Our project, "ProBabyotics," is founded on a systematic and iterative engineering strategy. We aimed to develop a modular platform to enhance infant nutrition by producing three key functional biomolecules: the human milk oligosaccharide
We rigorously applied the
Figure 1: Abstract figure for the three DBTL systems
This page documents our engineering journey, demonstrating how each DBTL cycle provided critical data and insights that informed the design of the subsequent iteration, leading to a robust and successful outcome for all three project goals.
Our primary engineering goal was to create a highly efficient microbial cell factory for the low-cost production of 2'-FL. We achieved this through a four-part DBTL cycle.
Our initial design was to establish a proof-of-concept for the 2'-FL biosynthetic pathway. We hypothesized that the first two enzymes, phosphomannomutase (manB) and mannose-1-phosphate guanylyltransferase (manC), would be sufficient to initiate the synthesis of 2'-FL by converting endogenous precursors and exogenous lactose(Li et al., 2018). We designed a minimal genetic circuit containing the manB and manC genes, cloned into a pET28a vector under the control of an inducible T7 promoter.
Figure 2: Construction of BL21-manB+manC. (A) The plasmid map of pET28a-manB+manC. (B) The gene circuit of BL21-manB+manC.
The manB and manC genes were synthesized and cloned into the pET28a vector. The resulting plasmid was transformed into E. coli BL21(DE3). Successful construction of the recombinant plasmid was confirmed by agarose gel electrophoresis, which showed a distinct band at the expected size of 1947 bp(Figure 3).
Figure 3: (A)Flow chart of bacterial culture; (B)the agarose gel electrophoresis analysis of the manB+manC gene fragment
Agarose gel electrophoresis analysis confirming the successful cloning of the manB+manC gene fragment (1947 bp).
The engineered BL21-manB+manC strain was cultured in M9 minimal medium with glycerol and yeast extract. Production was induced with 0.5 mM IPTG and 8 g/L lactose. After 72 hours of fermentation, the 2'-FL concentration in the supernatant was quantified using a commercial detection kit. The engineered strain was compared against control strains (wild-type BL21 and BL21 with an empty pET28a vector)(Figure 4, A).
Figure 4: Comparison of 2'-FL yield in BL21-manB+manC after 72-hour fermentation
The BL21-manB+manC strain produced 8.10 ± 1.03 mg/L, while control strains showed negligible production(Figure 4, B).
The results provided a crucial proof-of-concept. The co-expression of manB and manC was sufficient to initiate 2'-FL synthesis. However, the modest yield strongly indicated that the downstream steps—the synthesis of the GDP-L-fucose donor and the final fucosylation reaction—were the primary metabolic bottleneck(Byun et al., 2006). This learning directly informed our next design iteration: to construct the complete synthesis pathway.
Based on the learnings from Cycle 1.1, we designed a complete de novo synthesis pathway. We hypothesized that adding the three downstream genes—GDP-D-mannose-4,6-dehydratase (gmd), GDP-L-fucose synthase (fcl), and α-1,2-fucosyltransferase (futC)—would overcome the previously identified bottleneck and significantly increase the 2'-FL yield(Bonin
et al., 1997). The new composite part (Part: BBa_25DDSHWK) integrates all five genes into a single operon under the control of the T7 promoter(Figure 5).
Figure 5: Construction of BL21-manB+manC+gmd+fcl+futC. (A) The plasmid map of pET28a-manB+manC+gmd+fcl+futC (B) The gene circuit of BL21-manB+manC+gmd+fcl+futC.
The gmd, fcl, and futC genes were sequentially cloned into the pET28a-manB+manC plasmid, creating the final five-gene construct. The plasmid was then transformed into E. coli BL21(DE3). The successful integration of the new gene fragments was confirmed by agarose gel electrophoresis, which showed distinct bands at the expected sizes for gmd (1119 bp), fcl (963 bp), and futC(1095 bp)(Figure 6).
Figure 6: (A)Flow chart of bacterial culture; (B) Agarose gel electrophoresis analysis confirming the successful cloning of the gmd, fcl, and futC gene fragments.
