Metabolic engineering can have a pivotal role in increasing the environmental sustainability of the transportation and chemical manufacturing sectors. The field has already developed engineered microorganisms that are currently being used in industrial-scale processes. However, it is often challenging to achieve the titres, yields and productivities required for commercial viability. The efficiency of microbial chemical production is usually dependent on the physiological traits of the host organism, which may either impose limitations on engineered biosynthetic pathways or, conversely, boost their performance. In this Review, we discuss different aspects of microbial physiology that often create obstacles for metabolic engineering, and present solutions to overcome them. We also describe various instances in which natural or engineered physiological traits in host organisms have been harnessed to benefit engineered metabolic pathways for chemical production.
Subscribe to Journal
Get full journal access for 1 year
only $8.25 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Tax calculation will be finalised during checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Lee, S. Y. et al. A comprehensive metabolic map for production of bio-based chemicals. Nat. Catal. 2, 18–33 (2019). This review provides a comprehensive metabolic map of hundreds of chemicals that can be synthesized via metabolic engineering.
Deparis, Q., Claes, A., Foulquié-Moreno, M. R. & Thevelein, J. M. Engineering tolerance to industrially relevant stress factors in yeast cell factories. FEMS Yeast Res. 17, fox036 (2017). This review outlines different stress factors often found in industrial production processes involving yeast cell factories.
Mukhopadhyay, A. Tolerance engineering in bacteria for the production of advanced biofuels and chemicals. Trends Microbiol. 23, 498–508 (2015).
Nicolaou, S. A., Gaida, S. M. & Papoutsakis, E. T. A comparative view of metabolite and substrate stress and tolerance in microbial bioprocessing: from biofuels and chemicals, to biocatalysis and bioremediation. Metab. Eng. 12, 307–331 (2010).
Lam, F. H., Ghaderi, A., Fink, G. R. & Stephanopoulos, G. Engineering alcohol tolerance in yeast. Science 346, 71–75 (2014).
Cardinale, S., Tueros, F. G., Otto, M. & Sommer, A. Genetic-metabolic coupling for targeted metabolic engineering. CellReports 20, 1029–1037 (2017).
Wang, W. et al. Genome shuffling enhances stress tolerance of Zymomonas mobilis to two inhibitors. Biotechnol. Biofuels 12, 288 (2019).
Liu, R. et al. Directed combinatorial mutagenesis of Escherichia coli for complex phenotype engineering. Metab. Eng. 47, 10–20 (2018).
George, K. W. et al. Integrated analysis of isopentenyl pyrophosphate (IPP) toxicity in isoprenoid-producing Escherichia coli. Metab. Eng. 47, 60–72 (2018).
Kuroda, K. et al. Critical roles of the pentose phosphate pathway and GLN3 in isobutanol-specific tolerance in yeast. Cell Syst. https://doi.org/10.1016/j.cels.2019.10.006 (2019).
Pereira, R. et al. Adaptive laboratory evolution of tolerance to dicarboxylic acids in Saccharomyces cerevisiae. Metab. Eng. 56, 130–141 (2019).
Kildegaard, K. R. et al. Evolution reveals a glutathione-dependent mechanism of 3-hydroxypropionic acid tolerance. Metab. Eng. 26, 57–66 (2014).
Fletcher, E. et al. Evolutionary engineering reveals divergent paths when yeast is adapted to different acidic environments. Metab. Eng. 39, 19–28 (2017).
Matsusako, T., Toya, Y., Yoshikawa, K. & Shimizu, H. Identification of alcohol stress tolerance genes of Synechocystis sp. PCC 6803 using adaptive laboratory evolution. Biotechnol. Biofuels 10, 307 (2017).
Caspeta, L. et al. Altered sterol composition renders yeast thermotolerant. Science 346, 75–78 (2014).
Zhu, Z. et al. Multidimensional engineering of Saccharomyces cerevisiae for efficient synthesis of medium-chain fatty acids. Nat. Catal. 3, 64–74 (2020).
Lennen, R. M. & Pfleger, B. F. Modulating membrane composition alters free fatty acid tolerance in Escherichia coli. PLoS ONE 8, e54031 (2013).
Thorwall, S., Schwartz, C., Chartron, J. W. & Wheeldon, I. Stress-tolerant non-conventional microbes enable next-generation chemical biosynthesis. Nat. Chem. Biol. 16, 113–121 (2020). This perspective discusses bioprocesses that benefit from non-conventional microorganisms and existing challenges for their adoption.
Crosby, J. R. et al. Extreme thermophiles as emerging metabolic engineering platforms. Curr. Opin. Biotechnol. 59, 55–64 (2019).
Fu, X. Z. et al. Development of Halomonas TD01 as a host for open production of chemicals. Metab. Eng. 23, 78–91 (2014).
Preiss, L., Hicks, D. B., Suzuki, S., Meier, T. & Krulwich, T. A. Alkaliphilic bacteria with impact on industrial applications, concepts of early life forms, and bioenergetics of ATP synthesis. Front. Bioeng. Biotechnol. 3, 75 (2015).
Zhang, X., Lin, Y. & Chen, G.-Q. Halophiles as chassis for bioproduction. Adv. Biosyst. 2, 1800088 (2018).
Zheng, Y., Chen, J. C., Ma, Y. M. & Chen, G. Q. Engineering biosynthesis of polyhydroxyalkanoates (PHA) for diversity and cost reduction. Metab. Eng. https://doi.org/10.1016/j.ymben.2019.07.004 (2019).
