Overview
We designed a green strategy for producing PA54 monomers (cadaverine and succinate) using a single cell. The cell grows and accumulates lysine under aerobic conditions; when switching to anaerobic conditions, it activates the expression of lysine decarboxylase to catalyze the synthesis of cadaverine from lysine, while simultaneously synthesizing succinate. This enables synchronous acid-base neutralization during the synthesis of cadaverine and succinate, and the CO2 released from decarboxylation is fixed for succinate synthesis. We selected the lysine-producing strain Escherichia coli NT1003 as the chassis cell. Firstly, we constructed and screened the switch circuit to achieve regulatable expression of lysine decarboxylase (CadA) for catalyzing lysine. The switch circuit was further optimized to enhance its accuracy by optimizing the promoters and integrating the RNA thermometers. Subsequently, based on AI-based design and rational design, we performed rational engineering on the key enzyme CadA to improve its catalytic efficiency. Additionally, we adopted cofactor regulation strategy combined the modification site identified by metabolic network model to enhance succinate synthesis. Ultimately, we successfully achieved simultaneous production of cadaverine and succinate from glucose using a single cell.
Module 1: Switch---Controllable expression of CadA
Cycle 1 Constructing a switch genetic circuit to dynamically regulate CadA for controllable synthesis of cadaverine
Design
In E.coli NT1003, lysine is a product of aerobic fermentation, while succinate only accumulates during anaerobic processes. To realize the concept of simultaneous production of cadaverine and succinate(Figure 1-1A), we first designed a switch genetic circuit that allowed lysine accumulation without the activity of lysine decarboxylase CadA during the aerobic growth phase, and activated CadA to exert its catalytic activity for converting lysine into cadaverine during anaerobic phase.
After literature research and expert interviews, we ultimately selected two switch systems for regulatory testing: the temperature-responsive system and the protein encapsulation system.
Temperature-responsive system:
This system was constructed using a temperature-sensitive switch. The temperature-responsive repressor protein TlpA39 is expressed under the control of the constitutive promoter J23100. When the temperature is below 39℃, TlpA39 forms a dimer that can bind to the promoter PtlpA, thereby repressing the expression of the downstream cadA gene. At temperatures exceeding 39°C, TlpA39 monomerizes and dissociates from PtlpA, switching the circuit from an OFF to an ON state and enabling cadA expression. This design ensures that CadA is suppressed during low-temperature aerobic phase to permite lysine accumulation. and CadA expression is activated by a temperature upshift during the anaerobic phase to drive the conversion of lysine to cadaverine (Figure 1-1B).
Protein encapsulation system:
This system uses an artificial intrinsically disordered protein (A-IDP), which encapsulates the CadA under aerobic conditions to inhibit its activity, thereby preventing lysine from being catalytically consumed. Under anaerobic conditions, specific proteases (TEV or TVMV protease) cleave the corresponding sites (TEV-site or TVMV-site), separating A-IDP from CadA and releasing CadA from encapsulation. This proteolytic cleavage releases CadA from encapsulation to enable CadA to catalyze lysine into cadaverine, representing the switch from OFF to ON ( Figure 1-1C).
Figure 1-1 (A) Metabolic diagram of E. coli NT1003 for PA54 production. (B) Mechanism diagram of the temperature-responsive system. (C) Mechanism diagram of the protein encapsulation system.
Build
To construct the temperature-responsive system, we first designed a switch genetic circuit composed of the parts, including J23100, RBS, TlpA39, and PtlpA. The J23100-RBS-TlpA39-PtlpA sequence was commercially synthesized and cloned into the pET28a(ΔlacI7) vector. Subsequently, the genes encoding green fluorescent protein (mWasabi) and lysine decarboxylase (CadA) were individually cloned in downstream of the PtlpA promoter to assemble the complete genetic circuits: PJ23100-RBS-TlpA39-PtlpA-RBS-mWasabi and PJ23100-RBS-TlpA39-PtlpA-RBS-CadA.
Figure 1-2 The PCR validation diagram. M: Marker. Lane 1,2:cadA fragment. Lane 3, 4:mWasabi fragment.
To construct the protein encapsulation system, we first conducted literature research and then commissioned the synthesis of the 0.5 and 1.0 size A-IDP gene fragments along with the corresponding TEV and TVMV protease cleavage sites. Subsequently, the genes encoding green fluorescent protein (mWasabi) and lysine decarboxylase (CadA) were individually cloned in upstream of A-IDP to construct the genetic circuits, including mWasabi-TEV-site-0.5A-IDP, mWasabi-TEV-site-1.0A-IDP, CadA-TEV-site-0.5A-IDP, CadA-TEV-site-1.0A-IDP, CadA-TVMV-site-0.5A-IDP, and CadA-TVMV-site-1.0A-IDP. They were inserted into the pCDFDuet-Trc vector to obtain recombinant plasmids.
