibited, or the enzyme activity was lost due to low thermostability. Furthermore, the influence of saccharides on the germination of B. subtilis should be studied in the future to determine if vegetative cells are formed from spores during the reaction time. Such germination could reduce the number of spores and, consequently, lower the total enzyme activity in the solution.

To characterize the enzymes immobilized on spores, their optimum temperature was determined. A DNS assay was carried out for 30 minutes with CMC as the substrate at varying temperatures ranging from 40 °C to 90 °C (see Fig. 40). The highest activity was observed at 60 °C, and all activity values were normalized to this value. In general, the activity profiles were similar for all linker variants, with BsEglS-L1 showing the highest activity values at all temperatures tested. Earlier, Aa et al. (1994) demonstrated that the maximum activity of BsEglS is reached at 65 °C, which is consistent with the results obtained using spore solutions in this study. Therefore, the immobilization of the enzyme did not have a notable influence on the optimal temperature.

Furthermore, enzyme thermostability was evaluated. Spore-containing solutions were incubated for 2 hours at different temperatures ranging from 40 °C to 90 °C and then cooled down to the room temperature prior to the assay (see Fig. 41). The measured absorbance values were background corrected and normalized to the corresponding absorbance values obtained with spore solutions stored at room temperature prior to the reaction. It was shown that the enzymes lost their activity when incubated at temperatures higher than 60 °C. This result correlated with earlier findings of Aa et al. (1994). They could demonstrate that the enzyme retains its activity up to 55 °C but loses functionality when exposed to temperatures above this threshold for 30 minutes. Thus, the immobilization of endoglucanases did not improve the enzyme's thermostability.

In the future, the influence of longer preincubation times on enzyme activity should be studied to assess the potential efficiency of applying BsEglS at an industrial scale. Furthermore, all experiments performed with DNS assay should be repeated in triplicates to ensure more accurate enzyme activity determination. Additionally, methods for estimating the enzyme amount on the spore cells should be developed to enable better comparability of the results with values reported in the literature.

Exoglucanases
To examine the functioning of AtCelO displaying spores (AtCelO-L2), a qualitative assay was performed using PASC-Agar and PASA-Agar plates following the protocol described on the Experiments page. Initially, the OD₆₀₀ of spore solution was adjusted to 0.2, and 20 µL of it were pipetted onto the plates. The plates were incubated at 50 °C for 24 hours (exemplary shown in Fig. 42). However, no activity could be detected afterwards. Different OD₆₀₀s, ranging from 0.5 to 4.0, were tested, as well as the incubation at room temperature and 65 °C, but without success, indicating that B. subtilis has struggles with the production of this enzyme.

β-Glucosidases
To evaluate the performance of β-glucosidases displayed on spores, activity tests were conducted for spore-displayed enzymes (BhBglA-L1, BhBglA-L2, BhBglA-L3, PpBglB-L1, PpBglB-L3) using pNPG and cellobiose as substrates. Enzymatic activity was measured by monitoring absorbance at 405 nm for pNPG and 560 nm for cellobiose using the glucose assay. This approach allowed us to assess the catalytic efficiency of different linker variants (L1, L2, L3) for the same enzyme, as well as compare the effectiveness of the two β-glucosidases, BhBglA and PpBglB.

First, we prepared spores displaying BhBglA, as this enzyme showed promising results in previous assays involving induced expression. We aimed to determine whether our glucose assay could effectively measure the glucose concentration resulting from the degradation of 50 mM cellobiose. The assay was performed according to the protocol described on the Experiments page, using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit. After a 24-hour incubation period, the glucose assay was carried out, and the absorbance was measured at 560 nm. The results are presented in Fig. 43.

As a control, 50 mM cellobiose, diluted in 1X reaction buffer, was used, and the substrate absorbance was subtracted from the measured values. All three linker variants (L1, L2, L3) exhibited comparable absorbance values of approximately 0.2, corresponding to a glucose concentration of 13.8 µM, which, considering the dilution factor, results in 27.6 µM in the reaction. These results suggest that there is no difference in glucose production between the three BhBglA linker variants, indicating similar catalytic efficiency for each. The glucose assay appears to be effective, although a relatively high background absorbance though unpurified cellobiose was observed (data not shown), which still allowed for distinguishing enzymatic activity from the control. In the future, we should further investigate cellobiose purification to reduce background signal of cellobiose.