Ma, H. et al. Rational flux-tuning of Halomonas bluephagenesis for co-production of bioplastic PHB and ectoine. Nat. Commun. 11, 1–12 (2020).
Zaky, A. S., Tucker, G. A., Daw, Z. Y. & Du, C. Marine yeast isolation and industrial application. FEMS Yeast Res. 14, 813–825 (2014).
Yue, H. et al. A seawater-based open and continuous process for polyhydroxyalkanoates production by recombinant Halomonas campaniensis LS21 grown in mixed substrates. Biotechnol. Biofuels 7, 108 (2014).
Zeldes, B. M. et al. Extremely thermophilic microorganisms as metabolic engineering platforms for production of fuels and industrial chemicals. Front. Microbiol. 6, 1209 (2015).
Elleuche, S., Schäfers, C., Blank, S., Schröder, C. & Antranikian, G. Exploration of extremophiles for high temperature biotechnological processes. Curr. Opin. Microbiol. 25, 113–119 (2015).
Lynd, L. R., Weimer, P. J., van Zyl, W. H. & Pretorius, I. S. Microbial cellulose utilization: fundamentals and biotechnology. Microbiol. Mol. Biol. Rev. 66, 506–577 (2002).
Shaw, A. J. et al. Metabolic engineering of a thermophilic bacterium to produce ethanol at high yield. Proc. Natl Acad. Sci. USA 105, 13769–13774 (2008).
Tian, L. et al. Metabolic engineering of Clostridium thermocellum for n-butanol production from cellulose. Biotechnol. Biofuels 12, 186 (2019).
Straub, C. T. et al. Quantitative fermentation of unpretreated transgenic poplar by Caldicellulosiruptor bescii. Nat. Commun. 10, 1–6 (2019).
Beri, D., York, W. S., Lynd, L. R., Peña, M. J. & Herring, C. D. Development of a thermophilic coculture for corn fiber conversion to ethanol. Nat. Commun. 11, 1–11 (2020).
Sharma, A., Kawarabayasi, Y. & Satyanarayana, T. Acidophilic bacteria and archaea: acid stable biocatalysts and their potential applications. Extremophiles 16, 1–19 (2012).
Dunbar, W. S. Biotechnology and the mine of tomorrow. Trends Biotechnol. 35, 79–89 (2017).
Gumulya, Y. et al. In a quest for engineering acidophiles for biomining applications: challenges and opportunities. Genes 9, 116 (2018).
Wernick, D. G., Pontrelli, S. P., Pollock, A. W. & Liao, J. C. Sustainable biorefining in wastewater by engineered extreme alkaliphile Bacillus marmarensis. Sci. Rep. 6, 1–10 (2016).
Chen, G. Q. & Jiang, X. R. Next generation industrial biotechnology based on extremophilic bacteria. Curr. Opin. Biotechnol. 50, 94–100 (2018).
Zoghlami, A. & Paës, G. Lignocellulosic biomass: understanding recalcitrance and predicting hydrolysis. Front. Chem. 7, 874 (2019).
Moreno, A. D., Carbone, A., Pavone, R., Olsson, L. & Geijer, C. Evolutionary engineered Candida intermedia exhibits improved xylose utilization and robustness to lignocellulose-derived inhibitors and ethanol. Appl. Microbiol. Biotechnol. 103, 1405–1416 (2019).
Ye, S., Kim, J. & Kim, S. R. Metabolic engineering for improved fermentation of L-arabinose. J. Microbiol. Biotechnol. 29, 339–346 (2019).
Liu, N., Santala, S. & Stephanopoulos, G. Mixed carbon substrates: a necessary nuisance or a missed opportunity? Curr. Opin. Biotechnol. 62, 15–21 (2020).
Nijland, J. G. et al. Engineering of an endogenous hexose transporter into a specific D-xylose transporter facilitates glucose-xylose co-consumption in Saccharomyces cerevisiae. Biotechnol. Biofuels 7, 168 (2014).
Fujiwara, R., Noda, S., Tanaka, T. & Kondo, A. Metabolic engineering of Escherichia coli for shikimate pathway derivative production from glucose–xylose co-substrate. Nat. Commun. 11, 1–12 (2020).
Papapetridis, I. et al. Laboratory evolution for forced glucose-xylose co-consumption enables identification of mutations that improve mixed-sugar fermentation by xylose-fermenting Saccharomyces cerevisiae. FEMS Yeast Res. 18, 56 (2018).
Verhoeven, M. D., de Valk, S. C., Daran, J.-M. G., van Maris, A. J. A. & Pronk, J. T. Fermentation of glucose-xylose-arabinose mixtures by a synthetic consortium of single-sugar-fermenting Saccharomyces cerevisiae strains. FEMS Yeast Res. https://doi.org/10.1093/femsyr/foy075 (2018).
Davis, K. & Moon, T. S. Tailoring microbes to upgrade lignin. Curr. Opin. Chem. Biol. 59, 23–29 (2020).
Niu, W. et al. Direct biosynthesis of adipic acid from lignin-derived aromatics using engineered Pseudomonas putida KT2440. Metab. Eng. 59, 151–161 (2020).
Araki, T. et al. Regulation of vanillate and syringate catabolism by a MarR-type transcriptional regulator DesR in Sphingobium sp. SYK-6. Sci. Rep. 9, 1–15 (2019).
Anthony, W. E. et al. Development of Rhodococcus opacus as a chassis for lignin valorization and bioproduction of high-value compounds. Biotechnol. Biofuels 12, 1–14 (2019).
Wu, W., Liu, F. & Singh, S. Toward engineering E. coli with an autoregulatory system for lignin valorization. Proc. Natl Acad. Sci. USA 115, 2970–2975 (2018).
