Name: Transketolase A (TktA)

Base Pairs: 1995 bp

Origin: Escherichia coli K12

Properties:

Transketolase A (TktA) is an important thiamine pyrophosphate (TPP)-dependent enzyme found in Escherichia coli and many other microorganisms. It functions in the non-oxidative branch of the pentose phosphate pathway, catalyzing the transfer of two-carbon units between sugar phosphates. With a molecular weight of approximately 72 kDa per subunit, TktA typically forms a homodimer, and its activity depends on cofactors such as thiamine pyrophosphate and magnesium ions. The enzyme plays a central role in linking glycolysis with the biosynthesis of aromatic amino acids, nucleotides, and certain vitamins, as it helps generate erythrose-4-phosphate (E4P), a key precursor for the shikimate pathway.

TktA is constitutively expressed in bacteria but can be upregulated in conditions requiring increased flux through the pentose phosphate pathway, such as during rapid growth or oxidative stress. The stability and broad catalytic activity of TktA make it a versatile metabolic enzyme. In metabolic engineering, its role in boosting precursor supply for aromatic compound biosynthesis is well recognized.

Figure 1. Gene maps of TktA

Usage and Biology

In cellular metabolism, TktA acts as a bridge connecting central carbon metabolism to biosynthetic pathways. By catalyzing reversible reactions that interconvert sugar phosphates, it helps maintain a balance between ribose-5-phosphate, sedoheptulose-7-phosphate, glyceraldehyde-3-phosphate, and erythrose-4-phosphate. The production of E4P is particularly significant because this metabolite is the starting point for the shikimate pathway, which ultimately leads to the synthesis of aromatic amino acids such as phenylalanine, tyrosine, and tryptophan.

From a biological perspective, TktA not only contributes to biomass production but also supports redox homeostasis by working in concert with the oxidative branch of the pentose phosphate pathway, which generates NADPH. This dual role highlights its importance in both growth and stress adaptation. In bacteria like E. coli, the enzyme complements its isozyme TktB, though TktA is generally considered the primary transketolase responsible for sustaining flux under normal conditions.

In applied research, TktA has become a key target for metabolic engineering strategies aimed at enhancing the biosynthesis of value-added compounds. Overexpression of tktA is frequently used to increase the supply of precursors for L-tyrosine, L-DOPA, and various aromatic derivatives. Its relatively stable expression, well-characterized catalytic mechanism, and compatibility with heterologous pathways make it an attractive component in synthetic biology. Moreover, understanding its regulation and structural properties provides insights into optimizing flux distribution in engineered strains.

Cultivation

We obtained the TktA gene through PCR amplification. After amplification, we subjected the amplified product and the DNA maker to agarose gel electrophoresis. As shown in Figure 2, the length of the TktA amplification was 1995bp, which was in the correct relative position compared with the maker. After that, we recovered the fragment through agarose gel and recombined the fragment with pCDFDuet by homologous recombination.

Figure 2. Electrophoresis results of TktA gene and pCDFDuet and two other target fragments (AroF and AroE)

Name: 3-Deoxy-D-arabino-heptulosonate-7-phosphate synthase, tyrosine-sensitive (DAHP synthase) (AroF)

Base Pairs: 1071 bp

Origin: Escherichia coli K12

Properties:

3-Deoxy-D-arabino-heptulosonate-7-phosphate synthase, tyrosine-sensitive (DAHP synthase, AroF) is a key, rate-limiting enzyme in the aromatic amino acid biosynthesis pathway in bacteria and plants. It belongs to the sugar phosphate synthase family and catalyzes the condensation of phosphoenolpyruvate (PEP) and erythrose-4-phosphate (E4P) to produce 3-deoxy-D-arabinoheptulose-7-phosphate (DAHP). This is the first step in the shikimate pathway and the starting point for the biosynthesis of all aromatic compounds. AroF typically has a molecular weight of 38.8 kDa and exists as a homotrimer or tetramer with a typical α/β folding pattern.

AroF is unique in its sensitivity to feedback inhibition by tyrosine: when intracellular tyrosine accumulates, it binds to the allosteric site of AroF, inhibiting enzyme activity and preventing excessive metabolite synthesis. This metabolic regulation mechanism is a crucial means for microorganisms to maintain aromatic amino acid homeostasis. Because tyrosine is a precursor for numerous biosynthetic pathways (such as neurotransmitters, hormones, and polyphenols), AroF plays a crucial role in cellular metabolic networks.

Clinically and industrially, AroF is not directly associated with human disease, but its value in metabolic engineering is significant. For example, by knocking out feedback inhibition or modifying AroF, the production of aromatic amino acids (such as L-tyrosine, L-phenylalanine, and L-tryptophan) can be significantly increased, thereby improving the efficiency of downstream synthesis of pharmaceuticals, pigments, flavors, and polyphenols.

Figure 3. Gene maps of AroF

Usage and Biology

In biological contexts, AroF is the gatekeeper of the shikimate pathway, a pathway widely found in bacteria, fungi, and plants, but lacking in animal cells. This makes it an ideal target for antibiotic and herbicide development. AroF's core function is to initiate the production of DAHP, which is then converted through multiple steps to shikimate, ultimately leading to the synthesis of three essential aromatic amino acids. Because these amino acids are crucial for protein synthesis and secondary metabolite production, AroF activity directly determines the strength of the aromatic metabolic flux.

