Main
The ability to exert precise and predictable control over cellular behavior is a central goal of synthetic biology. In metabolic engineering, this control is often achieved by coordinating the activity of multiple transcription units (TUs), each consisting of a gene and its regulatory elements. Synthetic metabolic pathways are frequently constructed by transplanting coding sequences (CDSs) from diverse species into orthogonal hosts, thereby endowing them with new metabolic capabilities. This approach has yielded notable successes, including the development of cell-based alternatives to industrial manufacturing processes, the stabilization of pharmaceutical supply chains previously dependent on seasonal plant sources and the capture of atmospheric carbon dioxide to generate industrially valuable biomass1,2,3. While these strategies have been extensively demonstrated in microbial chassis, examples of metabolic engineering at this pathway scale in mammalian cells remain rare4,5, despite broad recognition that improved control of mammalian cell function could enhance cell therapies, produce increasingly complex biologics and improve the scalable manufacture of viral vectors6,7,8. Modern DNA synthesis technologies now make it possible to incorporate sequences from across the phylogenetic tree or de novo designed sequences, vastly expanding the functional landscape accessible to mammalian cells.
Despite this promise, the limited examples of mammalian metabolic engineering can in part be attributed to the lack of foundational knowledge needed to guide metabolic pathway design. Given a target metabolic pathway and a target host cell type, one is faced with a series of complex design decisions, including the selection of CDSs, the configuration of regulatory elements, the intracellular localization of each encoded protein and the relative expression levels and stoichiometry of each TU required for optimal function (Fig. 1a). These variables create a vast combinatorial design space, yet there is little mechanistic understanding to guide optimal choices.
a, Metabolic engineering benefits from increasing diversity across the different modules that make up TUs, including promoters, OLSs, enzymatic function, stoichiometry and copy number. b, Classical DBT frameworks in which pathways are tested in succession are inefficient in mammalian cells and other host chassis, which exhibit long doubling times or difficulties associated with engineering or delivering long DNAs that encode more complex biosynthetic pathways. c, SGE increases the scale of metabolic engineering by delivering barcoded individual TUs at high MOI such that each cell carries a unique pathway combination, thereby sampling much more pathway diversity. Following selection, cells displaying the desired phenotype can be sequenced to reveal TU combinations that conferred the desired function. Created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
In fast-growing, genetically tractable microbial systems such as Escherichia coli (20-min doubling time) or Saccharomyces cerevisiae (90-min doubling time), the lack of mechanistic knowledge can often be mitigated through the design–build–test (DBT) framework, where metabolic pathways are rapidly constructed, tested and iteratively refined until satisfactory performance is achieved9,10 (Fig. 1b). Applying similarly complex metabolic engineering strategies in mammalian cells is far more challenging, primarily because of much longer doubling times (typically over 20 h), which drastically slows the pace of iteration.
This limitation becomes increasingly severe as pathway complexity grows or as multiple functions are integrated into a single cell. While the final implementation of such complex functions may rely on recent advances in the synthesis and delivery of large DNA constructs to mammalian cells > 100 kb (ref. 11), these approaches are poorly suited for high-throughput screening, as delivering large DNA constructs into mammalian cells is highly inefficient. Modular combinatorial strategies, such as CLASSIC, offer a solution by enabling screening of ~105 constructs at the ~10-kb scale12 (2–3 TUs); however, there is a strong inverse relationship between DNA size and delivery success; it becomes exponentially more difficult to deliver larger constructs, rendering brute-force screening of complex synthetic designs impractical13.
To overcome the limitations of experimental screening in mammalian systems, some metabolic engineering approaches incorporate computational screening of candidate interventions before testing in the lab12,14. Sophisticated transcriptomic databases and metabolic models have been developed for commonly used mammalian cell types, enabling the identification of epistatic metabolic functions and the design of complex multiple-knockout cell lines that improve bioprocessing and manufacturability of antibodies15,16,17. However, these models are typically trained on observational data shaped by natural evolution and are optimized for a narrow range of species and cell types commonly used in biomanufacturing. As a result, they are poorly suited for generative design, including scenarios involving the introduction of heterologous or entirely synthetic TUs. The design of TU arrays that comprise synthetic metabolic pathways adds yet another layer of complexity, beyond what these models can currently handle. Thus, synthetic data are needed—systematic, large-scale metabolic perturbations that can be used to train machine learning models capable of predictive and design-oriented applications18.
Here, we introduce shotgun genetic engineering (SGE), a multiplexed approach to mammalian metabolic engineering that enables rapid discovery of complex pathway solutions and the creation of large-scale synthetic datasets for training predictive models. SGE leverages high-multiplicity integration of barcoded individual TUs from a pooled library, allowing distinct pathway variants to be randomly assembled within each cell. Because small DNA constructs are exponentially easier to synthesize, assemble and deliver, this design turns each cell into an independent experiment and enables millions of combinations to be screened in parallel. By selecting for cells with desired phenotypes and sequencing their associated barcodes, SGE bypasses the limitations of sequential DBT cycles and dramatically accelerates discovery timelines (Fig. 1c).
As proof of concept, we apply SGE to screen millions of pathway combinations for engineering two essential amino acid (EAA) biosynthetic pathways in mammalian cells. In prior work, we enabled Chinese hamster ovary (CHO) cells to proliferate without valine by introducing four E. coli genes, achieving a valine-free doubling time of 3.8 days5. However, we were unable to confer isoleucine independence, despite overlapping biosynthetic steps. Here, we use SGE to both reconstitute and optimize valine prototrophy in CHO cells, yielding clones with a near-wild-type doubling time of just 1.1 days. We also discover a functional six-gene solution enabling isoleucine prototrophy in CHO cells. In both cases, we saw strong signatures favoring mitochondrial localization of biosynthetic enzymes for optimized pathway functionality, indicating that organellar localization is an underexplored variable in mammalian metabolic engineering. Extending this approach, we introduce valine prototrophy to a human T lymphocyte (Jurkat) cell line—an initial demonstration of metabolic pathway expansion in immune cells, with potential applications in engineering stress-resilient cell therapies. Lastly, we show that SGE enables training of a machine learning classifier to identify genetic features predictive of pathway function. Together, these results establish SGE as a scalable framework for pathway-scale metabolic engineering in mammalian cells.
Results
Design of SGE
A central premise of SGE is that individual TUs are exponentially easier to synthesize, assemble and deliver than large multigene constructs. A pooled library of individually barcoded TUs is delivered at high multiplicity of infection (MOI), such that each cell receives a unique combination of pathway components and serves as an independent experiment19. This strategy avoids constructing and delivering each complete pathway individually, enabling scalable exploration of pathway composition, stoichiometry, expression and subcellular localization. To generate TU libraries, barcoded CDSs are synthesized with flanking type IIS restriction sites and directionally cloned into pooled vectors containing barcoded promoters and, where appropriate, N-terminal organellar localization signals (OLSs). Varying promoter strength enables optimization of pathway flux and mitigation of toxicity from pathway components or intermediates20,21,22,23,24, while organellar targeting can increase local enzyme concentrations or provide a favorable biochemical environment1,25. The CDS, promoter and OLS are assembled by Golden Gate cloning into lentiviral-compatible expression vectors, with identifying barcodes positioned within approximately 100 bp of one another in the 3′ untranslated region (UTR) for recovery by a single PCR amplicon.
The resulting library is delivered at high MOI, generating cells that vary in TU composition, copy number and stoichiometry. Cells are subjected to a selection or functional screen and enriched pathway configurations are identified by barcode sequencing (Fig. 1c). Although we use growth-based selection here, SGE is compatible with biosensors, fluorescence-activated cell sorting and other functional readouts. Libraries can also incorporate additional CDSs, promoter–OLS pairs, host factor perturbations or alternative delivery mechanisms (for example, transposons or recombinases).
We initially selected lentivirus because of the ease of titrating MOI and its broad host range, which allowed us to deploy the same libraries across multiple cell types. However, lentiviral delivery also comes with several practical limitations, including the need to perform packaging after library cloning, constraints on cargo size and potential template switching that can lead to barcode swapping26. Therefore, for some applications, especially when working with larger constructs or when maintaining barcode linkage is essential, a transposon-based strategy (for example, piggyBac) may be more suitable. Additionally, the incorporation of dual 5′ plus 3′ barcoding strategies as used in massively parallel reporter assays27 or direct long-read readouts may help mitigate barcode swapping concerns if lentivirus is to be used.
SGE valine/isoleucine library curation and construction
To confirm the feasibility of SGE as a metabolic engineering approach, we focused on reconstituting EAA biosynthesis in mammalian cells. This challenge offers a stringent test case; EAA biosynthesis was lost in the mammalian lineage over 500 million years ago and mammals must obtain nine amino acids from the environment. In contrast, many microorganisms retain the capacity to synthesize these compounds28,29. Therefore, engineering EAA pathways into mammalian cells presents a powerful model system to explore the boundaries of metabolic rewiring, given its complexity, biotechnological value and experimental tractability. Critically, this system offers a simple and effective selection strategy; cells are grown in the absence of the target amino acid and only those with functional biosynthetic pathways are able to grow5.
We initially focused on valine and isoleucine biosynthesis in CHO cells. In previous work, using rational design principles, we found that E. coli ilvN, ilvB, ilvC and ilvD conferred valine prototrophy when expressed at a fixed 1:1:1:1 stoichiometry5(Fig. 2a). However, this same gene set was insufficient to confer isoleucine biosynthesis to CHO cells despite its overlap in enzymatic steps with valine biosynthesis (Fig. 2b). We hypothesized that this might be because of (1) insufficient availability of an isoleucine pathway-specific substrate, 2-oxobutanoate; (2) feedback inhibition of early steps in the pathway by pathway products; or (3) suboptimal catalytic properties of the acetohydroxy acid synthase I (AHAS I) complex encoded by catalytic subunit, ilvB, and regulatory subunit, ilvN.
a, pMTIV pathway designed in a previous study5 to successfully confer valine prototrophy to CHO cells. All genes are derived from E. coli and encoded in a single open reading frame using 2A ribosomal-skipping peptide sequences. ilvN, ilvB, ilvC and ilvD are the subset of genes relevant to valine biosynthesis and were shown to be sufficient to enable valine prototrophy in CHO. b, Pathway map detailing enzymatic steps to enable valine/isoleucine biosynthesis in mammalian cells. Black arrows constitute native mammalian genes. Orange arrows constitute E. coli enzymes previously imported into CHO to enable valine biosynthesis5. Gene names listed in red are other E. coli-derived genes hypothesized to be helpful toward conferring biosynthetic function and that were included in the SGE library. Valine biosynthetic pathway intermediates: pyruvate, (S)-2-acetolactate, (R)-2,3-dihydroxy-3-methylbutanoate, 2-oxoisovalerate and valine. Isoleucine biosynthetic pathway intermediates: threonine, 2-oxobutanoate, (S)-2-aceto-2-hydroxybutanoate, (R)-2,3-dihydroxy-3-methylpentanoate, (S)-3-methyl-2-oxopentanoate and isoleucine. c, BCAA pathways are localized to different intracellular compartments in extant prototrophic species. Created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
Therefore, we constructed a library containing ilvN, ilvB, ilvC and ilvD, together with five additional pathway components (Fig. 2b and Table 1). Specifically, we added E. coli-derived ilvA to address the hypothesized limitation in the availability of the isoleucine-specific substrate 2-oxobutanoate. To address the second challenge of feedback inhibition, we included two mutant variants of ilvA with partial (ilvAL481F) or complete (ilvAL447F) insensitivity to negative feedback inhibition mediated by isoleucine itself30,31,32. In our previous work, we observed that rationally engineered valine prototrophic CHO cells (pMTIV) grew faster in media with reduced isoleucine concentrations, which we reasoned reflected strong feedback inhibition within the pathway5. Including these variants allowed us to test whether modulating feedback inhibition in a heterologous context could improve pathway function. Lastly, to potentially improve the catalytic properties of the pathway, we added ilvG and ilvM, isozymes of ilvB and ilvN, respectively. Together, ilvG and ilvM form an AHAS II complex, similar to that formed by ilvB and ilvN (AHAS I) but with different catalytic properties—most notably a pronounced (57–180-fold) preference for the isoleucine biosynthetic substrate, 2-oxobutanoate, over the shared isoleucine/valine biosynthetic substrate, pyruvate33,34. AHAS II is furthermore insensitive to feedback inhibition by isoleucine and valine, unlike AHAS I, which responds to inhibition by both amino acids. Lastly, a green fluorescent protein (GFP) CDS was included as a passenger control.