Figure1-3 (A) Genetic circuit diagram.(B) The colony PCR validation diagram. M: Marker. line l~9:mWasabi-TEV-0.5A-IDP. line 10~20: mWasabi-TEV-1.0A-IDP.
All recombinant plasmids were first transformed into E. coli DH5α. Positive clones identified by colony PCR (based on the correct band size) were subjected to commercial sequencing. Sequencing results confirmed the successful construction of the plasmids. These plasmids were then transformed into the E. coli NT1003 strain for functional characterization.
Test
For the functional assessment of the temperature-responsive system, we selected low temperature (33℃) and high temperature (39℃) as test conditions. The green fluorescent protein group measured fluorescence intensity at 6, 10, 24, and 34 h, as illustrated in Figure 1-4 A and B. Fluorescence intensity was consistently low at 33°C but exhibited a marked increase at 39°C. After 24 hours, the signal at 39°C was 2.56-fold higher than that at 33°C.
To further investigate the function on CadA expression, lysine and glucose of the CadA-expressing E. coli were analyzed using a biosensor analyzer at 0, 6, 10, and 24 h under 33℃ and 39℃. At the inducing temperature (39°C), lysine was rapidly consumed, maintaining low levels, whereas at 33°C, it steadily accumulated, confirming the thermal control of CadA (Figure 1-4 C and D).
Figure 1-4 Fluorescence intensity of the engineered strain E. coli NT1003 .(A) mWasabi (33℃). (B) mWasabi (39℃) Glucose consumption and lysine accumulation of the engineered strain E. coli NT1003. (C) CadA (33℃). (D) CadA (39℃).
To evaluate the protein encapsulation system, we first analyzed the fluorescence signal of recombinant strains expressing the green fluorescent protein mWasabi at 0, 8, and 21 h. Both the 0.5A-IDP and 1.0A-IDP groups produced detectable signals, with the 1.0A-IDP group showing significantly lower intensity than the 0.5A-IDP group. In CadA-expressing strains, both A-IDP groups completely suppressed lysine accumulation despite normal glucose consumption, unlike the empty plasmid control where lysine steadily increased.
Figure 1-5 (A) Glucose consumption and lysine accumulation of the engineered E. coli (CadA-0.5A-IDP) and (CadA-1.0A-IDP). (B)Fluorescence intensity of the engineered E. coli (mWasabi-0.5A-IDP) and (mWasabi-1.0A-IDP).
Learn
The simultaneous synthesis of cadaverine and succinate in a single cell constitutes a green and low-carbon process. To achieve the process, we learned that different switchs could be employed to construct regulatory systems for regulating the key enzyme CadA. In this cycle, we successfully constructed the Temperature-responsive system and the Protein encapsulation system.
In temperature-responsive system, our temperature-sensitive switch successfully regulated the timed expression of CadA, as evidenced by contrasting phenotypes of gene repression (low fluorescence, lysine accumulation) at 33°C and gene activation (high fluorescence, lysine consumption) at 39°C. This functional shift is mediated by the temperature-dependent binding of TlpA39 to the PtlpA promoter, which blocks transcription at low temperatures and releases it upon thermal induction.
The current protein encapsulation system failed to prevent lysine consumption by CadA in both 0.5A-IDP and 1.0A-IDP, suggesting insufficient enzyme encapsulation. The fluorescent result showed 1.0A-IDP conferred significantly greater repression than 0.5A-IDP.
Following Professor Liming Liu's suggestion, CadA may require larger A-IDP constructs for effective encapsulation and shielding, Future work will focus on engineering larger A-IDP variants to achieve complete enzyme encapsulation.
Cycle 2 Optimization of protein encapsulation system
Design
According to Prof. Liu's suggestion, 0.5 A-IDP and 1.0 A-IDP are relatively small with insufficient encapsulation efficiency. Therefore, we constructed the larger 2.0 A-IDP using 1.0 A-IDP as a template.
Figure 2-1 Construction process of pCDFDuet-Ptrc-CadA-TEV/TVMV-site-2.0A-IDP.
Build
We used the pCDFDuet-Trc-CadA-TEV-site-1.0A-IDP plasmid as a template to amplify the 1.0A-IDP gene fragment via PCR. The amplified product was inserted into pCDFDuet-Trc-CadA-TEV/TVMV-site-1.0A-IDP between Kpn I and Sal I, constructing the recombinant plasmid pCDFDuet-Trc-CadA-TEV-site-2.0A-IDP (Figure 2-2A). After transforming into E. coli DH5α, positive clones were screened by colony PCR, and then sent for sequencing verification.
Figure 2-2 (A) Genetic circuit diagram containing 2.0A-IDP. (B) colony PCR.M:Marker, Lane 1-24: products containing 2.0A-IDP.
Test
Figure2-3 Glucose consumption and lysine accumulation in the 2.0A-IDP group.
The recombinant plasmid was then transformed into the E. coli NT1003 strain for functional characterization. A SBA biosensor was used to determine glucose and lysine at 0 and 26 h. The results showed that glucose was consumed normally, but lysine was still not detected.