Jönsson, L. J. & Martín, C. Pretreatment of lignocellulose: Formation of inhibitory by-products and strategies for minimizing their effects. Bioresour. Technol. 199, 103–112 (2016).
OECD. Meeting Policy Challenges for a Sustainable Bioeconomy (OECD, 2018).
Claassens, N. J., Sousa, D. Z., Dos Santos, V. A. P. M., De Vos, W. M. & Van Der Oost, J. Harnessing the power of microbial autotrophy. Nat. Rev. Microbiol. 14, 692–706 (2016). This article discusses recent advances in engineering microorganisms to increase the efficiency of autotrophic production platforms.
Liu, Z., Wang, K., Chen, Y., Tan, T. & Nielsen, J. Third-generation biorefineries as the means to produce fuels and chemicals from CO2. Nat. Catal. 3, 274–288 (2020).
Chistoserdova, L., Kalyuzhnaya, M. G. & Lidstrom, M. E. The expanding world of methylotrophic metabolism. Annu. Rev. Microbiol. 63, 477–499 (2009).
Humphreys, C. M. & Minton, N. P. Advances in metabolic engineering in the microbial production of fuels and chemicals from C1 gas. Curr. Opin. Biotechnol. 50, 174–181 (2018).
Meyer, F. et al. Methanol-essential growth of Escherichia coli. Nat. Commun. 9, 1–10 (2018).
Yu, H. & Liao, J. C. A modified serine cycle in Escherichia coli coverts methanol and CO2 to two-carbon compounds. Nat. Commun. 9, 3992 (2018).
Kim, S. et al. Growth of E. coli on formate and methanol via the reductive glycine pathway. Nat. Chem. Biol. https://doi.org/10.1038/s41589-020-0473-5 (2020).
Chen, F. Y. H., Jung, H. W., Tsuei, C. Y. & Liao, J. C. Converting Escherichia coli to a synthetic methylotroph growing solely on methanol. Cell 182, 933–946.e14 (2020). This is the first example of an engineered E. coli strain capable of growing in methanol as the sole carbon source.
Antonovsky, N. et al. Sugar synthesis from CO2 in Escherichia coli. Cell 166, 115–125 (2016).
Bang, J. & Lee, S. Y. Assimilation of formic acid and CO2 by engineered Escherichia coli equipped with reconstructed one-carbon assimilation pathways. Proc. Natl Acad. Sci. USA 115, E9271–E9279 (2018).
Gleizer, S. et al. Conversion of Escherichia coli to generate all biomass carbon from CO2. Cell 179, 1255–1263.e12 (2019). This is the first example of an engineered E. coli strain capable of growing in CO2 as the sole carbon source.
Gassler, T. et al. The industrial yeast Pichia pastoris is converted from a heterotroph into an autotroph capable of growth on CO2. Nat. Biotechnol. https://doi.org/10.1038/s41587-019-0363-0 (2019). This article provides a remarkable example of the conversion of the industrial yeast P. pastoris into an autotroph capable of growing in CO2 as the sole carbon source.
Novak, K. & Pfï, S. Towards biobased industry: acetate as a promising feedstock to enhance the potential of microbial cell factories. FEMS Microbiol. Lett. 365, 226 (2018).
Do, D. T. H., Theron, C. W. & Fickers, P. Organic wastes as feedstocks for non-conventional yeast-based bioprocesses. Microorganisms 7, 229 (2019).
Fei, Q. et al. The effect of volatile fatty acids as a sole carbon source on lipid accumulation by Cryptococcus albidus for biodiesel production. Bioresour. Technol. 102, 2695–2701 (2011).
Xu, J., Liu, N., Qiao, K., Vogg, S. & Stephanopoulos, G. Application of metabolic controls for the maximization of lipid production in semicontinuous fermentation. Proc. Natl Acad. Sci. USA 114, E5308–E5316 (2017).
Yang, J. & Nie, Q. Engineering Escherichia coli to convert acetic acid to β-caryophyllene. Microb. Cell Fact. 15, 74 (2016).
Wei, N., Quarterman, J., Kim, S. R., Cate, J. H. D. & Jin, Y. S. Enhanced biofuel production through coupled acetic acid and xylose consumption by engineered yeast. Nat. Commun. 4, 1–8 (2013).
Spagnuolo, M., Hussain, M. S., Gambill, L. & Blenner, M. Alternative substrate metabolism in Yarrowia lipolytica. Front. Microbiol. 9, 1077 (2018).
Huang, X. F. et al. Culture strategies for lipid production using acetic acid as sole carbon source by Rhodosporidium toruloides. Bioresour. Technol. 206, 141–149 (2016).
Gong, Z. et al. Efficient conversion of acetate into lipids by the oleaginous yeast Cryptococcus curvatus. Biotechnol. Biofuels 8, 189 (2015).
Katre, G., Ajmera, N., Zinjarde, S. & RaviKumar, A. Mutants of Yarrowia lipolytica NCIM 3589 grown on waste cooking oil as a biofactory for biodiesel production. Microb. Cell Fact. 16, 176 (2017).
Chen, Z. & Liu, D. Toward glycerol biorefinery: metabolic engineering for the production of biofuels and chemicals from glycerol. Biotechnol. Biofuels 9, 205 (2016).
Mano, J., Liu, N., Hammond, J. H., Currie, D. H. & Stephanopoulos, G. Engineering Yarrowia lipolytica for the utilization of acid whey. Metab. Eng. 57, 43–50 (2020).
