<figure><img src="image_1.png" /></figure>

# Amine-Tolerant *E. coli* Strains Generated via Adaptive Evolution for Sustainable Synthesis of Chiral Amines

Josemarco Mendoza-Avila, Volker Döring, Madeleine Bouzon, Ivan Dubois, Tanja Knaus, Louis M. M. Mouterde, Anne Zaparucha, Francesco G. Mutti,* and Carine Vergne-Vaxelaire*

<figure><img src="image_2.png" /></figure>

Cite This: ACS Sustainable Chem. Eng. 2024, 12, 14435–14445

<figure><img src="image_3.png" /></figure>

Read Online

## ACCESS

<figure><img src="image_4.png" /></figure>

Metrics & More

<figure><img src="image_5.png" /></figure>

Article Recommendations

<figure><img src="image_6.png" /></figure>

Supporting Information

**ABSTRACT:** The biocatalytic synthesis of chiral amines from carbonyl compounds and ammonia is a major advance in sustainable synthetic chemistry. Using whole cells for bioamination reactions is advantageous given their low preparation cost and direct applicability; however, amine toxicity limits the reaction when living cells are used. Herein, we adapted *Escherichia coli* BL21(DE3) cells to grow in the presence of 100 mM hexan-2-amine *via* directed evolution in continuous culture, obtaining six times more tolerant strains than the wild-type. The adapted strains also displayed superior tolerance for structurally different amines. Coexpression of genes encoding for amine dehydrogenase (AmDH) and formate dehydrogenase activities in the adapted strains enabled the stereoselective bioamination (ee > 99%) of different prochiral ketones with up to 80% conversion at high substrate loading (up to 200 mM) without exogenous cofactor addition. The adapted cells displayed longer survival and higher population density during the reactions. The present biotechnological *E. coli* system contributes to the development of more robust biocatalysis for amine production.

**KEYWORDS:** biotransformations, amination, adaptive evolution, amine toxicity, continuous culturing, biotechnological E. coli system

<figure><img src="image_7.png" /></figure>

## INTRODUCTION

Chiral amines are high-value molecules, mostly serving as building blocks for the preparation of pharmaceutical and agrochemical active ingredients. Moreover, in stereoselective organic synthesis, they are applied as ligands for asymmetric catalysis and as resolving agents of racemic mixtures through diastereomeric salt formation.1,2 Therefore, the preparation of these amines by introducing nitrogen into a commodity chemical with total chemo- and enantiocontrol *via* a simple and direct reaction is greatly desirable.3 Additionally, as we intend to transition our manufacturing technologies to sustainable processes, we should enable these synthetic reactions by green approaches.

Biocatalysis enables the direct synthesis of enantiopure chiral amines from prochiral ketones using ammonia as an inexpensive nitrogen source and generating water as the sole byproduct. This is possible through NAD(P)H-dependent enzymes performing reductive amination, i.e., the one-pot conversion of a ketone or aldehyde to an amine *via* an intermediate imine.4,5 These enzymes—namely amine dehydrogenases (AmDHs), some imine reductases (IREDs) active toward carbonyl-containing substrates, and reductive aminases (RedAms), a subclass of IREDs⁶—complete the other enzyme families applied in the synthesis of optically active amines, i.e., ω-transaminases (ω-TAs),⁷,⁸ hydrolases,⁹,¹⁰ ammonia lyases,¹¹

amine oxidases,¹² Pictet–Spenglerases,¹³ and engineered cytochrome P450s.¹⁴

Enzymes can be utilized, immobilized or not, in purified form, expressed in cell-free extracts, or in whole-cell systems. This latter approach is advantageous in terms of increased enzyme stability, direct applicability, and low preparation cost. Indeed, the use of whole cells circumvents the expenses of cell lysis (35–100 vs 250–1000 euros kg⁻¹, respectively)¹⁵ and enzyme purification, whereas the protective nature of the cellular environment stabilizes the biocatalyst. Moreover, often, the use of living whole cells eliminates the need for cofactor supplementation by exploiting the cellular metabolism.¹⁶,¹⁷

Therefore, whole cell systems have been applied in amine synthesis based on redox reactions that generally require the addition of enzymatic cofactors when cell-free preparations are employed. We can mention, for example, the synthesis of cyclic chiral amines by *in vivo* cascade reactions starting from carboxylic acids,¹⁸ the formation of (S)-4-phenyl-2-butylamine

Received: May 27, 2024

Revised: September 2, 2024

Accepted: September 4, 2024

Published: September 16, 2024

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>
from the corresponding ketone by ω-TA-catalyzed reactions,¹⁹ and the fermentative production of aliphatic amines from amino acid precursors by the valine dehydrogenase from *Streptomyces viridifaci*²⁰—the active enzymes being heterologously expressed in *Escherichia coli* (*E. coli*) cells.

*E. coli* is extensively used as a host in whole-cell biotransformations, given its relatively easy genetic manipulation and high heterologous enzyme overexpression rates. Resting cells—i.e., nongrowing but still metabolically active cells—are often used for bioamination reactions in vivo, especially when the optimum temperature, pH value, or media composition for the bioconversion differ from the values allowing the best growth conditions. Nonetheless, the toxicity of substrates, intermediates, or products is a potential drawback when living cells are used. For example, when assessing the bioamination of alcohols by combining an alcohol dehydrogenase (ADH) with an AmDH in vivo using *E. coli* cells, we observed that moderate concentrations (from ca. 2 mM up to 20 mM, depending on the compound) of amine products caused toxicity and cell death, hence limiting the amine titers to a maximum of 15 mM for the best converted substrate.²¹ Accordingly, in the aforementioned examples of amine synthesis, reactions were performed at low or moderate substrate loadings (i.e., 3–25 mM) to match the toxicity profile of the related products.¹⁸⁻²⁰ The addition of a water-immiscible cosolvent, which acts as a reservoir of the toxic compound, can partially alleviate this problem. However, conventional organic solvents are themselves toxic for the cells, while biocompatible solvents are limited and usually more expensive. Therefore, strategies to protect living *E. coli* cells for biocatalytic applications have been recently proposed, including, for example, artificial sporulation by coating the cells with polydopamine,²² and cell embedding into alginate hydrogels.²³ Alternatively, the introduction of global genetic regulators can coordinate the responses of various genes to environmental changes in *E. coli*, which has led to cells with higher organic solvent resistance and improved biocatalytic behavior.²⁴

In this work, we aimed at evolving new *E. coli* strains, starting from conventional *E. coli* strain BL21(DE3), that have increased tolerance toward amines. The engineered strains were employed for the recombinant expression of AmDH and formate dehydrogenase (FDH) enzymes and studied in biotransformations using resting cells to synthesize chiral amines from ketones 1–7a (Figure 1). Based on our previous work, we focused on increasing the tolerance toward amines because the corresponding ketone precursors are generally less toxic.²¹ In this regard, we adapted the *E. coli* BL21(DE3) cells to grow in the presence of high amine concentrations via directed evolution in a continuous culture. Moreover, we compared the performance of the adapted cells with that of the nonadapted cells, showing that higher amine production could be achieved with the former in the absence of any external addition of nicotinamide adenine dinucleotide cofactor (NAD⁺).

## EXPERIMENTAL SECTION

Commercial chemicals, enzymes, strains, and growth media composition used in this study are given in *Supporting Information*.

**Continuous Cultures.** Adaptive directed evolution experiments were conducted in GENEMAT (GM3) devices for automated continuous culture. The volume of the culture was maintained constant (16 mL at 37 °C) by diluting with a pulse of fresh medium

<figure><img src="image_10.png" /><figcaption>**Figure 1.** (A) Representation of an adapted *E. coli* cell expressing AmDH and FDH activities. The former enables the reductive amination of prochiral ketones to chiral amines, the latter allows for NADH recycling. (B) Ketones studied in this work.</figcaption></figure>

and discarding the same volume of the cell suspension. Depending on the programmed growth regime, this occurred in response to a comparison of the optical density (OD) of the culture—automatically read every 10 min—to a fixed OD threshold (OD₆₀₀ nm 0.4). Constant aeration of the culture was guaranteed by a continuous flow of air, equivalent to one volume of culture per minute, which also prevented cell sedimentation. The formation of biofilms (i.e., wall growth) was avoided by enabling the transfer of the culture twice per day between two identical growth vessels, which were alternately sterilized. In this study, two growth regimes were used: turbidostat and medium swap. In the former regime, the dilution rate was dependent on the cell growth, and the dilutions with fresh medium (1.6 mL pulse) occurred when the OD of the culture exceeded the set threshold. In the latter regime, the medium to dilute “swapped” between permissive and stressing media, depending on whether the measured OD stayed below (permissive dilution pulse) or exceeded the threshold (stressing dilution pulse). Under the medium swap regime, dilutions of fixed volume were triggered every 10 min, hence imposing a generation time on the culture.