Learn
In this cycle, we learned that larger encapsulating proteins titer better encapsulation efficiency. However, a larger-sized encapsulating protein 2.0A-IDP failed to perfectly encapsulate CadA and inhibit its activity. Compared with the temperature-responsive system in lysine accumulation, we decided to adopt the temperature-sensitive switch for subsequent experiments.
Cycle3 The temperature-responsive characterization of temperature-responsive system
Design
By comparision, the Temperature-responsive system was adopted to control the expression of CadA. To quantify its regulatory precision, we then profiled the system's induction response across a spectrum of temperatures from 33 to 42°C.
Build
pET28a(ΔI7)-PJ23100-RBS-TlpA39-PtlpA-mWasabi and pET28a(ΔI7)-PJ23100-RBS-TlpA39-PtlpA-CadA were respectively transformed into E. coli NT1003 and cultured aerobically at 33, 35, 37, 39, and 42℃.
Figure 3-1 Strategy diagram
Test
The two engineered strains were aerobically cultured at different temperatures. Samples collected at 0, 6, 10, and 23 h were analyzed using a Thermo fluorescence analyzer and a biosensor to determine fluorescence intensity(reporter for mWasabi expression), lysine, and glucose, respectively. The mWasabi reporter strain showed low fluorescence at ≤37°C but strong induction at ≥39°C (Figure 3-2A). For the CadA group, lysine accumulated to 4.27 g/L, 2.6 g/L, and 2.4 g/L at 33°C, 35°C, and 37°C, respectively (Figure 3-2 B, C, D), but was not detected at 39°C and 42°C (Figure 3-2 E, F).
Figure 3-2 (A) Fluorescence detection diagram; Glucose consumption and lysine accumulation of the engineered E. coli strain. (B) CadA (33℃). (C) CadA (35℃). (D) CadA (37℃). (E) CadA (39℃). (F) CadA (42℃).
Learn
In this cycle, we learned that the temperature-responsive system was a clear temperature-dependent switch. Low fluorescence with lysine accumulation at ≤37°C confirmed the "OFF" state, while high fluorescence with no lysine accumulation at ≥39°C confirmed the "ON" state. It defined a clear activation threshold near 39°C, successfully enabling the timed induction of downstream gene through a simple temperature shift.
Cycle 4 Screening different intensity promoters to reduce leaky expression of CadA
Design
From Cycle 3, we found that lysine accumulation gradually decreased from 33°C to 37°C. Professor Guannan Liu hypothesized that insufficient repression stringency resulted in leaky expression of cadA and pointed out that the promoter strength affected the stringency and sensitivity. To further enhance the effect of TlpA39, we selected promoters of different strengths (BBa_J23106 and BBa_J23119) from the Part Registry and compared them with the original promoter (BBa_J23100). By varying TlpA39 expression via these promoters, we intend to adjust its repression capacity at PtlpA and thus improve control of CadA expression..
Figure 4-1 Promoter replacement mechanism diagram
Build
Based on the plasmid of pET28a (ΔI7)-PJ23100-RBS-TlpA39-PtlpA-CadA, we designed promoter mutation primers and cloned the whole plasmid using PCR. The linearized plasmid was isolated and purified by agarose gel electrophoresis, and then transferred into E. coli DH5α to obtain the targeted plasmids of pET28a (ΔI7)-PJ23106-RBS-TlpA39-PtlpA-CadA and pET28a (ΔI7)-PJ23119-RBS-TlpA39-PtlpA-CadA. Positive colonies were verified by sequencing. Then the plasmids were transferred into E. coli NT1003 respectively to evaluate the effect of promoters.
Figure 4-2 The colony PCR validation diagram.
Test
Three engineered strains with different promotes were cultured aerobically at 33℃ for 22 h, then shifted them to anaerobic culture at 42℃. A biosensor was used to measure the lysine and glucose during the aerobic phase (0 - 23h) and anaerobic phase (after 23h). PJ23106 and PJ23119 led to more lysine accumulation compared to PJ23100. PJ23106 was best, resulting in accumulating 5.1 g/L at 22h of the aerobic phase. Subsequently, during the high-temperature anaerobic phase, lysine was gradually consumed completely.
Figure 4-3 (A) Schematic Diagram; Glucose consumption and lysine accumulation of the engineered E. coli. (B) PJ23100-CadA (33℃), (C) PJ23106-CadA (33℃), (D) PJ23119-CadA(33℃).
Learn
In this cycle, we learned that promoters of different strengths determine the expression intensity of downstream genes. The strain with the PJ23106 promoter accumulated the highest lysine accumulation during the aerobic phase, which proved that PJ23106 effectively reduced CadA leaky expression. Additionally, preliminary experiments combining aerobic-anaerobic culture with high-low temperature switching showed that the Temperature-responsive system effectively supported lysine accumulation by repressing CadA during low-temperature aerobic growth of E. coli NT1003; while triggering its rapid consumption by inducing CadA expression upon switching to anaerobic conditions with temperature upshift.