In microbiological research, AroF is often used as a model to explore metabolic regulation. Its tyrosine sensitivity provides a paradigm for studying the mechanisms of allosteric enzymes, particularly the significance of feedback inhibition in metabolic homeostasis. Furthermore, researchers have also exploited site-directed mutagenesis or gene editing to remove feedback inhibition in AroF and enhance host cell metabolic productivity. For example, engineered AroF mutants of Escherichia coli have been used in industrial fermentation to synthesize aromatic amino acids and their derivatives, which are widely used in the pharmaceutical, food, and cosmetics industries.

At the application level, AroF is an important "tool enzyme" in synthetic biology and metabolic engineering. By regulating AroF expression levels or altering its allosteric properties, scientists can control the flux of aromatic metabolic pathways. This not only increases the production of basic amino acids but also promotes the biosynthesis of high-value-added compounds such as polyphenolic antioxidants, plant secondary metabolites, and some novel bio-based chemicals. Therefore, AroF shows significant potential in green manufacturing and sustainable chemical production.

Cultivation

We also obtained the AroF fragment by PCR amplification using E. coli K12 as a template. As shown in Figure 2, we can see that the AroF fragment is 1071bp long. The electrophoresis results show that it was successfully amplified. We then recovered and purified it from the gel and finally recombined it with other fragments into the pCDFDuet vector.

Name: 4-Hydroxyphenylacetate 3-monooxygenase, oxygenase component (HpaB)

Base Pairs: 1563 bp

Origin: Escherichia coli K12

Properties:

4-Hydroxyphenylacetate 3-monooxygenase, oxygenase component (HpaB) is a key protein in a class of flavin-dependent, two-component monooxygenase systems, typically acting in conjunction with its partner, the reductase HpaC. HpaB primarily catalyzes the hydroxylation of 4-hydroxyphenylacetate (4-HPA) with molecular oxygen to form 3,4-dihydroxyphenylacetic acid (homogentisate), a crucial step in the catabolism of aromatic compounds. HpaB proteins typically have a molecular weight of approximately 58.8 kDa and possess a typical monooxygenase fold and a flavin (FAD or FMN) binding pocket to complete the oxidation reaction.

Compared to other aromatic oxygenases, HpaB exhibits high substrate specificity and reaction efficiency, effectively overcoming the chemical inertness of aromatic rings and enabling further metabolism of recalcitrant aromatic compounds. This ability is crucial for bacteria to adapt to diverse carbon sources and degrade aromatic pollutants in the environment. The enzymatic characterization of HpaB reveals that it requires an electron donor (such as NADH or NADPH) and completes the catalytic cycle through electron transfer with the reductase HpaC.

In the natural environment, HpaB is primarily found in Escherichia coli and other Gram-negative bacteria, where it participates in the decomposition of aromatic compounds and energy harvesting. Because it directly processes benzene rings, HpaB is of great significance in environmental microbiology, pollutant degradation, and metabolic engineering.

Figure 4. Gene maps of HpaB

Usage and Biology

In biological processes, HpaB's primary function is to serve as the catalytic core of an oxidase complex, hydroxylating aromatic compounds and thus paving the way for downstream metabolism. The conversion of 4-HPA to homogenous black acid, mediated by HpaB, is a key step in the utilization of aromatic compounds as carbon and nitrogen sources by many bacteria. Due to the extreme chemical stability of aromatic rings, few enzymes in the natural environment can efficiently degrade these compounds. Therefore, the presence of HpaB significantly enhances the cellular metabolism of complex organic compounds.

In research, HpaB is often used to elucidate aromatic metabolic pathways, particularly the mechanism of aromatic hydroxylation. As a two-component monooxygenase, it provides an ideal model for understanding the coupling between electron transfer and oxidation reactions.

In terms of applications, HpaB exhibits numerous potentials:

1. Environmental bioremediation: HpaB is capable of degrading a variety of aromatic pollutants, such as certain phenols and substituted benzene derivatives, and is therefore often considered a key enzyme in microbial pollutant degradation. Modifying or optimizing its catalytic efficiency could accelerate the removal of recalcitrant organic pollutants.
2. Metabolic Engineering and Synthetic Biology: Due to HpaB's ability to perform targeted hydroxylation on benzene rings, scientists are exploring its integration into synthetic pathways to produce catechol derivatives, catechol compounds, and other high-value-added chemicals. These products can serve as precursors for the synthesis of pharmaceuticals, dyes, or materials.

3. Industrial Biocatalysis: HpaB also exhibits strong catalytic performance in vitro. When combined with suitable electron donor systems, it can be used in green chemical synthesis, replacing some traditional chemical reactions and reducing energy consumption and pollution.

From an evolutionary and ecological perspective, the widespread distribution and functional diversity of HpaB suggest that it plays a crucial role in microbial adaptation to complex ecological environments. HpaB differs in sequence and structure among different bacteria. Furthermore, in-depth analysis of HpaB will help us understand how bacteria utilize limited genetic resources to efficiently utilize diverse carbon sources.

Cultivation

We used E. coli K12 as a template and PCR amplified our target gene HpaB. The gene is 1563 bp long, as shown in Figure 5. Comparison with DNA maker showed that the amplification results were correct. We then purified the target gene by gel recovery. Finally, this fragment was homologously recombined with the other two fragments and the pETDuet backbone to form a complete plasmid.

