We describe an iterative approach for creating protein switches involving the in vitro recombination of two nonhomologous genes. We demonstrate this approach by recombining the genes coding for TEM1 ␤-lactamase (BLA) and the Escherichia coli maltose binding protein (MBP) to create a family of MBP-BLA hybrids in which maltose is a positive or negative effector of ␤-lactam hydrolysis. Some of these MBP-BLA switches were effectively ''on-off'' in nature, with maltose altering catalytic activity by as much as 600-fold. The ability of these switches to confer an effector-dependent growth͞no growth phenotype to E. coli cells was exploited to rapidly identify, from a library of 4 ؋ 10 6 variants, MBP-BLA switch variants that respond to sucrose as the effector. The transplantation of these mutations into wild-type MBP converted MBP into a ''sucrose-binding protein,'' illustrating the switches potential as a tool to rapidly identify ligand-binding proteins.allostery ͉ ␤-lactamase ͉ maltose binding protein R egulation of protein activity is fundamental to cellular function.One of the mechanisms a cell uses to modulate the level of protein activity is regulation of the amount of a protein present in a cell. Accordingly, many strategies for engineering control of cellular protein activity have focused on modulation of protein production and degradation, often through small-moleculedependent switches that regulate transcription, translation, localization, degradation, or protein splicing (1) or through the engineering of artificial gene-regulatory networks (2). The key strength of many of these approaches is that they are easily generalized. For example, a switch that regulates transcription of one gene can easily be adapted to regulate transcription of an arbitrary gene. However, the significant limitation of these approaches is the slow dynamics stemming from the indirect nature of the regulation (i.e., protein activity is regulated by controlling the amount of protein and not by regulating the protein's specific activity directly). In addition, modulation is only feasible in the context of the cell and cannot be easily transferred to an in vitro setting.A more satisfying approach is to modulate the protein activity directly, but this presents a considerable design problem. Inhibitors, if they can be found, can only can be used to down-regulate activity, and such a strategy suffers from the fact that one is not free to choose the signal that modulates the activity: The signal must be an inhibitor of the protein. One clever way around this limitation is to engineer the inhibitor such that a third molecule can regulate the inhibitor's affinity for the regulated protein, as was demonstrated for RNA aptamer inhibitors that could be regulated by an organic small molecule (3). Natural allosteric proteins solve this problem by having spatially distinct regulatory and active sites. Ligand binding or covalent modifications at one site affects the output function at a distant site through a conformational change. This mechanism has ...

Current methods for producing immunoglobulin G (IgG) antibodies in engineered cells often require refolding steps or secretion across one or more biological membranes. Here, we describe a robust expression platform for biosynthesis of full-length IgG antibodies in the Escherichia coli cytoplasm. Synthetic heavy and light chains, both lacking canonical export signals, are expressed in specially engineered E. coli strains that permit formation of stable disulfide bonds within the cytoplasm. IgGs with clinically relevant antigen- and effector-binding activities are readily produced in the E. coli cytoplasm by grafting antigen-specific variable heavy and light domains into a cytoplasmically stable framework and remodelling the fragment crystallizable domain with amino-acid substitutions that promote binding to Fcγ receptors. The resulting cytoplasmic IgGs—named ‘cyclonals'—effectively bypass the potentially rate-limiting steps of membrane translocation and glycosylation.

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The human gut is an ecosystem comprising trillions of microbes interacting with the host. The composition of the microbiota and their interactions play roles in different biological processes and in the development of human diseases. Close relationships between dietary modifications, microbiota composition and health status have been established. This review focuses on prebiotics, or compounds which selectively encourage the growth of beneficial bacteria, their mechanisms of action and benefits to human hosts. We also review advances in synthesis technology for human milk oligosaccharides, part of one of the most well-characterized prebiotic–probiotic relationships. Current and future research in this area points to greater use of prebiotics as tools to manipulate the microbial and metabolic diversity of the gut for the benefit of human health.

