A versatile CRISPR-Cas13d platform for multiplexed transcriptomic regulation and metabolic engineering in primary human T cells.

CRISPR technologies have begun to revolutionize T cell therapies; however, conventional CRISPR-Cas9 genome-editing tools are limited in their safety, efficacy, and scope. To address these challenges, we developed multiplexed effector guide arrays (MEGA), a platform for programmable and scalable regulation of the T cell transcriptome using the RNA-guided, RNA-targeting activity of CRISPR-Cas13d. MEGA enables quantitative, reversible, and massively multiplexed gene knockdown in primary human T cells without targeting or cutting genomic DNA. Applying MEGA to a model of CAR T cell exhaustion, we robustly suppressed inhibitory receptor upregulation and uncovered paired regulators of T cell function through combinatorial CRISPR screening. We additionally implemented druggable regulation of MEGA to control CAR activation in a receptor-independent manner. Lastly, MEGA enabled multiplexed disruption of immunoregulatory metabolic pathways to enhance CAR T cell fitness and anti-tumor activity in vitro and in vivo. MEGA offers a versatile synthetic toolkit for applications in cancer immunotherapy and beyond.

Spatial engineering of E. coli with addressable phase-separated RNAs.

Biochemical processes often require spatial regulation and specific microenvironments. The general lack of organelles in bacteria limits the potential of bioengineering complex intracellular reactions. Here, we demonstrate synthetic membraneless organelles in Escherichia coli termed transcriptionally engineered addressable RNA solvent droplets (TEARS). TEARS are assembled from RNA-binding protein recruiting domains fused to poly-CAG repeats that spontaneously drive liquid-liquid phase separation from the bulk cytoplasm. Targeting TEARS with fluorescent proteins revealed multilayered structures with composition and reaction robustness governed by non-equilibrium dynamics. We show that TEARS provide organelle-like bioprocess isolation for sequestering biochemical pathways, controlling metabolic branch points, buffering mRNA translation rates, and scaffolding protein-protein interactions. We anticipate TEARS to be a simple and versatile tool for spatially controlling E. coli biochemistry. Particularly, the modular design of TEARS enables applications without expression fine-tuning, simplifying the design-build-test cycle of bioengineering.

Biofuels for a sustainable future.

Rapid increases of energy consumption and human dependency on fossil fuels have led to the accumulation of greenhouse gases and consequently, climate change. As such, major efforts have been taken to develop, test, and adopt clean renewable fuel alternatives. Production of bioethanol and biodiesel from crops is well developed, while other feedstock resources and processes have also shown high potential to provide efficient and cost-effective alternatives, such as landfill and plastic waste conversion, algal photosynthesis, as well as electrochemical carbon fixation. In addition, the downstream microbial fermentation can be further engineered to not only increase the product yield but also expand the chemical space of biofuels through the rational design and fine-tuning of biosynthetic pathways toward the realization of "designer fuels" and diverse future applications.

Converting Escherichia coli to a Synthetic Methylotroph Growing Solely on Methanol.

Methanol, being electron rich and derivable from methane or CO2, is a potentially renewable one-carbon (C1) feedstock for microorganisms. Although the ribulose monophosphate (RuMP) cycle used by methylotrophs to assimilate methanol differs from the typical sugar metabolism by only three enzymes, turning a non-methylotrophic organism to a synthetic methylotroph that grows to a high cell density has been challenging. Here we reprogrammed E. coli using metabolic robustness criteria followed by laboratory evolution to establish a strain that can efficiently utilize methanol as the sole carbon source. This synthetic methylotroph alleviated a so far uncharacterized hurdle, DNA-protein crosslinking (DPC), by insertion sequence (IS)-mediated copy number variations (CNVs) and balanced the metabolic flux by mutations. Being capable of growing at a rate comparable with natural methylotrophs in a wide range of methanol concentrations, this synthetic methylotrophic strain illustrates genome editing and evolution for microbial tropism changes and expands the scope of biological C1 conversion.