Nielsen, J. It Is all about metabolic fluxes. J. Bacteriol. 185, 7031–7035 (2003).
Chen, X., Li, S. & Liu, L. Engineering redox balance through cofactor systems. Trends Biotechnol. 32, 337–343 (2014). This is a detailed review on the manipulation of cofactor systems to improve microbial redox balance.
Park, J. & Choi, Y. Cofactor engineering in cyanobacteria to overcome imbalance between NADPH and NADH: a mini review. Front. Chem. Sci. Eng. 11, 66–71 (2017).
Lim, J. H., Seo, S. W., Kim, S. Y. & Jung, G. Y. Model-driven rebalancing of the intracellular redox state for optimization of a heterologous n-butanol pathway in Escherichia coli. Metab. Eng. 20, 56–62 (2013).
Qiao, K., Wasylenko, T. M., Zhou, K., Xu, P. & Stephanopoulos, G. Lipid production in Yarrowia lipolytica is maximized by engineering cytosolic redox metabolism. Nat. Biotechnol. 35, 173–177 (2017).
Claassens, N. J., Sánchez-Andrea, I., Sousa, D. Z. & Bar-Even, A. Towards sustainable feedstocks: a guide to electron donors for microbial carbon fixation. Curr. Opin. Biotechnol. 50, 195–205 (2018). This article compares relevant electron donors to improve microbial CO2 fixation.
Karthikeyan, R., Singh, R. & Bose, A. Microbial electron uptake in microbial electrosynthesis: a mini-review. J. Ind. Microbiol. Biotechnol. 46, 1419–1426 (2019).
Gong, Z., Yu, H., Zhang, J., Li, F. & Song, H. Microbial electro-fermentation for synthesis of chemicals and biofuels driven by bi-directional extracellular electron transfer. Synth. Syst. Biotechnol. 5, 304–313 (2020).
Schievano, A. et al. Electro-fermentation – merging electrochemistry with fermentation in industrial applications. Trends Biotechnol. 34, 866–878 (2016).
Schuchmann, K. & Müller, V. Autotrophy at the thermodynamic limit of life: a model for energy conservation in acetogenic bacteria. Nat. Rev. Microbiol. 12, 809–821 (2014).
Lo, J. et al. Engineering electron metabolism to increase ethanol production in Clostridium thermocellum. Metab. Eng. 39, 71–79 (2017).
Liew, F. et al. Metabolic engineering of Clostridium autoethanogenum for selective alcohol production. Metab. Eng. 40, 104–114 (2017).
Hou, Y. et al. Metabolic engineering of cofactor flavin adenine dinucleotide (FAD) synthesis and regeneration in Escherichia coli for production of α-keto acids. Biotechnol. Bioeng. 114, 1928–1936 (2017).
Black, W. B. et al. Engineering a nicotinamide mononucleotide redox cofactor system for biocatalysis. Nat. Chem. Biol. 16, 87–94 (2020). This article describes the development and application of an orthogonal redox cofactor capable of supporting redox chemistries.
Hara, K. Y. & Kondo, A. ATP regulation in bioproduction. Microb. Cell Fact. 14, 198 (2015). This is a review of the role of ATP in bioproduction and strategies to alter its supply.
Meadows, A. L. et al. Rewriting yeast central carbon metabolism for industrial isoprenoid production. Nature 537, 694–697 (2016).
Man, Z. et al. Improvement of the intracellular environment for enhancing L-arginine production of Corynebacterium glutamicum by inactivation of H2O2-forming flavin reductases and optimization of ATP supply. Metab. Eng. 38, 310–321 (2016).
Wu, M. et al. Enhanced succinic acid production under acidic conditions by introduction of glutamate decarboxylase system in E. coli AFP111. Bioprocess. Biosyst. Eng. 40, 549–557 (2017).
Tao, S. et al. Regulation of ATP levels in Escherichia coli using CRISPR interference for enhanced pinocembrin production. Microb. Cell Fact. 17, 147 (2018).
Kok, S., Kozak, B. U., Pronk, J. T. & Maris, A. J. A. Energy coupling in Saccharomyces cerevisiae: selected opportunities for metabolic engineering. FEMS Yeast Res. 12, 387–397 (2012).
Zahoor, A., Messerschmidt, K., Boecker, S. & Klamt, S. ATPase-based implementation of enforced ATP wasting in Saccharomyces cerevisiae for improved ethanol production. Biotechnol. Biofuels 13, 185 (2020).
Hädicke, O., Bettenbrock, K. & Klamt, S. Enforced ATP futile cycling increases specific productivity and yield of anaerobic lactate production in Escherichia coli. Biotechnol. Bioeng. 112, 2195–2199 (2015).
Luo, Z., Zeng, W., Du, G., Chen, J. & Zhou, J. Enhanced pyruvate production in candida glabrata by engineering ATP futile cycle system. ACS Synth. Biol. 8, 787–795 (2019).
Lan, E. I. & Liao, J. C. ATP drives direct photosynthetic production of 1-butanol in cyanobacteria. Proc. Natl Acad. Sci. US. 109, 6018–6023 (2012).
Causey, T. B., Zhou, S., Shanmugam, K. T. & Ingram, L. O. Engineering the metabolism of Escherichia coli W3110 for the conversion of sugar to redox-neutral and oxidized products: Homoacetate production. Proc. Natl Acad. Sci. USA 100, 825–832 (2003).
Nielsen, J. & Keasling, J. D. Engineering cellular metabolism. Cell 164, 1185–1197 (2016).
Nielsen, J. Synthetic biology for engineering acetyl coenzyme a metabolism in yeast. mBio https://doi.org/10.1128/mBio.02153-14 (2014).