Each CDS was paired with either the strong EF1a or medium-strength PGK promoter, with or without a mitochondrial targeting sequence (MTS). Mitochondrial localization was included because compartmentalization can improve pathway function and branched-chain amino acid (BCAA) biosynthesis occurs in the mitochondria of extant eukaryotic prototrophs25,35 (Fig. 2c). Together, the ten CDSs, two promoters and two localization states produced a 40-member TU library. Each promoter–OLS combination and CDS was represented by three barcodes each to distinguish independent integration events.
We assembled the library using golden gate cloning (Methods) and performed amplicon-seq to monitor distribution of TUs before viral packaging. There was no greater than a twofold difference in read count between any two TUs within the library, indicating that the library distribution was well balanced with regard to component parts (Extended Data Fig. 1a). The library was packaged in lentivirus and 2.42 million CHO cells were infected at an MOI of 8.8 as estimated by qPCR following infection (Extended Data Fig. 1b).
Our choice of ten CDSs, two promoters and two OLSs (40 total TUs) reflected a balance between biological relevance and practical samplability. As we knew four genes to be sufficient to establish a valine prototrophic phenotype, we at a minimum wanted to sample all possible four-TU combinations, of which there are 123,410 from a 40-member library. However, we were interested in sampling variation in copy number as well, motivating a target of sampling all five-TU combinations of which there are 1,086,008. This design space can be sampled within a single 10-cm dish, which we reasoned was appropriate for a pilot-scale demonstration of the approach. Even modest expansions to the library, for example, adding one additional promoter or OLS to yield a 60-TU library, would increase the number of possible five-TU combinations to ~7 million, making comprehensive sampling substantially more challenging in practice. The effective extent to which this combinatorial space is explored can be modeled using a Monte Carlo simulation, as described below.
After infection, we performed amplicon-seq to monitor TU distribution and observed underrepresentation of EF1a promoters both with and without MTS (Extended Data Fig. 1a). Because head-to-head comparisons of PGK and EF1a in pooled lentiviral libraries have not been systematically examined before, the basis of this effect remains unclear. In our pooled setting, several mechanisms may contribute to the underrepresentation of EF1a-driven TUs. First, because EF1a is a stronger promoter than PGK, its recruitment of transcriptional machinery could interfere with transcription of the viral genome during packaging, thereby reducing viral titer36,37,38. Second, EF1a contains an intron that is absent from PGK and, although this intron is typically retained during lentiviral packaging, interactions between the intron’s splice sites and the viral splicing and packaging machinery could still modestly impair packaging efficiency39. Lastly, high-level expression driven by EF1a may impose a fitness cost when multiple pathway components are present in the same cell, leading to depletion of EF1a-linked TUs upon infection of the target cells. We also found lesser representation of ilvAL447F (2.6% of reads) and ilvG (1.2% of reads) compared to other CDSs.
Monte Carlo simulation to estimate scale of diversity introduced
This imbalance in TU representation prompted us to perform a Monte Carlo simulation to determine whether pathway combinations involving underrepresented TUs were likely to be captured in our infection experiment. We simulated randomized choices of TUs on the basis of the barcode distribution measured for the CHO-Div population at an MOI range of 2–14, assuming a Poisson distribution for integration events. At an MOI of 8, the simulation indicated that we would observe 1.56 million different pathway combinations across the 2.42 million infected cells, with 12.4% of cells encompassing the gene content of our valine biosynthetic pathway (ilvN, ilvB, ilvC and ilvD), while 2.0% of cells would have all four genes localized to the cytoplasm and 1.6% would have all four genes localized to the mitochondria (Extended Data Fig. 1c). We reasoned that this likely constituted sufficient presence of true pathway solutions for us to discover functional valine biosynthetic pathways after selection and proceeded to selection on a valine-restricted medium.
More broadly, these simulations illustrate how key experimental parameters such as library size, number of infected cells and MOI jointly determine the fraction of the combinatorial design space that can be sampled in a single screen. Therefore, the achievable coverage can be predicted for any SGE experiment using the same simulation framework, providing a useful starting point for selecting library complexity, MOI and culture scale when designing SGE screens.
Reconstitution and optimization of valine prototrophy in CHO
CHO-Div was subjected to reduced (4.25 µM) valine medium to favor outgrowth of cells carrying combinations of TUs that together comprised biosynthetic pathways for valine (Fig. 3a). In parallel, we subjected a control population (CHO-GFP) that was transduced with a control virus carrying only GFP to the same selection regimen. After 15 days in low-valine medium, we found clonal outgrowths of cells in the experimental population (CHO-Val) but not in the control CHO-GFP population (Fig. 3b). We picked 16 CHO-Val clones and passaged each individually in a valine-replete (170 µM) medium before characterization using an amino acid dropout assay. As a proxy for cell number, metabolic activity was measured using a resazurin-based reagent40 and valine biosynthetic function was estimated by measuring the ratio of valine-free to valine-replete metabolic activity (Val score). We considered a clone valine prototrophic if its Val score was at least twice that of the parental control. All selected clones (16/16) met this threshold (Fig. 3c). To verify that this functional assay accurately captures valine prototrophic function, we further characterized two clones, CHO-Val-D1 and CHO-Val-D3 (naming convention detailed in the Methods). We cultured both clones in valine-free medium over 10 days, measuring their absolute growth rate. While control cells died off by day 7 in valine-free medium, clones CHO-Val-D1 and CHO-Val-D3 instead proliferated over the 10 days of valine-free culture at rates of 1.14 and 1.05 days per doubling, respectively (Fig. 3d), corresponding to 83% and 80% of their growth rates in valine-replete medium (Extended Data Fig. 2a). This is a significant improvement in valine-free growth rate compared to the 3.77 days per doubling growth rate exhibited by pMTIV cells that we engineered using rational design in our previous study5, which corresponded to just 26% of their valine-replete growth rate.
a, Schematic detailing SGE workflow from infection to phenotyping and barcode readout for adherent cells. b, Representative images of clonal outgrowths after 15 days of selection on low-valine RPMI medium from cell populations infected with either GFP or the SGE library. Scale bar, 500 µm. c, PrestoBlue amino acid dropout assay of 16 individual clones selected on low-valine RPMI and the parental control cell line. Metabolic activity of each clone was measured in triplicate after growth in valine-free medium for 3 days and after growth in valine-replete medium for 3 days. Valine prototrophy was defined as a twofold improvement in Val score relative to the nonprototrophic parental cell line. Strong valine prototrophy was arbitrarily defined as a Val score of 0.75. Error bars represent the s.d. across triplicate wells. d, Growth curve of clones CHO-Val-D1 and CHO-Val-D3 cultured in valine-free RPMI and compared to the parental cell line, as well as a pMTIV cell line engineered in a previous study using rational design principles5. Error bars represent the s.d. across triplicate wells. e, Endogenous 13C valine labeling in cells cultured on RPMI with 13C6 glucose and 13C3 sodium pyruvate. Error bars represent the s.d. across triplicate wells for clones CHO-Val-D1, CHO-Val-D3 and parental control cells. pMTIV is a single replicate from a previous study5. f, Percentage of all valine prototrophic clones (Val score > 0.28) and all strongly valine prototrophic clones (Val score > 0.75) that contain each depicted CDS as measured by amplicon-seq. g, Composite barcode fingerprint across strongly valine prototrophic clones. For each TU within a sample, the percentage of barcodes assigned to that TU (relative to total reads for that sample) was calculated. The median percentage representation of each TU across all clones is displayed as a composite heat map. h, Relative abundance of TUs, which comprise the valine prototrophic pathway solution and their nine underlying barcode combinations (three per promoter–OLS, three per CDS) across all strongly valine prototrophic clones (Val score > 0.75). i, Comparison of representation of ilvN, ilvB, ilvC and ilvD in clones selected on low-valine medium relative to simulated expectations according to random TU integration at MOI = 8. j, The SGE-informed pathway was constructed by commercial synthesis of ~3-kb DNA fragments, which were assembled using homologous recombination in S. cerevisiae and delivered to CHO-Flp cells using Flp-In recombination. k, Growth curves of CHO cells harboring the SGE-informed pathway design or the pMTIV pathway engineered in previous work5 cultured in valine-free RPMI medium. Panels a, g, j created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
To further confirm that cells were endogenously synthesizing valine, we cultured CHO-Val-D1 and CHO-Val-D3 in a valine-free ‘heavy’ RPMI medium (Extended Data Fig. 2b and Supplementary Note 1). Both clones exhibited considerably increased 13C2–5 valine labeling at 96.5% and 96.6% respectively, compared to pMTIV cells at 32.2%, indicating strong improvement of the valine biosynthetic phenotype compared to the rationally designed metabolic pathway (Fig. 3e). Indeed, 13C2–5 valine levels detected in clones Val-D1 and Val-D3 matched 13C2–3 alanine levels, suggesting that the engineered biosynthesis of valine in these clones is on par with the native mammalian cellular ability to biosynthesize the non-EAA alanine, which is also biosynthesized from pyruvate.
We performed amplicon sequencing to identify the TUs underlying the valine prototrophy phenotype and to characterize genotypic differences between all 16 clones. To distinguish true integration events from background noise, we applied a z-score threshold to TU read counts (Extended Data Fig. 3a). Within each clone, TU barcode counts were grouped into discrete abundance levels consistent with differences in integration copy number, allowing us to estimate that CHO-Val-D1 and CHO-Val-D3 harbor nine and six TU integrations, respectively (Extended Data Fig. 3b). This corresponds to ~37 and ~23 kb of integrated DNA, respectively, or ~19 and ~11 kb of TU modules. In both clones, we found barcodes corresponding to ilvN, ilvB, ilvC and ilvD, consistent with the CDS content of the rationally designed pMTIV pathway with CHO-Val-D3 additionally exhibiting barcode signatures for ilvM, an isozyme of ilvN (Extended Data Fig. 3b). To validate these assignments, we performed long-read sequencing and RNA-seq on CHO-Val-D1 and CHO-Val-D3. Long-read sequencing confirmed TU identities for 38 of 40 calls (Extended Data Fig. 3c,d), while RNA-seq demonstrated that barcode-defined genotypes were largely concordant with transgene expression (Extended Data Fig. 3b,e).
