Part:BBa_K5117039

PcotYZ-BsRBS-BhBglA-L2-CotY-B0014

This part serves as transcriptional unit composed of:

• promoter PcotYZ of Bacillus subtilis (BBa_K5117021)
• ribosome binding site of Bacillus subtilis (BBa_K5117000)
• bglA gene of Bacillus halodurans encoding a beta-glucosidase (EC 3.2.1.21),
addition of a long flexible linker (L2) downstream of the coding sequence encoding the amino acids (GGGGS)₄ (BBa_K5117027)
• cotY gene of Bacillus subtilis (BBa_K5117022)
• bidirectional terminator B0014 (BBa_B0014)

Biosafety level: S1

Target organism: Bacillus subtilis

Main purpose of use: Immobilization of BhBglA on the spore crust of B. subtilis (spore surface display)

Potential application: Degradation of cellobiose

Sequence and Features

Assembly Compatibility:

• 10
  COMPATIBLE WITH RFC[10]
• 12
  INCOMPATIBLE WITH RFC[12]

  Illegal NheI site found at 764
  
• 21
  COMPATIBLE WITH RFC[21]
• 23
  COMPATIBLE WITH RFC[23]
• 25
  INCOMPATIBLE WITH RFC[25]

  Illegal NgoMIV site found at 1624
  
• 1000
  COMPATIBLE WITH RFC[1000]

Enzyme characterization according to literature

The characterization of the enzyme included in this composite part can be found on the basic part page (BBa_K5117027) of the enzyme.

Construct design

For compatibility with the BioBrick RFC[10] standard, the restriction sites EcoRI, XbaI, SpeI, PstI and NotI were removed from the coding sequence (CDS). To make the part compatible with the Type IIS standard, BsaI and SapI sites were removed as well. This was achieved by codon exchange using the codon usage table of Bacillus subtilis (Codon Usage Database Kazusa).

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studies, this promoter has so far provided the highest activity for spore surface display (Bartels et al. 2018, unpublished data of Elif Öztel). The promoter was followed by the ribosome binding site (RBS) for the host B. subtilis with a 7 bp spacer.

To anchor the target enzyme on the spore surface, it was 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).

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).

Within the framework of the TU Dresden iGEM 2024 Team, three linkers have been tested: 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).

The composite part documented in this page contains the long flexible linker (GGGGS)₄.

Following the CDS of cotY, a spacer consisting of 10 bp of the natural genome sequence downstream from the cotYZ operon was inserted. This creates space before the terminator and ensures that the ribosome is able to read the full length of the CDS. The 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 the BioBrick assembly standard and restriction-ligation-cloning. The vector pBS1C from the Bacillus BioBrickBox was used as an integrative plasmid backbone enabling genomic integration into the amyE locus of B. subtilis (Radeck et al. 2013).

Construction of spore display plasmids

First, all biological parts including the enzyme candidate (Fig. 1) as well as the promoter P_cotYZ, the terminator B0014 and the anchor gene cotY (Fig. 2) were amplified by PCR and purified using the HiYield® PCR Clean-up/Gel Extraction Kit (SLG, Germany). The RBS was added by oligonucleotides. The plasmid pSB1C3-BhBglA generated in the subcloning phase served as template for PCR of the enzyme candidate (see BBa_K5117017). The linker was added by oligonucleotides. The promoter and anchor gene were amplified from genomic DNA of B. subtilis W168 and the terminator from a plasmid provided by the laboratory collection of Prof. Thorsten Mascher (General Microbiology, TU Dresden).

Fig. 1: Agarose gel electrophoresis: PCR of BhBglA-L2. Oligonucleotides for amplification can be found on the Experiments page. The correct PCR product has a size of 1418 bp. 1 kb Plus DNA Ladder (NEB) served as marker (M). The PCR product was purified by gel extraction resulting in a DNA concentration of 138.5 ng/µl.