Reprogramming Yeast Metabolism from Alcoholic Fermentation to Lipogenesis.

Engineering microorganisms for production of fuels and chemicals often requires major re-programming of metabolism to ensure high flux toward the product of interest. This is challenging, as millions of years of evolution have resulted in establishment of tight regulation of metabolism for optimal growth in the organism's natural habitat. Here, we show through metabolic engineering that it is possible to alter the metabolism of Saccharomyces cerevisiae from traditional ethanol fermentation to a pure lipogenesis metabolism, resulting in high-level production of free fatty acids. Through metabolic engineering and process design, we altered subcellular metabolic trafficking, fine-tuned NADPH and ATP supply, and decreased carbon flux to biomass, enabling production of 33.4 g/L extracellular free fatty acids. We further demonstrate that lipogenesis metabolism can replace ethanol fermentation by deletion of pyruvate decarboxylase enzymes followed by adaptive laboratory evolution. Genome sequencing of evolved strains showed that pyruvate kinase mutations were essential for this phenotype.

Systems Biology of Metabolism.

Metabolism is highly complex and involves thousands of different connected reactions; it is therefore necessary to use mathematical models for holistic studies. The use of mathematical models in biology is referred to as systems biology. In this review, the principles of systems biology are described, and two different types of mathematical models used for studying metabolism are discussed: kinetic models and genome-scale metabolic models. The use of different omics technologies, including transcriptomics, proteomics, metabolomics, and fluxomics, for studying metabolism is presented. Finally, the application of systems biology for analyzing global regulatory structures, engineering the metabolism of cell factories, and analyzing human diseases is discussed.

Precise Editing at DNA Replication Forks Enables Multiplex Genome Engineering in Eukaryotes.

We describe a multiplex genome engineering technology in Saccharomyces cerevisiae based on annealing synthetic oligonucleotides at the lagging strand of DNA replication. The mechanism is independent of Rad51-directed homologous recombination and avoids the creation of double-strand DNA breaks, enabling precise chromosome modifications at single base-pair resolution with an efficiency of >40%, without unintended mutagenic changes at the targeted genetic loci. We observed the simultaneous incorporation of up to 12 oligonucleotides with as many as 60 targeted mutations in one transformation. Iterative transformations of a complex pool of oligonucleotides rapidly produced large combinatorial genomic diversity >105. This method was used to diversify a heterologous ß-carotene biosynthetic pathway that produced genetic variants with precise mutations in promoters, genes, and terminators, leading to altered carotenoid levels. Our approach of engineering the conserved processes of DNA replication, repair, and recombination could be automated and establishes a general strategy for multiplex combinatorial genome engineering in eukaryotes.

Engineering Cellular Metabolism.

Metabolic engineering is the science of rewiring the metabolism of cells to enhance production of native metabolites or to endow cells with the ability to produce new products. The potential applications of such efforts are wide ranging, including the generation of fuels, chemicals, foods, feeds, and pharmaceuticals. However, making cells into efficient factories is challenging because cells have evolved robust metabolic networks with hard-wired, tightly regulated lines of communication between molecular pathways that resist efforts to divert resources. Here, we will review the current status and challenges of metabolic engineering and will discuss how new technologies can enable metabolic engineering to be scaled up to the industrial level, either by cutting off the lines of control for endogenous metabolism or by infiltrating the system with disruptive, heterologous pathways that overcome cellular regulation.

Engineering peroxisomal biosynthetic pathways for maximization of triterpene production in Yarrowia lipolytica.