Having validated our barcode-based genotyping approach, we next defined genetic configurations underlying valine prototrophy by identifying CDSs present across all prototrophic clones (Val score > 0.28). We found ilvB, ilvC and ilvD presence in every clone exhibiting prototrophy, indicating their necessity in conferring valine biosynthetic function to CHO cells (Fig. 3f); ilvN appeared in 94% (15/16) of clones after selection, suggesting that it strongly supports valine biosynthetic function. This is consistent with prior reports that ilvN acts as a regulatory subunit that enhances ilvB catalytic activity and substrate specificity; ilvB alone retains activity but operates at ~15% of the ilvBN holoenzyme41. In the one clone that lacked ilvN, CHO-Val-D8, cells instead carried its isozyme, ilvM (Supplementary Fig. 1a), suggesting a possible role for a hybrid AHAS holoenzyme ilvBM, which has previously been demonstrated to be functional and demonstrates differential catalytic and feedback properties compared to its canonical AHAS I and AHAS II counterparts41,42.
We next sought to determine the optimal promoter and localization context for each CDS by examining TU barcode distributions across promoter–OLS combinations in strongly prototrophic clones (Val score > 0.75). Across 13 such clones, we generated a composite barcode ‘fingerprint’ using the median representation of each TU. Within this fingerprint, mitochondrially localized ilvB, ilvC, ilvD and ilvN expressed from a PGK promoter stood out prominently, defining a pathway ‘solution’ not previously identified through rational design (Fig. 3g). Given that each TU has nine underlying combinatorial barcodes (three barcodes per promoter–OLS combination, three barcodes per CDS), we were able to examine the underlying barcode patterns for each TU within our pathway solution. We found a diversity of barcode patterns underlying each TU among strongly prototrophic valine clones, suggesting that the different clones favored during low-valine selection were derived from multiple independent integration events (Fig. 3h). Across expanded clones, we observed no instances in which ilvN, ilvB, ilvC and ilvD were collectively cytoplasmic, while mitochondrial localization of all four genes was enriched 48-fold relative to expectation according to simulated random TU combinations at an MOI of 8 (Fig. 3i).
Although mitochondrially localized PGK-driven ilvAL481F was also represented in the composite barcode fingerprint, we excluded it from the pathway solution because of its overrepresentation in the starting CHO-Div population (Extended Data Fig. 1a). Simulations at MOI = 8 predicted its presence in 59% of clones (Supplementary Fig. 1b), which correlates well with its presence across all valine prototrophic clones, indicating that it is not overrepresented relative to expectation in our screen following selection in reduced valine conditions. Its absence from strongly prototrophic clones CHO-Val-D1 and CHO-Val-D3 further supports this conclusion (Extended Data Fig. 3b).
On the basis of these results, we attempted to reconstitute this optimized pathway on a single DNA construct. This design matched our previous pathway, pMTIV, using the same regulatory context and 2A ribosome-skipping peptide sequences (Fig. 2a), but contained mitochondrially localized versions of ilvN, ilvB, ilvC and ilvD. The pathway was synthesized in ~3-kb chunks, assembled using homologous recombination in S. cerevisiae43 and delivered to CHO cells using Flp-In recombination44 (Fig. 3j). The SGE-informed design enabled CHO cells to grow in valine-free RPMI at 1.25 days per doubling, corresponding to 74% of their growth rate in valine-replete conditions, substantially outperforming pMTIV cells (Fig. 3k). Together, these results demonstrate that SGE can uncover functional pathway architectures within vast combinatorial search spaces for complex metabolic engineering challenges and enable their reconstitution on a single DNA construct to confer desired cellular functions.
Discovery of metabolic pathway solutions to encode isoleucine prototrophy in CHO
To identify TU combinations that confer isoleucine prototrophy—an outcome we were unable to achieve through rational design—we subjected CHO-Div cells to reduced isoleucine RPMI conditions (1.5, 3 and 6 µM, respectively). In parallel, we subjected a control CHO-GFP population to the same selection regimens. After 15 days in low-isoleucine medium, we found large clonal outgrowths of cells in the experimental populations (CHO-Ile) across all conditions but not in the control CHO-GFP populations. Across the three isoleucine conditions, we picked 46 clones and passaged each individually in an isoleucine-replete (60 µM) medium before performing functional characterization. As previously, we estimated the isoleucine biosynthetic capacity of each clone by measuring the metabolic activity of each clone cultured in isoleucine-free medium and relativizing to metabolic activity in isoleucine-replete medium (Ile score). In total, 41% of clones (19/46) exhibited an improved Ile score relative to the control (Fig. 4a). We further characterized two clones, CHO-Ile-10-H1 and CHO-Ile-10-H2 (naming convention detailed in the Methods), which exhibited strong and intermediate prototrophy, respectively. We cultured both clones in isoleucine-free medium over 10 days and measured absolute cell numbers during this time. While control cells died off by day 6, clone CHO-Ile-10-H1 proliferated over 10 days in isoleucine-free medium at a rate of 1.69 days per doubling, corresponding to 48% of its growth in isoleucine-replete medium, while CHO-Ile-10-H2 initially exhibited a similar growth rate in isoleucine-free medium but tapered off after the eighth day of culture for an overall proliferation rate of 3.04 days per doubling over the 10 days of culture, corresponding to 33% of its growth rate in isoleucine-replete medium (Fig. 4b and Extended Data Fig. 4a).
a, PrestoBlue amino acid dropout assay of 46 individual clones selected on low-isoleucine RPMI and the parental control cell line. Metabolic activity of each clone was measured in triplicate after growth in isoleucine-free medium for 3 days and isoleucine-replete medium for 3 days. Isoleucine prototrophy was defined as a twofold improvement in Ile score relative to the nonprototrophic parental cell line. Strong isoleucine prototrophy was arbitrarily defined as Ile score > 0.75. Error bars represent the s.d. across triplicate wells. b, Growth curve of clones CHO-Ile-10-H1 and CHO-Ile-10-H2 cultured in isoleucine-free RPMI compared to the parental cell line. Error bars represent the s.d. across triplicate wells. c, Extracted ion chromatogram showing detection of M + 0 leucine and M + 2 isoleucine in Clone CHO-Ile-10-H1 cultured on isoleucine-free medium with 13C substrates. By comparison, nonprototrophic control cells cultured on isoleucine-replete medium with 13C substrates show only M + 0 leucine and M + 0 isoleucine. d, MS1 peaks corresponding to M + 2 and M + 6 isoleucine were detected in clone CHO-Ile-10-H1. IS, 13C6/15N1 isoleucine internal standard. e, Endogenous 13C isoleucine labeling in clones CHO-Ile-10-H1 and CHO-Ile-10-H2, as well as parental control cells, cultured on RPMI with 13C6 glucose and 13C3 sodium pyruvate. Error bars represent the s.d. across triplicate wells. f, Percentage of all isoleucine prototrophic clones (Ile score > 0.29) and all strongly isoleucine prototrophic clones (Ile score > 0.75) that contain each depicted CDS, as measured by amplicon-seq. g, Composite barcode fingerprint across strongly isoleucine prototrophic clones. For each TU within a sample, the percentage of barcodes assigned to that TU (relative to total reads for that sample) was calculated. The median percentage representation of each TU across all clones is displayed as a composite heat map. h, Comparison of representation of ilvM, ilvG, ilvB, ilvC, ilvD and ilvAL481F in clones selected on low-isoleucine medium relative to simulated expectations according to random TU integration at MOI = 8. Panel g created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
To further confirm that cells were endogenously synthesizing isoleucine, we cultured CHO-Ile-10-H1 and CHO-Ile-10-H2 in heavy RPMI medium (Extended Data Fig. 4b). In both clones, extracted ion chromatography, mass spectrometry (MS1 and MS2) analysis revealed signatures consistent with M + 2 and M + 6 isoleucine, the isotopologs we expect to detect if isoleucine is synthesized endogenously (Fig. 4c,d, Extended Data Fig. 4c and Supplementary Note 2). For clones CHO-Ile-10-H1 and CHO-Ile-10-H2, 78.5% and 39.5% of isoleucine was 13C2–6, respectively, with the latter exhibiting significant variability in labeling across replicates consistent with its less effective prototrophic function (Fig. 4e). Together, these clones represent a demonstration of isoleucine prototrophy in mammalian cells, a metabolic function last seen in the mammalian lineage over 500 million years ago.
As with the valine prototrophic clones described above, we performed amplicon sequencing on all 46 clones (prototrophic and auxotrophic) to identify the TU combinations underlying isoleucine prototrophy. Applying a z-score threshold to distinguish true integrations from background (Extended Data Fig. 5a), we detected ilvB, ilvC, ilvD, ilvG and ilvM in both clones, with CHO-Ile-10-H1 additionally carrying ilvA and ilvAL481F and CHO-Ile-10-H2 carrying ilvN (Extended Data Fig. 5b). On the basis of relative barcode abundance levels, we estimated that CHO-Ile-10-H1 and CHO-Ile-10-H2 harbor 13 and ten TU integrations, respectively, corresponding to ~52 kb and ~41 kb of integrated DNA, of which 26 and 21 kb were directly made up by TU modules, respectively.
To validate our barcode-based genotyping approach, we performed long-read sequencing and discovered concordance with amplicon-seq for 36/40 and 31/40 TU calls in clones CHO-Ile-10-H1 and CHO-Ile-10-H2, respectively (Extended Data Fig. 5c,d). RNA-seq measurements of transgene expression were largely consistent with barcode-defined genotypes in CHO-Ile-10-H1 but less so in CHO-Ile-10-H2 (Extended Data Fig. 5b,e). These differences are consistent with clonal heterogeneity, as additional low-abundance TUs detected by long-read sequencing were present only at background levels in amplicon-seq (Extended Data Fig. 5b–e) and may contribute to the variability observed in growth and isotope labeling across replicates. A comprehensive characterization of integration structure, clonal composition and global transcriptomic impact across CHO clones is provided in Supplementary Note 3.
We next sought to determine isoleucine pathway composition by identifying CDSs that were universally present across all strongly isoleucine prototrophic clones (Ile score > 0.75). Strikingly, 100% of strongly prototrophic clones exhibited barcodes corresponding to ilvB, ilvC, ilvD, ilvG, ilvM and ilvAL481F (Fig. 4f). To examine the promoter and localization context of these CDSs, we generated a composite barcode fingerprint using the median TU representation across strongly prototrophic clones. Consistent with the optimal valine pathway configuration, all CDSs were predominantly present in their mitochondrially localized configurations, with ilvAL481F, ilvM, ilvB, ilvC and ilvD favored under the PGK promoter and ilvG under the EF1a promoter (Fig. 4g). Notably, inclusion of ilvA, which catalyzes the first step of the isoleucine pathway (conversion of threonine to 2-oxobutanoate), was essential for optimal pathway performance. In particular, the enrichment of the partially feedback-resistant variant ilvAL481F across all strongly prototrophic clones suggests that alleviation of feedback inhibition is a key determinant of pathway efficiency.