Fig. 2: Agarose gel electrophoresis: PCR of P_cotYZ, cotY and B0014. Oligonucleotides for amplification can be found on the Experiments page. Amplification of P_cotYZ results in a band of 238 bp, L2-cotY displays a band of 544 bp and B0014 140 bp. 1 kb Plus DNA Ladder (NEB) served as marker (M). PCR products were purified resulting in DNA concentrations of 155.7 ng/µl (P_cotYZ), 143.8 ng/µl (L2-cotY), and 52.2 ng/µl (B0014).

The composite part was subsequently assembled via Overlap PCR by complementary overhangs, which were designed and added by oligonucleotides (Fig. 3).

Fig. 3: Agarose gel electrophoresis: Overlap PCR of the spore display construct P_cotYZ-BhBglA-L2-cotY-B0014. Oligonucleotides for amplification can be found on the Experiments page. The correct PCR product has a size of 2265 bp. Other weak bands are probably caused by unspecific binding of oligonucleotides. 1 kb Plus DNA Ladder (NEB) served as marker (M). The Overlap PCR product was purified by gel extraction resulting in a DNA concentration of 135.4 ng/µl.

Afterwards, the Overlap PCR product was cloned into the vector backbone pBS1C enabling genome integration into the amyE locus in B. subtilis (Radeck et al. 2013). For that purpose, the insert as well as pBS1C were digested with EcoRI and PstI and purified by PCR clean up (DNA concentration: 31.1 ng/µl and 25 ng/µl respectively) using the HiYield® PCR Clean-up/Gel Extraction Kit (SLG, Germany).

After ligation, the plasmid was transformed into chemically competent E. coli DH10β cells and transformants were selected by ampicillin resistance (100 μg/ml ampicillin) encoded on the vector backbone. As the vector was purified via PCR clean up, the vector backbone might re-ligate with the original RFP insert, but colonies with re-ligated plasmids appear pink on the plate and can therefore be distinguished from correct transformants.

The negative control containing no DNA showed no growth. The positive control containing pBS1C resulted in a pink bacterial lawn. The re-ligation control containing the digested vector pBS1C led to growth of approximately 400 pink and 20 white colonies. The transformation with the spore display plasmid pBS1C-P_cotYZ-BhBglA-L2-cotY-B0014 resulted in ≈ 300-400 colonies, both pink and white ones.

Selection plates of the E. coli DH10β transformation are shown below (Fig. 4).

Fig. 4: E. coli DH10β transformants on LB + ampicillin plates.

White colonies transformed with the spore display plasmid were analyzed by Colony PCR and agarose gel electrophoresis (Fig. 5). Colonies with a band at the correct size of the insert were chosen for plasmid isolation according to the HiYield® Plasmid Mini DNA Kit (SLG, Germany). The correct insert sequence of the plasmid was verified via sequencing by Microsynth Seqlab GmbH. Consequently, the spore display plasmid pBS1C-P_cotYZ-BhBglA-L2-cotY-B0014 was successfully constructed (DNA concentration: 212.4 ng/µl).

Fig. 5: Agarose gel electrophoresis: Insert amplification of P_cotYZ-BhBglA-L2-cotY-B0014 by Colony PCR of transformed E. coli DH10β cells. Oligonucleotides for amplification can be found on the Experiments page. Numbers 1-4 correspond to chosen colonies. The correct PCR product has a size of 2362 bp. Two colonies of the re-ligation control (RC) were tested as well, with RC1 appearing white on the plate containing an unknown insert, and RC2 appearing pink on the plate with a probable RFP insert. The negative control displayed no bands. 1 kb Plus DNA Ladder (NEB) served as marker (M). Colony 1 was verified by sequencing and contained the correct insert sequence.

Generation of Bacillus subtilis strains

Ultimately, this plasmid was transformed into the B. subtilis wildtype W168. Transformants were selected by chloramphenicol resistance (5 μg/ml) encoded on the pBS1C vector backbone. The negative control containing no DNA showed no growth. The transformation with the spore display plasmid pBS1C-P_cotYZ-BhBglA-L2-cotY-B0014, which was linearized by restriction with BsaI prior to transformation, resulted in ≈ 400-500 white colonies.

Selection plates of the B. subtilis W168 transformation are shown below (Fig. 6).