We questioned whether a 24-hour incubation period was beneficial, given the low absorbance observed, which suggested that the enzyme activity might be inhibited by the accumulation of glucose in the reaction medium. Therefore, we decided to discontinue the 24-hour incubation and instead assessed enzyme activity over a shorter time frame of 30 minutes, collecting samples at 10-minute intervals (three samples in total). Additionally, we included spores displaying PpBglB-L3 alongside spores displaying BhBglA-L2 to compare their activity. The glucose assay was performed according to the protocol (see Experiments page), and the results are presented in Fig. 44.

W168 control showed no absorbance at all time points, indicating no glucose production and confirming the absence of enzymatic activity in the control spores. BhBglA-L2 displayed increasing absorbance values over time, with the highest absorbance at 20 minutes (approximately 0.18) and slightly lower at 30 minutes. This suggests effective enzymatic activity, though the slight decrease could indicate either substrate saturation or variability in the measurements due to the lack of triplicates. PpBglB-L3 showed no absorbance, comparable to the W168 control, indicating no enzymatic activity under these conditions. These results suggest that BhBglA-L2 effectively degrades cellobiose and produces glucose within the 30-minute incubation period, while PpBglB-L3 shows no detectable activity. The peak absorbance of 20 minutes for BhBglA-L2 is likely to reflect experimental fluctuations, as triplicates were not performed.

Based on previous results and the understanding that cellobiose might be difficult for these enzymes to degrade efficiently, we decided to use pNPG as a substrate to validate the findings from the glucose assay. The assay was performed according to the protocol outlined in the Experiments section. We tested spores displaying BhBglA-L1, BhBglA-L2, BhBglA-L3, as well as PpBglB-L1 and PpBglB-L3, in two biological replicates, to assess and compare their enzymatic activities using pNPG as a more accessible substrate. The results are shown in Fig. 45.

W168 control shows no absorbance, confirming the absence of enzymatic activity and serving as a baseline for comparison. BhBglA-L1 exhibited the highest absorbance (around 2.2), indicating enzymatic activity when pNPG was used as a substrate. BhBglA-L2 showed slightly lower activity compared to BhBglA-L1, with an absorbance of approx. A₄₀₅ = 1.8. BhBglA-L3 displayed a high absorbance like BhBglA-L1, suggesting comparable activity between these two linkers. Both PpBglB-L1 and PpBglB-L3 showed no absorbance, indicating no enzymatic activity under the tested conditions.

Overall, these results indicate that BhBglA-L1 and BhBglA-L3 exhibited the highest activity among the tested variants, while PpBglB variants showed no activity. The use of pNPG as a substrate effectively demonstrated differences in enzyme performance among the linkers and between the two enzymes.

Based on the results of previous assays, PpBglB was excluded from further testing due to the lack of observable activity. Therefore, we focused on BhBglA with three different linkers (L1, L2, L3). The optimal temperature for these was determined by evaluating enzyme activity across a temperature range of 40 °C to 90 °C, using pNPG as the substrate to identify the temperature at which each construct displayed maximal catalytic performance (as shown in Fig. 46). The assay was conducted following the standard procedure, with the temperature varied between 40 °C and 90 °C (see Experiments page).

The W168 control shows no activity at any of the tested temperatures. At 40 °C, all three BhBglA linkers (L1, L2, L3) exhibit relatively high activity, with L2 showing the highest activity, followed by L1 and L3. The maximum activity for L1 and L2 is observed at 40 °C, while for L3, the maximum activity is at 50 °C. There is a slight decrease in activity for L1 and L2 at 50 °C. Generally, L3 shows higher activity than L1 and L2 at elevated temperatures. At 60 °C, the activity decreases for all variants, but L3 retains the highest residual activity, followed by L1 and L2. At higher temperatures (70 °C, 80 °C, and 90 °C), all immobilized enzymes show minimal activity, indicating a sharp decline in enzyme performance, likely due to denaturation. These results suggest that the optimal temperature for BhBglA activity varies between 40 °C and 50 °C depending on the linker, with activity diminishing above 60 °C. BhBglA-L3 appears to retain slightly higher stability at 60 °C compared to the other enzymes that are linked with L1 or L2.