Figure 5. Electrophoresis results of HpaB gene and HpaC gene

Name: 4-Hydroxyphenylacetate 3-monooxygenase, reductase component (HpaC)

Base Pairs: 513 bp

Origin: Escherichia coli K12

Properties:

4-Hydroxyphenylacetate 3-monooxygenase, reductase component (HpaC) is the electron transfer subunit of a two-component flavin-dependent monooxygenase system, typically acting in conjunction with the oxidase component, HpaB. HpaC's primary function is to transfer reducing equivalents (typically from NADH or NADPH) to HpaB, thereby providing the necessary electrons for aromatic ring hydroxylation. It belongs to the flavin reductase family and binds flavin mononucleotide (FMN) as a cofactor, maintaining the catalytic reaction through a redox cycle.

HpaC has a relatively small molecular weight, typically around 18.5 kDa, making it significantly lighter than HpaB. Its structural features include a typical flavin binding pocket and a conserved electron transfer motif, enabling efficient binding to NAD(P)H and electron donation. Compared to other similar reductases, HpaC's advantage lies in its highly specific interaction with HpaB, ensuring the directionality and efficiency of electron transfer, thereby driving the hydroxylation of aromatic compounds. Although HpaC does not directly catalyze the aromatic ring hydroxylation reaction, it is essential in the overall reaction system. Without HpaC's electron supply, HpaB cannot complete the oxidation reaction, and therefore HpaC is considered the core of this two-component system.

Figure 6. Gene maps of HpaC

Usage and Biology

In a biological context, HpaC functions primarily as an electron donor. It binds NADH/NADPH and transfers electrons to FMN, which then passes the reduced flavin to HpaB, thereby driving the hydroxylation of 4-hydroxyphenylacetic acid (4-HPA). This reaction is a crucial step in the bacterial breakdown of aromatic compounds. The final product, 3,4-dihydroxyphenylacetic acid (homohenoic acid), can be further utilized in metabolic pathways. Therefore, HpaC plays a key role in the bacterial utilization of aromatic compounds as energy and carbon sources.

In research applications, HpaC is often used as a representative electron transfer protein. Its collaboration with HpaB provides a good example for studying the electron transfer mechanism of two-component monooxygenases. By analyzing the crystal structure and kinetic characteristics of HpaC, scientists have revealed the electron transfer pathway from NAD(P)H → FMN → HpaB, providing a deep molecular foundation for understanding flavin-dependent redox reactions.

At the application level, the research and engineering of HpaC have multiple implications:

1. Environmental remediation: Because HpaC-driven HpaB activity can degrade recalcitrant aromatic pollutants, HpaC is considered an essential component of the pollutant biodegradation system. Engineering more efficient HpaC could help enhance the environmental purification capabilities of bacteria.

2. Metabolic engineering: In synthetic biology, modifying the electron transfer efficiency of HpaC and HpaB can improve the conversion efficiency of aromatic compounds, thereby enabling the production of catechol derivatives, catechols, or other high-value chemicals.

3. Industrial biocatalysis: The system formed by combining HpaC and HpaB can be applied to green chemical synthesis, providing specific hydroxylation of benzene ring structures and replacing some traditional high-energy chemical processes. By modifying HpaC, electron supply can be optimized, making the reaction more promising for industrial applications.

At the ecological and evolutionary levels, the existence of HpaC reflects microbial adaptation strategies to complex organic metabolism. HpaC differs in sequence and structure across species, and these variations influence electron transfer efficiency and cofactor dependence. Comparative genomic and functional analyses enable researchers to better understand how microorganisms optimize metabolic networks in diverse environments.

Overall, while HpaC is not directly involved in aromatic ring hydroxylation, as a provider of energy and electrons, it is a key component of the entire HpaB/HpaC two-component system. It is not only important for basic scientific research but also serves as a valuable tool in environmental microbiology, metabolic engineering, and industrial biocatalysis.

Cultivation

We used E. coli K12 as a template to amplify the target gene, HpaC, by PCR. The full length of the gene is 513 bp, as shown in Figure 5. Comparison with DNA Maker confirmed the correct amplification. We then purified the target gene by gel extraction. Finally, this fragment was homologously recombined with two other fragments and the pETDuet backbone to form a complete plasmid.

Name: prephenate/arogenate dehydrogenase (SO clade)

Base Pairs: 1140 bp

Origin: Sulfolobus

Properties:

The prephenate dehydrogenase ( PDH, EC 1.3.1.13 ) encoded by the TyrA ( SO ) gene belongs to the archaeal branch of the TyrA oxidoreductase superfamily that specifically uses prephenate as the preferred substrate. Typically, it is derived from the thermophilic archaea Sulfolobus tokodaii and S.acidocaldarius. The gene encodes about 295 amino acids. The protein is folded into a classic ' Rossmann-fold + NAD ( P ) binding domain ' double domain sandwich structure. The 222nd glycine and the 162nd hydrophobic residue form a relatively loose hydrophobic pocket, so that the C1-carboxyl group and the C4-hydroxyl group of the substrate prephenate can be anchored at the same time, thereby giving the enzyme a strict prephenate dehydrogenation activity and almost rejecting arogenate.

Transcriptome and chromosome collinearity analysis further revealed that TyrA ( SO ) usually forms a compact thermoregulatory operon with AroF ( SO ), TyrC ( SO ) and AroQ ( SO ) encoding chorismate mutase. Its promoter region contains typical TATA-box and CCR1 regulatory elements, which can be activated by Sulfolobus-specific transcription factor Tgr1 under heat shock or carbon restriction conditions, thus ensuring rapid replenishment of aromatic amino acid requirements in high temperature environments.