With the full pathway constructed, we conducted a comprehensive series of tests to quantify yield, verify product identity, and understand the system's dynamics.
Figure 6: (A)Flow chart of 2'-FL content assay; (B)Yield comparison of 2 '-fucosyllactose after 72 hours of fermentation of BL21 recombinant strain
The strain with the complete pathway (BL21-manB+manC+gmd+fcl+futC) produced significantly more 2'-FL than the initial strain (BL21-manB+manC) and controls(Figure 6).
Figure 7: (A)Mechanism of HPLC; (B)HPLC analysis of fermentation products of BL21.
Figure 8: (A)(B)(C)Experimental Flowchart; (D) Changes in 2'-FL yield, Lactose Content, and Cell Dry Weight Over Time in BL21-manB+manC+gmd+fcl+futC.
This round of experiments successfully transformed our proof-of-concept into a fully functional and validated production pathway. We gained three critical insights:
With the product identity confirmed and its synthesis from lactose established, our analysis shifted towards metabolic efficiency. The dynamic fermentation data showed continuous lactose consumption, prompting the question:
In analyzing the metabolism of our E. coli BL21(DE3) chassis, we identified a critical flaw: the native β-galactosidase enzyme, encoded by the lacZ gene, directly competes for our lactose substrate by hydrolyzing it into glucose and galactose(chin et al., 2021). We hypothesized that this substrate competition was a major bottleneck. To test this, we designed an experiment to compare our system's performance in BL21 versus E. coli DH5α, a common lab strain that is naturally lacZ-deficient(Figure 9).
Figure 9: Methods for Exploring the effect of two different strains(BL21/DH5α) on 2'FL yield. (A)Experimental Methods; (B)Experimental Flowchart
The pET28a-manB+manC-gmd-fcl-futC plasmid containing our complete five-gene operon was transformed into chemically competent E. coli DH5α cells, creating a new engineered strain for a direct head-to-head comparison.
We conducted a comparative fermentation experiment, culturing both the DH5α-based strain and the original BL21-based strain under identical conditions. After 72 hours, the final 2'-FL concentration was quantified.
Figure 10: Comparison of 2'-FL production in BL21 vs. DH5α chassis.
The lacZ-deficient DH5α strain produced 158.26 ± 21.67 mg/L, a ~19-fold improvement over the 8.23 ± 1.86 mg/L from the BL21 strain(Figure 10).
The dramatic ~19-fold increase in yield powerfully validated our hypothesis. Substrate competition from the native lacZ gene was a major limiting factor. By simply switching to a more suitable chassis, we successfully redirected metabolic flux towards our product. This critical learning established DH5α as our optimized chassis for all subsequent work and highlighted the importance of host selection in metabolic engineering.
Having optimized the chassis, we focused on the final rate-limiting enzyme, futC, which is of human origin. We hypothesized that its heterologous expression in E. coli led to poor solubility and misfolding, limiting the amount of active enzyme(Du et al., 2021). To solve this, we designed a new construct where a Thioredoxin A (TrxA) solubility-enhancing tag was fused to the N-terminus of futC(Figure 11).
Figure 11: Methods for the effect of TrxA tag fusion futC on 2 '-fucosyllactose yield. (A)Experimental Methods; (B)Experimental Flowchart; (C) The gene circuit of DH5α-manB+manC+gmd+fcl+TrxA-futC.
Using standard molecular cloning, we replaced the original futC gene in our plasmid with the newly designed TrxA-futC fusion gene. The resulting plasmid was transformed into our optimized E. coli DH5α chassis.
A comparative fermentation was conducted between the DH5α strain expressing the untagged futC and the new strain expressing the TrxA-futC fusion. Final 2'-FL yields were measured after 72 hours.
Figure 12: Comparison of 2'-FL production with and without the TrxA fusion tag.
The TrxA-futC strain yielded 279.47 ± 22.09 mg/L, a 56% increase over the untagged version (179.27 ± 11.58 mg/L)(Figure 12).