Fig. 6: B. subtilis W168 transformants on LB + chloramphenicol plates.

The successful genomic integration was verified via starch assay (Fig. 7). Four colonies 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 of the engineered B. subtilis strain were chosen for cryo-conservation, with clone 1 being used for subsequent activity tests.

Fig. 7: Starch assay of transformed B. subtilis W168 cells. Numbers 1-4 correspond to chosen colonies. W168 served as control and displayed a bright halo. In contrast, no halo was visible for the W168 strain with the integrated spore display construct.

Spore preparation

Spores were prepared by culturing cells in LB medium with chloramphenicol until they reached the exponential growth phase (OD₆₀₀ of 0.4–0.6). After washing and resuspension in DSM, the culture was incubated at 37 °C for 24 hours to induce sporulation. The cells were then 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 p-nitrophenyl-β-D-glucopyranoside (pNPG) assay (Zhang et al. 2017, Korotkova et al. 2009) and the p-nitrophenyl acetate (pNPAc) assay (Xi et al. 2021, Kademi et al. 2000) to achieve a final OD₆₀₀ of 0.2 in the reaction. The reaction was conducted using pNPG and pNPAc as substrates, followed by absorbance measurement at 405 nm to detect the formation of p-nitrophenol (pNP), which produces a yellow color. Further details are available on the Experimentspage.

Glucose assay for determination of glucose concentration after degradation of cellobiose

We first prepared spores displaying BhBglA, as this enzyme showed promising results in previous assays involving induced expression (see BBa_K5117017). Our aim was to determine whether the glucose assay could effectively measure the glucose concentration resulting from the degradation of 50 mM cellobiose by the immobilized enzymes. The assay was performed according to the protocol described on the Experimentspage, using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit. After a 24-hour incubation period, the glucose assay was carried out, and absorbance was measured at 560 nm. The results are presented in Fig. 8.

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 enzymes 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 (see calibration of glucose for glucose assay on Resultspage). These results suggest that there is almost no difference in glucose production between the enzymes, indicating similar catalytic efficiency. The glucose assay appears effective, although a relatively high background absorbance was observed with unpurified cellobiose (data not shown), which still allowed differentiation of enzymatic activity from the control. In the future, we should investigate cellobiose purification further to reduce the background signal.

Fig. 8: Glucose concentration determination after degradation of 50 mM cellobiose by spores displaying β-glucosidases (BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039), BhBglA-L3 ( BBa_K5117040)) (see Experimentspage). The reaction with 50 mM cellobiose was incubated for 24 hours at 50 °C. Following incubation, the glucose assay was performed using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit, and absorbance was measured at 560 nm. Spores from the W168 strain were used as a negative control, while 50 mM cellobiose was used as a substrate control. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction. The measured value from the cellobiose control was subtracted from the enzyme activity measurements to account for the background signal of unpurified cellobiose. The assay was performed in a single biological replicate (N = 1).

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 ( BBa_K5117043) alongside spores displaying BhBglA-L2 ( BBa_K5117039) to compare their activity. The glucose assay was performed according to the protocol (see Experimentspage), and the results are presented in Fig. 9.

Fig. 9: Glucose concentration determination following degradation of 50 mM cellobiose by spores displaying β-glucosidases (BhBglA-L2 ( BBa_K5117039) and PpBglB-L3 ( BBa_K5117043)) (see Experimentspage). The reaction with 50 mM cellobiose was incubated for 30 minutes at 50 °C, with samples collected every 10 minutes. After incubation, the glucose concentration was determined using the Amplex™ Red Glucose/Glucose Oxidase Assay Kit, with absorbance measured at 560 nm. Spores from the W168 strain were used as a negative control, and 50 mM cellobiose was used as a substrate control. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The measured value from the cellobiose control was subtracted from the enzyme activity measurements to account for the background signal of unpurified cellobiose.