Figure 7. Gene maps of TyrA(SO)

Usage and Biology

The prephenate dehydrogenase encoded by the TyrA(SO) gene plays a ' carbon flow gate ' role in the cellular metabolic network. Its biological function is not only limited to providing L-tyrosine precursors for protein synthesis, but also coordinating the balance of aromatic amino acids, quinone cofactors and various secondary metabolites by regulating the redox state of the prephenate node. In the natural habitat of the thermophilic archaeon Sulfolobus, temperature fluctuations and low pH environments lead to a significant increase in the demand for protein and nucleic acid repair. TyrA(SO) rapidly introduces NADPH generated by glycolysis and pentose phosphate pathway into the prephenate oxidation step through a heat-inducible operon structure. On the one hand, it supplements the tyrosine pool to support the synthesis of heat shock proteins and cell wall polymers. On the other hand, by increasing the intracellular concentration of 4-hydroxyphenylpyruvate (4-HPP), it provides sufficient precursors for the synthesis of downstream electron carriers such as ubiquinone and plastid quinone. So as to maintain the stability of the respiratory chain at high temperature.

Recent isotope tracing experiments have further revealed that the oxidative decarboxylation step catalyzed by TyrA(SO) has an ' irreversible ' characteristic in carbon flow distribution, so that prephenate is directed to the tyrosine pathway once it enters the channel, avoiding carbon competition with the phenylalanine or tryptophan synthesis pathway. This ' one-way valve ' effect is decisive for cell growth in an extreme environment with limited carbon sources.

Cultivation

The pETDuet1 plasmid was provided by the strain library of our unit, and the target genes TyrA(SO), HpaB and HpaC were all fragments amplified from the genome using primers using Escherichia coli K12 as templates. The three target genes were constructed together with the vector using the homologous recombination method.

Name: prephenate/arogenate dehydrogenase (HI clade)

Base Pairs: 1140 bp

Origin: Haloarchaea

Properties:

The TyrA(HI) gene was cloned from the tyrosine operon center of the extreme halophilic archaea Halobacterium salinarum and Haloferax volcanii, encoding a prephenate / arogenate dehydrogenase of about 303 amino acids. The sequence extends a 'salt adaptation plug-in 'rich in acidic residues ( D / E ) and cysteine at the N-terminus, which can maintain the rigid structure of the catalytic domain through surface negative charge shielding and dynamic disulfide bond exchange in 2-4 M KCl or NaCl environment. The enzyme showed a strict NAD preference for coenzymes, KM(NAD+)≈22 μM, KM( NADP+) > 1 mM, which matched the redox environment with a high NAD+/NADH ratio in halophilic archaea. More importantly, the C-terminal tail of TyrA(HI) contains a histidine-proline-rich HPH repeat, which can partially close the active pocket by conformational contraction when the intracellular tyrosine concentration increases to 3 mM, achieving a reversible weak feedback inhibition, which not only avoids excessive consumption of carbon flow, but also ensures the continuous supply of aromatic amino acids under extreme osmotic pressure.

Although TyrA(SO) and TyrA(HI) belong to the TyrA family, there are significant differences in substrate specificity, coenzyme preference and environmental adaptability. TyrA(SO) is more suitable for conventional metabolic engineering, while TyrA(HI) has potential in extreme environmental bio-manufacturing. Both of them are important models for studying the evolution and functional diversity of TyrA enzyme.

Figure 8. Gene maps of TyrA(HI)

Usage and Biology

TyrA ( HI ) acts as a ' salt-adapted aromatic carbon flow switch ' in halophilic archaea. Its enzymatic dimorphism allows cells to preferentially oxidize arogenate to tyrosine under 3 M NaCl, and simultaneously release NADH, providing reducing power and 4-HPP precursor for the synthesis of purpurin, and consuming intracellular reactive oxygen species through the tyrosine-quinone-melanin branch to maintain redox and osmotic balance in a strong light, hypertonic, and hypoxic salt lake environment. The weak feedback mechanism of its HPH tail ensures that the carbon flow rebounds rapidly after stress relief, supporting protein repair and compatible solute resynthesis.

With the characteristics of high salt-high enzyme activity, NAD+ preference and reversible self-inhibition, the gene has been implanted into Halomonas to construct ' Salt Lake Factory ', and L-DOPA was continuously produced in 3 M NaCl. The 150-column volume of the cell-free immobilization system still retained 80 % activity. Its tail was transformed into a high-salt tyrosine sensor after fluorescence modification, coupled with a light-violet membrane regeneration module to realize seawater-based photoenzymatic co-production of aromatic chemicals, providing core catalytic and regulatory elements for extreme environmental biomanufacturing and direct seawater fermentation.

Cultivation

The pETDuet1 plasmid was provided by the strain library of our unit, and the target genes TyrA(SO) and TyrA(HI) were all fragments amplified from the genome using primers using Escherichia coli K12 as templates. The target genes were constructed together with the vector using the homologous recombination method.

Composition: pETDuet1 backbone; TyrA(K12)-HpaB-HpaC gene fragment.

Apparatus used: pETDuet1 plasmid, TyrA(K12)-HpaB-HpaC gene fragment, Homologous recombination enzyme.