Cycle 5 Combining with RNA thermometers for precise temperature-controlled CadA expression
Design
After reviewing the literature, we identified high-efficiency RNA thermometers. At low temperatures, they facilitates the formation of a stem-loop structure at the ribosome-binding site (RBS), blocking ribosome binding and inhibiting translation. Upon temperature increase, the structure unfolds, restoring normal translation. To refine the system's temperature sensitivity and accuracy, we incorporated RNA thermometers U7 and U8 into the plasmid pET28a(ΔI7)-PJ23106-RBS-TlpA39-PtlpA-CadA, adding translational control to the existing transcriptional regulatory circuit.
Build
We constructed the plasmids pET28a(ΔI7)-PJ23106-RBS-TlpA39-PtlpA-U7-CadA and pET28a(ΔI7)-PJ23106-RBS-TlpA39-PtlpA-U8-CadA using methods consistent with cycle4. Subsequently, the plasmids were transformed into E. coli NT1003 to evaluate the effect of RNA thermometers.
Figure 5-1 RNA thermometer mechanism diagram.
Test
Two engineered strains were cultured aerobically at 33℃ for 23 h, then shifted them to anaerobic culture at 42℃. A biosensor was used to measure the lysine and glucose during the aerobic phase (0 - 23 h) and anaerobic phase (after 23h). The results are shown in Figure 5-2.
The engineered E. coli with the U7 and U8 RNA thermometers exhibited higher lysine accumulation (8.7 g/L and 10.1 g/L) at 37℃, while at 33℃, the levels increased by 2-fold and 2.3-fold, respectively, compared to the previous round(Figure 5-2).
Figure 5-2 Glucose consumption and lysine accumulation of the engineered E. coli. (A) PJ23106-U7-CadA (33℃). (B) PJ23106 -U7-CadA (35℃). (C) PJ23106 -U7-CadA (37℃). (D) PJ23106 -U8-CadA (33℃). (E) PJ23106 -U8-CadA (35℃). (F) PJ23106-U8-CadA (37℃).
Learn
In this cycle, we learned that incorporating RNA thermometers into the original temperature-responsive system enables dual-level regulation of CadA expression—at both transcriptional and translational levels via temperature. The optimized system could minimize CadA leakage at low temperatures—maximizing lysine accumulation—and permit accurate induction upon temperature upshift, enabling staged and precise control.
Module 2: Engineering CadA---Synthesis of cadaverine
Cycle 6 AI-guided design of CadA via multi-site combinatorial mutagenesis
Design
CadA is the key enzyme for the synthesis of cadaverine. To improve its catalytic efficiency, we adopted a computationally guided strategy. Molecular docking simulations was performed using the structure of CadA (PDB No. 6YN6) Hydrogen bonds, salt bridges, and hydrophobic interactions between the enzyme and its substrate were systematically analyzed. Based on the interaction analysis, we identified seven residues located near the active pocket that might make key contributions to substrate binding and catalysis: I182, H245, K246, W333, E526, K527 and Y652. These candidate sites were subsequently evaluated using a site-directed mutagenesis prediction model for virtual amino acid substitutions, and their catalytic efficiencies (Kcat) were predicted. The model recommended several pentuple mutants with predicted high enzymatic activity (as shown in Table 6-1).
Table 6-1 Prediction of enzymatic activity for combined mutations
| Mutation type |
Enzymatic activity Kcat（s⁻¹） |
| Wild type |
3.484 |
| H245Q, W333E, E526N, K527S, G655F |
13.6055 |
| H245Q, W333H, E526N, K527S, G655F |
13.6008 |
| H245Q, W333H, E526N, K527S, G655K |
13.4973 |
| H245Q, W333E, E526N, K527S, G655K |
13.4772 |
| H245F, W333E, E526N, K527S, G655F |
13.2887 |
Build
According to the prediction results, DNA fragment containing five target site-directed mutation sites was amplified by overlap extension PCR (overlap-PCR) using the original CadA plasmid pCDFDuet-Ptrc-CadA as the template and site-directed mutagenesis primers. Subsequently, the homologous recombination method was used to ligate the gene fragment with the plasmid backbone, obtaining the mutant plasmids, including pCDFDuet-Ptrc-CadAH245Q,W333E,E526N,K527S,G655F, pCDFDuet-Ptrc-CadAH245Q,W333H,E526N,K527S,G655F, pCDFDuet-Ptrc-CadAH245Q,W333H,E526N,K527S,G655K, pCDFDuet-Ptrc-CadAH245Q,W333E,E526N,K527S,G655K, pCDFDuet-Ptrc-CadAH245F,W333E,E526N,K527S,G655F, These plasmids were then transformed into competent E. coli DH5α. Positive clones were screened by colony PCR, and the successful construction was finally verified by sequencing.
Figure 6-1 Construction of combined mutations. (A) strategy diagram of combined mutations. (B) PCR validation diagram of fragment. (D) PCR of Vector. (E) The colony PCR validation diagram.