Zhang, S. et al. Metabolic engineering for efficient supply of acetyl-CoA from different carbon sources in Escherichia coli. Microb. Cell Fact. 18, 130 (2019).
Liu, H., Marsafari, M., Wang, F., Deng, L. & Xu, P. Engineering acetyl-CoA metabolic shortcut for eco-friendly production of polyketides triacetic acid lactone in Yarrowia lipolytica. Metab. Eng. 56, 60–68 (2019).
Hirokawa, Y., Kubo, T., Soma, Y., Saruta, F. & Hanai, T. Enhancement of acetyl-CoA flux for photosynthetic chemical production by pyruvate dehydrogenase complex overexpression in Synechococcus elongatus PCC 7942. Metab. Eng. 57, 23–30 (2020).
Liu, Q. et al. Rewiring carbon metabolism in yeast for high level production of aromatic chemicals. Nat. Commun. 10, 4976 (2019).
Burg, J. M. et al. Large-scale bioprocess competitiveness: the potential of dynamic metabolic control in two-stage fermentations. Curr. Opin. Chem. Eng. 14, 121–136 (2016).
Dinh, C. V. & Prather, K. L. Layered and multi-input autonomous dynamic control strategies for metabolic engineering. Curr. Opin. Biotechnol. 65, 156–162 (2020).
Shen, X., Wang, J., Li, C., Yuan, Q. & Yan, Y. Dynamic gene expression engineering as a tool in pathway engineering. Curr. Opin. Biotechnol. 59, 122–129 (2019).
Yu, T. et al. Reprogramming yeast metabolism from alcoholic fermentation to lipogenesis. Cell 174, 1549–1558.e14 (2018). This article describes a major reprogramming of yeast metabolism resulting in strains displaying high production levels of lipids.
Ruiz, B. et al. Production of microbial secondary metabolites: regulation by the carbon source. Crit. Rev. Microbiol. 36, 146–167 (2010).
Risdian, C., Mozef, T. & Wink, J. Biosynthesis of polyketides in streptomyces. Microorganisms 7, 124 (2019).
Keasling, J. D. Manufacturing molecules through metabolic engineering. Science 330, 1355–1358 (2010).
Paddon, C. J. et al. High-level semi-synthetic production of the potent antimalarial artemisinin. Nature 496, 528–532 (2013).
Galanie, S., Thodey, K., Trenchard, I. J., Interrante, M. F. & Smolke, C. D. Complete biosynthesis of opioids in yeast. Science 349, 1095–1100 (2015).
Srinivasan, P. & Smolke, C. D. Biosynthesis of medicinal tropane alkaloids in yeast. Nature https://doi.org/10.1038/s41586-020-2650-9 (2020). This article describes the production of tropane alkaloids in yeast, achieved by incorporating a metabolic pathway across six subcellular compartments.
Zhao, M. et al. Pathway engineering in yeast for synthesizing the complex polyketide bikaverin. Nat. Commun. 11, 1–10 (2020).
Milne, N. et al. Metabolic engineering of Saccharomyces cerevisiae for the de novo production of psilocybin and related tryptamine derivatives. Metab. Eng. 60, 25–36 (2020).
Komatsu, M., Uchiyama, T., Omura, S., Cane, D. E. & Ikeda, H. Genome-minimized Streptomyces host for the heterologous expression of secondary metabolism. Proc. Natl Acad. Sci. USA 107, 2646–2651 (2010).
Jiang, M., Zhang, H., Park, S. H., Li, Y. & Pfeifer, B. A. Deoxysugar pathway interchange for erythromycin analogues heterologously produced through Escherichia coli. Metab. Eng. 20, 92–100 (2013).
Nielsen, J. C. & Nielsen, J. Development of fungal cell factories for the production of secondary metabolites: Linking genomics and metabolism. Synth. Syst. Biotechnol. 2, 5–12 (2017).
Bond, C., Tang, Y. & Li, L. Saccharomyces cerevisiae as a tool for mining, studying and engineering fungal polyketide synthases. Fungal Genet. Biol. 89, 52–61 (2016).
Anyaogu, D. C. & Mortensen, U. H. Heterologous production of fungal secondary metabolites in aspergilli. Front. Microbiol. 6, 77 (2015).
Gomez-Escribano, J. P. & Bibb, M. J. Streptomyces coelicolor as an expression host for heterologous gene clusters. in Methods in Enzymology Vol. 517, 279–300 (Academic Press Inc., 2012).
Weber, T. et al. Metabolic engineering of antibiotic factories: New tools for antibiotic production in actinomycetes. Trends Biotechnol. 33, 15–26 (2015).
Palazzotto, E., Tong, Y., Lee, S. Y. & Weber, T. Synthetic biology and metabolic engineering of actinomycetes for natural product discovery. Biotechnol. Adv. 37, 107366 (2019).
Watve, M. G., Tickoo, R., Jog, M. M. & Bhole, B. D. How many antibiotics are produced by the genus Streptomyces? Arch. Microbiol. 176, 386–390 (2001).
Liu, R., Deng, Z. & Liu, T. Streptomyces species: ideal chassis for natural product discovery and overproduction. Metab. Eng. 50, 74–84 (2018).
Zabala, D., Braña, A. F., Flórez, A. B., Salas, J. A. & Méndez, C. Engineering precursor metabolite pools for increasing production of antitumor mithramycins in Streptomyces argillaceus. Metab. Eng. 20, 187–197 (2013).