We examined the nine underlying barcode possibilities for the favored pathway solution and found barcode diversity across the seven strongly prototrophic clones, suggesting that clones arose from independent integration events (Supplementary Fig. 2a). We further examined how often ilvC, ilvD, ilvB, ilvG, ilvM and ilvAL481F were collectively observed to be cytoplasmically or mitochondrially localized in isolated clones, as well as how often they were observed irrespective of intracellular location and compared each to simulated expectation. We found that mitochondrial localization of these TUs was dramatically selected for at 217× expectation while cytoplasmic localization of all six CDSs was not observed at all across the 46 clones (Fig. 4h).
Lastly, we looked at CDS representation amongst isoleucine auxotrophic clones (Supplementary Fig. 2b). Intriguingly, we found presence of ilvB, ilvC and ilvD across 100% of auxotrophic clones indicating that there was selection for partial metabolic pathways. This is consistent with a model in which infected cells are resource sharing and metabolic intermediates are available in the medium allowing cells encoding only partial biosynthetic pathways a selective advantage. Such a mechanism would favor CDSs that encode later enzymatic steps in the biosynthetic pathway, which is what we observe with ilvD and ilvC being most overrepresented relative to simulated expectation followed by ilvM, ilvG and ilvB.
Together, these results demonstrate that SGE can uncover not only functional metabolic pathway architectures for previously intractable metabolic engineering problems but also provide insight into the mechanistic basis for their function.
Double isoleucine/valine prototrophy in CHO-Val and CHO-Ile clones
Having established that we could achieve valine and isoleucine prototrophy individually using SGE, we next asked whether these behaviors could coexist within the same cells given the substantial overlap between pathway components. To this end, we cultured all CHO-Val clones and CHO-Ile clones in a double isoleucine/valine dropout medium to assess whether dual prototrophy could arise without direct selection. Indeed, 37.5% (6/16) of CHO-Val clones and 23.9% (11/46) of CHO-Ile clones surpassed the threshold for double isoleucine/valine prototrophy (2× the metabolic activity of the parental control line; Ile/Val score > 0.28), although none met the strong double prototrophy threshold (Ile/Val score > 0.75) (Extended Data Fig. 6a). A comprehensive characterization of the barcode signatures underlying single versus double prototrophic clones is provided in Supplementary Note 4.
These results illustrate both the diversity of phenotypes accessible through SGE and the insights such diversity can provide for pathway optimization. Importantly, they underscore a key strength of the SGE framework: the optimal genetic solution for a single metabolic trait (for example, valine or isoleucine prototrophy) is not necessarily the optimal solution for combined traits (for example, valine and isoleucine prototrophy). Traditional DBT-based engineering approaches often struggle with such multiobjective tradeoffs but the scale and combinatorial depth of SGE allow for simultaneous selection across multiple traits, enabling discovery of genotypes that balance or even reconcile conflicting demands across pathways.
Engineering valine prototrophic human T cells
To examine the versatility of SGE across cell types, we wanted to ascertain whether it was possible to engineer Jurkat cells to become prototrophic for valine and isoleucine. Unlike CHO cells, Jurkat cells grow in suspension, allowing for seamless isolation, expansion and characterization of large numbers of individual clones after selection through either serial dilution or cell sorting. Using the same lentivirus library from previous experiments, we infected 3 million Jurkat cells (Jurkat-Div) and observed a similar barcode distribution to the distribution we previously observed in CHO-Div cells (Extended Data Fig. 7a). The MOI of the infection was estimated by qPCR to be 3.4 (Extended Data Fig. 7b).
Given the lower MOI relative to CHO cells, we performed Monte Carlo simulations at a range of MOIs (2–8) to determine the likelihood of finding known solutions given the measured barcode distribution (Extended Data Fig. 7c). At MOI = 4, our simulation indicated that we should see 5.9 × 105 different pathways within the 3 million infected cells, of which 2.31% of cells should carry some combination of ilvN, ilvB, ilvC and ilvD, while 0.19% of cells should carry all four TUs localized to the mitochondria and 0.23% should carry all TUs localized to the cytoplasm. On the other hand, only 0.06% of Jurkat-Div cells were expected to carry any combination of our discovered isoleucine biosynthetic pathway consisting of ilvM, ilvG, ilvB, ilvC, ilvD and ilvAL481F, with 0.00% of cells expected to carry entirely mitochondrially or entirely cytoplasmically localized versions of all six genes (Extended Data Fig. 7d). These estimates suggested that valine prototrophy would be readily accessible, whereas isoleucine prototrophy would be rare in the Jurkat-Div population.
Accordingly, Jurkat-Div cells were cultured on low-valine medium (0, 1.7 or 4.25 µM) alongside control cells infected only with GFP (Jurkat-GFP) (Extended Data Fig. 8a). At all valine concentrations, both cell populations saw a reduction in cell number over 14 days. By day 21, however, Jurkat-Div cells saw an increase in cell number across all conditions, whereas Jurkat-GFP cells did not recover. We serially diluted the resulting selected Jurkat-Div cell populations (Jurkat-Val) to isolate and expand 119 individual clones in valine-replete conditions before characterization using our amino acid dropout assay (Fig. 5a). As before, we considered a clone prototrophic if its Val score was at least twofold that of a parental control (Val score > 0.23), and 71% (85/119) of clones met this threshold (Fig. 5b). We further characterized two clones exhibiting valine prototrophy, Jurkat-Val-31 and Jurkat-Val-32, culturing both clones in valine-free medium over 11 days and measuring their absolute growth rate. While control cells exhibited stagnant cell numbers over 11 days of culture, clones Jurkat-Val-31 and Jurkat-Val-32 proliferated at 2.02 and 1.87 days per doubling, respectively, corresponding to 50% and 52% of their growth rates in valine-replete medium (Fig. 5c and Extended Data Fig. 8b).
a, Schematic detailing SGE workflow from infection to phenotyping and barcode readout for suspension cells. b, PrestoBlue amino acid dropout assay of 119 individual clones selected on low-valine RPMI and the parental control cell line. Metabolic activity of each clone was measured in triplicate after growth in valine-free medium for 3 days and after growth in valine-replete medium for 3 days. Valine prototrophy was defined as a twofold improvement in Val score relative to the nonprototrophic parental cell line. Strong valine prototrophy was arbitrarily defined as a Val score of 0.75. Error bars represent the s.d. across triplicate wells. c, Growth curve of clones Jur-Val-31 and Jur-Val-32 cultured in valine-free RPMI and compared to the parental cell line. Error bars represent the s.d. across triplicate wells. d, Percentage of all valine prototrophic clones (Val score > 0.23) and all strongly valine prototrophic clones (Val score > 0.75) that contain each depicted CDS as measured by amplicon-seq. e, Composite barcode fingerprint across strongly valine prototrophic clones. For each TU within a sample, the percentage of barcodes assigned to that TU (relative to total reads for that sample) was calculated. The median percentage representation of each TU across all clones is displayed as a composite heat map. f, Comparing representation of ilvN, ilvB, ilvC and ilvD in clones selected on low-valine medium relative to simulated expectations according to random TU integration at MOI = 4. g, The random forest model classifies outcomes using a majority vote across many decision trees trained on subsets of data and features. h, Fourfold cross-validation was used to ensure stability of the random forest model. Metrics (left), top ten features (middle) and classifications (right) across the four folds are summarized. Error bars represent the s.d. across the four cross-validation folds. Panels a, e, g created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
We next asked whether isoleucine prototrophy could similarly be achieved in Jurkat cells. Jurkat-Div cells were cultured in low-isoleucine medium (0, 6 or 12 µM) alongside Jurkat-GFP controls. Across all conditions and 32 days of culture, Jurkat-Div cells failed to outcompete control cells, indicating that isoleucine prototrophic phenotypes were not present in the population (Extended Data Fig. 8c).
Together, these results highlight that simulated pathway diversity, determined using TU distributions and MOI, can be used to predict and guide pathway discovery across cell types.
Characterization of valine prototrophic human T cell clones
To identify the TU combinations underlying valine prototrophy in Jurkat cells, we performed amplicon-seq on all isolated clones, applying a z-score threshold to distinguish signal from background (Extended Data Fig. 9a). Clones Jurkat-Val-31 and Jurkat-Val-32 exhibited similar TU profiles, comprising mitochondrially localized ilvM, ilvB, ilvN, ilvC and ilvD expressed from PGK promoters, along with cytoplasmically localized ilvB in the same promoter context (Extended Data Fig. 9b,c). This configuration closely mirrors the optimal mitochondrially localized pathway architecture we identified in CHO cells.
On the basis of relative barcode abundance, we estimated that each clone harbored nine TU integrations corresponding to ~32 kb of integrated DNA, of which ~15 kb was directly made up by TU modules. We additionally performed long-read sequencing for clone Jurkat-Val-31 and discovered concordance between long-read TU calls and amplicon-seq TU calls for 40/40 TUs (Extended Data Fig. 9d,e). Barcodes mapped to their assigned promoter–OLS in 95.5% of reads and to their assigned CDS in 100% of reads (Extended Data Fig. 9f and Supplementary Table 1). Transcripts from SGE transgenes measured by RNA-seq were in good agreement with the TU barcodes detected in amplicon-seq (Extended Data Fig. 9c,g).
Next, to define the pathway underlying valine prototrophy in Jurkat cells, we identified CDSs that were universally present across all prototrophic clones. ilvB, ilvC, ilvD and ilvN were present in all Jurkat valine prototrophic clones, matching the CDS content of the solution previously identified for the CHO-Div population (Fig. 5d). To determine the optimal promoter and localization context for each CDS, we generated a composite barcode fingerprint on the basis of the median representation of each TU across clones. This analysis revealed that ilvC, ilvD and ilvN were favored in their mitochondrially localized context under control of the PGK promoter while ilvB appeared to be favored under control of a PGK promoter in both the cytoplasm and the mitochondria (Extended Data Fig. 10a).
However, when looking at the three promoter barcodes and three CDS barcodes underlying these five TUs, we noticed that many clones exhibited barcode signatures shared with other clones suggesting that multiple sequenced samples had arisen from the same founder clone (Extended Data Fig. 10b). We expect to observe such clonal ‘siblings’ if particular pathway solutions confer a strong selective advantage and undergo expansion during selection.
To quantify this effect, we examined global similarity in TU profiles across clones. We identified 69 clones with at least one sibling and 50 unique ‘singleton’ clones (Extended Data Fig. 10c). Using graph-based clustering, we found ten separate sibling groups varying in size (2–24 clones) and sibling group size did not correlate to average Val score (Extended Data Fig. 10d). Inspection of the largest sibling groups (2, 4, 9 and 5, consisting of 24, 12, ten and seven clones, respectively) revealed a consistent pattern; ilvN, ilvC and ilvD were uniformly localized to mitochondria, while ilvB was observed localized to the mitochondria, cytoplasm or both compartments (Extended Data Fig. 10e). To avoid bias from clonal expansion, we removed duplicate siblings and analyzed CDS usage across the population consisting only of unique clones, which yielded a largely unchanged CDS usage profile (Extended Data Fig. 10f; compare to Fig. 5d). We also recomputed a composite barcode fingerprint, which yielded a pathway solution consisting only of mitochondrially localized ilvN, ilvB, ilvC and ilvD (Fig. 5e). Consistent with this, the collective presence of ilvN, ilvB, ilvC and ilvD was enriched 31-fold relative to expectation, while their colocalization to mitochondria was enriched 283-fold (Fig. 5f).