Test
We separately transformed the plasmids verified to be correct by sequencing—including pCDFDuet-Ptrc-CadAH245Q,W333E,E526N,K527S,G655F, pCDFDuet-Ptrc-CadAH245Q,W333H,E526N,K527S,G655F, pCDFDuet-Ptrc-CadAH245Q,W333H,E526N,K527S,G655K, pCDFDuet-Ptrc-CadAH245Q,W333E,E526N,K527S,G655K, pCDFDuet-Ptrc-CadAH245F,W333E,E526N,K527S,G655F, as well as the original control plasmid pCDFDuet-Ptrc-CadA into the E. coli NT1003. Following protein expression, a whole-cell catalytic assay was performed to detect lysine conversion. Regrettably, we found that all CadA mutants carrying 5 mutations had completely lost their enzymatic activity and were unable to consume the substrate lysine.
Learn
Through this round of cycles, we successfully constructed CadA mutants with 5 mutation sites; however, their enzymatic activity was completely lost. It is speculated that these mutation sites included key amino acid residues in the enzyme's active center or residues related to the binding of the cofactor PLP. It is likely due to structural alterations that disrupt the active site and/or cofactor binding. Therefore, in the next cycle, we chose to perform single-point mutations based on the sites provided by the in silico experiment group to identify the key amino acid residues .
Cycle 7 Engineering key enzyme CadA through single point mutation
Design
Since multi-site mutations resulted in a complete loss of enzymatic activity, we conducted a systematic single-site mutation analysis. Based on autodock result anlysis, the key sites (I182, H245, K246, W333, E526, K527 and Y652) were identified to assess their individual functional importance (Figure7-1).
Figure 7-1 Interactions of key amino-acid residues
First, we performed alanine scanning mutagenesis on all predicted sites. Alanine (A) has a side chain consisting of only a simple methyl group, which can effectively eliminate the specific interactions (e.g., hydrogen bonds, hydrophobic interactions, charge interactions) of the original side chain while minimizing disturbances to the protein backbone conformation and overall structure. Through this step, we quickly screened out the key amino acid residues. Subsequently, the sites with modification potential were mutated into five amino acids of diverse physicochemical properties—glutamic acid (E), leucine (L), serine (S), phenylalanine (F), and valine (V) to explore their functional roles. Specifically: E (negatively charged, hydrophilic, large side chain) is used to probe charge interactions; L and V (hydrophobic, neutral, branched side chains with different sizes) are used to probe hydrophobic packing and spatial volume tolerance; F (hydrophobic, neutral, large-volume rigid aromatic ring) is used to probe π-π stacking interactions and sensitivity to large side chains; S (polar, hydrophilic, small side chain) is used to probe the ability to form hydrogen bonds and tolerance to small side chains.
Build
Based on the plasmid of pCDFDuet-Ptrc-CadA, we designed point mutation primers and cloned the whole plasmid using PCR. The linearized plasmid was isolated and purified by agarose gel electrophoresis, and then transferred into E. coli DH5α to obtain the targeted plasmids of pCDFDuet-Ptrc-CadAI182A, pCDFDuet-Ptrc-CadAH245A, pCDFDuet-Ptrc-CadAK246A, pCDFDuet-Ptrc-CadAW333A, pCDFDuet-Ptrc-CadAE526A, pCDFDuet-Ptrc-CadAK527A, pCDFDuet-Ptrc-CadAY652A.
Verified by sequencing, all the plasmids were successfully constructed.
For the potential modification sites identified after alanine scanning—I182, H245, and K246—we mutated each of these three sites into glutamic acid (E), leucine (L), serine (S), phenylalanine (F), and valine (V) respectively using the same method. Verified by sequencing, the construction of pCDFDuet-Ptrc-CadAK246V failed, whereas the remaining fourteen mutant plasmids were successfully constructed.
Figure 7-2 Construction of single-point mutations. (A) Strategy diagram of single-point mutations. (B) The colony PCR validation diagram of alanine mutations. (C) The colony PCR validation diagram of mutations.
Test
The mutant plasmids and the unmutated original plasmid were separately transformed into the E. coli NT1003. After culturing for expression, a whole-cell catalytic assay was performed to determine the catalytic activity towards the substrate lysine. Under the condition of pH=7, the activity of the unmutated wild type (WT) was set as 100%, and the relative activities of other mutants are shown in Figure 7-3.
Figure 7-3 Enzymatic activities of single-point mutations. (A) Enzymatic activities of alanine mutations. (B) Mutation results of I182, H245, and K246 Sites (WT activity set as 100%).
Learn
In this cycle, we constructed alanine scanning at positions I182, H245, K246, W333, E526, K527, and Y652. It was found that the activity was almost completely lost after mutation at the W333, E526, K527, and Y652, indicating that these amino acids might be functionally critical residues directly related to substrate binding or the catalytic active site. It is conducive to further understanding the catalytic mechanism of the enzyme in subsequent studies.