Coze, F., Gilard, F., Tcherkez, G., Virolle, M.-J. & Guyonvarch, A. Carbon-flux distribution within streptomyces coelicolor metabolism: a comparison between the actinorhodin-producing strain M145 and its non-producing derivative M1146. PLoS ONE 8, e84151 (2013).
Wang, W. et al. Harnessing the intracellular triacylglycerols for titer improvement of polyketides in Streptomyces. Nat. Biotechnol. https://doi.org/10.1038/s41587-019-0335-4 (2019). This article describes a multi-omics approach to increase stationary phase production of polyketides in Streptomyces species.
Nielsen, J. C. et al. Comparative transcriptome analysis shows conserved metabolic regulation during production of secondary metabolites in filamentous fungi. mSystems https://doi.org/10.1128/mSystems.00012-19 (2019).
Kohlwein, S. D., Veenhuis, M. & van der Klei, I. J. Lipid droplets and peroxisomes: key players in cellular lipid homeostasis or a matter of fat-store’em up or burn’em down. Genetics 193, 1–50 (2013).
Liu, G. S. et al. The yeast peroxisome: A dynamic storage depot and subcellular factory for squalene overproduction. Metab. Eng. 57, 151–161 (2020).
Zhu, Z. et al. Enabling the synthesis of medium chain alkanes and 1-alkenes in yeast. Metab. Eng. 44, 81–88 (2017).
DeLoache, W. C., Russ, Z. N. & Dueber, J. E. Towards repurposing the yeast peroxisome for compartmentalizing heterologous metabolic pathways. Nat. Commun. 7, 1–11 (2016).
Duran, L., Montaño López, J. & Avalos, J. L. ¡Viva la mitochondria!: harnessing yeast mitochondria for chemical production. FEMS Yeast Res. https://doi.org/10.1093/femsyr/foaa037/5863938 (2020).
Avalos, J. L., Fink, G. R. & Stephanopoulos, G. Compartmentalization of metabolic pathways in yeast mitochondria improves the production of branched-chain alcohols. Nat. Biotechnol. 31, 335–341 (2013).
Yee, D. A. et al. Engineered mitochondrial production of monoterpenes in Saccharomyces cerevisiae. Metab. Eng. 55, 76–84 (2019).
Li, S., Liu, L. & Chen, J. Compartmentalizing metabolic pathway in Candida glabrata for acetoin production. Metab. Eng. 28, 1–7 (2015).
Xua, P., Qiao, K., Ahn, W. S. & Stephanopoulos, G. Engineering Yarrowia lipolytica as a platform for synthesis of drop-in transportation fuels and oleochemicals. Proc. Natl Acad. Sci. USA 113, 10848–10853 (2016).
Arendt, P. et al. An endoplasmic reticulum-engineered yeast platform for overproduction of triterpenoids. Metab. Eng. 40, 165–175 (2017).
Kim, J. E. et al. Tailoring the Saccharomyces cerevisiae endoplasmic reticulum for functional assembly of terpene synthesis pathway. Metab. Eng. 56, 50–59 (2019).
Yang, K. et al. Subcellular engineering of lipase dependent pathways directed towards lipid related organelles for highly effectively compartmentalized biosynthesis of triacylglycerol derived products in Yarrowia lipolytica. Metab. Eng. 55, 231–238 (2019).
Zhang, Y. et al. Xylose utilization stimulates mitochondrial production of isobutanol and 2-methyl-1-butanol in Saccharomyces cerevisiae. Biotechnol. Biofuels 12, 223 (2019).
Grewal, P. S., Samson, J. A., Baker, J. J., Choi, B. & Dueber, J. E. Peroxisome compartmentalization of a toxic enzyme improves alkaloid production. Nat. Chem. Biol. 17, 96–103 (2021).
Kerfeld, C. A., Aussignargues, C., Zarzycki, J., Cai, F. & Sutter, M. Bacterial microcompartments. Nat. Rev. Microbiol. 16, 277–290 (2018). This is a review of the composition, assembly and applications of bacterial microcompartments.
Kerfeld, C. A. & Sutter, M. Engineered bacterial microcompartments: apps for programming metabolism. Curr. Opin. Biotechnol. 65, 225–232 (2020).
Lawrence, A. D. et al. Solution structure of a bacterial microcompartment targeting peptide and its application in the construction of an ethanol bioreactor. ACS Synth. Biol. 3, 454–465 (2014).
Flamholz, A. I. et al. Functional reconstitution of a bacterial CO2 concentrating mechanism in e. Coli. eLife 9, 1–57 (2020).
Nichols, T. M., Kennedy, N. W. & Tullman-Ercek, D. Cargo encapsulation in bacterial microcompartments: Methods and analysis. in Methods in Enzymology Vol. 617, 155–186 (Academic Press Inc., 2019).
Kalnins, G. et al. Encapsulation mechanisms and structural studies of GRM2 bacterial microcompartment particles. Nat. Commun. 11, 1–13 (2020).
Hagen, A., Sutter, M., Sloan, N. & Kerfeld, C. A. Programmed loading and rapid purification of engineered bacterial microcompartment shells. Nat. Commun. 9, 1–10 (2018).
Wei, S. P. et al. Formation and functionalization of membraneless compartments in Escherichia coli. Nat. Chem. Biol. https://doi.org/10.1038/s41589-020-0579-9 (2020).
Zhao, E. M. et al. Light-based control of metabolic flux through assembly of synthetic organelles. Nat. Chem. Biol. 15, 589–597 (2019).