Constructing efficient cell factories for product synthesis is frequently hampered by competing pathways and/or insufficient precursor supply. This is particularly evident in the case of triterpenoid biosynthesis in Yarrowia lipolytica, where squalene biosynthesis is tightly coupled to cytosolic biosynthesis of sterols essential for cell viability. Here, we addressed this problem by reconstructing the complete squalene biosynthetic pathway, starting from acetyl-CoA, in the peroxisome, thus harnessing peroxisomal acetyl-CoA pool and sequestering squalene synthesis in this organelle from competing cytosolic reactions. This strategy led to increasing the squalene levels by 1,300-fold relatively to native cytosolic synthesis. Subsequent enhancement of the peroxisomal acetyl-CoA supply by two independent approaches, 1) converting cellular lipid pool to peroxisomal acetyl-CoA and 2) establishing an orthogonal acetyl-CoA shortcut from CO2-derived acetate in the peroxisome, further significantly improved local squalene accumulation. Using these approaches, we constructed squalene-producing strains capable of yielding 32.8 g/L from glucose, and 31.6 g/L from acetate by employing a cofeeding strategy, in bioreactor fermentations. Our findings provide a feasible strategy for protecting intermediate metabolites that can be claimed by multiple reactions by engineering peroxisomes in Y. lipolytica as microfactories for the production of such intermediates and in particular acetyl-CoA-derived metabolites.

Peroxisomal metabolic coupling improves fatty alcohol production from sole methanol in yeast.

Methanol is an ideal feedstock for chemical and biological manufacturing. Constructing an efficient cell factory is essential for producing complex compounds through methanol biotransformation, in which coordinating methanol use and product synthesis is often necessary. In methylotrophic yeast, methanol utilization mainly occurs in peroxisomes, which creates challenges in driving the metabolic flux toward product biosynthesis. Here, we observed that constructing the cytosolic biosynthesis pathway resulted in compromised fatty alcohol production in the methylotrophic yeast Ogataea polymorpha. Alternatively, peroxisomal coupling of fatty alcohol biosynthesis and methanol utilization significantly improved fatty alcohol production by 3.9-fold. Enhancing the supply of precursor fatty acyl-CoA and cofactor NADPH in the peroxisomes by global metabolic rewiring further improved fatty alcohol production by 2.5-fold and produced 3.6 g/L fatty alcohols from methanol under fed-batch fermentation. We demonstrated that peroxisome compartmentalization is helpful for coupling methanol utilization and product synthesis, and with this approach, constructing efficient microbial cell factories for methanol biotransformation is feasible.

Engineering a universal and efficient platform for terpenoid synthesis in yeast.

Engineering microbes for the production of valuable natural products is often hindered by the regulation of native competing metabolic networks in host. This is particularly evident in the case of terpenoid synthesis in yeast, where the canonical terpenoid precursors are tightly coupled to the biosynthesis of sterols essential for yeast viability. One way to circumvent this limitation is by engineering product pathways less connected to the host native metabolism. Here, we introduce a two-step isopentenol utilization pathway (IUP) in Saccharomyces cerevisiae to augment the native mevalonate pathway by providing a shortcut to the synthesis of the common terpenoid precursors, isopentenyl diphosphate (IPP) and dimethylallyl diphosphate (DMAPP). As such, the IUP was capable of elevating the IPP/DMAPP pool by 147-fold compared with the native pathway. We further demonstrate that cofeeding isoprenol and prenol enhances geranyl diphosphate (GPP) content for monoterpene biosynthesis. More importantly, we established a synthetic three-step route for efficient synthesis of di-and tetraterpene precursor geranylgeranyl diphosphate (GGPP), circumventing the competition with farnesyl diphosphate (FPP) for sterol biosynthesis and elevating the GGPP level by 374-fold. We combine these IUP-supported precursor-forming platforms with downstream terpene synthases to harness their potential and improve the production of industrially relevant terpenoids by several fold. Our exploration provides a universal and effective platform for supporting terpenoid synthesis in yeast.

A versatile in situ cofactor enhancing system for meeting cellular demands for engineered metabolic pathways.