We also examined CDS representation amongst 24 valine auxotrophic Jurkat clones carrying at least one integration (Supplementary Fig. 3). Unlike the auxotrophic CHO clones selected on low-isoleucine medium, which all carried a partial isoleucine biosynthetic pathway, selection for partial biosynthetic pathways in Jurkat clones selected on low-valine medium was much less pronounced with slight CDS overrepresentation of ilvD, ilvC and ilvB relative to expectation. This may reflect the lower MOI used to infect Jurkat cells or that clonal resource sharing is less pronounced in suspension cell culture.
These results highlight the generalizability of SGE across cell types including rodent and human cell lines and its ability to uncover pathway solutions that converge on shared design principles despite differences in cellular context.
Machine learning to predict prototrophy phenotypes from SGE-generated data
Lastly, we asked whether machine learning could be used to predict prototrophy phenotypes directly from the TU composition of each clone. We decided to apply a random forest model for classification because of its strength in interpreting nonlinear relationships between features, its robustness to overfitting and its ability to provide feature importance scores (Fig. 5g). As input features, we used the proportion of reads assigned to each TU within a clone (analogous to the clone-level TU abundance heat maps). Each clone’s phenotypic performance was binarized as prototrophic or auxotrophic using the Val score threshold of 0.23. The samples were partitioned into four mutually exclusive class-balanced folds to perform stratified fourfold cross-validation (Supplementary Fig. 4). For each fold, the model was trained on the other three folds and evaluated on the held-out fold for four separate 75:25 train–test splits. When looking at metrics across folds, the model successfully classified samples into prototrophic or auxotrophic with a mean accuracy of 0.942 ± 0.005, a mean area under the receiver operating characteristic (ROC) curve of 0.993 ± 0.006 and a mean area under the precision–recall curve of 0.997 ± 0.002 across folds, indicating strong discriminative ability and stability across different train–test sample compositions (Fig. 5h).
In total, 112/119 clones were correctly classified and the top four features driving classification were mitochondrially localized ilvN, ilvC, ilvD and ilvB under control of a PGK promoter, matching the pathway solutions that we discovered by manual analysis and interpretation. This classification accuracy also held true when removing duplicate sibling clones from the dataset before model training; the same top four features were discovered and correct classifications were produced for 53/59 clones (Supplementary Fig. 5a,b).
Five auxotrophic clones were misclassified as prototrophic; two (Jurkat-Val-55 and Jurkat-Val-104) carried all four CDSs that form our discovered pathway solution and likely reflect examples of gene silencing or barcode swapping and two clones (Jurkat-Val-79 and Jurkat-Val-113) lacked ilvN only, potentially reflecting insufficient AHAS activity for prototrophic function, while the final clone (Jurkat-Val-110) lacked both ilvB and ilvD (Supplementary Fig. 6a). On the other hand, two prototrophic clones were misclassified as auxotrophic. Clone Jurkat-Val-60 (Val score: 0.31) appears to be clonally impure, carrying very lowly abundant ilvC and ilvD, which the model may not consider sufficient to generate a prototrophic phenotype. Clone Jurkat-Val-68 (Val score: 1.05) carried ilvN, ilvC and ilvD under control of PGK-MTS, as found in our discovered optimal pathway solution, while ilvB was under the control of EF1a-MTS, which is a much less represented possibility in our starting SGE library and may, therefore, not have been classified as contributing to the valine prototrophic phenotype (Supplementary Fig. 6b).
These results demonstrate that SGE-generated data are well suited for machine-learning-based modeling and such models can effectively extract biologically meaningful features, enabling pathway-scale analysis for solutions to complex metabolic engineering problems.
Discussion
Here, we introduce SGE, a method for efficiently exploring vast combinatorial design spaces comprising multiple interacting TUs in mammalian cells. By leveraging single cells as independent experiments and using random high-copy integration of individual TUs per cell, SGE enables high-throughput assembly and screening of millions of synthetic metabolic pathways. This approach bypasses the inefficiencies associated with building and delivering large DNA constructs, substantially accelerating pathway discovery and optimization.
Using this approach, we generated and screened millions of synthetic metabolic pathways to engineer EAA prototrophy in mammalian cells. We achieved (1) near-wild-type growth in valine-free medium and (2) isoleucine prototrophy in mammalian cells. The functional solutions involved integration of 23–52 kb of synthetic DNA—construct sizes impractical to screen using conventional methods. We also provide initial evidence that leveraging organellar compartmentalization can enhance engineered pathway functionality in mammalian cells. Lastly, we demonstrate the versatility of SGE across mammalian cell types (species, in suspension or adhesion) and present a pilot integration with machine learning to predict functional pathway combinations.
A consistent theme across the metabolic engineering efforts described here was the preference for mitochondrial localization of BCAA pathway enzymes. This was unexpected given that the genes in our library were sourced from E. coli, a prokaryote lacking subcellular compartments. However, mitochondria are thought to be of bacterial origin and share notable protein composition with bacteria, which may help explain this preference45. Equivalent BCAA biosynthesis genes in extant eukaryotic BCAA prototrophic microbes such as fungi are also localized to mitochondria, providing a clue that there may be biochemical advantages associated with mitochondrial localization, such as increased pyruvate concentrations relative to cytosol46,47 or greater availability of relevant pathway cofactors (for example, iron–sulfur clusters and thiamine diphosphate)25,48,49. Similarly, in plants, these BCAA biosynthetic enzymes are targeted to chloroplasts, which are also an abundant source of pyruvate50. Another possibility is that BCAA biosynthetic enzymes simply benefit from being much more concentrated in a limited organellar physical space, thereby boosting pathway flux and function, as previously demonstrated for other pathways localized to the mitochondria in S. cerevisiae35. This finding highlights the utility of incorporating organellar diversity in the SGE screen and encourages incorporation of yet more organellar targeting signals in future engineering efforts in mammalian cells.
Use of SGE comes with several limitations, most notably the requirement for a selectable phenotype. Essential nutrients provide a built-in live–dead screen that effectively identifies cells carrying optimal solutions. However, when the desired phenotype is not linked to cell survival, alternative selection strategies such as coupling pathway activity to fluorescent biosensors, surface marker expression or engineered genetic circuits that connect pathway output to a selectable reporter may be applied51,52. Another limitation of SGE as currently implemented is the use of lentivirus for random integration. We observed substantial biases in TU representation introduced during viral packaging. Specifically, EF1a-driven constructs were markedly underrepresented compared to those with PGK promoters and similar biases were noted for certain CDSs in ways not attributable solely to size. Transitioning to transposon-based delivery systems could alleviate this issue. Lastly, while amplicon-seq can inform what TUs have been integrated in clones of interest, it provides little information about which genes are expressed. In cell lines prone to silencing, this may confound interpretation. Pairing SGE with either bulk or single-cell RNA-seq would provide expression-level information, enabling more accurate assessment of functional pathway configurations.
While SGE excels at rapidly exploring vast design spaces through random integration, successful multi-TU solutions identified in this manner may need to be reconstituted into compact, single-vector formats for practical deployment. However, TUs may behave differently when assembled into condensed architectures53. A hybrid strategy may be effective in this case, using SGE for high-throughput discovery, followed by CLASSIC12 or related methods for validation and optimization in compact constructs in lower throughput.
Despite its current limitations, SGE represents a powerful strategy for the scalable engineering of increasingly complex biosynthetic traits in mammalian cells. In this study, we demonstrated the discovery of a functional six-gene isoleucine biosynthesis pathway from a 40-member TU library. We anticipate that SGE can be extended to larger libraries by proportionally increasing the MOI during integration, enabling access to even more complex phenotypes. These may include prototrophy for additional EAAs, growth factors and vitamins, enhanced tolerance to environmental stressors such as hypoxia, shear stress and metabolic byproducts or rewiring of metabolic programs toward alternative cell states or identities. The inclusion of elements that modulate host factors (CRISPRa/i machinery) would further expand the design space explored by this approach.
Ultimately, the ability to engineer mammalian cells with complex pathways has broad implications across basic research, biotechnology and medicine. For instance, a mammalian cell engineered to be prototrophic for all amino acids would require a reduced number of nitrogen sources, opening the door to using stable isotope labeling to chart presently unknown corners of the mammalian metabolic map54. Meanwhile, mammalian cells engineered to become independent of specific nutrients could lower media costs and simplify formulations for biomanufacturing applications such as cultivated meat and viral vector production55,8. Lastly, cell therapies may be metabolically augmented to better withstand the hostile tumor microenvironment, which is marked by nutrient deprivation, acidosis, oxidative stress and inflammation, with potential for improving the performance of cell-based therapeutics used in the treatment of solid tumors56,57,58,59.
Methods
Cell lines, media, cell counting and images
CHO Flp-In cells (Thermo Fisher, R75807) and Jurkat cells (M. Pacold lab, New York University) were used in all experiments. All cell lines tested negative for Mycoplasma. For growth assays involving amino acid dropout formulations, medium was prepared from an amino-acid-free RPMI 1640 powder base (US Biological, R8999-20) and custom combinations of amino acids were added back in as needed to match the standard amino acid concentrations for RPMI 1640 except for isoleucine, which was considered 100% supplemented at 0.06 mM. Additionally, RPMI medium was supplemented with 2 mM sodium pyruvate as previously described5. Custom amino acid dropout medium was adjusted to a pH of 7.3 with HCl, sterile-filtered and supplemented with 10% dialyzed FBS (Fisher Scientific, 26400044) and penicillin–streptomycin (100 U per ml) before use. For metabolomics experiments, medium was prepared from an amino-acid-free, glucose-free RPMI 1640 powder base (US Biological, R9010-01) and custom combinations of amino acids were added in to match the standard concentrations for RPMI 1640. In the place of glucose, isotopically heavy 13CU glucose (Cambridge Isotope Labs, CLM-1396-PK) and 13C3 sodium pyruvate (Millipore Sigma, 490717) were added at 11.1 mM and 2 mM, respectively. Cell counts were performed using a Countess 3 FL. Images of clones were captured on an EVOS M5000 using a ×10 objective and scale bars were added using Fiji.
SGE library construction
Promoters and OLSs were PCR-amplified and allotted unique barcodes using PCR primers encompassing said barcodes and BsmBI restriction sites. Each barcoded promoter–OLS combination was cloned into an assembly vector with lentiviral long terminal repeats using Gibson assembly. CDSs of interest were codon-optimized and synthesized commercially. Each CDS was allotted three unique barcodes by PCR amplification with primers encompassing said barcodes along with BsmBI restriction sites to facilitate downstream Golden Gate cloning. Each PCR-amplified, barcoded CDS was cloned into a pCR-Blunt-II-TOPO vector. Concentrations of each TOPO-cloned CDS were measured using a Qubit fluorometer before pooling of all CDSs of interest at equimolar concentration. Concentrations of each promoter–OLS plasmid were similarly measured using a Qubit fluorometer before pooling of all promoter–OLS plasmids at equimolar concentration. The pooled CDS mixture was spiked into the Golden Gate reaction at 5× the molar concentration of the pooled assembly vector mix, which was used at 100 ng. We added 1.5 μl of 10× NEB T4 ligase buffer, 0.15 μl of BSA, 1.1 μl of BsmBI-v2 (10,000 U per ml), 1 μl of T4 ligase (20,000 U per ml) and then supplemented with H2O to a total volume of 15 μl. Reactions were run for 90 cycles of 42 °C for 3 min and 16 °C for 4 min before one cycle of 50 °C for 5 min and 80 °C for 5 min. Resulting libraries were transformed into ElectroMAX Stbl4 E. coli (Invitrogen, 11635-018) for amplification before library extraction and amplicon-seq to measure library distributions. Promoter–OLS and CDS sequences are shown in Supplementary Table 8.