The significant 56% increase in yield conclusively demonstrated that the solubility of the futC enzyme was indeed a major bottleneck. Our protein engineering strategy was successful. This final optimization completed our DBTL cycle for the 2'-FL module, resulting in a robust, high-yielding production strain and a clear, logical engineering workflow.
This system involved a complete "gene-to-function" engineering cycle to produce an active protease for improving protein digestion.
Our goal was to produce active trypsin. To avoid the toxicity of an active protease to the host cell, we designed a system to express its inactive precursor (Choi & Lee et al., 2004), trypsinogen (
Figure 13: Construction of BL21-TRYP. (A) The plasmid map of pET28a-TRYP; (B) The gene circuit of BL21-TRYP; (C) The flow chart of Construction of BL21-TRYP; (D)the agarose gel electrophoresis analysis of the TRYP gene fragment
The pET-28a-TRYP vector was successfully constructed and transformed into E. coli BL21(DE3). High-level expression resulted in the formation of insoluble inclusion bodies. We therefore developed and optimized a multi-step downstream process(Figure 14):
Figure 14: This is a flow chart of "Trypsin Expression and Refolding".
We validated our system at both the protein and functional levels.
Figure 15: (A)The flow chart of Western Blot; (B)SDS-PAGE analysis of recombinant TRYP overexpression in E. coli
Figure 16: (A)The flow chart of "
The results provided a powerful demonstration of a complete gene-to-function workflow. We successfully expressed the target gene, established an effective protocol to recover active protein from inclusion bodies, and confirmed the high catalytic activity of the final product. This validated our design and provided a functional enzyme ready for its intended application in hydrolyzing casein.
This cycle focused on producing and validating the function of β-galactosidase to address lactose intolerance(Kumar et al., 2022).
Our objective was to produce active β-galactosidase (lacZ) using the existing BioBrick part
Figure 17: Construction of BL21-LacZ. (A) The plasmid map of pET28a-LacZ; (B) The gene circuit of BL21-LacZ; (C)The flow chart of Construction of BL21-LacZ; (D) the agarose gel electrophoresis analysis of the lacZ gene fragment
The lacZ gene was successfully cloned into the pET-28a vector and transformed into E. coli BL21(DE3).
We validated our system at both the protein and functional levels.
Figure 18: (A) This is a flow chart of Western Blot; (B) SDS-PAGE analysis of recombinant lacZ overexpression in E. coli.
Figure 19: (A) This is a flow chart of measurement of the lactose content lacZ breaks down; (B) Efficacy of recombinant β-galactosidase in lactose degradation
Efficacy of recombinant β-galactosidase in lactose degradation. The enzyme successfully degraded 80% of the initial lactose substrate (from 10 mM to ~2 mM) within 8 hours(Figure 19).
The results clearly demonstrated that our engineered and purified lacZ protein is highly active and functional. The successful in vitro validation confirms that our recombinant enzyme is a viable candidate for its intended application: breaking down lactose in dairy products. This work also provides valuable new characterization data for the existing BioBrick part BBa_I732005.
We plan to use Escherichia coli Nissle 1917 as the host strain to construct a stable and efficient engineered strain for large-scale production of 2′-FL, trypsin, and lactose. As a GRAS-certified probiotic, Nissle 1917 has good safety and metabolic engineering potential, making it ideal for food production. By optimizing key metabolic pathways, we aim to improve the synthesis efficiency and yield of 2′-FL, trypsin, and lactose, and facilitate the transition from laboratory-scale to industrial production. The goal is to achieve high-purity, cost-effective products that meet the nutritional needs of infants and promote gut health. This platform will also provide a scalable route for the synthesis of other human milk oligosaccharides (HMOs). Currently, we are using BL21 and DH5α strains for our experiments. Though the probiotic strain is safe for food use, like other Gram-negative bacteria, it may produce endotoxins. To ensure the safety of infant formulas, we will reduce endotoxin levels by modifying related genes and optimizing purification steps, establishing a safe and efficient system for the production of 2′-FL, trypsin, and lactose for nutrition and health applications.