Cofactor imbalance obstructs the productivities of metabolically engineered cells. Herein, we employed a minimally perturbing system, xylose reductase and lactose (XR/lactose), to increase the levels of a pool of sugar phosphates which are connected to the biosynthesis of NAD(P)H, FAD, FMN, and ATP in Escherichia coli. The XR/lactose system could increase the amounts of the precursors of these cofactors and was tested with three different metabolically engineered cell systems (fatty alcohol biosynthesis, bioluminescence light generation, and alkane biosynthesis) with different cofactor demands. Productivities of these cells were increased 2-4-fold by the XR/lactose system. Untargeted metabolomic analysis revealed different metabolite patterns among these cells, demonstrating that only metabolites involved in relevant cofactor biosynthesis were altered. The results were also confirmed by transcriptomic analysis. Another sugar reducing system (glucose dehydrogenase) could also be used to increase fatty alcohol production but resulted in less yield enhancement than XR. This work demonstrates that the approach of increasing cellular sugar phosphates can be a generic tool to increase in vivo cofactor generation upon cellular demand for synthetic biology.

Verbascum species as a new source of saffron apocarotenoids and molecular tools for the biotechnological production of crocins and picrocrocin.

Crocins are glucosylated apocarotenoids present in flowers and fruits of a few plant species, including saffron, gardenia, and Buddleja. The biosynthesis of crocins in these plants has been unraveled, and the enzymes engineered for the production of crocins in heterologous systems. Mullein (Verbascum sp.) has been identified as a new source of crocins and picrocrocin. In this work, we have identified eight enzymes involved in the cleavage of carotenoids in two Verbascum species, V. giganteum and V. sinuatum. Four of them were homologous to the previously identified BdCCD4.1 and BdCCD4.3 from Buddleja, involved in the biosynthesis of crocins. These enzymes were analyzed for apocarotenogenic activity in bacteria and Nicotiana benthamiana plants using a virus-driven system. Metabolic analyses of bacterial extracts and N. benthamiana leaves showed the efficient activity of these enzymes to produce crocins using ß-carotene and zeaxanthin as substrates. Accumulations of 0.17% of crocins in N. benthamiana dry leaves were reached in only 2 weeks using a recombinant virus expressing VgCCD4.1, similar to the amounts previously produced using the canonical saffron CsCCD2L. The identification of these enzymes, which display a particularly broad substrate spectrum, opens new avenues for apocarotenoid biotechnological production.

Ectopic expression of a bacterial thiamin monophosphate kinase enhances vitamin B1 biosynthesis in plants.

Plants and bacteria have distinct pathways to synthesize the bioactive vitamin B1 thiamin diphosphate (TDP). In plants, thiamin monophosphate (TMP) synthesized in the TDP biosynthetic pathway is first converted to thiamin by a phosphatase, which is then pyrophosphorylated to TDP. In contrast, bacteria use a TMP kinase encoded by ThiL to phosphorylate TMP to TDP directly. The Arabidopsis THIAMIN REQUIRING2 (TH2)-encoded phosphatase is involved in TDP biosynthesis. The chlorotic th2 mutants have high TMP and low thiamin and TDP. Ectopic expression of Escherichia coli ThiL and ThiL-GFP rescued the th2-3 mutant, suggesting that the bacterial TMP kinase could directly convert TMP into TDP in Arabidopsis. These results provide direct evidence that the chlorotic phenotype of th2-3 is caused by TDP rather than thiamin deficiency. Transgenic Arabidopsis harboring engineered ThiL-GFP targeting to the cytosol, chloroplast, mitochondrion, or nucleus accumulated higher TDP than the wild type (WT). Ectopic expression of E. coli ThiL driven by the UBIQUITIN (UBI) promoter or an endosperm-specific GLUTELIN1 (GT1) promoter also enhanced TDP biosynthesis in rice. The pUBI:ThiL transgenic rice accumulated more TDP and total vitamin B1 in the leaves, and the pGT1:ThiL transgenic lines had higher TDP and total vitamin B1 in the seeds than the WT. Total vitamin B1 only increased by approximately 25-30% in the polished and unpolished seeds of the pGT1:ThiL transgenic rice compared to the WT. Nevertheless, these results suggest that genetic engineering of a bacterial vitamin B1 biosynthetic gene downstream of TMP can enhance vitamin B1 production in rice.