The applied growth regimes and culture media (final pH of 7.2) were as follows. For adaptation of *E. coli* BL21(DE3), (a) turbidostat regime: MS-glucose 0.2% w/v with 15 mM of *rac*-hexan-2-amine (*rac*-2a); (b) medium swap regime: permissive medium MS-glucose 0.2% w/v with 15 mM of *rac*-2a, stressing medium MS-glucose 0.2% w/v with 30 mM of *rac*-2a; (c) turbidostat regime: MS-glucose 0.2% w/v with 30 mM of *rac*-2a. For adaptation of *E. coli* JHX 1.2.3, (a) turbidostat regime: MS-glucose 0.2% w/v with 30 mM of *rac*-2a; (b) medium swap regime: permissive medium MS-glucose 0.2% w/v with 30 mM of *rac*-2a, stressing medium MS-glucose 0.2% w/v with 100 mM of *rac*-2a; (c) turbidostat regime: MS-glucose 0.2% w/v with 100 mM of *rac*-2a.

**E. coli Growth Profiles.** Bacterial growth was characterized using a Microbiology Reader Bioscreen C (Thermo Fisher Scientific) for incubation and automated OD monitoring. *E. coli* was precultured to an OD₆₀₀ nm between 1.0 and 1.2 in MS-glucose 0.2% w/v medium and inoculated into fresh culture medium at 1:100 dilution. 200 µL aliquots of the resulting medium were transferred into honeycomb 100-well plates, which were incubated at 37 °C under continuous agitation. *E. coli* growth curves were generated by OD₆₀₀ nm readings every 15 min. Each test was performed in triplicate.
<figure><img src="image_11.png" /><figcaption>**Figure 2.** Topology and fluidics of a GENEMAT (GM3) continuous culture device functioning under a medium swap culture regime. A constant flow of compressed air aliments the fluidic system, assuring aeration of the culture growing in the left chamber (yellow volume), dilutions from either a permissive (A) or a stressing medium (B) reservoir, and the concomitant transfer of an equivalent culture volume to waste (denoted with W). At fixed intervals, the culture is transferred to the right chamber (C) and the left chamber is purged with cleaning agent (5 N NaOH, in gray). The left chamber is then rinsed with water and the culture transferred back, followed by a cleaning of the right chamber (not shown). Twin culture chambers and fluidic reservoirs are connected through silicon tubes. Electric valves (indicated by red-contoured circles) ensure synchronized flows to execute a programmed storyboard. Cell density measurement by a diode-photocell attached to the left chamber (not shown) enables comparison with a set cell density threshold conditioning the opening and closing of the valves to trigger dilutions according to the chosen culture regime.</figcaption></figure>

**Minimum Inhibitory Concentration (MIC) Assays.** *E. coli* BL21(DE3) strain and evolved derivatives were inoculated in MS-glucose 0.2% w/v medium and cultured overnight at 37 °C and 150 rpm. The following day, fresh MS-glucose 0.2% w/v medium was inoculated with the overnight culture and incubated under the same conditions to an OD₆₀₀ nm between 0.5 and 0.9. A series of culture tubes containing MS-glucose 0.2% w/v medium and the compound to be tested at varied concentrations was prepared by combining 2750 µL of fresh MS-glucose 0.2% w/v medium and 250 µL of amine or ketone in appropriate concentration (3 mL final volume, final pH 7.2). The tubes were inoculated with 30 µL of culture within the aforementioned OD range and incubated for 18 h at 37 °C and 150 rpm. The highest concentration of amine or ketone that permitted cell growth and the lowest concentration that completely inhibited it defined the toxicity range of each tested compound. Based on these toxicity ranges, a second round of toxicity experiments was conducted by following the same method. The MIC was defined as the lowest concentration of tested compound that, under the studied conditions, prevented visible cell growth, as determined by visual assessment. Each compound was tested in triplicate. For each tested compound, the concentrations assayed in both rounds of toxicity experiments are given in Table S1.

**Engineering Recombinant E. coli Cells.** Ch1-AmDH and Cb-FDH were cloned into pET plasmid vectors by employing standard molecular biology techniques. When the pETDuet-1 system was used, Ch1-AmDH and Cb-FDH were cloned into the first and second multiple cloning sites, respectively. The resulting genetic constructs were denoted as pET28b_NHis6_Ch1-AmDH, pETDuet-1_Ch1-AmDH_Cb-FDH, and pETDuet-1_NHis6_Ch1-AmDH_Cb-FDH, where NHis6 denotes the presence of an N-terminal hexa histidine tag. Transformation of these constructs into chemically competent *E. coli* cells resulted in strains named His-Ch1, Ch1/Cb, and His-Ch1/Cb, respectively. Lastly, cotransformation of pET28b_NHis6_Ch1-AmDH and pET15b_NHis6_Cb-FDH resulted in the strains His-Ch1/His-Cb. Details about the preparation of chemically competent cells and transformation are given in the Supporting Information.

**General Procedure for E. coli Culturing and Protein Expression.** Recombinant *E. coli* strains were inoculated in LB medium containing the appropriate antibiotic and precultured overnight at 37 °C and 150 rpm. Kanamycin (50 µg L⁻¹) was used for maintaining pET28b derivatives; ampicillin (100 µg L⁻¹) was used for maintaining pETDuet-1 derivatives; and both antibiotics were used for maintaining the replicons of both kinds. The next day, fresh LB medium with antibiotic was inoculated with the overnight culture at 1:100 dilution and grown at 37 °C and 150 rpm to an OD₆₀₀ nm between 0.6 and 0.9. Recombinant protein expression was induced by adding 0.5 mM IPTG (final concentration), followed by cultivation at

25 °C and 150 rpm for an additional 18 h. Cells were then harvested by centrifugation at 4 °C and 4500 rpm for 10 min. Protein expression was verified by 12% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) or 12% native PAGE (Figure S1).

**General Procedure for Resting Cell Preparation and In Vivo Bioamination of Ketones.** Freshly harvested *E. coli* cells were washed with precooled ammonium formate buffer (1 M HCOONH₄/NH₄OH, pH 8.5), recovered by centrifugation at 4 °C and 4500 rpm for 10 min, and resuspended in new precooled buffer to a final concentration ≥75 mg⁻¹ cell wet weight (cww). The cells in the resulting suspension were referred to as resting cells. Unless otherwise stated, bioamination reactions were performed in 1 M HCOONH₄/NH₄OH (pH 8.5, final volume 1 mL) containing resting cells (60 mg mL⁻¹ cww) and the specified concentration of ketone (1–7a). Reactions were incubated at 30 °C and 170 rpm in a Cole-Parmer orbital shaker for 24 h. Each reaction was performed in triplicate using resting cells prepared from independent cultures grown in parallel.

Biotransformation analysis was initiated by quenching the reactions (1 mL) with 10 M KOH (200 µL), followed by extraction with ethyl acetate (2 × 500 µL) containing 20 mM toluene as an internal standard. The combined organic phases were dried over MgSO₄ anhydrous, and the analytical yield was calculated based on the areas of substrate and product determined by GC-FID using a DB-1701 column (Agilent). The enantiomeric excess (ee) was determined by GC-FID using a Varian Chrompack Chirasil Dex-CB column (Agilent) after derivatization with acetic anhydride and 4-dimethylaminopyridine. Commercial standards (1–7a and 1–6b), dissolved in ethyl acetate containing 20 mM toluene, were used as a reference. Reference amine 7b was synthesized biocatalytically, as described in the Supporting Information (Figure S2). Details about the GC-FID analysis (Figures S3–S9) and derivatization procedure are given in the Supporting Information.

**Cell Survivability Assays.** A 100 µL aliquot from the resting cell sample to be tested was gently washed with fresh LB medium. 50 µL of an appropriate dilution in LB medium was plated on LB agar plates that were incubated overnight at 37 °C. Plates with kanamycin (50 µg L⁻¹) and ampicillin (100 µg L⁻¹) were used for His-Ch1/His-Cb cells. The next day, the colony forming units (cfu) per mL of the sample were estimated by counting the colonies on the plate.