Multiplexed genome engineering approaches can be used to generate targeted genetic diversity in cell populations on laboratory timescales, but methods to track mutations and link them to phenotypes have been lacking. We present an approach for tracking combinatorial engineered libraries (TRACE) through the simultaneous mapping of millions of combinatorially engineered genomes at single-cell resolution. Distal genomic sites are assembled into individual DNA constructs that are compatible with next-generation sequencing strategies. We used TRACE to map growth selection dynamics for Escherichia coli combinatorial libraries created by recursive multiplex recombineering at a depth 10(4)-fold greater than before. TRACE was used to identify genotype-to-phenotype correlations and to map the evolutionary trajectory of two individual combinatorial mutants in E. coli. Combinatorial mutations in the human ES2 ovarian carcinoma cell line were also assessed with TRACE. TRACE completes the combinatorial engineering cycle and enables more sophisticated approaches to genome engineering in both bacteria and eukaryotic cells than are currently possible.

A heterotropic allosteric effect involves an effector molecule that is distinct from the substrate or ligand of the protein. How heterotropic allostery originates is an unanswered question. We have previously created several heterotropic allosteric enzymes by recombining the genes for TEM1 b-lactamase (BLA) and maltose binding protein (MBP) to create BLAs that are positively or negatively regulated by maltose. We show here that one of these engineered enzymes has ;10 6 M À1 affinity for Zn 2+, a property that neither of the parental proteins possesses. Furthermore, Zn 2+ is a negative effector that noncompetitively switches off b-lactam hydrolysis activity. Mutagenesis experiments indicate that the Zn 2+ -binding site does not involve a histidine or a cysteine, which is atypical of natural Zn 2+ -binding sites. These studies also implicate helices 1 and 12 of the BLA domain in allosteric signal propagation. These results support a model for the evolution of heterotropic allostery in which effector affinity and allosteric signaling emerge simultaneously.

In this study, we present a minimal template design and accompanying methods to produce assayable quantities of custom sequence proteins within 24 hr from receipt of inexpensive gene fragments from a DNA synthesis vendor. This is done without the conventional steps of plasmid cloning or cell‐based amplification and expression. Instead the linear template is PCR amplified, circularized, and isothermally amplified using a rolling circle polymerase. The resulting template can be used directly with cost‐optimized, scalably‐manufactured Escherichia coli extract and minimal supplement reagents to perform cell‐free protein synthesis (CFPS) of the template protein. We demonstrate the utility of this template design and 24 hr process with seven fluorescent proteins (sfGFP, mVenus, mCherry, and four GFP variants), three enzymes (chloramphenicol acetyltransferase, a chitinase catalytic domain, and native subtilisin), a capture protein (anti‐GFP nanobody), and 2 antimicrobial peptides (BP100 and CA(1–7)M(2–9)). We detected each of these directly from the CFPS reaction using colorimetric, fluorogenic, and growth assays. Of especial note, the GFP variant sequences were found from genomic screening data and had not been expressed or characterized before, thus demonstrating the utility of this approach for phenotype characterization of sequenced libraries. We also demonstrate that the rolling circle amplified version of the linear template exhibits expression similar to that of a complete plasmid when expressing sfGFP in the CFPS reaction. We evaluate the cost of this approach to be $61/mg sfGFP for a 4 hr reaction. We also detail limitations of this approach and strategies to overcome these, namely proteins with posttranslational modifications.

An array of genetic screens and selections has been developed for reporting protein folding and solubility in the cytoplasm of living cells. However, there are currently no analogous folding assays for the bacterial periplasm, despite the significance of this compartment for the expression of recombinant proteins, especially those requiring important posttranslational modifications (e.g., disulfide bond formation). Here, we describe an engineered genetic selection for monitoring protein folding in the periplasmic compartment of Escherichia coli cells. In this approach, target proteins are sandwiched between an N-terminal signal recognition particle (SRP)-dependent signal peptide and a C-terminal selectable marker, TEM-1 b-lactamase. The resulting chimeras are localized to the periplasmic space via the cotranslational SRP pathway. Using a panel of native and heterologous proteins, we demonstrate that the folding efficiency of various target proteins correlates directly with in vivo b-lactamase activity and thus resistance to ampicillin. We also show that this reporter is useful for the discovery of extrinsic periplasmic factors (e.g., chaperones) that affect protein folding and for obtaining folding-enhanced proteins via directed evolution. Collectively, these data demonstrate that our periplasmic folding reporter is a powerful tool for screening and engineering protein folding in a manner that does not require any structural or functional information about the target protein.