Conversion of CO2 into organic acids by engineered autotrophic yeast.

The increase of CO2 emissions due to human activity is one of the preeminent reasons for the present climate crisis. In addition, considering the increasing demand for renewable resources, the upcycling of CO2 as a feedstock gains an extensive importance to establish CO2-neutral or CO2-negative industrial processes independent of agricultural resources. Here we assess whether synthetic autotrophic Komagataella phaffii (Pichia pastoris) can be used as a platform for value-added chemicals using CO2 as a feedstock by integrating the heterologous genes for lactic and itaconic acid synthesis. 13C labeling experiments proved that the resulting strains are able to produce organic acids via the assimilation of CO2 as a sole carbon source. Further engineering attempts to prevent the lactic acid consumption increased the titers to 600 mg L-1, while balancing the expression of key genes and modifying screening conditions led to 2 g L-1 itaconic acid. Bioreactor cultivations suggest that a fine-tuning on CO2 uptake and oxygen demand of the cells is essential to reach a higher productivity. We believe that through further metabolic and process engineering, the resulting engineered strain can become a promising host for the production of value-added bulk chemicals by microbial assimilation of CO2, to support sustainability of industrial bioprocesses.

Biosynthesis of tetrahydropapaverine and semisynthesis of papaverine in yeast.

Tetrahydropapaverine (THP) and papaverine are plant natural products with clinically significant roles. THP is a precursor in the production of the drugs atracurium and cisatracurium, and papaverine is used as an antispasmodic during vascular surgery. In recent years, metabolic engineering advances have enabled the production of natural products through heterologous expression of pathway enzymes in yeast. Heterologous biosynthesis of THP and papaverine could play a role in ensuring a stable supply of these clinically significant products. Biosynthesis of THP and papaverine has not been achieved to date, in part because multiple pathway enzymes have not been elucidated. Here, we describe the development of an engineered yeast strain for de novo biosynthesis of THP. The production of THP is achieved through heterologous expression of two enzyme variants with activity on nonnative substrates. Through protein engineering, we developed a variant of N-methylcoclaurine hydroxylase with activity on coclaurine, enabling de novo norreticuline biosynthesis. Similarly, we developed a variant of scoulerine 9-O-methyltransferase capable of O-methylating 1-benzylisoquinoline alkaloids at the 3' position, enabling de novo THP biosynthesis. Flux through the heterologous pathway was improved by knocking out yeast multidrug resistance transporters and optimization of media conditions. Overall, strain engineering increased the concentration of biosynthesized THP 600-fold to 121 µg/L. Finally, we demonstrate a strategy for papaverine semisynthesis using hydrogen peroxide as an oxidizing agent. Through optimizing pH, temperature, reaction time, and oxidizing agent concentration, we demonstrated the ability to produce semisynthesized papaverine through oxidation of biosynthesized THP.

Methanol biotransformation toward high-level production of fatty acid derivatives by engineering the industrial yeast Pichia pastoris.

Methanol-based biorefinery is a promising strategy to achieve carbon neutrality goals by linking CO2 capture and solar energy storage. As a typical methylotroph, Pichia pastoris shows great potential in methanol biotransformation. However, challenges still remain in engineering methanol metabolism for chemical overproduction. Here, we present the global rewiring of the central metabolism for efficient production of free fatty acids (FFAs; 23.4 g/L) from methanol, with an enhanced supply of precursors and cofactors, as well as decreased accumulation of formaldehyde. Finally, metabolic transforming of the fatty acid cell factory enabled overproduction of fatty alcohols (2.0 g/L) from methanol. This study demonstrated that global metabolic rewiring released the great potential of P. pastoris for methanol biotransformation toward chemical overproduction.

Engineering nonphotosynthetic carbon fixation for production of bioplastics by methanogenic archaea.