**Preparative Biotransformation of Heptan-2-one 3a.** *E. coli* (His-Ch1/His-Cb) was cultured (2 L per strain), induced, and harvested, as described above. The preparative biotransformations consisted of 70 mL of resting cell suspension (60 mg mL⁻¹ cww) in ammonium formate buffer 1 M HCOONH₄/NH₄OH (pH 8.5) containing 1.6 g (14 mmol) of ketone 3a in a 250 mL glass bottle. The biotransformations were performed in duplicate per strain. The reactions were incubated at 30 °C and 170 rpm for 24 h. The
analytical yield was determined by GC-FID using a 1 mL aliquot (worked-up, as described above). The preparative reaction mixture was acidified to pH 2–4 through the addition of a concentrated HCl solution. The reaction was extracted with methyl *tert*-butyl ether (3 × 25 mL) to remove the unreacted ketone substrate. The aqueous phase was then basified to pH 11–13 through the addition of concentrated KOH solution and extracted with methyl *tert*-butyl ether (3 × 25 mL). The organic fractions were combined and dried with anhydrous MgSO₄ anhydrous. After filtration and evaporation of the solvent, amine (R)-3b was obtained as a yellow oil. The authenticity of the product was confirmed by ¹H NMR (300 MHz, CDCl₃, Figure S10).

## RESULTS AND DISCUSSION

**E. coli Adaptive Evolution to rac-Hexan-2-amine.**
Inspired by our previous work,²¹ we first determined the toxicity of *rac*-hexan-2-amine (*rac-2b*) on *E. coli* BL21(DE3) cells. In liquid MS-glucose medium, the presence of 14 mM *rac-2b* caused retarded cell growth, whereas at 18 mM cell growth was totally inhibited even after 48 h of cultivation (Figure S11). This indicates that *rac-2b* is toxic to *E. coli* BL21(DE3) at relatively low concentrations, hence limiting the applicability of these living whole-cells for the bioamination of the corresponding ketone 2a to low substrate loadings. We therefore set out to adapt *E. coli* BL21(DE3) cells to grow in the presence of high concentrations of *rac-2b*. To attain this goal, we used the GENEMAT device, which maintains a cell population in continuous culture under defined growth conditions without biofilm formation. Figure 2 shows the general scheme of a GENEMAT (GM3) functioning in medium swap mode and depicts its self-cleaning principle based on a twin chamber setup. Depending on the culture regime, the cells were diluted with fresh growth medium, either periodically (conditional medium swap regime) or triggered by the growth kinetics (turbidostat regime).²⁵ Thus, this technology enables bacteria to proliferate over unlimited periods of time with concomitant accumulation of genetic variants—natural selection—which are progressively more adapted to the chosen culture conditions.²⁶

The first adaptation was initiated with a turbidostat of *E. coli* BL21(DE3), referred to as a wild-type strain, in MS-glucose medium containing 15 mM *rac-2b* (Figure 3A). Under this regime, the culture was diluted with fresh medium every time a fixed cell density threshold (OD₆₀₀ nm 0.4) was exceeded, thus selecting the fastest-growing cells [generation time (*t*<sub>g</sub>) 1.6 h, day 3]. To further increase the tolerance to *rac-2b*, the adapted cell population was subsequently cultivated under a conditional medium swap regime for gradual adaptation to grow in a stressing medium containing 30 mM *rac-2b* (Figure 3A). Under this regime, dilutions were conducted at a fixed dilution rate, thus imposing a *t*<sub>g</sub> on the culture. Accordingly, the culture was diluted with fresh stressing medium every time the above-mentioned cell density threshold was exceeded. Otherwise, the dilutions were performed with a permissive medium containing 15 mM *rac-2b*. Consequently, the lowest tolerable concentration of *rac-2b* was maintained over the passing generations. After only 30 h of cultivation in the conditional medium swap regime, the proportion of dilutions with the stressing medium reached 100%, which was indicative that the cells grew in 30 mM *rac-2b* (Figure 3A). The short duration of medium swap cultivation suggested that the adaptation was likely to occur during the initial turbidostat culture. To stabilize and further decrease the population's *t*<sub>g</sub> (1.2 h, day 9), the cell population was ultimately grown in the stressing medium under the

<figure><img src="image_12.png" /><figcaption>**Figure 3.** (A) Adaptation of *E. coli* BL21(DE3) to growth in 30 mM *rac-2b* in continuous culture. (B) Adaptation of isolate JHX 1.2.3 to growth in 100 mM *rac-2b* in continuous culture. Black curves show the average daily generation time (*t*<sub>g</sub>), while blue curves show the ratio of the number of diluting pulses with stressing medium to the total number of diluting pulses for 24 h. *t*<sub>g</sub> is the time it took the bacterial population to double in number. Blue and orange zones denote periods of cultivation under the turbidostat and conditional medium swap regimes, respectively; additionally, areas with diagonal stripes indicate the days in which the cultivation regime was changed. Applied *rac-2b* concentrations in the culture media are given above each graph. During periods of cultivation under the medium swap regime, a *t*<sub>g</sub> of 3.2 (A) and 2.4 h (B), was set by the volume of the medium pulses injected at regular time intervals. Isolated strains are shown with vertical arrows. Note: in (A), given the short duration of conditional medium swap cultivation, the average daily *t*<sub>g</sub> on day 5 was <3.2 h, which was the effective *t*<sub>g</sub> imposed by the fixed dilution rate.</figcaption></figure>

turbidostat regime (Figure 3A). This adapted *E. coli* population was named JHX 1.2.

Three isolated strains from this population displayed different tolerances to *rac-2b* (Figure S12). The best-growing isolate, strain JHX 1.2.3, was chosen to inoculate a second continuous culture for further adaptation to 100 mM *rac-2b*. The cells were first grown under a turbidostat regime in MS-glucose containing 30 mM *rac-2b* until day 3 (Figure 3B). Thereafter, from day 3 to 10, the adapting cells were cultivated in a conditional medium swap alimented with MS-glucose 30 mM *rac-2b* (permissive) and MS-glucose 100 mM *rac-2b* (stressing) media. On day 10, the proportion of dilutions with
<figure><img src="image_13.png" /><figcaption>**Figure 4.** (A) Plate reader growth profile of *E. coli* cells in MS-glucose medium containing *rac*-2b (30 mM) during incubation at 37 °C and constant agitation. (B) Same as A with *rac*-2b (100 mM). (C) MIC of amines *rac*-2b, *rac*-3b, and 8b on *E. coli* cells. (D) MIC of enantiopure amines (S)-2b and (R)-2b on *E. coli* cells. (E) MIC of ketones 2–3a, 7a, and 8a on *E. coli* cells. (F) Survival (logarithmic scale) of *E. coli* resting cells (60 mg mL⁻¹ cell wet weight, cww) over time in HCOONH₄/NH₄OH (1 M, pH 8.5) containing *rac*-2b (100 mM) after incubation at 30 °C and 170 rpm for 24 h. In (A,B), each curve represents the average of three measurements. In (C–E), each compound was tested in triplicate; error bars show the toxicity range for each compound, determined by the highest tested concentration that permitted cell growth and the second-lowest concentration that completely inhibited it. In (F), error bars show the standard deviation of three experiments.</figcaption></figure>

the stressing medium reached 100% (Figure 3B). Lastly, a period of cultivation under a turbidostat regime was used to consolidate the population ($t_g$ 1.2 h, day 16). This final population was referred to as JHX 4.2. As observed with the previous isolates, clonal variance in terms of adaptation level occurred for this population (Figure S13). The best-growing isolate, strain JHX 4.2.8, was selected for further experiments.

**Growth Profiles, Survivability, and Tolerance of Adapted Cells toward Amines.** We characterized the growth profile of the adapted strains in the MS-glucose medium containing *rac*-2b. Isolates JHX 1.2.3 and 4.2.8 grew in 30 mM *rac*-2b, whereas the nonadapted strain BL21(DE3) did not; additionally, growth acceleration was similar between both adapted strains but lagged ca. 1 h for JHX 4.2.8 (Figure 4A). As expected, solely JHX 4.2.8 grew in 100 mM *rac*-2b, albeit with a long lag phase (ca. 10 h) (Figure 4B). To discard metabolic consumption of the exogenous amine product—which would hamper the biocatalytic application of the adapted strains—the cells were cultivated with *rac*-2b (10–100 mM), and the amine concentration in the growth medium was monitored over time (Table S2). The results showed that the *rac*-2b concentration remained roughly constant after 24 h of incubation, thus excluding amine metabolism as the underlying adaptation mechanism.