Additionally, the thermostability of the BhBglA variants was assessed by pre-incubating spore solutions at temperatures ranging from 40 °C to 90 °C for 2 hours, followed by measuring residual enzyme activity with pNPG (as shown in Fig. 47). This approach allowed us to evaluate the thermal stability of the immobilized enzymes on spores fused with different linkers by assessing how well each retained activity following heat exposure.

The relative activity of BhBglA (L1, L2, and L3) was analyzed after pre-incubation at various temperatures ranging from 40 °C to 90 °C, followed by a reaction with pNPG for 10 minutes at 50 °C. The measured absorbance values were background corrected and normalized to the corresponding absorbance values obtained with spore solutions stored at room temperature prior to the reaction. After pre-incubation at 40 °C, all three tested immobilized enzymes exhibited some relative activity (25 % to 30 %), with BhBglA-L1 and BhBglA-L3 showing the highest values. In contrast, no activity was observed for any of enzymes after pre-incubation at temperatures from 50 °C to 90 °C, suggesting that the enzymes underwent heat-induced denaturation or lost their functional capacity at these higher temperatures.

These findings indicate that BhBglA-L1 and BhBglA-L3 might be more thermostable than BhBglA-L2, as they retain higher activity after exposure to 40 °C. However, above 40 °C, all tested enzymes lose their activity, suggesting a temperature limit for functional stability due to heat-induced denaturation. The W168 strain, used as a control, showed no activity across all temperatures tested, confirming the specificity of the enzymatic function observed. These results differ from previous literature, which reported an optimal temperature of 45 °C for BhBglA activity, with the enzyme retaining 80% of its activity after incubation at 45 °C for 1 hour (Naz et al., 2010). The discrepancy might be attributed to differences in enzyme immobilization and experimental setups, such as exposure time to heat.

The findings obtained also explain why we observed only minimal glucose production in the glucose assay conducted over 24 hours. The rapid decline in enzyme activity at temperatures above 40 °C likely limited the efficiency of cellobiose degradation, resulting in a low glucose yield. Therefore, for future experiments, the degradation of cellobiose should be carried out at 40 °C to ensure optimal enzyme performance. Additionally, the thermostability assessment should be extended by incubating the spore solution at 40 °C for longer periods prior to the reaction. This approach would provide a more comprehensive evaluation of enzyme stability and suitability for industrial applications at this temperature. By understanding the enzyme's long-term stability at 40 °C, it would be possible to determine whether it can sustain the desired catalytic activity over prolonged industrial processes. It would also be useful to investigate if the enzyme is active at lower temperatures and assess its stability under these conditions. Evaluating the enzyme's activity and stability at temperatures below 40 °C could help determine if the enzyme remains effective in milder conditions.

To strengthen the reliability of these findings, it is essential to perform these experiments in triplicates to confirm the observed trends and ensure reproducibility of the results.

Further, we tested whether the spores interact with pNPAc, a substrate typically used to determine PETase activity, to rule out any potential intraspecific enzymatic properties of the spores themselves. This was done to ensure that any observed activity was due to the specific β-glucosidase enzymes displayed on the spores and not inherent spore-associated activity (as shown in Fig. 48).

With pNPAc, the absorbance values are low and similar for all constructs, suggesting limited interaction, which confirms that the activity measured is specific to the β-glucosidase enzymes and not due to the inherent properties of the spores. Thus, these results indicate that spores displaying BhBglA exhibit specific enzymatic activity towards pNPG, while interaction with pNPAc is minimal, ruling out intraspecific enzymatic properties of the spores themselves. This supports the conclusion that the observed activity is due to the expressed β-glucosidase enzymes.