The conversion of CO2 to value-added products allows both capture and recycling of greenhouse gas emissions. While plants and other photosynthetic organisms play a key role in closing the global carbon cycle, their dependence on light to drive carbon fixation can be limiting for industrial chemical synthesis. Methanogenic archaea provide an alternative platform as an autotrophic microbial species capable of non-photosynthetic CO2 fixation, providing a potential route to engineered microbial fermentation to synthesize chemicals from CO2 without the need for light irradiation. One major challenge in this goal is to connect upstream carbon-fixation pathways with downstream biosynthetic pathways, given the distinct differences in metabolism between archaea and typical heterotrophs. We engineered the model methanogen, Methanococcus maripaludis, to divert acetyl-coenzyme A toward biosynthesis of value-added chemicals, including the bioplastic polyhydroxybutyrate (PHB). A number of studies implicated limitations in the redox pool, with NAD(P)(H) pools in M. maripaludis measured to be <15% of that of Escherichia coli, likely since methanogenic archaea utilize F420 and ferredoxins instead. Multiple engineering strategies were used to precisely target and increase the cofactor pool, including heterologous expression of a synthetic nicotinamide salvage pathway as well as an NAD+-dependent formate dehydrogenase from Candida boidinii. Engineered strains of M. maripaludis with improved NADH pools produced up to 171 ± 4 mg/L PHB and 24.0 ± 1.9% of dry cell weight. The metabolic engineering strategies presented in this study broaden the utility of M. maripaludis for sustainable chemical synthesis using CO2 and may be transferable to related archaeal species.

Toward a glycyl radical enzyme containing synthetic bacterial microcompartment to produce pyruvate from formate and acetate.

Formate has great potential to function as a feedstock for biorefineries because it can be sustainably produced by a variety of processes that don't compete with agricultural production. However, naturally formatotrophic organisms are unsuitable for large-scale cultivation, difficult to engineer, or have inefficient native formate assimilation pathways. Thus, metabolic engineering needs to be developed for model industrial organisms to enable efficient formatotrophic growth. Here, we build a prototype synthetic formate utilizing bacterial microcompartment (sFUT) encapsulating the oxygen-sensitive glycyl radical enzyme pyruvate formate lyase and a phosphate acyltransferase to convert formate and acetyl-phosphate into the central biosynthetic intermediate pyruvate. This metabolic module offers a defined environment with a private cofactor coenzyme A that can cycle efficiently between the encapsulated enzymes. To facilitate initial design-build-test-refine cycles to construct an active metabolic core, we used a "wiffleball" architecture, defined as an icosahedral bacterial microcompartment (BMC) shell with unoccupied pentameric vertices to freely permit substrate and product exchange. The resulting sFUT prototype wiffleball is an active multi enzyme synthetic BMC functioning as platform technology.

Yeast has evolved to minimize protein resource cost for synthesizing amino acids.

Proteins, as essential biomolecules, account for a large fraction of cell mass, and thus the synthesis of the complete set of proteins (i.e., the proteome) represents a substantial part of the cellular resource budget. Therefore, cells might be under selective pressures to optimize the resource costs for protein synthesis, particularly the biosynthesis of the 20 proteinogenic amino acids. Previous studies showed that less energetically costly amino acids are more abundant in the proteomes of bacteria that survive under energy-limited conditions, but the energy cost of synthesizing amino acids was reported to be weakly associated with the amino acid usage in Saccharomyces cerevisiae Here we present a modeling framework to estimate the protein cost of synthesizing each amino acid (i.e., the protein mass required for supporting one unit of amino acid biosynthetic flux) and the glucose cost (i.e., the glucose consumed per amino acid synthesized). We show that the logarithms of the relative abundances of amino acids in S. cerevisiae's proteome correlate well with the protein costs of synthesizing amino acids (Pearson's r = -0.89), which is better than that with the glucose costs (Pearson's r = -0.5). Therefore, we demonstrate that S. cerevisiae tends to minimize protein resource, rather than glucose or energy, for synthesizing amino acids.