We then studied the amine tolerance of the adapted strains through MIC assays (Figure 4C and Table S1). We defined MIC as the lowest amine concentration that precludes visible cell growth in MS-glucose medium after 18 h of incubation at 37 °C. The *rac*-2b MIC of intermediate strain JHX 1.2.3 augmented from 16 mM (nonadapted cells) to 46 mM. Strain

JHX 4.2.8 displayed even higher tolerance to *rac*-2b (MIC 96 mM), which amounted for an overall six-times MIC increase. Notably, JHX 4.2.8 showed a high tolerance to both enantiomers of 2b (Figure 4D and Table S1). To our delight, we found that the *E. coli* adaptation in continuous culture using *rac*-2b also resulted in greater tolerance to other amines. Indeed, with MIC values of 40 and 134 mM for amines *rac*-3b and 8b, respectively, JHX 4.2.8 displayed four to five times higher amine tolerance contrasted to the BL21(DE3) strain (Figure 4C). Additional toxicity experiments revealed that JHX 4.2.8 cells possess increased resistance to structurally different amines bearing, for example, aromatic, hydroxy, or unsaturated substituents, including secondary amines (Figure S14). Overall, these results suggest the existence of common mechanisms of toxicity mitigation for amine compounds, which are based on mutations that occurred during the *E. coli* adaptation to growth on high *rac*-2b concentrations.

The toxicity of selected ketones on the cells was also explored (Figure 4E and Table S1). Nearly the same ketone tolerance was displayed by JHX 4.2.8 and BL21(DE3) cells. Among the assayed compounds, only 8a was found to show a minor MIC difference (4 mM) between both strains. Notably, in BL21(DE3) cells, the MIC of ketones 2–3a and 8a was roughly twice that of the related amines 2–3b and 8b. These results are in line with our previous finding that amines were considerably more toxic compared to their corresponding ketone precursors.²¹

We next studied the influence of a higher amine tolerance on the survivability of the adapted cells under bioproduction conditions. We incubated BL21(DE3) and JHX 4.2.8 resting
<figure><img src="image_14.png" /><figcaption>**Figure 5.** (A) Bioamination of **2a** (20–50 mM) by *E. coli* resting cells bearing different plasmids for coexpression of Ch1-AmDH and Cb-FDH. (B) Bioamination of **2–3a** and **7a** at varied substrate concentrations (50–300 mM). (C) Progress of the bioamination of **2a** (100 mM). (D) Survival (logarithmic scale) of the cells over time in reaction samples at 100 mM of **2a**. (E) Bioamination of **2a** (100–200 mM) at varied cell concentrations (60–100 mg mL⁻¹ cww). (F) Bioamination of **3–4a** and **7a** (200 mM) at 100 mg mL⁻¹ cww. In (A–D), cell loading is 60 mg mL⁻¹ cww. In (A,B,E,F), the reaction time is 24 h. In (A–F), the reactions were performed in 1 mL HCOONH₄/NH₄OH (1 M, pH 8.5) and incubated at 30 °C and 170 rpm; error bars show the standard deviation of three experiments.</figcaption></figure>

cells in aqueous ammonium formate buffer (1 M, pH 8.5) containing 100 mM rac-2b, and we estimated the cell survival over time as measures of colony-forming units per milliliter of sample (cfu mL⁻¹) (Figure 4F). This buffer solution was chosen since it is commonly used for the biocatalytic amination of ketones employing oxidoreductases.²⁷,²⁸ As expected, the survivability of the adapted cells was superior to that of the nonadapted cells. After 24 h, the cfu mL⁻¹ count for the JHX 4.2.8 cells was 6 orders of magnitude greater compared with the BL21(DE3) cells. Additionally, when the resting cells were incubated in the same buffer solution without rac-2b, after 24 h, the survival for both wild-type and JHX 4.2.8 cells remained high (i.e., > 10⁷ cfu mL⁻¹) and, interestingly, the cfu mL⁻¹ count after 18 and 24 h was approximately one or 2 orders of magnitude higher for the adapted cells (Figure S15). These findings suggest that, in the resting cell state, both strains tolerate ammonium formate buffer (1 M, pH 8.5) to a sufficient extent to be used in biocatalytic applications in vivo, employing this buffer as reaction medium.

**Biotransformations with Adapted *E. coli* Cells.** A chimeric AmDH obtained *via* domain shuffling of two first-generation variants (Ch1-AmDH),²⁹ previously shown to catalyze the amination of ketones **1–4a** and **7a**, was chosen to test the potential biocatalytic applicability of our adapted cells. This NADH-dependent oxidoreductase is a robust enzyme known to be active on acetophenone and derivatives thereof,²⁷,²⁹ phenylacetone derivatives,²⁷ and aliphatic ketones.²¹,²⁷,³⁰ His-tagged Ch1-AmDH was cloned into the pET28b plasmid, and the resulting construct transformed into chemically competent *E. coli* BL21(DE3) cells and the adapted derivatives JHX 1.2.3 and JHX 4.2.8. The whole-cell protein

expression profile of the resulting strains, named WT His-Ch1, J123 His-Ch1, and J428 His-Ch1, respectively (Table S3, entries 1–3), demonstrated that Ch1-AmDH was efficiently expressed with comparable expression levels in all strains (Figure S1).

The bioamination of the studied ketones was assayed using resting cells harvested from induced cultures of these strains expressing AmDH activity. The biocatalytic reactions were performed using a cell loading of 60 mg mL⁻¹ cww in ammonium chloride buffer (1 M, pH 8.7) and monitored by GC-FID (Tables S4 and S5). Two structurally different ketones, **2a** and **7a** (20 mM), were initially investigated. The noncommercially available amine **7b** was biocatalytically synthesized to quantify the formation of **7b** in the studied reactions (Figure S2). Independent of the strain being tested, **2a** was not converted into the related amine, while **7a** was converted to a very limited extent (<2% analytical yield, Table S6). We speculated that the bioamination reaction could be restricted by an insufficient amount of NADH in the cells. In *E. coli*, the aerobic metabolism of *D*-glucose causes the net and gradual formation of NADH *via* glycolysis and the tricarboxylic acid cycle.³¹ Thus, we tested the bioamination of **2–3a** and **7a** (20–50 mM) by adding *D*-glucose. However, the same previously unsatisfactory results were observed (Table S7). Neither the external supplementation of NAD⁺ or NADH (1 mM) nor the addition of glycerol (1–10 equiv)—a carbon source for *E. coli*, more reduced than glucose—resulted in different outcomes (Tables S8 and S9).

Overall, these findings indicated that under the tested conditions NADH could not be sufficiently regenerated *via* *E. coli* metabolism regardless of the amount of *D*-glucose or
glycerol added and the addition of a catalytic amount of the nicotinamide cofactor. This NADH insufficiency may stem from the cellular stress to which the resting cells are subjected during the reaction, resulting in an oxidative cell environment that favors the presence of NAD⁺ over NADH. Indeed, depending on environmental factors such as temperature, medium, or aeration, NAD⁺ is predominant in *E. coli* with a NAD⁺/NADH ratio ranging from 8 to 43.³²⁻³⁵ Moreover, the enzymes of the central metabolism in resting cells are still active, and thus an introduced biosynthetic pathway may compete with the native enzymes for energy and even substrates.¹⁷ Finally, the cofactor availability could be limited due to partial membrane permeabilization under bioproduction conditions.³⁶ Limitations linked to ketone toxicity were unlikely, as, for example, the loaded concentration of 2a in most of our reactions was lower than its respective MIC on adapted and nonadapted cells (i.e., 32 mM).