Finally, we evaluated the reusability of the spore-displayed enzymes for BhBglA fused with three different linkers. The spores were used in five reaction cycles, with a washing step between each cycle. The number of spores in the first cycle was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The washing step involved removing the reaction products by ensuring the spores settled at the bottom of the reaction tube through centrifugation for 5 minutes at 13,000 rpm. The supernatant was discarded, followed by the addition of 1 ml of dH₂O, another centrifugation step, and subsequent removal of the water. 100 µl of fresh dH₂O were added, and the spores were stored until the next usage (20 minutes later). The reaction was conducted with pNPG as the substrate for 15 minutes instead of the usual 10 minutes. After completing the final fifth cycle, the reaction mixture was measured, then incubated for an additional 1 hour, followed by another measurement (indicated as 5* in Fig. 49).

The normalization was set to the first cycle (100 %) for each linker, and the subsequent cycles were normalized relative to the first cycle for each respective linker. The normalization for the control was set to BhBglA-L1 to ensure effective comparison. The W168 control showed no activity, confirming the absence of inherent enzymatic function. In the second cycle, a decrease in activity was observed for all linkers, with L1 retaining the highest activity (approx. 60 %), followed by L3 (approx. 50 %) and L2 (approx. 40 %). Activity continued to decline across subsequent cycles, with L1 maintaining slightly higher activity in the third cycle compared to L2 and L3. By the fourth and fifth cycles, enzyme activity was minimal for all linkers, indicating a loss of catalytic performance after repeated use. After the additional 1-hour incubation in the fifth cycle (5*), BhBglA-L1 showed an increase in activity (up to 40 %), suggesting some enzymatic potential during prolonged incubation. This may indicate slower catalytic degradation of pNPG due to the partial loss of spores.

These results indicate that while spores with immobilized BhBglA enzymes are reusable, there is a noticeable decline in activity with each reuse, likely due to enzyme inactivation or loss during the washing steps. BhBglA-L1 demonstrated relatively better reusability, which might be coincidental, possibly due to reduced enzyme loss during washing.

The reusability of spore-displayed BhBglA with different linkers was assessed across five reaction cycles. Although enzyme activity decreased with each subsequent cycle, BhBglA-L1 showed relatively better performance, with some recovery observed after extended incubation in the fifth cycle. This suggests that while the spores are reusable, enzyme inactivation or loss during washing impacts catalytic performance.

However, the OD₆₀₀ of the spore solution was not measured during these experiments, which may have influenced the results. In future experiments, it would be advisable to measure the OD₆₀₀ and start with a higher value than the OD₆₀₀ = 0.2 used in our experiments. This would allow for normalization to OD₆₀₀ and better management of spore loss during the washing steps, leading to more reliable and comparable results across different reaction cycles.

PETase
The PETase activity of spores displaying BhrPET with three different linkers (L1, L2, L3) was evaluated using an activity assay based on the method described by Xi et al. (2021). The reaction mixture contained phosphate buffer, spores with a final OD₆₀₀ of 0.2, pNPAc at a final concentration of 1 mM, and deionized water, and was incubated at 40 °C for 10 minutes. Detailed protocol can be found on the Experiments page. Following incubation, the reaction was centrifuged at 13,000 rpm for 1 minute to avoid the spores interfering with the absorbance measurement. After centrifugation, 100 µl of the supernatant was transferred into a 96-well plate for absorbance measurement at 405 nm, performed in triplicates.

The PETase activity assay results for spores displaying BhrPET with different linkers are summarized in Fig. 50. The controls included reactions without spore solution and with W168 spores, which did not display enzymes, serving as negative controls. The absorbance values at 405 nm, with the control subtracted, indicate the enzyme activity. W168 control showed negligible absorbance, confirming the absence of enzymatic activity. Among the variants, BhrPET-L3 exhibited the highest activity, with an absorbance of approximately 0.9, followed closely by BhrPET-L1 at around 0.8. BhrPET-L2 showed a lower activity compared to the other linker variants, with an absorbance of about 0.5. These results suggest that the choice of linkers has a substantial effect on the enzyme's performance, with BhrPET-L3 being the most effective under the tested conditions.

