Bacteria

To achieve our goal of developing a long-term, low-stress solution to gluten cross-contamination, selecting a suitable probiotic host was critically important. The ideal bacterium had to be safe for human consumption (Generally Recognized As Safe, GRAS), capable of colonizing the gut, resilient under gastrointestinal conditions, and able to efficiently produce and secrete enzymes.

For context, one enzyme-based supplement available in Canada is GluteGuard, which contains caricain, a papaya-derived enzyme that can help reduce symptoms of accidental gluten exposure [1]. However, this supplement must be taken before every meal, placing a significant burden on users. Our aim with a probiotic host was to provide continuous protection in the gut, allowing the bacteria to act throughout the day while requiring only a single daily dose.

Initial Cloning and Proof of Concept

For initial development, cloning was performed in Escherichia coli DH5α due to its genetic stability and reliability for molecular cloning [2]. As a model organism, E. coli was ideal for proof-of-concept studies due to its well-characterized genetics and competence.

However, choosing a gram-negative bacterium like E. coli introduced technical challenges. Unlike gram-positive bacteria, which are more commonly used in probiotics and are naturally better at secreting proteins, E. coli has an outer membrane that limits secretion efficiency. These challenges required careful consideration of secretion tags, promoter sequences, and selection markers, with ampicillin or kanamycin resistance used to maintain stable cultures.

Considering Alternative Chassis

The team explored several alternative probiotic chassis, including gram-positive strains such as Lactiplantibacillus plantarum, which are generally recognized for their higher secretion efficiency. Switching to a gram-positive chassis would have required redesigning the genetic constructs, including promoters, secretion tags, ribosome-binding sites, and selection markers. Given the project timeline, this approach was considered too time-intensive and risky.

In parallel, the team explored isolating strains from the probiotic blend Webber Natural Digestion Probiotic 30 Billion containing eight different strains. Initial inoculation and plating produced a dense bacterial lawn, which later serial dilutions allowed for some distinguishable colonies. Differentiating all eight strains without strain-specific selective media proved too complex and time-consuming, so this part of the project was eventually deprioritized. The effort did, however, inform future considerations for selective agents and experimental design.

The tables below summarize the strains considered, outlining the advantages and disadvantages of each potential probiotic chassis.

Final Choice

Ultimately, the team decided to remain with the E. coli, selecting E. coli Nissle 1917, a strain with a long history of safe human use. This choice required addressing specific technical considerations: designing type-specific secretion tags for extracellular proteins, choosing compatible promoter sequences and selection markers, and preparing optimized media for E. coli physiology. Although other gram-positive strains were evaluated, E. coli Nissle 1917 provided the best balance between safety, feasibility, and project constraints.

Enzymes

Working towards our goal of engineering this probiotic, we initially searched for enzymes capable of targeting gluten or its analogs. Our review of enzyme treatments for Celiac disease revealed several promising candidates for oral enzyme therapy. For our project, we focused on AN-PEP, a prolyl endopeptidase derived from Aspergillus niger, while the evaluation of other enzymes was performed entirely in silico. At first, we examined these enzymes individually, but as we progressed in our research, we began exploring combinations of complementary enzymes to potentially enhance gluten degradation efficiency (see below).

AN-PEP

Prolyl endopeptidase (PEP) derived from Aspergillus niger (AN-PEP) has been shown to effectively degrade immunogenic gluten peptides. In a dynamic, multi‐compartmental gastrointestinal in vitro model, AN‐PEP was shown to degrade almost all immunogenic gluten epitopes from gluten‐containing meals into non‐immunogenic fragments during passage through the stomach compartment [29].

FVp-P

Fvp-P, a gluten-degrading enzyme from Flammulina velutipes, demonstrated the ability to cleave all Celiac disease–active peptides at multiple sites with greater efficiency than AN-PEP. These findings indicate that Fvp-P represents a promising candidate for gluten degradation in real food matrices [30]. The enzyme was also utilized by the Macau iGEM team in 2023 [31], where it showed the most promising performance among the enzymes tested.

Kuma030

Kuma030 is a rationally designed synthetic enzyme developed to specifically degrade immunogenic gluten peptides. One study showed that treatment of gliadin with Kuma030 eliminates its ability to trigger a T cell response. In simulated gastric digestion assays, Kuma030 degraded over 99% of the immunogenic gliadin fraction within physiologically relevant time frames, reducing it below the toxic threshold for celiac patients and demonstrating strong potential as an oral therapeutic candidate for Celiac disease [32]. It was also evaluated by the Macau iGEM team in 2023 [31], although they were unable to successfully express the enzyme.