While the application of whole-cell catalysts without additional enzymes for cofactor recycling may be possible, enhanced biocatalytic activities have been reported for *E. coli* resting cells coexpressing an enzymatic system for nicotinamide cofactor recycling.⁴⁹,³⁷,³⁸ We chose to coexpress Ch1-AmDH and the FDH from *Candida boidinii* (Cb-FDH)³⁹ that catalyzes the NAD⁺-dependent oxidation of formate to hydrogen carbonate for the regeneration of NADH. Both enzymes were cloned into *E. coli* BL21(DE3) and JHX 4.2.8 cells employ either two single expression vectors or a duet vector harboring both genes. Strains harboring the duet vector (Table S3, entries 4–7) showed efficient expression of Cb-FDH; however, the expression of Ch1-AmDH was low (Figure S1). Conversely, the coexpression from two single vectors (Table S3, entries 8–9) resulted in the preferential expression of Ch1-AmDH over Cb-FDH. Indeed, we observed that Ch1-AmDH was efficiently expressed in *E. coli* only when the related gene bore an N-terminal His-tag. Strains coexpressing both oxidoreductases all showed a similar expression pattern between *E. coli* BL21(DE3) and JHX 4.2.8 (Figure S1). The biocatalytic amination of carbonyl compounds using purified AmDHs and Cb-FDH *in vitro* necessitated a higher molar concentration of the former over the latter enzyme to achieve high conversions.²⁷,⁴⁰ However, at this stage, all of the constructed recombinant strains were tested for best bioconversion.

We initially assayed the reactivities of the strains carrying each of the different plasmid systems used for coexpression, namely, strains Ch1/Cb, His-Ch1/Cb, and His-Ch1/His-Cb. The reactions were performed using resting cells (60 mg mL⁻¹ cww) and ketone 2a (20–50 mM) in aqueous ammonia/ ammonium formate buffer (1 M, pH 8.5), which acts both as formate and nitrogen source (Figure 5A). The analytical yield and enantiomeric excess were determined by GC-FID (Figures S3–S9). Notably, the use of the new recombinant strains resulted in the formation of amine product 2b, without detectable formation of the related alcohol, a potential side product of the *E. coli* metabolism. As expected, regardless of the substrate loading and use of adapted or nonadapted cells, the highest analytical yields (75–95%) occurred with His-Ch1/His-Cb cells—preferentially expressing Ch1-AmDH over Cb-FDH—while only moderate (27–66%) and low (13–30%) analytical yields were obtained with His-Ch1/Cb and Ch1/Cb cells, respectively. We attributed the observed reactivities to the different expressions of both oxidoreductases resulting from the employed plasmid systems and the presence

of a His-tag on the Ch1-AmDH gene. In effect, as determined by densitometry of native polyacrylamide gels, the average Ch1-AmDH to Cb-FDH ratio measured for His-Ch1/His-Cb cells was 10.2, whereas that for Ch1/Cb and His-Ch1/Cb cells was only 0.3 and 0.2, respectively (Figure S1). For comparison, the ratios used for *in vitro* reactions (i.e., using purified enzymes) range between 2.1 and 9.3.²⁷ Based on bioconversion results obtained for ketones 2–3a and 7a, His-Ch1/His-Cb strains were preferred over His-Ch1/Cb strains in further experiments (Figure S16).

Figure 5A,B depicts that, in general, the J428 His-Ch1/His-Cb strain led to higher conversions compared to the nonadapted counterpart. However, the analytical yield, but not the amount, of amine decreased for both strains as the ketone substrate concentration was augmented from 50 to 300 mM. Interestingly, each of the three tested ketones (2–3a and 7a) exhibited a different concentration-dependent bioconversion pattern. In fact, the greater gains in bioamination activity with adapted cells were observed in reactions at 100 and 200 mM 2a and 3a, respectively. Moreover, for the adapted cells, the conversion of 2a was elevated up to 100 mM of substrate and then decreased at higher substrate concentrations. The elevated tolerance of strain JHX 4.2.8 to amine 2b (MIC 96 mM) might be directly linked to this result, but the complexity of whole-cell transformations and still limited understanding of the associated microbial physiology make any statement of correlation hypothetical.⁴¹ Aromatic ketone 7a, however, was poorly aminated (i.e., <20% analytical yield) at 200 mM concentration, regardless of the tested strain.

To examine the progress of the bioamination and to test whether the reaction suffers from environmental effects, we monitored the formation of 2b over time and measured the survival rate of the cells in the reaction mixture at each time point. The bioamination of 2a (100 mM) with J428 His-Ch1/His-Cb cells plateaued approximately at 70% yield after 16 h, while the reaction catalyzed by nonadapted His-Ch1/His-Cb cells merely reached about 20% yield in 2b after the same reaction time (Figure 5C). Remarkably, at 2 h of reaction, a sharp and similar decrease in cell survival was observed for both strains (Figure 5D). Subsequently, the cell survival continued decreasing until complete loss of viability for both adapted and nonadapted cells after 18 h. Notably, after 4 h of reaction, the survivability of the adapted cells was superior to that of the nonadapted, with a cfu mL⁻¹ count higher by approximately 1 order of magnitude. Factually, this higher population density and longer survival correlate somehow with the accumulation of amine 2b through amination of the ketone substrate. Strain JHX 4.2.8 is considerably more tolerant to amine 2b than its nonadapted counterpart. But many parameters may come into play, including increased cell permeability or higher stability of the internal cofactor in the adapted cells. Indeed, besides the ability to survive in the reaction medium, an efficient whole-cell biocatalyst should allow the efficient transport of both substrate(s) and product(s) across the cell membrane, should not consume either of them in side reactions, and should be able to provide the necessary cofactors.⁴² The results depicted in Figure 5C,D revealed that reaching a conversion plateau coincided with the complete decline in cell survival observed for both strains.

Next, we evaluated whether higher cell loadings could improve the conversion of 2a (Figure 5E). Reactions at 100 mM 2a showed minimum analytical yield variations regardless of the cell concentration of each tested strain; moreover, the
<figure><img src="image_15.png" /><figcaption>**Figure 6.** (A) Bioamination of 1–2a and 4a at varied substrate concentrations (50–300 mM) in HCOONH₄/NH₄OH (1 M, pH 8.5) with 1 mM NAD⁺. (B) Bioamination of 2a (400–500 mM) in HCOONH₄/NH₄OH (1 M, pH 8.5) with (+) and without (−) 1 mM NAD⁺. In (A,B), cell loading is 60 mg mL⁻¹ cww, reaction volume is 1 mL, and incubation was done at 30 °C and 170 rpm for 24 h. Error bars indicate the standard deviation of three experiments.</figcaption></figure>

<table><caption>**Table 1.** Maximum Amine Analytical Yields at the Specified Substrate Concentration for the Bioamination of Each Ketone Assayed in This Study Using Adapted and Non-adapted Cells</caption><thead><tr><th>entry</th><th>strain</th><th>substrate</th><th>substrate concentration (mM, g L<sup>-1</sup>)</th><th>analytical yield (%)<sup>a</sup></th><th>product concentration (g L<sup>-1</sup>)</th><th>STY (g L<sup>-1</sup> h<sup>-1</sup>)<sup>b</sup></th><th>ee %<sup>c</sup></th></tr></thead><tbody><tr><td>1</td><td>WT His-Ch1/His-Cb</td><td>1a</td><td>50, 4.3</td><td>24 ± 1</td><td>0.9 ± 0.1</td><td>0.06</td><td>&gt;98.3 (R)</td></tr><tr><td>2</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>21 ± 1</td><td>0.8 ± 0.1</td><td>0.05</td><td>&gt;98.8 (R)</td></tr><tr><td>3</td><td>WT His-Ch1/His-Cb</td><td>2a</td><td>100, 10</td><td>23 ± 1</td><td>1.9 ± 0.1</td><td>0.12</td><td>&gt;99.6 (R)</td></tr><tr><td>4</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>75 ± 7</td><td>7.5 ± 0.7</td><td>0.47</td><td>&gt;99.9 (R)</td></tr><tr><td>5</td><td>WT His-Ch1/His-Cb</td><td>3a</td><td>200, 22.8</td><td>33 ± 3</td><td>6.5 ± 0.5</td><td>0.40</td><td>&gt;99.9 (R)</td></tr><tr><td>6</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>75 ± 5</td><td>15.2 ± 1.0</td><td>0.95</td><td>&gt;99.9 (R)</td></tr><tr><td>7</td><td>WT His-Ch1/His-Cb</td><td>4a</td><td>100, 12.8</td><td>55 ± 14</td><td>5.2 ± 1.4</td><td>0.33</td><td>&gt;99.8 (R)</td></tr><tr><td>8</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>82 ± 7</td><td>8.5 ± 0.8</td><td>0.53</td><td>&gt;99.9 (R)</td></tr><tr><td>9</td><td>WT His-Ch1/His-Cb</td><td>5a</td><td>100, 10</td><td>16 ± 1</td><td>1.5 ± 0.1</td><td>0.09</td><td>&gt;99.5 (R)</td></tr><tr><td>10</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>43 ± 4</td><td>3. ± 0.3</td><td>0.23</td><td>&gt;99.8 (R)</td></tr><tr><td>11</td><td>WT His-Ch1/His-Cb</td><td>6a</td><td>100, 11.4</td><td>69 ± 4</td><td>7.8 ± 0.3</td><td>0.49</td><td>&gt;99.9 (R)</td></tr><tr><td>12</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>98 ± 1</td><td>11.2 ± 0.2</td><td>0.70</td><td>&gt;99.9 (R)</td></tr><tr><td>13</td><td>WT His-Ch1/His-Cb</td><td>7a</td><td>50, 7.6</td><td>83 ± 4</td><td>6.5 ± 0.3</td><td>0.41</td><td>&gt;99.9 (R)</td></tr><tr><td>14</td><td>J428 His-Ch1/His-Cb</td><td></td><td></td><td>96 ± 1</td><td>7.4 ± 0.2</td><td>0.46</td><td>&gt;99.9 (R)</td></tr></tbody></table>