Apel, A. R., Ouellet, M., Szmidt-Middleton, H., Keasling, J. D. & Mukhopadhyay, A. Evolved hexose transporter enhances xylose uptake and glucose/xylose co-utilization in Saccharomyces cerevisiae. Sci. Rep. 6, 19512 (2016).
Wang, C. et al. Identification of important amino acids in Gal2p for improving the L-arabinose transport and metabolism in saccharomyces cerevisiae. Front. Microbiol. 8, 1391 (2017).
Hu, M. L. et al. Enhanced bioconversion of cellobiose by industrial Saccharomyces cerevisiae used for cellulose utilization. Front. Microbiol. 7, 241 (2016).
de Sales, B. B. et al. Cloning novel sugar transporters from Scheffersomyces (Pichia) stipitis allowing d-xylose fermentation by recombinant Saccharomyces cerevisiae. Biotechnol. Lett. 37, 1973–1982 (2015).
Leandro, M. J., Gonçalves, P. & Spencer-Martins, I. Two glucose/xylose transporter genes from the yeast Candida intermedia: first molecular characterization of a yeast xylose-H+ symporter. Biochem. J. 395, 543–549 (2006).
Li, J. et al. Functional analysis of two L-arabinose transporters from filamentous fungi reveals promising characteristics for improved pentose utilization in Saccharomyces cerevisiae. Appl. Environ. Microbiol. 81, 4062–4070 (2015).
Galazka, J. M. et al. Cellodextrin transport in yeast for improved biofuel production. Sci. 330, 84–86 (2010).
Young, E. M., Comer, A. D., Huang, H. & Alper, H. S. A molecular transporter engineering approach to improving xylose catabolism in Saccharomyces cerevisiae. Metab. Eng. 14, 401–411 (2012).
Hara, K. Y. et al. Transporter engineering in biomass utilization by yeast. FEMS Yeast Res. https://doi.org/10.1093/femsyr/fox061 (2017).
Park, S. H., Kim, S. & Hahn, J. S. Improvement of isobutanol production in Saccharomyces cerevisiae by increasing mitochondrial import of pyruvate through mitochondrial pyruvate carrier. Appl. Microbiol. Biotechnol. 100, 7591–7598 (2016).
Steiger, M. G., Rassinger, A., Mattanovich, D. & Sauer, M. Engineering of the citrate exporter protein enables high citric acid production in Aspergillus niger. Metab. Eng. 52, 224–231 (2019).
Darbani, B., Stovicek, V., Van Der Hoek, S. A. & Borodina, I. Engineering energetically efficient transport of dicarboxylic acids in yeast Saccharomyces cerevisiae. Proc. Natl Acad. Sci. USA 116, 19415–19420 (2019).
Pérez-García, F. & Wendisch, V. F. Transport and metabolic engineering of the cell factory Corynebacterium glutamicum. FEMS Microbiol. Lett. https://doi.org/10.1093/femsle/fny166 (2018).
Wang, G. et al. Transportome-wide engineering of Saccharomyces cerevisiae. Metab. Eng. 64, 52–63 (2021).
Hadadi, N. & Hatzimanikatis, V. Design of computational retrobiosynthesis tools for the design of de novo synthetic pathways. Curr. Opin. Chem. Biol. 28, 99–104 (2015).
Lin, G. M., Warden-Rothman, R. & Voigt, C. A. Retrosynthetic design of metabolic pathways to chemicals not found in nature. Curr. Opin. Syst. Biol. 14, 82–107 (2019).
Jones, D. T. & Woods, D. R. Acetone-butanol fermentation revisited. Microbiol. Mol. Biol. Rev. 50, 484–524 (1986).
Tracy, B. P., Jones, S. W., Fast, A. G., Indurthi, D. C. & Papoutsakis, E. T. Clostridia: the importance of their exceptional substrate and metabolite diversity for biofuel and biorefinery applications. Curr. Opin. Biotechnol. 23, 364–381 (2012).
Li, S., Huang, L., Ke, C., Pang, Z. & Liu, L. Pathway dissection, regulation, engineering and application: Lessons learned from biobutanol production by solventogenic clostridia. Biotechnol. Biofuels 13, 1–25 (2020).
Wan, N., Sathish, A., You, L., Tang, Y. J. & Wen, Z. Deciphering Clostridium metabolism and its responses to bioreactor mass transfer during syngas fermentation. Sci. Rep. 7, 1–11 (2017).
Zu, T. N. K., Liu, S., Gerlach, E. S., Mojadedi, W. & Sund, C. J. Co-feeding glucose with either gluconate or galacturonate during clostridial fermentations provides metabolic fine-tuning capabilities. Sci. Rep. 11, 29 (2021).
Jones, S. W. et al. CO2 fixation by anaerobic non-photosynthetic mixotrophy for improved carbon conversion. Nat. Commun. 7, 1–9 (2016).
Nguyen, N. P. T., Raynaud, C., Meynial-Salles, I. & Soucaille, P. Reviving the Weizmann process for commercial n-butanol production. Nat. Commun. 9, 3682 (2018).
Senger, R. S. & Papoutsakis, E. T. Genome-scale model for Clostridium acetobutylicum: Part I. Metabolic network resolution and analysis. Biotechnol. Bioeng. 101, 1036–1052 (2008).
Mahamkali, V. et al. Redox controls metabolic robustness in the gas-fermenting acetogen Clostridium autoethanogenum. Proc. Natl Acad. Sci. USA 117, 13168–13175 (2020).
Haas, T., Krause, R., Weber, R., Demler, M. & Schmid, G. Technical photosynthesis involving CO2 electrolysis and fermentation. Nat. Catal. 1, 32–39 (2018).