EP-B2 and SC-PEP

EP-B2, a glutamine-specific cysteine endoprotease from germinating barley seeds, and SC-PEP, a prolyl endopeptidase from Sphingomonas capsulata, have been studied together as a combination therapy for Celiac disease. SC-PEP specifically cleaves the proline residues. In vitro and in vivo evaluation of this enzyme combination in rats demonstrates its ability to retain its enzymatic activity in acidic environments comparable to the stomach and duodenum [33]. However, a phase II clinical trial later showed that the combination did not significantly improve histologic or symptom scores compared to placebo, likely due to limited enzymatic activity and degradation within the gastric environment [34].

EP-B2 and AN-PEP

The combination of EP-B2 and AN-PEP offers the advantage of maintaining enzymatic activity and stability in the acidic environment of the stomach. In this pair, EP-B2 initially cleaves large gluten proteins at glutamine residues, producing smaller oligopeptides that can still trigger inflammation. AN-PEP then further hydrolyzes these peptides at proline residues, effectively breaking down gliadin into non-immunogenic fragments [35].

Peptides

Gliadin, one of the two major protein components of gluten, is a key trigger of the immune response in individuals with celiac disease. This protein contains several epitope-binding regions that can provoke adverse immune reactions. To address this, we considered developing a peptide capable of shielding or blocking these reactive sites as a potential strategy to mitigate gluten’s immunogenic effects [36].

Our next step was to identify peptides that could bind to the immunogenic 33-mer amino acid region of the gliadin sequence—potentially preventing recognition of its epitope-rich areas. Previous research using phage display libraries (12-mer and 7-mer amino acids) identified over 160 unique gliadin-binding peptide sequences. Among these, p61 and p64 were the most frequently isolated peptides [37]. These peptides could function either independently or in conjunction with gluten-degrading enzymes.

Assays

To test our project, we selected three complementary assays to evaluate the enzymatic breakdown of gluten. Additionally, we implemented two assays to verify that our peptide caps functioned as intended.

Enzymatic breakdown of gluten

Gluten and Gliadin Plate Assays

The first two assays utilized agar plates containing either gluten or gliadin. Both operated on the same principle—assessing proteolytic activity, specifically the ability of our expressed enzymes to degrade gluten or gliadin, as indicated by the formation of a clear zone around bacterial colonies.

To perform the assay, transformed E. coli cultures were plated with IPTG to induce the lac operator, triggering expression and secretion of the target enzymes. This induction allowed colonies capable of degrading the substrate to form visible zones of clearance.

Although gluten plates are typically used to identify bacteria that naturally degrade gluten, in our case they served to confirm that our plasmid-encoded gluten-degrading enzyme was functioning as expected. E. coli strains cannot degrade gluten or gliadin naturally without a genetically modified plasmid.

Previous studies have identified several bacterial species with inherent gluten-degrading capabilities, including Streptococcus salivarius, Bacillus pumilus, Rothia dentocariosa, Bacillus subtilis, and Staphylococcus epidermidis—all isolated from human saliva [38]. These could serve as positive controls, with Bacillus subtilis being the most accessible candidate for use in Level 1 microbiology labs. Negative controls consisted of untransformed E. coli strains such as DH5α, which lack gluten-degrading capacity.

Colorimetric AN-PEP Activity Assay

The third assay was a colorimetric assay adapted from the method of [39]. It measures the activity of the enzyme AN-PEP by monitoring optical density at 410 nm during incubation with gluten and Z-Gly-Pro-pNA (5 mM) in K phosphate buffer (pH 7.0, 0.1 M). The reaction is stopped using acetate buffer (pH 4.0, 1 M) containing 10% Triton X-100, with a final volume of 250 µL per well in a 96-well plate.

Z-Gly-Pro-pNA is a synthetic substrate of prolyl endopeptidases; upon cleavage, it releases a yellow chromophore measurable at 410 nm. This assay allowed us to evaluate two variables:

• Degradation time: Determining the time required for AN-PEP to reduce gluten concentration to a “safe” threshold. Reactions were run at 5-minute intervals for 95 minutes, informed by our survey results indicating most individuals with celiac disease experience symptoms within 0–3 hours of gluten exposure.
• Degradation efficiency: Measuring gluten degradation at varying initial concentrations (1000 ppm to 10 ppm). The lower value was selected after consultation with Dr. Therrien, considering that the Canadian standard for gluten-free products is below 20 ppm [40].

Our aim was to establish standard curves using the commercial probiotic GluteGuard before testing our own enzyme AN-PEP, allowing for a direct comparison of degradation efficiency between our probiotic and products currently available on the market.

Peptide Cap Evaluation

ELISA

The second set of assays aimed to evaluate the effectiveness of the peptide caps using ELISA-based technology. An ELISA (enzyme-linked immunosorbent assay) is a biochemical technique that detects the presence of a target molecule—such as a protein or antigen—through specific antibody binding, producing a measurable color change. In this context, ELISA detects gluten or gliadin based on their ability to bind specific antibodies. If our peptide caps successfully masked the epitope regions of gliadin, the antibodies would bind less effectively, resulting in a negative or reduced signal.