Lentiviral packaging and infection
Lentivirus was packaged by plating 4 × 106 HEK293T cells on 10-cm plates and incubating cells overnight at 37 °C. Cells were transfected with a plasmid mix consisting of 3.5 µg of the SGE library, 6.0 µg of psPAX2 (Addgene, 12260) and 3.0 µg of pMD2.G (Addgene, 12259) using Lipofectamine 2000 (Thermo Fisher Scientific, 11668019) in accordance with the manufacturer’s instructions. Transfected HEK293T cells were incubated for 48 h, before medium was collected and replenished. Cells were then incubated for an additional 24 h before medium was collected again. Collected medium was centrifuged at 200g for 5 min and the resulting supernatant was filtered using a 0.45-µm filter. Virus was concentrated using Millipore Amicon Ultra-15 centrifugal filter units (Millipore Sigma, UFC903024) for 30 min at 4,000g before storage at −80 °C until use. For CHO infections, cells were plated at 1.5 M cells per 10 cm and incubated overnight before infection. At the time of infection, the medium was replaced with fresh medium containing polybrene at 8 µg ml−1 and concentrated virus. Cells were incubated with the virus for 24 h before replenishing with fresh medium. For Jurkat infections, 3 million cells were resuspended in 6 ml of medium containing polybrene at 8 μg ml−1 and concentrated virus before centrifugation at 931g for 2 h at 30 °C. An additional 6 ml of medium (not containing polybrene) was subsequently added to cells dropwise before 24 h of incubation at 37 °C, after which medium/virus was replenished with fresh medium.
MOI estimation by qPCR
gDNA was extracted from cells using the QIAamp DNA mini kit (Qiagen, 51304), in accordance with the manufacturer’s protocol. qPCR reactions were performed as 10-μl reactions with 0.5 µl of 5 µM primer mix using SYBR green master I on a QuantStudio 7 Pro. Primers were designed to amplify amplicons 60–120 bp in size. MOI was estimated by measuring amplification of a sequence universally found in each TU found in the SGE library and relativizing to amplification of CHO targets, Dhfr and Hprt (expected to be single copy), and Jurkat targets, RPP30 and ALB (expected to be found in two copies), using the ∆Ct method. All reactions were performed in triplicate.
Monte Carlo simulation of TU integrations across variable MOIs
To model the distribution of TU combinations integrated into a cell population, we performed Monte Carlo simulations assuming Poisson-distributed MOIs. For CHO cells, we simulated 2,420,000 cells at MOI values of 2, 4, 6, 8, 10, 12 and 14, spanning a range around the estimated MOI of 8.8. For Jurkat cells, 3,000,000 cells were simulated at MOIs of 2, 4, 6 and 8, centered around the estimated MOI of 3.4. Integration events were simulated by first drawing the number of integrations per cell from a Poisson distribution. For cells with one or more integrations, TUs were sampled with replacement according to empirically defined distributions. The resulting sets of integrated TUs were sorted and recorded as combinations, irrespective of integration order. For each MOI, the frequency of unique TU combinations was tallied and used to benchmark the probability of observing specific TU sets. These simulations were used not only to quantify diversity in our study but also to provide a general framework for designing SGE experiments. By varying the library size, MOI and number of simulated cells, one can estimate the achievable coverage of TU combinations under their specific experimental parameters. The analysis code used to run these simulations is available in the GitHub repository associated with this paper.
Clonal isolation and amino acid prototrophy screens
After selection on the indicated amino acid dropout medium, CHO clones were manually isolated using a P200 pipette tip, transferred to a 96-well plate and expanded before phenotypic screening. For each selection condition (CHO-Val in 4.25 µM valine RPMI; CHO-Ile in 1.5, 3 and 6 µM isoleucine RPMI), clones were distributed across two separate 10-cm dishes to allow sampling of all infected cells but minimize risk of clones merging. Clones from each dish were assigned to a separate row of the 96-well plate and named according to plate position (for example, C1–C8 and D1–D8). CHO-Ile clones were further labeled with numeric identifiers corresponding to the isoleucine concentration used for selection (2.5%, 5% and 10% of 0.06 mM). For instance, CHO-Ile-10-XX refers to a CHO clone subject to 10% Ile RPMI or RPMI with 6 µM Ile. For Jurkat cells, clones were isolated by serial dilution and numbered sequentially.
We screened for amino acid prototrophic clones using PrestoBlue HS cell viability reagent (Thermo Fisher, P50200), a resazurin-based viability reagent that fluoresces in response to cellular reducing activity. Fluorescence intensity is proportional to the number of metabolically active (viable) cells in the medium. For each amino acid dropout condition, we relativized triplicate wells for each clone in the amino acid dropout condition to triplicate wells in an amino-acid-replete condition to control for differences in overall growth rate and seeding density. Amino-acid-replete medium was made by making amino acid dropout medium and adding in the missing amino acid. Fluorescence measurements were conducted on an Agilent Biotek Synergy Neo2.
Amplicon-seq library preparation
gDNA from clones of interest was prepared using the Zymo Quick-DNA 96 Plus kit (Zymo, D4070) in accordance with the manufacturer’s protocol. PCR1 (UMI addition) was performed using primers annealing to a universal primer sequence found in the 3′ UTR across all SGE TU vectors. PCR1 used 250 ng of gDNA template, 2.5 µl of each 10 µM primer and Q5 polymerase in a total reaction volume of 50 µl. PCR1 cycling conditions were as follows: 98 °C for 5 min, followed by four cycles of 98 °C for 20 s, 65 °C for 20 s and 72 °C for 30 s, with a final extension at 72 °C for 60 s. PCR cleanup was performed using AmpureXP beads (0.8×) and eluted in 15 µl of 0.1× TE. For PCR2 (sample indexing and sequencing adaptor addition), 10 µl of eluate from the previous reaction was used in another 50-µl reaction using Q5 polymerase and 5 µl of a 5 µM indexing primer mix. SYBR green dye was also added to the reaction mix to allow us to monitor the reaction progression and stop reactions before saturation (typically 16–19 cycles). PCR2 cycling conditions were as follows: 98 °C for 5 min, followed by 16 cycles of 98 °C for 10 s, 65 °C for 10 s and 72 °C for 20 s. PCR cleanup was again performed using AmpureXP beads (0.8×) and eluted in 15 µl of 0.1× TE. Sequencing was performed on a NextSeq500 75 cycle kit to yield paired-end 36-bp reads.
Amplicon-seq analysis
FASTQ files from paired ends were merged using pear (version 0.9.11). Stitched FASTQ files were processed to recover known barcode combinations identifying promoter–OLS combinations and CDSs to quantify TU integration events. Reads were parsed to identify two universal priming regions flanking the barcode segments. For reads containing both universal primers, two 8-bp barcodes were extracted and concatenated to form a unique identifier for each TU.
Each extracted barcode combination was matched against a known barcode reference library for exact matches. However, we noticed that a subset of clones exhibited a large proportion of reads that did not match any in the reference library but exhibited a 1-bp mismatch relative to reads that were found in the reference library. As no two barcodes in the library exhibited a Hamming distance < 2 and 99.9% of possible barcode pairings exhibited a Hamming distance ≥ 3, we opted to allow for 1-bp mismatches if possible to do so unambiguously. Briefly, if no exact barcode match was found, sequences were compared to a precomputed lookup table containing all possible 1-bp mismatch variants of the reference barcodes. Reads with a unique (unambiguous) 1-bp mismatch were assigned to the corresponding reference barcode, while ambiguous 1-bp mismatches (matching multiple barcodes) were excluded from downstream quantification. Samples with fewer than 50,000 matching reads were further excluded from downstream analysis.
For analysis of CDS presence within each clone, read counts for each TU were used to calculate a z score. Given three barcodes per promoter–OLS combination and three barcodes per CDS, each TU had nine underlying barcodes. Read counts for each TU were aggregated and log-transformed before calculating a z score for each TU on a per sample basis to separate true signal from background. For generation of per-clone heat maps, the percentage of reads assigned to a given TU within a sample was calculated, normalized to total reads per sample. For generation of heat maps to identify optimal pathways, the median percentage representation of each TU across all indicated clones was displayed as a composite heat map. Raw read counts are shown in Supplementary Table 9 and TU aggregated read counts are shown in Supplementary Table 10.
Metabolomics
Cells were cultured in RPMI medium containing 13Cglucose and 13Csodium pyruvate with or without valine/isoleucine (as indicated) before cell harvest. Cell pellets were generated by trypsinization, followed by low-speed centrifugation, and the pellet was frozen at −80 °C until further processing. A metabolite extraction was carried out on each sample with an extraction ratio of 1 × 106 cells per ml (80% methanol containing internal standards, 500 nM). The liquid chromatography (LC) column was a Millipore ZIC-pHILIC (2.1 × 150 mm, 5 μm) coupled to a Dionex Ultimate 3000 system and the column oven temperature was set to 25 °C for the gradient elution. A flow rate of 100 μl min−1 was used with the following buffers: (A) 10 mM ammonium carbonate in water (pH 9.0) and (B) neat acetonitrile. The gradient profile was as follows: 80–20% B (0–30 min), 20–80% B (30–31 min) and 80–80% B (31–42 min). The injection volume was set to 1 μl for all analyses (42 min of total run time per injection). MS analyses were carried out by coupling the LC system to a Thermo Q Exactive HF MS instrument operating in heated electrospray ionization mode. MS was performed for 30 min with a polarity switching data-dependent top 3 method for both positive and negative modes, with targeted MS2 scans for the monoisotopic, 13CU and 13CU/15NU valine m/z values. The spray voltage for both positive and negative modes was 3.5 kV and the capillary temperature was set to 320 °C with a sheath gas rate of 35, auxiliary gas rate of 10 and maximum spray current of 100 μA. The full MS scan for both polarities were conducted at 120,000 resolution with an automatic gain control (AGC) target of 3 × 106 and a maximum injection time (IT) of 100 ms; the scan range was 67–1,000 m/z. Tandem MS spectra for both positive and negative mode used a resolution of 15,000, AGC target of 1 × 105, maximum IT of 50 ms, isolation window of 0.4 m/z, isolation offset of 0.1 m/z, fixed first mass of 50 m/z and three-way multiplexed normalized collision energies of 10, 35 and 80. The minimum AGC target was 1 × 104 with an intensity threshold of 2 × 105. All data were acquired in profile mode. All valine/isoleucine data were processed using Thermo XCalibur Qualbrowser for manual inspection and annotation of the resulting spectra and peak heights referring to authentic valine/isoleucine standards and labeled internal standards as described.