The W168 control showed no absorbance at any time point, indicating no glucose production and confirming the absence of enzymatic activity in the control spores. BhBglA-L2 ( BBa_K5117039) displayed increasing absorbance over time, peaking at 20 minutes and slightly decreasing at 30 minutes. This suggests effective enzymatic activity, though the slight decrease could be due to substrate saturation or measurement variability, as triplicates were not performed. PpBglB-L3 showed no absorbance, similar 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 at 20 minutes for BhBglA-L2 likely reflects experimental fluctuations, as no triplicates were performed.

pNPG assay for β-glucosidase activity determination

Based on previous results (see pBS0EX-BhBglA/pBS0EX-BhBglA ( BBa_K5117017) 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 page. We tested spores displaying BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039), BhBglA-L3 ( BBa_K5117040)) as well as PpBglB-L1 ( BBa_K5117041) and PpBglB-L3 ( BBa_K5117043) in two biological replicates (N = 2), to assess and compare their enzymatic activities using pNPG as a more accessible substrate. The results are shown in Fig. 10.

Fig. 10: Evaluation of enzymatic activity of spore-displayed β-glucosidases (BhBglA-L1 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117043)). The reaction was conducted at 40 °C for 10 minutes, followed by absorbance measurement at 405 nm to indicate the formation of pNP. Spores of the W168 strain were used as a control, and additional control without spore solution was included. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The assay was performed in two biological replicates (N = 2). The absorbance from this control was subtracted from the measured values to account for background signal.

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-L1 and PpBglB-L3 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, immobilized PpBglB on spores was excluded from further testing due to the lack of observable activity. Therefore, we focused on BhBglA which was immobilized on spores fused with three different linkers (BhBglA-L1 , BhBglA-L2 and BhBglA-L3).

Determination of optimal temperature

The optimal temperature for BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040) 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 exhibited maximal catalytic performance, as shown in Fig. 11. The assay was conducted according to the standard procedure, with the incubation temperature varied from 40 °C to 90 °C (see Experimentspage).

Fig. 11: Determination of optimal temperature for spores displayed glucosidases (BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040)) using pNPG as substrate (see Experimentspage). The reaction was conducted at various temperatures ranging from 40 °C to 90 °C for 10 minutes, followed by absorbance measurement at 405 nm to indicate pNP formation. Spores from the W168 strain and an additional control without spore solution were used as controls. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The assay was performed with one biological replicate (N = 1). Absorbance from the control was subtracted from the measured values to account for background signal. The relative activity is shown, with the highest value used for normalization, which was obtained from BhBglA-L2 at its maximum activity at 40 °C.

The W168 control shows no activity at any of the tested temperatures. At 40 °C, all three enzymes BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040) exhibit relatively high activity, with BhBglA-L2 showing the highest activity, followed by BhBglA-L1 and BhBglA-L3. The maximum activity for BhBglA-L1 and BhBglA-L2 is observed at 40 °C, while BhBglA-L3 reaches its maximum activity at 50 °C. A slight decrease in activity is seen for BhBglA-L1 and BhBglA-L2 at 50 °C. Generally, BhBglA-L3 shows higher activity than BhBglA-L1 and BhBglA-L2 at elevated temperatures.

At 60 °C, the activity decreases for all variants, but BhBglA-L3 retains the highest residual activity, followed by BhBglA-L1 and BhBglA-L2. At higher temperatures (70 °C, 80 °C, and 90 °C), all immobilized enzymes exhibit minimal activity, indicating a sharp decline in 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 BhBglA-L1 and BhBglA-L2.

Assessment of thermostability of spore-displayed β-glucosidases

Additionally, the thermostability of the BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040) 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. 12. 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.

Fig. 12: Determination of the thermostability of spores displaying glucosidases (BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040)) using pNPG as substrate (see Experimentspage). Before the reaction with pNPG, spores were pre-incubated at different temperatures ranging from 40 °C to 90 °C for 2 hours. Following pre-incubation, the reaction with pNPG was conducted for 10 minutes at 50 °C, and pNP formation was measured by absorbance at 405 nm. Spores from the W168 strain and a control without spores were used as controls. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The assay was performed with one biological replicate (N = 1). The absorbance from the control without spores was subtracted from the measured values to account for the background signal. The measured values were background-corrected and normalized to the corresponding values obtained without pre-incubation at higher temperatures.