The initial PETase activity assay showed fluctuations in activity between the constructs, which may have been influenced by the very low yield of spores in the experiment. To address this issue, the spore preparation was repeated twice for subsequent tests to ensure consistent results. The procedure was repeated as previously described. Unfortunately, the yield of BhrPET-L2 spores was too low to perform any experiments. The Fig. 51 displays the mean absorbance values of two replicates, with the control (substrate without spores) subtracted. The results indicate that BhrPET-L1 exhibited the highest enzyme activity, with a maximum absorbance of approximately 2.5, while BhrPET-L3 showed slightly lower activity. Due to the higher performance, all subsequent experiments testing thermostability and optimal temperature were conducted using BhrPET-L1, as its spore yield was also the highest (data not shown).

Additionally, after the spore preparation, the spores appeared impure, which suggests there may have been issues with the sporulation of the PET constructs. This means that the actual enzyme activity might be higher than what was observed.

The optimal temperature for BhrPET-L1 was determined by conducting the enzyme activity assay at varying temperatures ranging from 40 °C to 90 °C for 10 minutes. The results of the temperature-dependent activity assay for BhrPET-L1 are shown in Fig. 52.

The graph illustrates the relative absorbance at 405 nm for W168 and BhrPET-L1 spores, with values normalized to the highest observed absorbance (A₄₀₅ = 2.35) for BhrPET-L1 at 50 °C, set as 100 %. The W168 control demonstrated negligible absorbance across all temperatures, confirming the absence of enzymatic activity. BhrPET-L1 showed nearly 90% of its maximum activity at 40 °C and around 80% at 60 °C. However, at 70 °C, enzyme activity sharply declined to approximately 10 %, and no activity was detected at 80 °C or 90 °C. These findings suggest that the optimal temperature range for BhrPET-L1 is between 40 °C and 50 °C. Temperatures above 60 °C impair the enzyme's stability and catalytic performance, likely due to the thermal instability of either the enzyme or the substrate.

Spore solutions were incubated at various temperatures ranging from 40 °C to 90 °C for 2 hours to evaluate the thermostability of the enzyme displayed on the spores. After heat treatment, the samples were allowed to cool to room temperature. Subsequently, the residual enzyme activity of BhrPET-L1 was determined using an assay previously described (for 10 minutes at 40 °C). The measured values were background corrected and normalized to the corresponding values obtained without the pre-incubation of the spore solution at higher temperatures. The results of this assessment are presented in Fig. 53.

BhrPET-L1 retained substantial activity up to 60 °C, indicating good thermostability under these conditions. However, enzyme activity declined above 60 °C, with minimal activity detected at 70 °C (20 %) and 80 °C (approx. 17 %) and almost no activity observed at 90 °C. This indicates that BhrPET-L1 loses both stability and catalytic function at higher temperatures. The W168 control showed negligible activity across all temperatures, confirming that the observed enzymatic activity in BhrPET-L1 is specific to the enzyme displayed on the spores.

BhrPET-L1 demonstrates promising activity within the 40 °C to 50 °C temperature range and maintains good thermostability up to 60 °C, making it suitable for applications within moderate temperature environments. However, the enzyme's stability and activity drastically diminish at temperatures above 60 °C.

Noteworthy, the obtained results contradict earlier findings from the work of Xi et al. (2021), in which almost a linear increase of activity was observed from 30 °C to 90 °C after assessment of the optimal temperature. In addition, with the thermostability experiments performed by Xi et al. (2021) it was shown that BhrPET could retain 80 % of its activity when pre-incubated at 80 °C for 2 hours, whereas in our project a drastic loss of BhrPET activity was detected after its pre-incubation for 2 hours at 70 °C and higher temperatures. The discrepancies in the results could be influenced by multiple factors. First, Xi et al. (2021) applied different assay conditions and used p-nitrophenyl-octanoate and not pNPAc as a substrate. Second, the solutions with spore displaying BhrPET contained some impurities as described above, which could have interfered with the applied assay. Finally, enzyme immobilization on the spore surface could resulted in a negative impact on the protein activity and stability.