<sup>a</sup>Reaction conditions: the indicated resting cells (60 mg mL⁻¹, cww), and the specified concentration of 1–7a in HCOONH₄/NH₄OH (1 M, pH 8.5); reaction volume 1 mL; incubation at 30 °C and 170 rpm for 24 h. Analytical yield determined by GC-FID using a DB-1701 column (Agilent).

<sup>b</sup>Space-time yield (STY) assuming that the conversions all plateaued after 16 h of reaction. <sup>c</sup>Determined by GC-FID using a Chrompack Chirasil-DEX CB column (Agilent). In all entries, the level of precision in the ee % calculation depends on the instrumental detection limit (i.e., 2 area units) rather than the intrinsic stereoselectivity of the biocatalyst, which was virtually perfect (the S-enantiomer was never detected).

nonadapted cells could not meet the performance of the adapted ones even at high cell concentrations. In contrast, at 200 mM 2a, the amine formation augmented as the cell loading was increased for both strains. The highest observed analytical yield occurred at 100 mg mL⁻¹ cww with the adapted cells, which outperformed the nonadapted ones by ca. 20% (net analytical yield increase). We next assayed the bioamination of additional substrates (3–4a, 200 mM) at a 100 mg mL⁻¹ cell loading (Figure 5F). Under these conditions, the J428 His-Ch1/His-Cb strain led to ca. 20% higher production of amines 3–4b compared to the WT His-Ch1/His-Cb strain, while the conversion of 7a by both strains was moderate (i.e., <30% analytical yield).

For a better understanding of our biocatalytic system, we conducted additional experiments in which external NAD⁺ (1

mM) was supplemented with the biotransformation medium. The addition of NAD⁺ resulted in elevated conversions with both adapted and nonadapted cells at substrate concentrations as high as 300 mM (Figure 6A). In the presence of external NAD⁺, elevated analytical yields (>70%) were obtained for ketones 2–3a (300 mM) with both adapted and nonadapted cells. However, the addition of cofactor provoked a significant increase in the bioconversion of 7a at high concentrations only when the adapted cells were used. Reactions performed at even higher substrate loadings (2a, 400–500 mM) still resulted in moderate analytical yields with both tested strains only when NAD⁺ was added (Figure 6B).

Given that nicotinamide adenine dinucleotide cofactors are not naturally transported through the E. coli cell membrane, we attributed these observations to a certain degree of cell lysis,
which occurred under the conditions of the bioamination (alkaline pH, salinity, temperature, and presence of the ketone substrate and, in nonadapted cells at least, of the amine product). Additional experiments suggested that there is an extracellular contribution to the observed amine formation (Figure S17), which was a lot more pronounced when NAD⁺ was added, as evidenced by the bioconversion results after 4 and 6 h (Figure S17A). Figure S17B, which was obtained by loading equivalent amounts of sample, also showed that the amount of extracellularly released enzymes gradually increased over time, in particular after 4 h.

The bioconversion results obtained with the addition of NAD⁺, although important for a more accurate characterization of our system, are not very conducive for the purpose of this study. The addition of NAD⁺ boosted the conversion promoted by Ch1-AmDH and Cb-FDH that were released into the medium upon cell death and lysis. Logically, the concentration of catalytically active NAD⁺ coming from the lysed cells was greatly diluted in the free medium. When extracellular NAD⁺ was added to the free medium, the released enzymes could perform the reductive amination of the substrate more efficiently, thereby leading to higher analytical yields. The occurrence of a cell lysis phenomenon in resting cell systems may have been exploited unwittingly in some studies. For example, the use of cell-free extracts of glucose dehydrogenase coupled to living whole-cells expressing heterologous dehydrogenases may have implied the release of intracellular materials to achieve substrate bioconversions.⁴³,⁴⁴ We remark that the major aim of the present study was to engineer *E. coli* cells that are tolerant to the toxic amine so that the intracellular reductive amination reaction would be enhanced, as illustrated in Figure 5. Future studies will aim at utilizing the adapted *E. coli* cells in a fermentative mode for which growing cells are used. As shown in the current study using resting *E. coli* cells, further engineering is needed to achieve this objective. However, it is already very remarkable that *E. coli* cells could be evolved as shown in this study to perform reductive amination at high substrate concentrations utilizing resting cells without the need for supplementation of exogenous NAD⁺. This represents a significant achievement considering the inherent cost of NAD(P)H cofactors and intrinsic enhancement of intracellular events related to the presence of high amine concentrations, enabling higher amine formation.

Table 1 summarizes the maximum amine product concentrations (in g L⁻¹) obtained at 60 mg mL⁻¹ cell loading without externally added NAD⁺ for each substrate assayed in this study. Except for ketone **1a**, the adapted cells resulted in up to three times higher analytical yields compared to the nonadapted ones. A substrate-dependent effect was observed—medium-size aliphatic ketones (2–4a and 6a) were preferentially aminated over shorter (1a and 5a) or aromatic (7a) substrates, presumably due to the cell adaptation being conducted with amine **2b**. Notably, our amine-tolerant cells resulted in superior amine titers (1–15.2 g L⁻¹) compared to various studies on whole-cell-catalyzed aminations employing living *E. coli* cells (1 g L⁻¹ range; Table S10). Our amine product concentrations were up to 12 mM for amine **1b**, 75 mM for amine **2b**, 150 mM for amine **3b**, 82 mM for amine **4b**, 43 mM for amine **5b**, 98 mM for amine **6b**, and 48 mM for amine **7b**. Remarkably, the stereoselectivity in all tested reactions with adapted and nonadapted cells was virtually

perfect (ee ≥ 99.3%, R), which matches the ees reported in *vitro* using Ch1-AmDH.²⁷,²⁹,³⁰

To demonstrate the potential applicability of the ketone bioamination with amine-tolerant *E. coli* cells (His-Ch1/His-Cb) for larger-scale production of enantiopure amines, we performed a preparative biotransformation with 200 mM **3a** (1.6 g, 14 mmol). In a 70 mL resting cell suspension, (R)-**3b** was obtained in 42 ± 4 and 65 ± 5% analytical yield employing nonadapted and adapted cells, respectively. As already reported, amine recovery from the reaction mixtures was only partial due to the high amine volatility, resulting in approximately 20% isolated yield for the nonadapted cells and 31% for the adapted ones, yet the product was obtained with high chemical (>97%, GC) and optical purity (ee > 99%, R) (Figure S10). This result is a significant advancement for reductive amination bioprocesses. In fact, our reference work by Houwman *et al*.²¹ reported bioamination of 20 mM of hexan-2-ol with 40% conversion at 250 mL scale, while the scarce reported biotransformations in fed-batch mode with living whole-cells harboring amine-forming enzymes are usually performed at <10 mL scale and/or low substrate concentration.¹⁹,⁴⁵

<figure><img src="image_16.png" /></figure>

## CONCLUSIONS

In this work, we developed amine-tolerant *E. coli* strains *via* adaptive directed evolution in continuous culture. These adapted strains exhibited significantly higher tolerance to various amines compared to that of the wild-type strain. Using an AmDH/FDH system, the adapted cells, in their resting state, enabled the bioamination of prochiral ketones with high chemical and optical purity, achieving product yields up to more than three times higher (up to 15 g L⁻¹) than those with nonadapted cells. Most tested ketones yielded at least 70% amine products, depending on the substrate concentration, without needing NAD⁺ cofactor supplementation. The adapted cells showed longer survival and higher population density, which directly or indirectly enabled a more sustained bioconversion. These amine-tolerant cells could be broadly applied in any whole-cell-based reaction involving amines. Future studies will identify the mutations responsible for this high amine tolerance and examine their impact on cell physiology and metabolism to understand the adaptation mechanisms.