Identification and analysis of ‘sibling’ clones
To detect samples with globally similar barcode profiles, all pairwise comparisons between samples were computed using a Pearson correlation coefficient. Pairs were retained if each sample had at least two nonzero barcode features (to avoid spurious correlations from sparse data) and the Pearson correlation coefficient between the sample profiles was ≥0.9. To organize putative sibling clones identified through barcode profile correlation into groups, we applied a graph-based clustering approach. The set of high-correlation sample pairs was used to construct an undirected graph in which each node represented a sample and each edge represented a strong global match using the NetworkX Python package, with edges connecting sample pairs that met the correlation threshold. Connected components within the graph were then identified, each representing a cluster of samples inferred to be derived from the same original clone or barcoded integration event. Samples not belonging to any component were defined as singletons.
Random forest classification
To identify TU features predictive of valine prototrophy, we trained a random forest classifier using relative barcode read counts as input features. Relative barcode read counts were calculated as the absolute read count for each TU per clone normalized by the total barcode read count for that clone. In total, 119 samples represented by 40 features (that is, TUs) were used to train the model. A binary phenotype label was defined by thresholding the Val score at 0.23, classifying samples as either prototrophic (Val score > 0.23) or nonprototrophic. For each clone, the random forest classifier output an estimated probability of being prototrophic or nonprototrophic, representing the averaged class likelihood produced across all decision trees. Discrete class predictions were obtained by assigning clones to the class with the higher predicted probability. We used stratified fourfold cross-validation to ensure balanced class distributions across splits. In each fold, a random forest classifier was trained on 75% of the data and evaluated on the remaining 25%. Model performance was assessed using confusion matrices, classification reports, area under ROC curves, area under precision–recall curves and average precision. Confusion matrices were plotted per fold and mean performance metrics were reported across folds with s.d. Feature importances were extracted from each trained model to identify the most predictive TUs. The top ten features were reported per fold and mean feature importances were calculated to identify features consistently ranked as important across folds.
RNA-seq
RNA-seq was performed commercially (Plasmidsaurus), yielding at least 10 million single-end reads per sample following transcript purification with poly(A) mRNA capture and cDNA synthesis with oligo-dT primers. Given that SGE transgenes were not polyadenylated (to facilitate use of lentivirus as a delivery method), detection of SGE transgenes using this methodology is underpowered and expected to yield lower reads per kilobase per million mapped reads (RPKM) than endogenous polyadenylated transcripts. Each sample type was submitted in duplicate. Resulting raw fastq files were subjected to adaptor trimming and quality control using trim_galore (version 0.6.10) and cutadapt (version 4.4) with default parameters. For transgene quantification, trimmed reads from CHO samples were aligned to a custom reference genome based on the Cricetulus griseus assembly CriGri-PICRH-1.0 (GCF_003668045.3) to which SGE transgene sequences (ilvA, ilvB, ilvC, ilvD, ilvG, ilvM, ilvN and EGFP) were appended as separate contigs. Trimmed reads from Jurkat samples were aligned to a custom human genome reference based on GRCh38/hg38, to which SGE transgene sequences were appended as separate contigs. Reads were aligned to custom genomes using STAR (version 2.7.11b) using default STAR parameters. For each sample, the number of reads mapped to each contig was extracted from the BAM files using SAMtools (version 1.21) idxstats. This produced per-contig counts along with contig lengths. To account for both sequencing depth and differences in transgene length, expression levels were quantified using RPKM, providing a length-normalized measure of transgene expression comparable across samples and transgenes.
For differential expression analysis, trimmed reads from CHO samples were quasi-mapped against the C. griseus transcriptome derived from assembly CriGri-PICRH-1.0 (GCF_003668045.3) to which SGE transgenes were appended. Trimmed reads from Jurkat samples were quasi-mapped against a Gencode v49 human transcriptome reference to which SGE transgenes were appended. Quasi-mapping was conducted using Salmon (version 1.10.1) with a k-mer size of 25. Transcript abundances were imported to R using tximport and differential expression analysis was performed using DESeq2. Genes with adjusted P < 0.05 and |log2FC (fold change)| > 1 were considered significantly differentially expressed.
For Gene Ontology (GO) enrichment analysis in CHO samples, functional enrichment was performed using mouse orthologs as a proxy. Gene symbols for genes with adjusted P < 0.05 and |log2FC| > 1 were mapped to mouse Entrez identifiers using clusterProfiler against the org.Mm.eg.db database. GO categories were considered significantly enriched at adjusted P < 0.05. For GO enrichment analysis in Jurkat samples, functional enrichment was performed for genes with adjusted P < 0.05 and |log2FC| > 1 using clusterProfiler against the org.Hs.eg.db database.
Long-read sequencing
gDNA was isolated from 3–6 million cells using either the QIAamp DNA mini kit (Qiagen) or the Monarch genomic purification kit (New England Biolabs), incorporating an RNase treatment phase in all instances. To achieve target fragment lengths, 2 µg of gDNA was diluted in 150 µl of nuclease-free water and sheared using g-TUBE (Covaris) at 3,000g. Following shearing, the DNA was purified using the Zymo DNA clean and concentrator kit and analyzed on an Agilent TapeStation 4150 (Genomic DNA ScreenTape) to verify the distribution.
Nanopore libraries were constructed using the Oxford Nanopore native barcoding kit 24 V14 (SQK-NBD114.24). Sample purity and concentration were validated by NanoDrop spectrophotometry and Qubit fluorometry, respectively. Sequencing was performed by loading 80–100 fmol of the library onto R10 MinION flow cells or 100–120 fmol onto PromethION flow cells. To optimize throughput, flow cells were washed and reloaded once pore activity reached approximately 30%.
Raw Oxford Nanopore POD5 files were processed using the Dorado analysis engine (https://github.com/nanoporetech/dorado/releases/tag/v1.0.0). The POD5 files were basecalled again using the super model (dna_r10.4.1_e8.2_400bps_sup@v5.2.0). The resulting BAM files were demultiplexed into a separate BAM file per barcode using Dorado demux. Each BAM file was aligned against its corresponding reference FASTA files using the Dorado align command. Per-base coverage depth tracks were generated using BEDTools (version 2.30.0)62.
BAM files were analyzed to assign TU identities on a per-read basis using contig alignments. Reads were filtered to retain alignments with MAPQ ≥ 50 and length ≥ 200 bp. Primary and supplementary alignments were included, whereas secondary alignments were excluded.
For transgene quantification, PGK-MTS (508-bp promoter + 108-bp MTS) and EF1a-MTS (1,187-bp promoter + 108-bp MTS) sequences were used as promoter contigs. Reads were classified as no-MTS or MTS promoter variants according to whether alignments to these contigs extended beyond positions 550 and 1230 for PGK and EF1a, respectively. For ilvA, ilvAL447F and ilvAL481F, the native ilvA sequence was used as the alignment contig and variant identity was resolved by scanning each read for 15-bp motif windows corresponding to native or mutant sequence states at contig positions 1341 and 1443, respectively (ilvA: native/native; ilvAL447F: mutant/native; ilvAL481F: native/mutant), allowing a Hamming distance of 1. After promoter and CDS assignment, only reads with exactly one promoter call and exactly one CDS call were retained for TU quantification.
For estimation of barcode swapping rates, we additionally filtered reads to retain those with high-confidence alignments to both DNA sequences flanking the 127-bp SGE amplicon allowing a maximal distance of 150 bp between the two alignments. Alignments were used to reconstruct full-length read sequences on the basis of hard clipping information contained within the CIGAR string. Reconstructed reads were parsed to identify the two universal priming regions flanking the 8-bp SGE barcodes, allowing a Hamming distance of 5, and the corresponding 8-bp barcodes were matched to their assigned promoter–OLS or CDS, allowing a Hamming distance of 1.
For identification of lentiviral integration sites, we looked for split-read junctions between integrated sequences (that is, TU modules and lentiviral sequences) and the host genome. Candidate junctions were retrieved from reads in which both alignments had MAPQ ≥ 50 and length ≥ 100 bp; one genomic breakpoint was assigned per read. Breakpoints were clustered by genomic position within a 10-kb window to define candidate integration sites. A candidate integration site was retained if its breakpoint cluster was supported by at least two reads. Each site was mapped to its relevant reference genome annotation (CriGri-PICRH-1.0 for CHO, GRCh38/hg38 for Jurkat) using ChIPseeker and classified by genomic feature (promoter, exon, intron or intergenic).
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
Amplicon, long-read sequencing data were deposited to the Sequence Read Archive under BioProject PRJNA1282762 (ref. 63). Metabolomics data were deposited to Zenodo (https://doi.org/10.5281/zenodo.21874813)64.
Code availability
Computational pipeline for long-read sequencing can be accessed from GitHub (https://github.com/FenyoLab/ont_analysis_pipeline). All other scripts written to analyze data and generate figures can also be accessed from GitHub (https://github.com/julietrolle/shotgun-genetic-engineering).
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Acknowledgements
We thank M. Pacold for sharing Jurkat cells and members of the J.D.B. lab for helpful discussions. Figure schematics were made using BioRender.com and ChatGPT provided assistance in generating code for data analysis. We thank M. Maurano and his team for help with DNA sequencing.
Funding
This study was supported in part by the National Human Genome Research Institute, National Institutes of Health (NIH) grant RM1-HG009491 to J.D.B., NIH grant DP5OD036167 to S.P. and Chan Zuckerberg Initiative grant 2024-349901 to S.P. J.T. is supported by a SeaBridge Fellowship funded by the Washington Research Foundation, Brotman Baty Institute and the Chan Zuckerberg Initiative.
Ethics declarations
Competing interests
Two patent applications related to this work have been filed with J.T., S.P. and J.D.B. listed as inventors on one or both applications (US Patent application nos. 63/533,483 and 63/828,744). J.D.B. is a founder and director of CDI Labs, a founder of and consultant to Opentrons LabWorks/Neochromosome and a founder of JATech and serves or served on the scientific advisory boards of CZ Biohub New York, Logomix, Rome Therapeutics, SeaHub, Tessera Therapeutics and the Wyss Institute.
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Peer review information
Nature Biotechnology thanks Caleb Bashor and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.
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Extended data
Extended Data Fig. 1 Transcription unit distribution in CHO pre-infection (DNA Library) and post-infection (CHO-Div) of SGE library.
(a) Transcription unit distribution prior to lentiviral packaging compared to transcription unit distribution after infection of CHO cells. Y-axis indicates promoter/OLS context while the X-axis indicates CDS. Numbers are percentages; that is, all TU numbers add up to 100%. (b) qPCR to determine copy number of TU integrants relative to two single copy genomic targets (Dhfr, Hprt) for estimation of MOI. (c) Probability of finding a known valine biosynthesis pathway combination in CHO-Div based on a Monte Carlo simulated infection of 2.42 M cells with the library distribution measured for CHO-Div cells in (A). Depicted is the probability of finding ilvN, ilvB, ilvC and ilvD irrespective of subcellular compartmentalization (black), all 4 genes localized to the cytoplasm (red), or all 4 genes localized to mitochondria (green) at a range of MOIs. Panel a created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
Extended Data Fig. 2 Characterization of clones selected on low valine medium, including 13C tracing to confirm endogenous biosynthesis of valine in Clone CHO-Val-D1.