Figure 9. Plasmid map of pETDuet1-TyrA(K12)-HpaB-HpaC

Engineering Principle:

The pETDuet1 plasmid was provided by the strain library of our unit, and the target genes TyrA(K12), HpaB and HpaC were all fragments amplified from the genome using primers using Escherichia coli K12 as templates. The three target genes were constructed together with the vector using the homologous recombination method.We used E.coli K12 as a template to amplify other target fragments. Firstly, the corresponding primers were designed, and then the three target genes of TyrA, hpaB and hpaC were amplified by using the cultured K12 strain as a template. The plasmid backbone was also obtained by reverse PCR. After the amplification reaction was completed, the product was identified by agarose gel electrophoresis. The results showed that the plasmid backbone and the three target genes were successfully amplified. The backbone length was 5420 bp, the TyrA gene was 1122 bp, the hpaB gene was 1563 bp, and the hpaC gene was 513 bp. The specific data are shown in Figure 10.

Figure 10. Electrophoresis results of pETDuet and TyrA(K12), hpaB and hpaC

After obtaining the correct size of the gene fragment, we need to obtain the purified vector and fragment by gel recovery technology. Subsequently, a complete recombinant plasmid was constructed by homologous recombination technology and transformed into the clone strain. After overnight culture, clones with good growth were selected for colony PCR identification. As shown in Figure 11, we selected six monoclonals and used the designed primers for amplification. The results showed that the length of the amplified product was 1500 bp, which was consistent with the expected length, indicating that the transformation process was successful.

Figure 11. Results of monoclonal plates and colony PCR after transformation

Finally, we inoculated the successful transformants into seed solution, cultured them, extracted the plasmids, and sent them to a biotechnology company for sequencing. The results are shown in Figure 12. The sequencing results show that the constructed recombinant plasmids have no base mutations or deletions.

Figure 12. Plasmid sequencing results

Cultivation, Purification and SDS-PAGE:

Before the first round of fermentation experiments, we performed protein expression tests on two plasmids ( pCDFDuet-tktA-aroF-aroE and pETDuet-TyrA ( K12 ) -hpaB-hpaC ) to verify the expression of the six target genes in E.coli BL21 ( DE3 ). The rapid protocol was 37 ℃, 220 rpm, 1 mM IPTG induction for 6 h. The results of SDS-PAGE showed that all the target bands were visible in 4 supernatants and 1 precipitate of plasmid 1 and 3 supernatants and 2 precipitate of plasmid 2, which confirmed that the six proteins were successfully expressed. Part of the protein of plasmid 2 exists in the form of inclusion bodies, which may be related to the collection operation. In view of the fact that we are strengthening the levodopa synthesis pathway, this expression has met expectations and can continue to promote subsequent fermentation experiments ( Figure 13 ).

Figure 13. Target protein expression test using plasmids 1 ( pCDFDuet-tktA-aroF-aroE ) and 2 ( pETDuet-TyrA(K12)-hpaB-hpaC ). Wells 1-4 contain the supernatant from plasmid 1, and well 5 contains the precipitate from plasmid 1. Wells 6-8 contain the supernatant from plasmid 2, and wells 9-10 contain the precipitate from plasmid 2.

Composition: pETDuet1 backbone; TyrA(SO)-HpaB-HpaC gene fragment.

Apparatus used: pETDuet1 plasmid, TyrA(SO)-HpaB-HpaC gene fragment, Homologous recombination enzyme.

Figure 14. Plasmid map of pETDuet1-TyrA(SO)-HpaB-HpaC

Engineering Principle:

The pETDuet1 plasmid was provided by the strain library of our unit, and the target genes TyrA(SO) is synthesized by a biological company, HpaB and HpaC were all fragments amplified from the genome using primers using Escherichia coli K12 as templates. The three target genes were constructed together with the vector using the homologous recombination method.

we constructed the pETDuet-TyrA(SO)-hpaB-hpaC plasmid. Compared with pETDuet-TyrA(K12)-hpaB-hpaC, the difference between the two plasmids is the source of TyrA. Therefore, we used homologous recombination to replace TyrA based on pETDuet-TyrA(K12)-hpaB-hpaC. We designed two pairs of primers, one to amplify the TyrA gene from the new source, and the other pair of primers to amplify the rest of the pETDuet-TyrA(K12)-hpaB-hpaC except TyrA(K12). After amplification, agarose gel electrophoresis was performed for identification, as shown in Figure 15A. As shown in Figure 15B , the plasmid backbone and TyrA(SO) amplification results after electrophoresis were consistent with the expected lengths: the plasmid backbone was 7535 bp long, and TyrA(SO) was 1152 bp long. The two fragments were then recovered and purified from the gel. After homologous recombination, the recombinant products were transformed into E. coli TOP10. As shown in Figure 15B , single clones emerged after overnight growth. To verify the correctness of the transformed single clones, we performed colony PCR identification. Twelve single clones were selected for identification. In Figure 15C , the colony PCR result showed a size of 2100 bp, demonstrating that the transformed single clones all contained the correct plasmid. The correct single clones were then inoculated into seed solution, and the plasmids were extracted and sent to a biotechnology company for sequencing. Sequencing results confirmed that the constructed plasmid was successful.

Figure 15 A. Electrophoresis results of plasmid backbone and target gene fragments, B. Plate of recombinant plasmid transformed into E. coli, C. Electrophoresis results of monoclonal colony after PCR, D. Sequencing results of recombinant plasmid

Composition: pETDuet1 backbone; TyrA(HI)-HpaB-HpaC gene fragment.