We selected I182, H245, and K246 that exhibited less severe activity loss, for subsequent mutations to E, L, S, F, and V. Although their activity was not improved, this information will provide guidance for follow-up research. This avoids the massive screening and sequencing workload caused by full saturation mutagenesis, greatly improving research efficiency.
Faced with the challenges of these results, we did not become discouraged. Instead, we actively reflected, extensively reviewed literature, and took the initiative to conduct in-depth exchanges with experts. Through sufficient discussions and cross-fertilization of ideas, we gradually formed a new hypothesis: rather than directly modifying catalytic residues that are highly sensitive to mutations, could we indirectly improve catalytic performance by enhancing the overall structural stability of the enzyme? Consequently, the in silico experiment group began to attempt re-predicting potential beneficial mutations from the perspective of decamer stability.
Cycle 8 Engineering key enzyme CadA through stabilizing the structure for improvement of enzyme activity
Design
Apart from the aforementioned key amino acid residues, we shifted our modification strategy to enhancing the stability of CadA's decameric structure, as its decameric form is essential for efficiently exerting decarboxylation function. Through computer simulation analysis combined with literature reports, we predicted several pairs of double-site mutations (F14C/K44C, V12C/K41C, F14C/L45C, L93C/E445C, Y13C/P36C, E104K) or single-site mutations (E104K) that may stabilize the decameric structure by forming disulfide bonds or other interactions. We also referenced mutation pairs from the literature (V12C/K44C, F14C/D41C, L89C/E445C).
Figure 8-1 The structure of CadA.
Build
Based on the prediction results, using the original CadA plasmid pCDFDuet-Ptrc-CadA as the template, site-directed mutagenesis primers were designed for the mutation sites. The gene fragments containing the mutation sites were amplified by PCR, and then ligated with the plasmid backbone DNA fragment via homologous recombination method, resulting in the mutant plasmids: pCDFDuet-Ptrc-CadAF14C/K44C, pCDFDuet-Ptrc-CadAV12C/K41C, pCDFDuet-Ptrc-CadAF14C/L45C, pCDFDuet-Ptrc-CadAL93C/E445C, pCDFDuet-Ptrc-CadAY13C/P36C, pCDFDuet-Ptrc-CadAE104K, pCDFDuet-Ptrc-CadAV12C/K44C, pCDFDuet-Ptrc-CadAF14C/D41C, and pCDFDuet-Ptrc-CadAL89C/E445C. These plasmids were transformed into competent E. coli DH5α. Positive clones were screened by colony PCR, and the successful construction was finally verified by sequencing.
Figure 8-2 The colony PCR validation diagram.
Test
Following cultivation and induced expression, the mutated recombinant cells were harvested and subjected to whole-cell catalysis. Under the condition of pH=7 and 20 g/L lysine as the substrate, the consumption of lysine was detected. The activity of the wild type (WT) was set as 100%, and the relative activities of other mutants are shown in Figure 8-3. The Y13C/P36C mutant exhibited a maximum increase of 113.26% in enzymatic activity. Additionally, the enzymatic activities of V12C/D41C, F14C/D41C, and L93C/E445C mutants were also higher than that of the wild type (WT).
Figure 8-3 Enzymatic activities of mutants modified for disulfide bond formation (WT activity set as 100%).
Learn
Through this cycle, we learned that modifications such as Y13C/P36C, V12C/D41C, F14C/D41C, and L93C/E445C promote the formation of disulfide bonds, thereby stabilizing the structure of CadA. We also recognized that the higher-order structure of an enzyme is the material basis for its activity, and maintaining the stability of the structure is one of the effective strategies to improve enzymatic activity.
Module3: Cofactor--- Improvement of succinate
Cycle 9 Overexpression of GAP to promote the synthesis of succinate
Design
For our other co-produced product, succinate, based on the metabolic flux analysis, we chose to overexpress glyceraldehyde-3-phosphate dehydrogenase (GAP) to enhance the supply of reducing power. Succinate is a reductive product that requires the consumption of excess reducing power during its synthesis. To boost succinate synthesis, we screened four NADP⁺-dependent glyceraldehyde-3-phosphate dehydrogenases (GAP) originated from Corynebacterium glutamicum ATCC 13032 (CggapC), Bacillus subtilis (BsrocG), Clostridium saccharobutylicum DSM 13864 (CsgapC), and Streptococcus mutans (SmgapN). These enzymes catalyze the conversion of D-glyceraldehyde 3-phosphate to 3-phospho-D-glycerate and simultaneously generate NADPH, thereby increasing intracellular reducing power.
Figure 9-1 The metabolic pathway catalyzed by glyceraldehyde-3-phosphate dehydrogenase.