MILESTONE 3:
Generation of B. subtilis spores with active immobilized enzymes
Endoglucanase: BsEglS
β-Glucosidase: BhBglA
PETase: BhrPET

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Conclusion

Project Summary

During this project, we successfully implemented our two strategies of induced expression and spore surface display. In the induced expression reference approach, all enzyme candidates were successfully cloned into both replicative and integrative expression plasmids, allowing for the overexpression of target genes.
Despite some difficulties during the initial activity assays, we ultimately developed functional assays for endoglucanases, exoglucanases, β-glucosidases, and PETases. Enzyme activity was detected for endoglucanases (BsEglS, BpEglA, AtCelA), PETase (BhrPET) and β-glucosidases (BhBglA). Even though we did not find any activity with β-glucosidase (PpBglB), we proceeded with spore surface display for this enzyme based on supporting literature. No activity was observed for the exoglucanases (AtCelO, AtCelS). The lack of activity for exoglucanases could be due to several factors, such as improper folding, insufficient expression levels, or instability of the enzyme under assay conditions. Additionally, the activity observed for enzymes expressed using replicative vectors was higher than that observed for those using integrative vectors; however, not many tests were conducted in this area due to time constraints.
Functional enzymes were chosen for our final spore surface display strategy. Selected enzymes were fused to an anchor protein within the Bacillus spore crust, whereby flexible (L1, L2) and rigid (L3) linkers were tested. Spore display plasmids of the endoglucanase (BsEglS), β-glucosidases (BhBglA and PpBglA, excluding PpBglB-L2) and PETase (BhrPET) were successfully generated. We were unable to construct exoglucanases in time (except AtCelO-L2) due to difficulties that occurred during cloning.

Ultimately, B. subtilis spores with immobilized enzymes were analyzed. The endoglucanase BsEglS, the β-glucosidase BhBglA, and the PETase BhrPET all displayed activity. Additionally, a reusability assay for the β-glucosidase BhBglA was conducted, providing promising results regarding the sustainability of our project.
The characterization of enzymes immobilized on spores revealed that the optimal temperature for activity varied among the studied enzymes. For BsEglS, the highest activity was observed at 60 °C, consistent with earlier findings that suggested an optimal temperature of 65 °C, indicating that immobilization did not affect the optimal temperature. Similarly, BhrPET and BhBglA demonstrated optimal activity at 40 °C to 50 °C.
The thermostability analysis showed that all enzymes experienced a decline in activity at temperatures above 60 °C, likely due to heat-induced denaturation. BhrPET-L1 retained activity up to 60 °C, while BhBglA variants lost activity after pre-incubation above 40 °C. The control strain W168 exhibited no activity, confirming the specificity of the enzymatic activity observed. These findings indicate that while the immobilization of endoglucanases and other enzymes on spores provided functional enzymes, their thermostability did not improve beyond their natural temperature limits.

All laboratory work is visualized on the Timeline page, providing an overview of all lab activities carried out during this summer.

Project Achievements

Although we encountered difficulties in the lab that led to some unsuccessful results, we also experienced many successes, all of which are summarized below.

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Outlook

Considerations for Replicating Experiments

In both of our strategies (induced expression and spore surface display), we tested both clones of our generated B. subtilis strains only once. Typically, the experiments were performed in a single replicate, with few exceptions (such as spore-displaying endoglucanases and β-glucosidases).
All constructed strains, however, were always stored in biological duplicates (clones 1 and 2), which provides the opportunity to test both clones in future experiments. Additionally, three independent biological experiments should be conducted for each strain to obtain three replicates. Furthermore, all measurements should have been performed in triplicates for better accuracy.
All information and protocols required for replicating experiments are documented on the Experiments page. If future iGEM teams would like to reproduce our experiments and need additional details, please do not hesitate to contact us: igem@tu-dresden.de or our principal investigator Prof. Thorsten Mascher: thorsten.mascher@tu-dresden.de.