Apparatus used: pETDuet1 plasmid, TyrA(HI)-HpaB-HpaC gene fragment, Homologous recombination enzyme.

Figure 16. Plasmid map of pETDuet1-TyrA(HI)-HpaB-HpaC

Engineering Principle:

The pETDuet1 plasmid was provided by the strain library of our unit, and the target genes TyrA(HI) is synthesized by a biological company, HpaB and HpaC were all fragments amplified from the genome using primers using Escherichia coli K12 as templates. The three target genes were constructed together with the vector using the homologous recombination method.

When constructing the pETDuet-TyrA(HI) -hpaB-hpaC vector, we adopted a similar strategy as pETDuet-TyrA(SO) -hpaB-hpaC. First, primers were designed to amplify the TyrA(HI) target gene, and a pair of primers were designed to amplify the backbone DNA. After the amplification was completed, the amplification effect was verified by agarose gel electrophoresis. As shown in Figure 17A, the length of the amplified vector and the target gene is consistent with the expected value, of which the skeleton DNA is 7535 bp and the target gene is 1146 bp. Subsequently, the two were ligated by homologous recombination and transformed into the corresponding cloned E.coli strains. After overnight culture (as shown in Figure 17B), 12 single colonies were screened for cloning PCR analysis to confirm the success of the transformants.

Figure 17 A. Plasmid backbone and target gene electrophoresis results. B.plate image of the transformed single colonies.

After screening the corresponding monoclonal colonies for PCR amplification, we analyzed the amplification products by electrophoresis. As shown in Figure 18A, the colony PCR results are in line with the expected 2000 bp length. Subsequently, we selected the monoclones that successfully obtained the colony PCR to be inoculated into the seed solution, and the plasmids were extracted and sent to the biotechnology company for sequencing. Figure 18B shows the sequencing results. The results show that the recombinant plasmid was successfully constructed, and no base mutation or deletion was found.

Figure 18 A. Plasmid backbone and target gene electrophoresis results. B.plate image of the transformed single colonies.

Composition: pETDuet1 backbone; TktA-AroF-AroE gene fragment.

Apparatus used: pETDuet1 plasmid, TktA-AroF-AroE gene fragment, Homologous recombination enzyme.

Figure 19. Plasmid map of pCDFDuet1-TktA-AroF-AroE

Engineering Principle:

The pCDFDuet1 plasmid was provided by the strain library of our unit, and the target genes TktA、AroF and AroE were all fragments amplified from the genome using primers using Escherichia coli K12 as templates. The three target genes were constructed together with the vector using the homologous recombination method.

We used homologous recombination to construct the pCDFDuet-tktA-aroF-aroE plasmid. First, using PCR technology, we designed corresponding primers and amplified the target fragments of tktA, aroF, and aroE using Escherichia coli K12 as a template. We also amplified the pCDFDuet plasmid backbone using a laboratory-preserved plasmid. As we can see from Figure 20A, the agarose gel electrophoresis results after amplification showed that the amplified lengths of the backbone and target fragments were all in line with the expected sizes. The pCDFDuet plasmid backbone is 3781 bp in length, tktA is 1995 bp in length, aroF is 1071 bp in length, and aroE is 819 bp in length.

Figure 20. A. Electrophoresis results of plasmid backbone and target fragment, B. Recombinant plasmid transformed into cloning strain TOP10, C. Electrophoresis results after single clone colony PCR, D. Plasmid sequencing results

After obtaining the empty vector and the corresponding length of the fragment, we recovered the agarose gel to obtain the purified vector and fragment, and determined the concentration of these gene fragments by a nanotiter. After obtaining the target concentration, we used homologous recombination technology to recombine the vector and fragment. Subsequently, the recombinant product was transformed into E.coli TOP10 clone strain. As shown in Figure 20B, after overnight culture after transformation, we observed single colonies that were successfully transformed.

However, whether the transformation is successful or not, it needs to be verified accordingly. First, we randomly selected four independent single colonies of the same size in the culture dish through colony PCR technology, and identified them using colony PCR primers. The designed amplified fragment length was 2102 bp (Figure 20C). After PCR amplification and agarose gel electrophoresis identification, it was found that the size of the amplified fragment was in line with expectations, so it was confirmed that these monoclonal clones had been successfully transformed into recombinant plasmids. Finally, it is necessary to amplify the correct clone, extract the plasmid, and send it to the biological sequencing company for sequencing. This step is designed to confirm the correct position and order of the plasmid and the ligase, and to detect whether there is a base deletion or mutation. As shown in Figure 20D, our construction results are fully in line with expectations.

Composition: pETDuet1 backbone;TyrA(K12)-RBS2-HpaB-HpaC gene fragment.

Apparatus used: pETDuet1 plasmid, TyrA(K12)-RBS2-HpaB-HpaC gene fragment, Homologous recombination enzyme.

Figure 21. Plasmid map of pETDuet1-TyrA(K12)-RBS2-HpaB-HpaC

Engineering Principle:

The construction of pETDuet1-TyrA(K12)-RBS2-HpaB-HpaC is based on pETDuet1-TyrA(K12)-HpaB-HpaC. It only needs to design the new RBS sequence on the primer and amplify it with the original plasmid as the template. The completed plasmid is pETDuet1-TyrA(K12)-RBS2-HpaB-HpaC with the RBS sequence replaced. We then sequenced the plasmid to ensure that there was no deletion or mutation at the replaced position.