Build
The gene sequences of glyceraldehyde-3-phosphate dehydrogenases (CggapC from BBa_25RYKWX3, CsgapC from BBa_25099LJK, and SmgapN from BBa_259YBJBJ) were synthesized by a company after codon optimization. The gene of BsrocG (from BBa_250WWUYW) was obtained by PCR amplification using the Bacillus subtilis genome as the template. The different gene fragments were cloned into the plasmid pCDFDuet under the control of the trc promoter via the In-Fusion cloning method, resulting in the plasmids pCDFDuet-Ptrc-CggapC, pCDFDuet-Ptrc-BsrocG, pCDFDuet-Ptrc-CsgapC, and pCDFDuet-Ptrc-SmgapN. These plasmids were transformed into E. coli NT1003 for expression.
Figure 9-2 (A) The schematic of plasmid pCDFDuet-Ptrc-BsrocG. (B) The schematic of plasmid pCDFDuet-Ptrc-CggapC. (C) The schematic of plasmid pCDFDuet-Ptrc-CsgapC. (D) The schematic of plasmid pCDFDuet-Ptrc-SmgapN.
Figure 9-3 The colony PCR validation diagram. (A) BsrocG. (B) CggapC. (C) CsgapC. (D) SmgapN.
Test
The function of NADP⁺-dependent glyceraldehyde-3-phosphate dehydrogenase was investigated based on the succinate titer produced by E. coli, as quantified by HPLC. The results showed that the expression of CggapC could effectively increase the titer of succinate by 24.5%. However, we also found that it exerted an negative effect on lysine during the aerobic phase, with a 40% decrease compared with the control.
Figure 9-4 (A) The effect of different glyceraldehyde-3-phosphate dehydrogenases on lysine. (B) The effect of different glyceraldehyde-3-phosphate dehydrogenases on succinate.
Learn
In this cycle, we learned that enzymes from different sources exhibit varying activities, which affect the titer of target products. We screened four NADP⁺-dependent glyceraldehyde-3-phosphate dehydrogenases from different organisms and found that the expression of CggapC achieved the optimal increase in succinate, but it simultaneously reduced lysine synthesis.
We reviewed literatures and discussed with other professors, who provided us with several suggestions. Dr. Xiaojie Guo recommended using a low-copy-number plasmid to mitigate the impact of excessive reducing power on cells and reduce cellular metabolic burden. He also showed strong interest in our modular design concept and suggested adjusting metabolic pathways to increase product titer.
Cycle 10 The coordination of lysine and succinate production by changing plasmid
Design
After consulting experts, we realized that using the pCDFDuet plasmid for expressing glyceraldehyde-3-phosphate dehydrogenase might not be the optimal choice. After in-depth internal team discussions, we decided to adopt the low-copy-number pACYCDuet plasmid to reduce the metabolic burden of the strain.
Build
Using pCDFDuet-Ptrc-CggapC, pCDFDuet-Ptrc-BsrocG, pCDFDuet-Ptrc-CsgapC, and pCDFDuet-Ptrc-SmgapN as templates, the CggapC, BsrocG, CsgapC, and SmgapN fragments were amplified by PCR. Each gene of CggapC, BsrocG, CsgapC, and SmgapN was separately inserted into the linearized pACYCDuet vector using In-Fusion cloning, titering four recombinant plasmids, pACYCDuet-CggapC, pACYCDuet-BsrocG, pACYCDuet-CsgapC, and pACYCDuet-SmgapN.
These plasmids were transformed into E. coli DH5α, and positive colonies were verified via PCR and sequencing. Subsequently, the plasmids with correct sequencing results were transformed into the lysine-producing strain E. coli NT1003, and direct fermentation was used to evaluate their impact on lysine and succinate production.
Figure 10-1 (A) The schematic of plasmid pACYCDuet-BsrocG. (B) The schematic of plasmid pACYCDuet-CggapC. (C) The schematic of plasmid pACYCDuet-CsgapC. (D) The schematic of plasmid pACYCDuet-SmgapN.
Figure 10-2 The colony PCR validation diagram. (A) BsrocG. (B) CggapC. (C) CsgapC. (D) SmgapN.
Test
The performance was evaluated by detecting lysine and succinate produced by recombinant E. coli via HPLC. The results showed that the expression of four dehydrogenase using a low-copy-number plasmid pACYCDuet,led to an increase in succinate. Among them, CggapC still exhibited the highest succinate titer, and no decrease in lysine titer (Figure 10-3). Therefore, the CggapC was selected in subsequent work.
Figure 10-3 (A) The effect of different glyceraldehyde-3-phosphate dehydrogenases on lysine using a low-copy-number plasmid. (B) The effect of different glyceraldehyde-3-phosphate dehydrogenases on succinate using a low-copy-number plasmid.
Learn
In this cycle, we have learned how to optimize gene expression for the increased product titer. Due to the difference in the replicon, different plasmid has different copy number in the strain, which would affect the gene expression level. Therefore, the selection of appropriate plasmid is also an effective strategy for improving strain producing ability and maintaining activity.