Due to time and financial constraints, a full quantitative ELISA kit could not be obtained. Instead, a commercially available rapid gluten detection test—capable of identifying gluten concentrations as low as 10 ppm—was used [41]. Although qualitative, this test provided a fast and affordable preliminary assessment, allowing us to gauge the potential effectiveness of the peptide caps while simulating the experience of someone living with celiac disease.

His-Tag Synthesis and Purification

Both the enzymes and peptide caps were synthesized to include a His-tag, enabling potential enzyme activity assays with purified samples. The His-tag, a short sequence of histidine residues, allows rapid and efficient purification of the target protein from a mixture of cellular proteins through immobilized metal affinity chromatography (IMAC). Nickel-binding beads selectively interact with the histidine residues, washing away unwanted proteins and permitting the elution of a highly purified enzyme under controlled conditions. These purified samples are suitable for downstream analyses, including activity assays or structural studies.

Sequencing

Samples for sequencing included both parts from the iGEM distribution kit and synthesized parts ordered from Twist Bioscience. The team used the Monarch Spin Plasmid Miniprep Kit (NEB T1110) protocol [42]. During the first few sequencing attempts, we encountered issues with low plasmid DNA concentrations, particularly with the distribution kit parts. After learning that several other iGEM teams were experiencing similar difficulties, we implemented a series of optimization steps to improve yield:

• Increased culture volume: Using the largest recommended overnight culture volume of 5 mL for mini preps.
• Sample concentration: After centrifugation, we removed most of the supernatant and resuspended the remaining pellet in approximately 100 µL of solution to maximize cell density.
• Reduced elution volume: DNA was eluted in only 30 µL to achieve higher concentration.

Despite these adjustments, some samples remained at the lower end of the acceptable concentration range. However, we opted to send slightly larger total volumes for sequencing, as this would facilitate the process even if concentrations were below the optimal range. We were pleased that the sequencing results for most of our samples came back as expected.

All parts were initially designed and verified in silico using SnapGene through simulated Golden Gate cloning. Once sequencing results were received, alignment analysis in SnapGene allowed us to easily confirm that our cloned sequences matched the expected constructs. In a few cases, minor variations such as single-nucleotide substitutions were observed, but since these occurred outside of our parts of interest, they were not considered problematic.

Both transformed parts and Golden Gate–assembled clones were sent for sequencing through Plasmidsaurus. Overall, sequencing verification confirmed that our assembly and cloning strategies were effective, validating both the design and the experimental workflow used throughout this stage of the project.

Troubleshooting AN-PEP Cloning and Unexpected Deletions

The cloning process proved crucial in uncovering a significant issue with the AN-PEP gene. During cloning using the Golden Gate assembly protocol, we employed green–white screening, as our plasmid backbone contained a fluorescent green cassette. Following successful cloning, this cassette would be replaced with our part insert.

When we first checked the transformation plates, we were thrilled to observe small white colonies—an early indication that cloning may have been successful. However, to confirm that the AN-PEP gene was properly inserted, two white colonies from each plate were picked, grown overnight in liquid culture, and mini-prepped. The DNA concentrations from these preps were satisfactory, and the samples were subsequently sent for sequencing.

The sequencing results, however, were not what we expected. In all eight of the samples, the AN-PEP insert was missing, with an approximately 1.5 kb deletion occurring in the region where the gene should have been. In most cases, the promoter, ribosome binding site (RBS), and terminator were also partially or completely deleted along with the AN-PEP sequence. The deletions varied slightly in both position and length, creating considerable confusion.

Adding to the uncertainty, one of the eight samples showed a plasmid size of approximately 7 kb, compared to the expected 4 kb, while the remaining samples were closer to 3 kb. We hypothesized that the larger plasmid might have resulted from an unexpected plasmid fusion event, though the mechanism behind this and its connection to the deletions remained unclear.

Our mentors suggested that the correct clone might still be present among our colonies and encouraged us to continue sampling to locate an intact version. Nonetheless, several members of the team argued that since all eight samples exhibited nearly identical deletions, and similar issues had reportedly been observed by the UT Austin iGEM team, the problem was likely linked to the coding sequence of AN-PEP itself or its behavior during cloning.

To further investigate, we performed agarose gel electrophoresis to visualize the plasmid sizes. Knowing the expected plasmid length, we digested the samples prior to running the gels to minimize the presence of supercoiled plasmids, which can complicate accurate size determination.

This phase of troubleshooting provided valuable insights into how certain coding sequences—particularly large or complex ones like AN-PEP which has multiple repetitive codon sequeces—may behave unpredictably during cloning. It also highlighted the importance of verifying results through multiple methods, including sequencing and gel electrophoresis, before drawing conclusions about cloning success.

After weeks of trying to isolate a clone with our designed plasmid without success, we opted to instead order the full transcription unit believing that it would be more time efficient to move forward with the project rather than continue to somewhat blindly test every possible colony.

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