To replace the RBS at the start of the TyrA gene in the pETDuet-TyrA(K12)-hpaB-hpaC plasmid, we used inverse PCR. We designed a pair of primers targeting the desired RBS sequence and amplified the plasmid using the original pETDuet-TyrA(K12)-hpaB-hpaC plasmid as a template. Compared to the original plasmid, only the RBS sequence was altered in the amplified plasmid. Because the newly amplified plasmid was essentially the same length as the original, we avoided agarose gel electrophoresis and instead performed transformation after amplification and extracted the plasmid for sequencing. The sequencing results in Figure 23 demonstrate the successful replacement of the new RBS sequence. The sequence on the left represents the replacement of RBS2.

Composition: pETDuet1 backbone;TyrA(K12)-RBS3-HpaB-HpaC gene fragment.

Apparatus used: pETDuet1 plasmid, TyrA(K12)-RBS3-HpaB-HpaC gene fragment, Homologous recombination enzyme.

Figure 22. Plasmid map of pETDuet1-TyrA(K12)-RBS3-HpaB-HpaC

Engineering Principle:

The construction of pETDuet1-TyrA(K12)-RBS3-HpaB-HpaC is based on pETDuet1-TyrA(K12)-HpaB-HpaC. It only needs to design the new RBS sequence on the primer and amplify it with the original plasmid as the template. The completed plasmid is pETDuet1-TyrA(K12)-RBS2-HpaB-HpaC with the RBS sequence replaced. We then sequenced the plasmid to ensure that there was no deletion or mutation at the replaced position.The construction of this step is consistent with the method of replacing RBS2，The sequencing results in Figure 23 demonstrate the successful replacement of the new RBS sequence. The sequence on the right represents the replacement of RBS3.

Figure 23. Sequencing results after replacing the RBS sequence

Functional Test

1. Target protein expression test

After the construction of plasmid 1 and three TyrA plasmids 2, two of them were transformed into new E.coli expression strains. Since the production of L-DOPA requires six genes to function simultaneously, we first transformed two plasmids into E.coli BL21 ( DE3 ) (Figure 24). After overnight culture, we observed that the double-transformed host bacteria showed corresponding monoclonal colony growth. Next, we will verify whether the double plasmid transformation is completed by colony PCR.

Figure 24. The corresponding single colony plate image after plasmid 1 and plasmid 2 were transformed into BL21 (DE3) together

After the completion of the double plasmid transformation, in order to verify whether the two plasmids exist at the same time, we designed specific primers for each plasmid and performed colony PCR on the monoclonal of three candidate strains ( strain 1-3 ). The expected band length : strain 1 ( plasmid 1 1000 bp, plasmid 2 800 bp ), strain 2 ( plasmid 1 800 bp, plasmid 2 1000 bp ), strain 3 ( both plasmids are 1000 bp ). The results of agarose gel electrophoresis showed that the size of all amplified bands was completely consistent with the theoretical value (Figure 25), which confirmed that the double plasmids had been successfully co-transformed and the experiment was successfully completed.

Figure 25. Electrophoresis of colony PCR results of three strains after double plasmid transformation

Before the first round of fermentation, the protein expression of the two plasmids was tested to verify whether the six genes could be normally translated in BL21 (DE3). After the transformation of pCDFDuet-tktA-aroF-aroE (plasmid 1) and pETDuet-TyrA (K12) -hpaB-hpaC (plasmid 2), the rapid induction scheme was adopted : 37 ℃, 220 rpm, 1 mM IPTG, induction for 6 h, and immediate sampling and detection.

After induction, we collected protein samples and divided them into supernatant and crude protein precipitation, and verified the expression effect of the target gene by SDS-PAGE detection. As shown in Figure 13, wells 1-4 were the supernatant of plasmid 1, well 5 was the precipitate of plasmid 1, wells 6-8 were the supernatant of plasmid 2, and wells 9-10 were the precipitate of plasmid 2. Plasmid 2 contained inclusion bodies, and all the proteins corresponding to the target genes were successfully expressed. Although the target protein corresponding to plasmid 2 has inclusion bodies during the collection process, this may be related to the protein collection operation steps. In addition, since we are enhancing the intracellular levodopa biosynthesis pathway, it is confirmed that the expression has reached the expected goal.

1. Detection of L-dopa production using an ELISA test kit

After confirming the correct expression of the target protein, we then carried out 48 h parallel fermentation of strains carrying three different sources of tyrosinase A (TyrA) to screen the optimal strain for high-yield levodopa (L-DOPA). The fermentation parameters were consistent with the protein expression stage : 37 ℃, 220 rpm, 1 mM IPTG induction, and 3 biological replicates per strain (Figure 26A). After the fermentation broth was pretreated (Figure 26B-D), the concentration of the product was determined by L-DOPA enzyme-linked immunosorbent assay ( ELISA ) kit. After the reaction was terminated, the absorbance was read at 450 nm ( Figure 26E ), and the data were directly used for yield comparison and strain evaluation.