Cycle 11 Constructing cofactor conversion to promote the synthesis of succinate and lysine
Design
Following Dr. Xiaojie Guo’s suggestion, we learned that certain transhydrogenases can also promote the rational distribution and flow of cofactors. Additionally, based on literature research, we ultimately selected the soluble pyridine nucleotide transhydrogenase (sthA) derived from E. coli MG1655.
Figure 11-1 The metabolic pathway catalyzed by soluble pyridine nucleotide transhydrogenase (sthA).
Build
The gene fragment of sthA was subcloned together with CggapC into linearized pACYCDuet, generating the plasmid pACYCDuet-CggapC-sthA. The plasmid was transformed into E. coli DH5α, and positive colonies were verified via PCR and sequencing analysis. Finally, the plasmid was transformed into the lysine-producing strain E. coliNT1003, and direct fermentation was used to evaluate its impact on lysine and succinate production.
Figure 11-2 (A) The schematic of plasmid pACYCDuet-CggapC-sthA. (B) The colony PCR validation diagram.
Test
Its performance was evaluated by detecting lysine and succinate produced by E. coli via HPLC. The results showed that the succinate titer of the strain harboring pACYCDuet-CggapC-sthA increased by 28% relative to control; meanwhile, the lysine titer reached 14.9 g/L. We decided to use this strain in subsequent experiments.
Figure 11-3 (A) Lysine production of the strain harboring pACYCDuet-CggapC-sthA. (B) Succinate production of the strain harboring pACYCDuet-CggapC-sthA.
Learn
In this cycle, we recognized the importance of constructing cofactor conversion. The demand for succinate synthesis is NADH. We previously focused mainly on enhancing the overall reducing power by GAP exression. After following Dr. Xiaojie Guo’s suggestion, we introduced soluble pyridine nucleotide transhydrogenase (sthA) to promote the rational distribution and flow of cofactors. As a result, the lysine titer increased to 14.9 g/L, and the succinate titer rose from 5.0 g/L to 6.4 g/L, achieving a further improvement in production. This also provides reference value for the work of other teams.
Module 4: An intelligent cell---Production of cadaverine-succinate
Cycle 12 Construction of a cell factory for synchronized synthesis of cadaverine-succinate
Design
Through preliminary experiments, we optimized the thermo-responsive switch to control the expression of cadA and the cofactor system. Subsequently, we introduced two plasmids into E. coli NT1003 to construct a cell factory capable of efficiently co-producing cadaverine and succinate..
Build
The plasmids pET28a(I7)-PJ23106-RBS-TlpA39-PtlpA-U7-CadA and pET28a(ΔI7)-PJ23106-RBS-TlpA39-PtlpA-U8-CadA were co-transformed with the plasmid pACYCDuet-CggapC-sthA into the lysine-producing strain E. coli NT1003 to construct a cell factory capable of co-producing cadaverine and succinate.
Figure 12-1 Construction of a cell factory for synchronized synthesis of cadaverine-succinate.
Test
The performance was assessed by detecting cadaverine and succinate via HPLC. According to the fermentation results, the recombinant strain successfully achieved the co-synthesis of cadaverine and succinate, with the highest titers reaching 5.7 g/L and 5.4 g/L, respectively. This indicates that the designed synthetic pathway is feasible in experiments, and a cell factory for the synchronized synthesis of cadaverine and succinate has been successfully developed.
Figure 12-2 Co-production of cadaverine and succinate in the cell factory.
Learn
In this cycle, we constructed a cell factory for the synchronized synthesis of cadaverine and succinate. Through precise metabolic flux regulation, the molar ratio of the two products is close to the ideal 1:1. It creates conditions for the direct generation of PA54 salt in the subsequent acid-base neutralization, reducing the demand for acids or bases, and minimizing costs. Secondly, from the perspective of carbon metabolism, the matching of the titers of the two products indicates a good balance between CO2 release and fixation, improving carbon atom economy and laying a robust foundation for the efficient and green production of the PA54.
Summary
The overall goal of the project was to develop an intelligent cell factory for the simultaneous biosynthesis of cadaverine and succinate—key monomers for bio-based PA54—using synthetic biology strategies. We designed and constructed a temperature-sensitive switch to enable lysine accumulation during the aerobic phase at low temperature and its efficient conversion to cadaverine under anaerobic conditions at high temperature, while simultaneously producing succinate.
To enhance the precision of the temperature switch, we optimized both transcriptional and translational controls, increasing lysine accumulation by 2.3-fold. We also engineered the key enzyme CadA through rational design, obtaining a mutant with 113% higher catalytic activity than WT. Furthermore, by introducing NADP⁺-dependent GAP dehydrogenase and balancing cofactors via soluble transhydrogenase expression, the results showed that the succinate titer increased by 28% relative to control; meanwhile, the lysine titer reached 14.9 g/L.
By integrating the temperature switch, optimized CadA, and cofactor modules, we successfully constructed an intelligent E. coli cell factory capable of co-producing 5.7 g/L cadaverine and 5.4 g/L succinate from glucose, laying a solid foundation for the green and low-carbon production of PA54.
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