The relative activity of BhBglA-L1, BhBglA-L2, and BhBglA-L3 was analyzed after pre-incubation at temperatures ranging from 40 °C to 90 °C, followed by a reaction with pNPG for 10 minutes at 50 °C. The absorbance values were background-corrected and normalized to the values obtained with spore solutions stored at room temperature before the reaction. After pre-incubation at 40 °C, all three 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 the 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 suggest that BhBglA-L1 and BhBglA-L3 may be more thermostable than BhBglA-L2, as they retained higher activity after exposure to 40 °C. However, above 40 °C, all tested enzymes lost their activity, indicating a temperature limit for functional stability due to heat-induced denaturation. The W168 strain, used as a control, showed no activity at any of the temperatures tested, confirming the specificity of the observed enzymatic function. 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 due to differences in enzyme immobilization and experimental conditions, 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.

Using pNPAc to evaluate non-enzymatic properties of spores

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. 13.

Fig. 13: Evaluation of spore interaction with substrates pNPG (dark purple) and pNPAc (light purple) to determine specific enzyme activity of spores displayed glucosidases (BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040)) to determine specific enzyme activity (see Experimentspage). The reactions were conducted at 40 °C for 10 minutes, followed by absorbance measurement at 405 nm to indicate the formation of pNP. Spores of the W168 strain were used as a control. The absorbance from the control without spores was subtracted from the measured values to account for background signal. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The assay was performed in one biological replicate (N = 1). The W168 control shows no with pNPAc or pNPG indicating no enzymatic activity. The absorbance from the control without spores was subtracted from the measured values to account for background signal. The measured values were background corrected and normalized to the corresponding values obtained without the pre-incubation of the spore solution at higher temperatures.

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 β-glucosidases (BhBglA-L1, BhBglA-L2, BhBglA-L3) 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.

Reusability

Finally, we evaluated the reusability of the spore-displayed enzymes for BhBglA (BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039), BhBglA-L3 ( BBa_K5117040). 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. 14).

Normalization was set to the first cycle (100%) for each enzyme fused to a different linker, and subsequent cycles were normalized relative to the first cycle for each respective enzyme. The control was normalized to BhBglA-L1 to allow 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 BhBglA-L1 showing a decline of approximately 40%. Activity continued to decrease across subsequent cycles for all enzymes, with BhBglA-L1 showing reduced performance. By the fourth and fifth cycles, enzyme activity was minimal for all linkers, indicating a loss of catalytic efficiency after repeated use. However, after an additional 1-hour incubation in the fifth cycle (5*), BhBglA-L1 showed an increase in activity (up to 40%), suggesting some residual enzymatic potential during prolonged incubation. This may indicate slower catalytic degradation of pNPG, potentially due to the partial loss of spores.

Fig. 14: Evaluation of the reusability of spore-displayedglucosidases (BhBglA-L1 ( BBa_K5117038), BhBglA-L2 ( BBa_K5117039) and BhBglA-L3 ( BBa_K5117040)) across five reaction cycles (see Experimentspage). Relative activity is shown, normalized to the first cycle (100%) for each linker, with BhBglA-L1 used as the reference for control normalization. Spores underwent five reaction cycles, each followed by washing steps involving centrifugation and resuspension in dH₂O. The reaction was conducted with pNPG as the substrate for 15 minutes at 40 °C, followed by absorbance measurement at 405 nm to indicate pNP formation. After the fifth cycle, an additional measurement was taken after a 1-hour incubation (indicated as 5*). Spores from the W168 strain were used as a control. The absorbance from the control without spores was subtracted from the measured values to account for background signal. The amount of spores was adjusted to achieve an OD₆₀₀ of 0.2 in the reaction mixture. The assay was performed in one biological replicate (N = 1).

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, possibly due to reduced enzyme loss during washing, with some recovery observed after extended incubation in the fifth cycle. This suggests that although the spores are reusable, enzyme inactivation or loss during washing negatively impacts catalytic performance.

However, the OD₆₀₀ of the spore solution was not measured during these experiments, which may have influenced the results. For 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.

References

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