Cheng, C., Bao, T. & Yang, S. T. Engineering Clostridium for improved solvent production: recent progress and perspective. Appl. Microbiol. Biotechnol. 103, 5549–5566 (2019).
Charubin, K., Bennett, R. K., Fast, A. G. & Papoutsakis, E. T. Engineering Clostridium organisms as microbial cell-factories: challenges & opportunities. Metab. Eng. 50, 173–191 (2018).
Claassens, N. J. A warm welcome for alternative CO2 fixation pathways in microbial biotechnology. Microb. Biotechnol. 10, 31–34 (2017).
Bar-Even, A., Noor, E., Lewis, N. E. & Milo, R. Design and analysis of synthetic carbon fixation pathways. Proc. Natl Acad. Sci. USA 107, 8889–8894 (2010).
Schwander, T., Von Borzyskowski, L. S., Burgener, S., Cortina, N. S. & Erb, T. J. A synthetic pathway for the fixation of carbon dioxide in vitro. Science 354, 900–904 (2016).
Satanowski, A. et al. Awakening a latent carbon fixation cycle in Escherichia coli. Nat. Commun. 11, 1–14 (2020).
He, H., Höper, R., Dodenhöft, M., Marlière, P. & Bar-Even, A. An optimized methanol assimilation pathway relying on promiscuous formaldehyde-condensing aldolases in E. coli. Metab. Eng. 60, 1–13 (2020).
Sánchez-Andrea, I. et al. The reductive glycine pathway allows autotrophic growth of Desulfovibrio desulfuricans. Nat. Commun. 11, 1–12 (2020).
Lu, X. et al. Constructing a synthetic pathway for acetyl-coenzyme A from one-carbon through enzyme design. Nat. Commun. 10, 1378 (2019).
Bogorad, I. W. et al. Building carbon-carbon bonds using a biocatalytic methanol condensation cycle. Proc. Natl Acad. Sci. USA 111, 15928–15933 (2014).
Siegel, J. B. et al. Computational protein design enables a novel one-carbon assimilation pathway. Proc. Natl Acad. Sci. USA 112, 3704–3709 (2015).
Chou, A., Clomburg, J. M., Qian, S. & Gonzalez, R. 2-Hydroxyacyl-CoA lyase catalyzes acyloin condensation for one-carbon bioconversion. Nat. Chem. Biol. 15, 900–906 (2019).
Claassens, N. J. et al. Replacing the Calvin cycle with the reductive glycine pathway in Cupriavidus necator. Metab. Eng. 62, 30–41 (2020).
Wang, Y., Fan, L., Tuyishime, P., Zheng, P. & Sun, J. Synthetic methylotrophy: a practical solution for methanol-based biomanufacturing. Trends Biotechnol. https://doi.org/10.1016/j.tibtech.2019.12.013 (2020).
Cotton, C. A., Claassens, N. J., Benito-Vaquerizo, S. & Bar-Even, A. Renewable methanol and formate as microbial feedstocks. Curr. Opin. Biotechnol. 62, 168–180 (2020). This is a review of the use of formate and methanol as microbial feedstocks.
Work in the laboratory of J.L.A. is supported by the US Department of Energy, Office of Science, Office of Biological and Environmental Research Genomic Science Program under award number DE-SC0019363; the US Department of Defense, Air Force Office of Scientific Research (MURI) under award number FA9550-20-1-0241; the NSF CAREER program under award CBET-1751840; The Pew Charitable Trusts; and The Henry & Camille Dreyfus Foundation.
The authors declare no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
An economy based on the use of biological renewable sources to produce value-added products, such as food, chemicals, materials and energy.
- Adaptive laboratory evolution
(ALE). The continuous culturing of an organism under constant selection pressure, yielding microbial strains with improved phenotypes.
- Genome shuffling
Methods that enable the recombination of genomes, generating populations with genetic diversity, in which some strains might possess beneficial traits.
- Catabolite repression
Regulatory mechanisms that inhibit the use of secondary nutrient sources when a preferred one is present.
Devices, apparatuses or systems for growing organisms under controlled conditions (for example, temperature, pH, oxygen and nutrient supply) for the synthesis of desired products.
Organisms that thrive in high salt concentrations.
Organisms that thrive at high temperatures (typically above 45 °C)
Organisms that can optimally grow at pH 3 or lower.
Organisms that can optimally grow at pH 9 or higher.
Organisms that can synthesize organic molecules from CO2 using energy and electrons derived from chemicals.
Organisms that can synthesize organic molecules from CO2 using light as an energy source and split water as a source of electrons.
Organisms that utilize reduced carbon substrates with no carbon–carbon bonds (for example, methane, methanol and formate) as their sole carbon and energy sources.
Organisms that derive their own biomass from organic matter produced by other organisms.
Organisms that use both CO2 and organic carbon sources to generate their biomass.
- Primary metabolism
Set of enzymatic reactions involved in pathways necessary for growth and development, typically active when nutrients are present in the medium. Primary metabolites, which are essential to sustain cell growth, are synthesized by primary metabolism.
- Secondary metabolism
Set of biosynthetic pathways that confer survival advantages on the organism but are not essential for cell growth. Secondary metabolism is typically active during the stationary phase in some microorganisms. Many bioactive compounds, referred to as ‘secondary metabolites’ or ‘natural products’ are synthesized by secondary metabolism.
About this article
Cite this article
Montaño López, J., Duran, L. & Avalos, J.L. Physiological limitations and opportunities in microbial metabolic engineering. Nat Rev Microbiol (2021). https://doi.org/10.1038/s41579-021-00600-0