Characterization of Enzymes

During this project, various methods and strategies were employed to characterize and improve the expression and activity of several enzymes. However, there are several areas that could be refined and improved in future studies to ensure more reliable and informative results.
The optimization of protein yield obtained after expression in B. subtilis is still possible. For that, e.g., the influence of different promoters on protein production should be explored. Furthermore, in the future, the expression of proteins should be repeated under different conditions, such as varying temperature and incubation times, to determine optimal expression time and settings.
The use of SDS-PAGE proved useful for determining the molecular weight of expressed proteins, as described in the results section. However, the SDS-PAGE experiments should have been repeated, as some gels were of poor quality. Furthermore, the purification of lysates and supernatants would likely lead to better and more distinct results in SDS-PAGE, enhancing the accuracy of our molecular weight assessments.

The optimal pH and pH stability of the spore-displaying enzymes should be determined to further characterize their performance. Testing these parameters will help understand how the enzymes behave under different environmental conditions, which is essential for potential industrial applications. Further optimization of assay conditions could be obtained by testing different OD₆₀₀ values as well as substrate concentrations. This could also help to determine enzyme substrate specificity (e.g., as K_m value). Additionally, determining the actual amount of enzymes displayed on the spore surface is crucial for better activity assessment and comparison with other studies.
The reusability assay of immobilized β-glucosidase BhBglA showed promising results, indicating the potential for sustainable application. However, a better protocol for purification and yield of spores is required to improve reproducibility. Starting with a higher OD₆₀₀ or adjusting substrate concentration may help optimize these experiments and yield better enzyme activity.
The glucose assay was used throughout this study; however, the assay has not been fully implemented due to high background signal from cellobiose. Attempts were made to purify cellobiose, but these were unsuccessful (see Engineering page). Tests were also conducted to optimize the assay using lysate, but without purification of the lysate, the results were unreliable. The purification of cellobiose using glucose oxidase could be considered, though issues with interference in the glucose assay must be addressed. Other coupled assays may provide more accurate insights into enzyme activity.

Overall, the experiments have shown promising results, especially for the enzymes BsEglS, BhBglA, and BhrPET. However, further optimization and detailed characterization are required to better understand and improve the efficiency of the enzymatic activity and spore immobilization processes, as well as to upscale this project for industrial applications.

Future Plans

Regarding cloning, all remaining spore display constructs must be generated, including PpBglB-L2, AtCelO-L1, AtCelO-L3 as well as all three AtCelS constructs. Moreover, we only constructed N-terminal fusions of the enzymes to the anchor protein. C-terminal fusions can also be tested in future experiments. All following transformation experiments and activity assays need to be performed.
The activity assay for exoglucanases should also be further developed, as we continue to encounter difficulties in detecting any activity from the enzyme candidates. If the tested exoglucanase candidates AtCelO and AtCelS prove to be non-functional in B. subtilis, alternative enzymes will need to be identified through literature research.

Finally, when one active enzyme from each category (endoglucanase, exoglucanase, β-glucosidase, PETase) will be identified, these enzymes can be immobilized on one spore by fusing them to different anchor proteins of the spore coat. Many spore coat proteins, e.g. CotY, CotB, CotC and OxdD, have already been used for protein immobilization (Zhang et al., 2019; Lin et al., 2020), making the B. subtilis spore a three-dimensional immobilization platform. However, it is still unknown whether this approach will retain functional enzymes due to possible interaction between the immobilized proteins resulting in mutual inhibition and loss of activity. Nevertheless, the enzymes BhBglA, BsEglS as well as BhrPET are promising candidates for this strategy.
Following the successful application of this approach, the degradation of actual textile fibers needs to be tested. For that purpose, extensive experiments must be performed regarding the pretreatment of textile waste and subsequent degradation ability of our spores.

To successfully upscale our project from the lab to an industrial level, we would need to ensure safety and compliance with regulations. While modifying the spores to lack the ability to germinate is not strictly necessary for industrial use, it would be required if we wanted to avoid an S1 (biosafety level 1) classification. To proceed with this approach, we would first need to consult with the ZKBS (Central Committee on Biological Safety) to determine their specific requirements. Combining approaches, we could consider using the non-germinating strains engineered by the LMU Munich iGEM 2012 team and transform them with our spore display constructs in future experiments.
If the creation of B. subtilis spores unable to germinate and carrying active textile degrading enzymes is successful at lab scale, our project could finally be transferred to industrial scale by testing these spores in large bioreactors (see Entrepreneurship page).

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References