(a) Absolute cell count of clones CHO-Val-D1 and CHO-Val-D3 cultured in valine-replete RPMI compared to the parental cell line, previously engineered pMTIV cells, and SGE-Informed Design. Error bars represent standard deviation across triplicate wells. (b) Schematic detailing 13C labeling strategy to detect biosynthesized valine. Prototrophic cells were cultured on valine-free 13C labeled medium while control cells were cultured in a 13C valine-replete equivalent. (c) Extracted ion chromatography shows detection of 13C Valine in Clone CHO-Val-D1, which matches the retention time of spiked-in internal standard 13C5/15N1 Valine. Detection of 13C Valine in parental control cells is minimal by comparison. (d) MS1 peak corresponding to 13C5 Valine is seen for Clone CHO-Val-D1 and not for the parental control cells. (e) MS2 confirms that fragmentation patterns for putative 12C5 valine, 13C5 valine and the 13C5/15N1 valine internal standard match expectation.
Extended Data Fig. 3 Genotypic characterization of clones CHO-Val-D1 and CHO-Val-D3.
(a) Log-transformed read counts were used to calculate a Z-score for each TU across all valine prototrophic clones. A Z-score threshold of 1.15 was used to define TU presence vs. TU absence across the 16 clones. (b) Percentage of amplicon-seq reads corresponding to each TU within clones CHO-Val-D1 and CHO-Val-D3. (c) Percentage of long reads corresponding to each TU in clones CHO-Val-D1 and CHO-Val-D3. (d) Concordance of TUs called by amplicon-seq and by long-read sequencing in clones CHO-Val-D1 and CHO-Val-D3. Amplicon-seq presence/absence calls utilize a Z-score threshold of 1.15 to separate signal from background, while no threshold was utilized for long-read sequencing. (e) Transgene quantification of SGE-introduced CDSs by RNA-seq in clones CHO-Val-D1 and CHO-Val-D3. Given near-identical CDS for ilvA, ilvA-L447F and ilvA-L481F, these could not be differentiated from each other in this analysis and reads mapping to any of these are presented collectively.
Extended Data Fig. 4 Characterization of clones selected on low isoleucine medium including 13C tracing to confirm endogenous biosynthesis of isoleucine in Clone CHO-Ile-10-H1.
(a) Absolute cell counts of clones CHO-Ile-10-H1 and CHO-Ile-10-H2 cultured on isoleucine-replete medium compared to the parental cell line. Error bars represent standard deviation across triplicate wells. (b) Schematic detailing 13C labeling strategy to detect biosynthesized isoleucine. Prototrophic cells were cultured on isoleucine-free 13C labeled medium while control cells were cultured in a 13C isoleucine-replete equivalent. (c) MS2 confirms that fragmentation patterns for putative 12C isoleucine, 13C2 isoleucine, 13C6 isoleucine and the 13C6/15N1 isoleucine internal standard match expectation.
Extended Data Fig. 5 Genotypic characterization of clones CHO-Ile-10-H1 and CHO-Ile-10-H2.
(a) Log-transformed read counts were used to calculate a Z-score for each TU across all isoleucine prototrophic clones. A Z-score threshold of 1.15 was used to define TU presence vs. TU absence across the 46 clones. (b) Percentage of amplicon-seq reads corresponding to each TU within clones CHO-Ile-10-H1 and CHO-Ile-10-H2. (c) Percentage of long reads corresponding to each TU in clones CHO-Ile-10-H1 and CHO-Ile-10-H2. (d) Concordance of TUs called by amplicon-seq and by long-read sequencing in clones CHO-Ile-10-H1 and CHO-Ile-10-H2. Amplicon-seq presence/absence calls utilize a Z-score threshold of 1.15 to separate signal from background, while no threshold was utilized for long-read sequencing. (e) Transgene quantification of SGE-introduced CDSs by RNA-seq in clones CHO-Ile-10-H1 and CHO-Ile-10-H2. Given near-identical CDS for ilvA, ilvA-L447F and ilvA-L481F, these could not be differentiated from each other in this analysis and reads mapping to any of these are presented collectively.
Extended Data Fig. 6 Sampling CHO-Val and CHO-Ile clones for double isoleucine/valine prototrophy.
(a) PrestoBlue amino acid dropout assay of 16 CHO-Val clones and 46 CHO-Ile clones cultured on isoleucine-free, valine-free RPMI. Metabolic activity was measured in triplicate following 3 days of growth on double isoleucine-free, valine-free medium. Error bars represent standard deviation across triplicate wells. (b) Percentage of double isoleucine/valine prototrophic clones that contain the indicated CDS. (c) CDS presence amongst double isoleucine/valine prototrophic CHO-Val clones and double isoleucine/valine prototrophic CHO-Ile clones. (d) Phenotype distribution amongst 16 CHO-Val clones and 46 CHO-Ile clones. (e) CDS presence by phenotype in CHO-Val (left) and CHO-Ile clones (right) as well as the expected distribution according to simulated data. Notable differences across phenotypes among CHO-Ile clones are highlighted in red boxes for ilvA, ilvA-L481F, and ilvG.
Extended Data Fig. 7 Transcription unit distribution in Jurkat pre-infection (DNA Library) and post-infection (Jurkat-Div) of SGE library.
(a) Transcription unit distribution prior to lentiviral packaging compared to transcription unit distribution after infection of Jurkat cells (b) qPCR of TU integrants relative to two double copy genomic targets (RPP30, ALB) for estimation of MOI. (c) Probability of finding a presumed valine biosynthesis pathway combination post-infection based on a simulated infection of 3 M Jurkat cells with the library distribution measured for Jurkat-Div cells in (A). Depicted is the probability of finding ilvN, ilvB, ilvC and ilvD irrespective of subcellular compartmentalization (black), all 4 genes localized to the cytoplasm (red), or all 4 genes localized to mitochondria (green) at a range of MOIs. (d) Probability of finding a presumed isoleucine biosynthesis pathway combination post-infection based on a simulated infection of 3 M Jurkat cells with the library distribution measured for Jurkat-Div cells in (A). Depicted is the probability of finding ilvM, ilvG, ilvB, ilvC, ilvD and ilvA-L481F irrespective of subcellular compartmentalization (black), all 6 genes localized to the cytoplasm (red), or all 6 genes localized to mitochondria (green) at a range of MOIs. Panel a created in BioRender; Trolle, J. https://biorender.com/43izsew (2026).
Extended Data Fig. 8 Long-term selection of Jurkat-Div cells led to identification of valine prototrophic Jurkat clones.
(a) Cell counts of Jurkat-Div and Jurkat-GFP control cells over 21 days of selection on a range of low valine RPMI conditions. (b) Growth curve of clones Jurkat-Val-31 and Jurkat-Val-32 cultured on valine-replete medium. Error bars represent standard deviation across triplicate wells. (c) Cell counts of Jurkat-Div and Jurkat-GFP control cells over 32+ days of selection on a range of low isoleucine RPMI conditions.
Extended Data Fig. 9 Characterization of Jurkat-Div clones selected on low valine medium.
(a) Log-transformed read counts were used to calculate a Z-score for each TU across all valine prototrophic clones. A Z-score threshold of 1.15 was used to define TU presence vs. TU absence across the 119 clones. (b) Percentage of amplicon-seq reads corresponding to each TU within clone Jurkat-Val-32. (c) Percentage of amplicon-seq reads corresponding to each TU within clone Jurkat-Val-31. (d) Percentage of long reads corresponding to each TU in clone Jurkat-Val-31. (e) Concordance of TUs called by amplicon-seq and by long-read sequencing in clone Jurkat-Val-31. Amplicon-seq presence/absence calls utilize a Z-score threshold of 1.15 to separate signal from background, while no threshold was utilized for long-read sequencing. (f) Frequency of barcode swapping in clone Jurkat-Val-31 as detected by long-read sequencing. (g) Transgene quantification of SGE-introduced CDSs by RNA-seq in clone Jurkat-Val-31. Given near-identical CDS for ilvA, ilvA-L447F and ilvA-L481F, these could not be differentiated from each other in this analysis and reads mapping to any of these are presented collectively.
Extended Data Fig. 10 Many clones arose from the same founder clones within the Jurkat-Val population.
(a) Composite barcode ‘fingerprint’ across strongly valine prototrophic clones. For each TU within a sample, the percentage of barcodes assigned to that TU was calculated. The median percentage representation of each TU across all clones is displayed as a composite heatmap. (b) Relative abundance of TUs, which comprise the valine prototrophic pathway ‘solution’ and their 9 underlying barcode combinations (3 per promoter/OLS, 3 per CDS) across all strongly valine prototrophic clones (Val Score > 0.75). (c) Distribution of clones that are part of sibling clonal groups and clones that are singletons. (d) Graph-based clustering identified 10 separate sibling groups in Jurkat clones selected on low valine medium ranging in size from 2–24 clones each. Simple linear regression indicates that sibling group size does not significantly correlate to an increase in average Val score (R2 = 0.15; 95% CI for slope: −0.007 to 0.021; DFn = 1; DFd = 8; P-value = 0.2736). (e) Representative heatmaps for each of the 4 largest sibling groups: Groups 2, 4, 9, and 5. (f) Percentage of all valine prototrophic clones (Val Score > 0.23) and all strongly valine prototrophic clones (Val Score > 0.75) that contain each depicted CDS as measured by amplicon-seq after removal of duplicate sibling clones.
Supplementary information
Reporting Summary (download PDF )
Peer Review file (download PDF )
Supplementary Tables 1–10 (download XLSX )
Supplementary Table 1: Long reads mapped to SGE transgenes in Jurkat-Val-31. Supplementary Table 2: Long reads mapped to SGE transgenes in CHO-Val-D1, CHO-Val-D3, CHO-Ile-10-H1 and CHO-Ile-10-H2. Supplementary Table 3: SGE transgene integration site mapping in CHO-Val-D1, CHO-Val-D3, CHO-Ile-10-H1 and CHO-Ile-10-H2. Supplementary Table 4: Differentially expressed genes across all pairwise comparisons between CHO parental cells (valine-replete medium), CHO-Val-D1 (valine-replete medium) and CHO-Val-D1 (valine-free medium), as well as between CHO parental cells (valine-replete medium) and CHO-Val-D3 (valine-free medium). Supplementary Table 5: Differentially expressed genes across all pairwise comparisons between CHO parental cells (isoleucine-replete medium), CHO-Ile-10-H1 (isoleucine-replete medium) and CHO-Ile-10-H1 (isoleucine-free medium), as well as between CHO parental cells (isoleucine-replete medium) and CHO-Ile-10-H1 (isoleucine-free medium). Supplementary Table 6: SGE transgene integration site mapping in Jurkat-Val-31. Supplementary Table 7: Differentially expressed genes across all pairwise comparisons between Jurkat parental cells (valine-replete medium), Jurkat-Val-31 (valine-replete medium) and Jurkat-Val-31 (valine-free medium). Supplementary Table 8: Sequences for promoters, OLSs and CDSs used in the SGE library. Supplementary Table 9: Raw barcode read counts from amplicon-seq across all sequenced clones and controls. Supplementary Table 10: TU aggregated barcode counts from amplicon-seq across all sequenced clones and controls.
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Trolle, J., Sessa, S., Wudzinska, A. et al. Highly multiplexed mammalian metabolic engineering with a shotgun approach. Nat Biotechnol (2026). https://doi.org/10.1038/s41587-026-03318-7
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DOI: https://doi.org/10.1038/s41587-026-03318-7