<figure><img src="image_17.png" /></figure>

## ASSOCIATED CONTENT

<figure><img src="image_18.png" /></figure>

### Supporting Information

The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acssuschemeng.4c04356.

GC chromatograms and ¹H NMR spectra of biocatalytically synthesized compounds and materials and methods
(PDF)

<figure><img src="image_19.png" /></figure>

## AUTHOR INFORMATION

### Corresponding Authors

Francesco G. Mutti — Van't Hoff Institute for Molecular Sciences, HIMS-Biocat, University of Amsterdam, Science Park 904, Amsterdam 1098 XH, The Netherlands;
● orcid.org/0000-0002-6771-5102; Email: F.Mutti@uva.nl

Carine Vergne-Vaxelaire — Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry 91057, France; ● orcid.org/
## Authors

Josemarco Mendoza-Avila — Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry 91057, France; Van 't Hoff Institute for Molecular Sciences, HIMS-Biocat, University of Amsterdam, Science Park 904, Amsterdam 1098 XH, The Netherlands

Volker Döring — Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry 91057, France

Madeleine Bouzon — Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry 91057, France; orcid.org/0000-0002-9581-6584

Ivan Dubois — Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry 91057, France

Tanja Knaus — Van 't Hoff Institute for Molecular Sciences, HIMS-Biocat, University of Amsterdam, Science Park 904, Amsterdam 1098 XH, The Netherlands; orcid.org/0000-0001-9942-9226

Louis M. M. Mouterde — URD Agro-Biotechnologies Industrielles, CEBB, AgroParisTech, Pomacle 51110, France; orcid.org/0000-0002-4096-1629

Anne Zaparucha — Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry 91057, France; orcid.org/0000-0003-2531-4148

Complete contact information is available at:
https://pubs.acs.org/10.1021/acssuschemeng.4c04356

## Author Contributions

C.V.V., F.M., and V.D. conceived the project. C.V.V., F.M., and L.M. managed it with input from V.D., M.B., T.K., and A.Z. I.D. and J.M.-A. performed the continuous cultures. J.M.-A. carried out the gene cloning, protein expression, the different cell characterizations, toxicity assays, and biotransformations with input from T.K. J.M.-A., C.V.V., and F.M. wrote the manuscript with input from V.D., L.M., M.B., and A.Z.

## Funding

This work was supported by the "ADI 2021" project funded by the IDEX Paris-Saclay, ANR-11-IDEX-0003-02.

## Notes

The authors declare no competing financial interest.

## ACKNOWLEDGMENTS

The authors thank Philippe Marlière, who historically conceived the experimental principle used in this publication. The GENEMAT automates were codeveloped by Commissariat à l'énergie atomique et aux énergies alternatives (CEA) and Philippe Marlière and are co-owned by CEA and ALTAR. The authors thank Alice Girolamo (University of Amsterdam) for her assistance in the laboratory and fruitful discussions. The authors acknowledge Florent Allais (AgroParisTech) for his comments on the manuscript. This work was supported by CEA, CNRS, and the University of Evry Val d'Essonne-University of Paris-Saclay.

## ABBREVIATIONS

<table><tr><td>NAD(P)(H)</td><td>(reduced form)</td></tr><tr><td>AmDH</td><td>amine dehydrogenase</td></tr><tr><td>IRED</td><td>imine reductase</td></tr><tr><td>RedAm</td><td>reductive aminase</td></tr><tr><td>TA</td><td>transaminase</td></tr><tr><td>ADH</td><td>alcohol dehydrogenase</td></tr><tr><td>FDH</td><td>formate dehydrogenase</td></tr><tr><td>E. coli</td><td>Escherichia coli</td></tr><tr><td>LB</td><td>Luria-Bertani</td></tr><tr><td>OD</td><td>optical density</td></tr><tr><td>MIC</td><td>minimum inhibitory concentration</td></tr><tr><td>IPTG</td><td>isopropyl β-D-1-thiogalactopyranoside</td></tr><tr><td>SDS-PAGE</td><td>sodium dodecyl sulfate polyacrylamide gel electrophoresis</td></tr><tr><td>GC-FID</td><td>gas chromatography coupled to flame ionization detector</td></tr><tr><td>cfu</td><td>colony forming units</td></tr></table>

## REFERENCES

(1) Turner, N. J.; Truppo, M. D. Biocatalytic Routes to Nonracemic Chiral Amines. In *Chiral Amine Synthesis*; Nugent, T. C., Ed.; Wiley VCH, 2010; pp 431–459.

(2) Abdelraheem, E.; Damian, M.; Mutti, F. G. Biocatalytic Amine Synthesis. In *Comprehensive Chirality*; Cossy, J., Ed.; Academic Press, 2022; pp 210–304.

(3) Nugent, T. C. Asymmetric Reductive Amination. In *Chiral Amine Synthesis*; Nugent, T. C., Ed.; Wiley, 2010; pp 225–245.

(4) Ducrot, L.; Bennett, M.; Grogan, G.; Vergne-Vaxelaire, C. NAD(P)H-Dependent Enzymes for Reductive Amination: Active Site Description and Carbonyl-Containing Compound Spectrum. *Adv. Synth. Catal.* **2021**, *363* (2), 328–351.

(5) Mutti, F. G.; Knaus, T. Enzymes Applied to the Synthesis of Amines. In *Biocatalysis for Practitioners*; De Gonzalo, G., Lavandera, I., Eds.; Wiley, 2021; pp 143–180.

(6) Sharma, M.; Mangas-Sanchez, J.; Turner, N. J.; Grogan, G. NAD(P)H-Dependent Dehydrogenases for the Asymmetric Reductive Amination of Ketones: Structure, Mechanism, Evolution and Application. *Adv. Synth. Catal.* **2017**, *359* (12), 2011–2025.

(7) Fuchs, M.; Farnberger, J. E.; Kroutil, W. The Industrial Age of Biocatalytic Transamination. *Eur. J. Org Chem.* **2015**, *2015*, 6965–6982.

(8) Guo, F.; Berglund, P. Transaminase biocatalysis: optimization and application. *Green Chem.* **2017**, *19* (2), 333–360.

(9) Busto, E.; Gotor-Fernández, V.; Gotor, V. Hydrolases in the Stereoselective Synthesis of N-Heterocyclic Amines and Amino Acid Derivatives. *Chem. Rev.* **2011**, *111* (7), 3998–4035.

(10) Verho, O.; Bäckvall, J. E. Chemoenzymatic Dynamic Kinetic Resolution: A Powerful Tool for the Preparation of Enantiomerically Pure Alcohols and Amines. *J. Am. Chem. Soc.* **2015**, *137* (12), 3996–4009.

(11) Parmeggiani, F.; Weise, N. J.; Ahmed, S. T.; Turner, N. J. Synthetic and Therapeutic Applications of Ammonia-lyases and Aminomutases. *Chem. Rev.* **2018**, *118* (1), 73–118.

(12) Turner, N. J. Enantioselective Oxidation of C–O and C–N Bonds Using Oxidases. *Chem. Rev.* **2011**, *111* (7), 4073–4087.

(13) Schmidt, N. G.; Eger, E.; Kroutil, W. Building Bridges: Biocatalytic C–C-Bond Formation toward Multifunctional Products. *ACS Catal.* **2016**, *6* (7), 4286–4311.

(14) Prier, C. K.; Zhang, R. K.; Buller, A. R.; Brinkmann-Chen, S.; Arnold, F. H. Enantioselective, intermolecular benzylic C–H amination catalysed by an engineered iron-haem enzyme. *Nat. Chem.* **2017**, *9* (7), 629–634.
(15) Hollmann, F.; Opperman, D. J.; Paul, C. E. Biocatalytic Reduction Reactions from a Chemist's Perspective. *Angew. Chem., Int. Ed.* **2021**, *60* (11), 5644–5665.