Figure 26. A. Fermentation test of strain 123 (three replicates), B. Sample preparation, C. Reaction, D. End of reaction, E. Absorbance test

We first plotted the standard curve ( as shown in Figure 27A ). The content of levodopa in the sample was then calculated according to the standard curve and absorbance data. As shown in Figure 27B, strain 1 had the highest levodopa production. This indicates that tyrosinase A derived from K12 exhibits better enzyme activity in the modified strain. For this modified strain, the genes from related species have stronger adaptability, so we selected this gene for subsequent RBS screening.

Figure 27. A. Standard curve of the ELISA test kit; B. L-DOPA production by three strains, where Strain 1 contains TyrA from K12, Strain 2 contains TyrA from SO, and Strain 3 contains TyrA from HI.

After the RBS sequence replacement was completed, fermentation testing was repeated to determine which RBS sequence performed best. Three strains were inoculated into LBG (20 g/L glucose) medium at an initial OD of 0.05. After fermentation, product testing was performed to identify the optimal RBS sequence for the TyrA gene and improve expression of the key enzyme. The fermentation testing steps were similar to those described above. Three replicates of different RBS strains were tested, as shown in Figure 28A. The assays were performed using a kit, with the reaction progressing and reaction completion times (Figure 28BC). Finally, the fermentation yields were calculated using a standard curve.

Figure 28. A Fermentation test of different RBS strains (three replicates), BC is divided into reaction progress and reaction completion

By substituting the absorbance value into the standard curve for calculation, we can obtain the L-dopa content produced by the strain under different RBS sequences. According to the histogram in Figure 29, three different RBS sequences all have a certain effect on gene expression. Among the RBS sequences we screened, the optimal expression level increased the strain yield by 16.4 % to 500.3 mg /L.

Figure 29. The yield of levodopa corresponding to different RBS sequences

The optimal TyrA source (K12) and RBS sequence (TCACACAGG) were locked. Subsequently, we systematically investigated the effect of initial glucose concentration on the synthesis of L-DOPA in BL21 (DE3). Five gradients of 10,20,30,40 and 50 g/L were set, and the remaining fermentation conditions were consistent with the previous stage. Figure 30 showed that the yield increased with the increase of sugar concentration, and reached a peak of 689.3 mg/L at 40 g/L ; continue to increase to 50 g/L, the product decreased, suggesting that high glucose load disrupted intracellular metabolic flow. In the next step, new variables such as IPTG concentration and induction time will be introduced to continue to explore the key factors limiting high yield.

Figure 30. Analysis of results at different initial sugar concentrations (optimal RBS sequence strain)

In order to further explore the production factors of levodopa, we simultaneously investigated the three key parameters of inducer concentration, glucose concentration and fermentation time. The concentration of glucose was set to 5 concentration gradients of 10-50 g/L, and the concentration of inducer was set to 6 concentration gradients of 0,0.2,0.4,0.6,0.8 and 1.0 mM. The fermentation time was set to 36 hours and 48 hours, respectively. After the strain was inoculated in LBG medium with an initial OD value of 0.05, the product was detected and analyzed after the fermentation was completed.

Figure 31. Effects of inducer concentration and glucose concentration on levodopa production at different times

Figure 31 shows the effects of inducer concentration and glucose concentration on levodopa production at different times. The overall trend is that when the inducer concentration is 0.8 mM and the glucose concentration is 40 g/L, the levodopa production is higher and significantly higher. When the inducer concentration is 0.2 mM, the production is higher without adding the inducer concentration than after adding it. This may be due to contamination during fermentation. In addition, by comparing the overall situation at 36 h and 48 h, we can see that the levodopa production at 48 h is higher.

The heat map in Figure 32 is a correlation map, and the response surface also tells us a trend. Finding the optimal inducer concentration and glucose concentration at different time points is consistent with the conclusion of the above histogram. We can see that the optimal glucose concentration is 40 g/L, and the inducer concentration is 0.8 mM.

Figure 32. Optimal inducer concentration and glucose concentration at different time points

Finally, we conducted a joint analysis of inducer concentration, glucose concentration and time. In the three-dimensional results of Figure 33, it can be seen that the fermentation effect of 0.8-1.0 mM, 40g / L, 36-48 h is better. Finally, we chose 0.8 mM, 40 g/L, 48 h conditions for fermentation of E.coli ECN.

Figure 33. Effects of different inducer concentrations, glucose concentrations, and time on levodopa

After determining the optimal IPTG concentration, glucose concentration and induction time in BL21(DE3), we co-transformed plasmid 1 and optimized plasmid 2 into the probiotic strain E.coli ECN to achieve L-DOPA probiotic production. Figure 34 shows that the double plasmids were successfully transformed. The 800 bp target band was obtained by colony PCR, which further verified that the transformation was correct. Subsequently, the correct monoclonal cells were selected for expansion and induced fermentation to detect the L-DOPA yield of ECN.

Figure 34. Plasmid 1 and plasmid 2 were transformed into E. coli ECN (left), and the electrophoresis results of colony PCR identification were shown (right).

1. Detection of L-dopa production using HPLC technology

Subsequently, the optimized culture conditions in BL21(DE3) were directly translated to ECN for L-DOPA fermentation verification. After fermentation, three parallel samples were quantitatively detected by HPLC. The chromatographic peaks were shown in Figure 35. The results of the three determinations were 1.064,1.065 and 1.069 g/L, with an average of 1.066 g/L, and the reproducibility was good. It was confirmed that ECN could still stably produce levodopa in the context of probiotics.

Figure 35. Peak diagram of levodopa production after HPLC detection of ECN fermentation