(16) Klatte, S.; Lorenz, E.; Wendisch, V. F. Whole cell biotransformation for reductive amination reactions. *Bioengineered* **2014**, *5* (1), 56–62.

(17) Lin, B.; Tao, Y. Whole-cell biocatalysts by design. *Microb. Cell Fact.* **2017**, *16* (1), 106.

(18) Hepworth, L. J.; France, S. P.; Hussain, S.; Both, P.; Turner, N. J.; Flitsch, S. L. Enzyme Cascades in Whole Cells for the Synthesis of Chiral Cyclic Amines. ACS Catal. **2017**, *7* (4), 2920–2925.

(19) Farnberger, J. E.; Lorenz, E.; Richter, N.; Wendisch, V. F.; Kroutil, W. In vivo plug-and-play: a modular multi-enzyme single-cell catalyst for the asymmetric amination of ketoacids and ketones. *Microb. Cell Fact.* **2017**, *16* (1), 132.

(20) Kim, D. I.; Chae, T. U.; Kim, H. U.; Jang, W. D.; Lee, S. Y. Microbial production of multiple short-chain primary amines via retrobiosynthesis. *Nat. Commun.* **2021**, *12* (1), 173.

(21) Houwman, J. A.; Knaus, T.; Costa, M.; Mutti, F. G. Efficient synthesis of enantiopure amines from alcohols using resting E. coli cells and ammonia. *Green Chem.* **2019**, *21* (14), 3846–3857.

(22) Sun, Z.; Hübner, R.; Li, J.; Wu, C. Artificially sporulated Escherichia coli cells as a robust cell factory for interfacial biocatalysis. *Nat. Commun.* **2022**, *13* (1), 3142.

(23) Gao, L.; Feng, L.; Sauer, D. F.; Wittwer, M.; Hu, Y.; Schiffels, J.; Li, X. Engineered living hydrogels for robust biocatalysis in pure organic solvents. *Cell Rep. Phys. Sci.* **2022**, *3* (10), 101054.

(24) Li, J.; Guo, Y.; Li, J.; Zhang, H.; Wang, L.; Liang, C. Improved stress tolerance of recombinant Escherichia coli strain expressing nitrile hydratase by combining Global Regulator IrrE. *Mol. Catal.* **2024**, *557*, 113904.

(25) Mutzel, R.; Marlière, P. Method and device for selecting accelerated proliferation of living cells in suspension. WO 2000034433 A1, 2000.

(26) Marlière, P.; Patrouix, J.; Döring, V.; Herdewijn, P.; Tricot, S.; Cruveiller, S.; Bouzon, M.; Mutzel, R. Chemical Evolution of a Bacterium's Genome. *Angew. Chem., Int. Ed.* **2011**, *50* (31), 7109–7114.

(27) Knaus, T.; Böhmer, W.; Mutti, F. G. Amine dehydrogenases: efficient biocatalysts for the reductive amination of carbonyl compounds. *Green Chem.* **2017**, *19* (2), 453–463.

(28) Ducrot, L.; Bennett, M.; André-Leroux, G.; Elisée, E.; Marynberg, S.; Fossey-Jouenne, A.; Zaparucha, A.; Grogan, G.; Vergne-Vaxelaire, C. Expanding the Substrate Scope of Native Amine Dehydrogenases through In Silico Structural Exploration and Targeted Protein Engineering. *ChemCatChem* **2022**, *14* (22), No. e202200880.

(29) Bommarius, B. R.; Schürmann, M.; Bommarius, A. S. A novel chimeric amine dehydrogenase shows altered substrate specificity compared to its parent enzymes. *Chem. Commun.* **2014**, *50* (95), 14953–14955.

(30) Mutti, F. G.; Knaus, T.; Scrutton, N. S.; Breuer, M.; Turner, N. J. Conversion of alcohols to enantiopure amines through dual-enzyme hydrogen-borrowing cascades. *Science* **2015**, *349* (6255), 1525–1529.

(31) Lodish, H.; Berk, A.; Matsudaira, P.; Kaiser, C. A.; Krieger, M. P.; Scott, M.; Zipursky, L.; Darnell, J. *Mol. Cell. Biol.*, 5th ed.; W. H. Freeman, 2003; pp 304–311.

(32) Bautista, J.; Satrüstegui, J.; Machado, A. Evidence suggesting that the NADPH/NADP ratio modulates the splitting of the isocitrate flux between the glyoxylic and tricarboxylic acid cycles, in Escherichia coli. *FEBS Lett.* **1979**, *105* (2), 333–336.

(33) Chassagnole, C.; Noisommit-Rizzi, N.; Schmid, J. W.; Mauch, K.; Reuss, M. Dynamic modeling of the central carbon metabolism of Escherichia coli. *Biotechnol. Bioeng.* **2002**, *79* (1), 53–73.

(34) Henry, C. S.; Broadbelt, L. J.; Hatzimanikatis, V. Thermodynamics-Based Metabolic Flux Analysis. *Biophys. J.* **2007**, *92* (5), 1792–1805.

(35) Bennett, B. D.; Kimball, E. H.; Gao, M.; Osterhout, R.; Van Dien, S. J.; Rabinowitz, J. D. Absolute metabolite concentrations and

implied enzyme active site occupancy in Escherichia coli. *Nat. Chem. Biol.* **2009**, *5* (8), 593–599.

(36) Walton, A. Z.; Stewart, J. D. Understanding and Improving NADPH-Dependent Reactions by Nongrowing Escherichia coli Cells. *Biotechnol. Prog.* **2004**, *20* (2), 403–411.

(37) Lee, W.-H.; Park, J.-B.; Park, K.; Kim, M.-D.; Seo, J.-H. Enhanced production of ε-caprolactone by overexpression of NADPH-regenerating glucose 6-phosphate dehydrogenase in recombinant Escherichia coli harboring cyclohexanone monooxygenase gene. *Appl. Microbiol. Biotechnol.* **2007**, *76* (2), 329–338.

(38) Jeon, H.; Yoon, S.; Ahsan, M. M.; Sung, S.; Kim, G.-H.; Sundaramoorthy, U.; Rhee, S.-K.; Yun, H. The Kinetic Resolution of Racemic Amines Using a Whole-Cell Biocatalyst Co-Expressing Amine Dehydrogenase and NADH Oxidase. *Catalysts* **2017**, *7* (9), 251.

(39) Schütte, H.; Flossdorf, J.; Sahm, H.; Kula, M.-R. Purification and Properties of Formaldehyde Dehydrogenase and Formate Dehydrogenase from Candida boidinii. *Eur. J. Biochem.* **1976**, *62* (1), 151–160.

(40) Tseliou, V.; Knaus, T.; Masman, M. F.; Corrado, M. L.; Mutti, F. G. Generation of amine dehydrogenases with increased catalytic performance and substrate scope from ε-deaminating L-Lysine dehydrogenase. *Nat. Commun.* **2019**, *10* (1), 3717.

(41) Kuhn, D.; Blank, L. M.; Schmid, A.; Bühler, B. Systems biotechnology — Rational whole-cell biocatalyst and bioprocess design. *Eng. Life Sci.* **2010**, *10* (5), 384–397.

(42) de Carvalho, C. C. R. Whole cell biocatalysts: essential workers from Nature to the industry. *Microb. Biotechnol.* **2017**, *10* (2), 250–263.

(43) Liu, J.; Li, Z. Enhancing cofactor recycling in the bioconversion of racemic alcohols to chiral amines with alcohol dehydrogenase and amine dehydrogenase by coupling cells and cell-free system. *Biotechnol. Bioeng.* **2019**, *116* (3), 536–542.

(44) Tong, F.; Qin, Z.; Wang, H.; Jiang, Y.; Li, J.; Ming, H.; Qu, G.; Xiao, Y.; Sun, Z. Biosynthesis of Chiral Amino Alcohols via an Engineered Amine Dehydrogenase in E. coli. *Front. Bioeng. Biotechnol.* **2022**, *9*, 778584.

(45) Leipold, F.; Hussain, S.; Ghislieri, D.; Turner, N. J. Asymmetric Reduction of Cyclic Imines Catalyzed by a Whole-Cell Biocatalyst Containing an (S)-Imine Reductase. *ChemCatChem* **2013**, *5* (12), 3505–3508.