生物工程

Chloroplast genomes present an alternative strategy for large-scale engineering of photosynthetic eukaryotes. Prior to our work, the chloroplast genomes of Chlamydomonas reinhardtii (204 kb) and Zea mays (140 kb) had been cloned using bacterial and yeast artificial chromosome (BAC/YAC) libraries, respectively. These methods lack design flexibility as they are reliant upon the random capture of genomic fragments during BAC/YAC library creation; additionally, both demonstrated a low efficiency (≤ 10%) for correct assembly of the genome in yeast. With this in mind, we sought to create a highly flexible and efficient approach for assembling the 117 kb chloroplast genome of Phaeodactylum tricornutum, a photosynthetic marine diatom. Our original article demonstrated a PCR-based approach for cloning the P. tricornutum chloroplast genome that had 90%–100% efficiency when screening as few as 10 yeast colonies following assembly. In this article, we will discuss this approach in greater depth as we believe this technique could be extrapolated to other species, particularly those with a similar chloroplast genome size and architecture.

Droplet microfluidic platforms have been broadly used to facilitate DNA transfer in mammalian and bacterial hosts via methods such as transformation, transfection, and conjugation, as introduced in our previous work. Herein, we recapitulate our method for conjugal DNA transfer between Bacillus subtilis strains in a droplet for increased conjugation efficiency and throughput of an otherwise laborious protocol. By co-incubating the donor and recipient strains in droplets, our method confines cells into close proximity allowing for increased cell-to-cell interactions. This methodology is advantageous in its potential to automate and accelerate the genetic modification of undomesticated organisms that may be difficult to cultivate. This device is also designed for modularity and can be integrated into a variety of experimental workflows in which fine-tuning of donor-to-recipient cell ratios, growth rates, and media substrate concentrations may be necessary.

Generating protein conjugates using the bioorthogonal ligation between tetrazines and trans-cyclooctene groups avoids the need to manipulate cysteine amino acids; this ligation is rapid, site-specific, and stoichiometric and allows for labeling of proteins in complex biological environments. Here, we provide a protocol for the expression of conjugation-ready proteins at high yields in Escherichia coli with greater than 95% encoding and labeling fidelity. This protocol focuses on installing the Tet2 tetrazine amino acid using an optimized genetic code expansion (GCE) machinery system, Tet2 pAJE-E7, to direct Tet2 encoding at TAG stop codons in BL21 E. coli strains, enabling reproducible expression of Tet2-proteins that quantitatively react with trans-cyclooctene (TCO) groups within 5 min at room temperature and physiological pH. The use of the BL21 derivative B95(DE3) minimizes premature truncation byproducts caused by incomplete suppression of TAG stop codons, which makes it possible to use more diverse protein construct designs. Here, using a superfolder green fluorescent protein construct as an example protein, we describe in detail a four-day process for encoding Tet2 with yields of ~200 mg per liter of culture. Additionally, a simple and fast diagnostic gel electrophoretic mobility shift assay is described to confirm Tet2-Et encoding and reactivity. Finally, strategies are discussed to adapt the protocol to alternative proteins of interest and optimize expression yields and reactivity for that protein.

Diatoms serve as a source for a variety of compounds with particularbiotechnological interest. Therefore, redirecting the flow to a specific pathwayrequires the elucidation of the gene’s specific function. The mostcommonly used method in diatoms is biolistic transformation, which is a veryexpensive and time-consuming method. The use of episomes that are maintained asclosed circles at a copy number equivalent to native chromosomes has become auseful genetic system for protein expression that avoids multiple insertions,position-specific effects on expression, and potential knockout of non-targetedgenes. These episomes can be introduced from bacteria into diatoms viaconjugation. Here, we describe a detailed protocol for gene expression thatincludes 1) the gateway cloning strategy and 2) the conjugation protocol for themobilization of plasmids from bacteria to diatoms.

While site-specific translational encoding of phosphoserine (pSer) into proteins in Escherichia coli via genetic code expansion (GCE) technologies has transformed our ability to study phospho-protein structure and function, recombinant phospho-proteins can be dephosphorylated during expression/purification, and their exposure to cellular-like environments such as cell lysates results in rapid reversion back to the non-phosphorylated form. To help overcome these challenges, we developed an efficient and scalable E. coli GCE expression system enabling site-specific incorporation of a non-hydrolyzable phosphoserine (nhpSer) mimic into proteins of interest. This nhpSer mimic, with the γ-oxygen of phosphoserine replaced by a methylene (CH₂) group, is impervious to hydrolysis and recapitulates phosphoserine function even when phosphomimetics aspartate and glutamate do not. Key to this expression system is the co-expression of a Streptomyces biosynthetic pathway that converts the central metabolite phosphoenolpyruvate into non-hydrolyzable phosphoserine (nhpSer) amino acid, which provides a > 40-fold improvement in expression yields compared to media supplementation by increasing bioavailability of nhpSer and enables scalability of expressions. This “PermaPhos” expression system uses the E. coli BL21(DE3) ∆serC strain and three plasmids that express (i) the protein of interest, (ii) the GCE machinery for translational installation of nhpSer at UAG amber stop codons, and (iii) the Streptomyces nhpSer biosynthetic pathway. Successful expression requires efficient transformation of all three plasmids simultaneously into the expression host, and IPTG is used to induce expression of all components. Permanently phosphorylated proteins made in E. coli are particularly useful for discovering phosphorylation-dependent protein–protein interaction networks from cell lysates or transfected cells.

Key features

• Protocol builds on the nhpSer GCE system by Rogerson et al. (2015), but with a > 40-fold improvement in yields enabled by the nhpSer biosynthetic pathway.

• Protein expression uses standard Terrific Broth (TB) media and requires three days to complete.

• C-terminal purification tags on target protein are recommended to avoid co-purification of prematurely truncated protein with full-length nhpSer-containing protein.

• Phos-tag gel electrophoresis provides a convenient method to confirm accurate nhpSer encoding, as it can distinguish between non-phosphorylated, pSer- and nhpSer-containing variants.

Graphical overview

Reconstitution of membrane proteins into large unilamellar vesicles is an essential approach for their functional analysis under chemically defined conditions. The orientation of the protein in the liposomal membrane after reconstitution depends on many parameters, and its assessment is important prior to functional measurements. Common approaches for determining the orientation of a membrane-inserted protein are based on limited proteolytic digest, impermeable labeling reagents for specific amino acids, or membrane-impermeable quenchers for fluorescent proteins. Here, we describe a simple site-specific fluorescent assay based on self-labeling enzyme tags to determine the orientation of membrane proteins after reconstitution, exemplified on a reconstituted SNAP-tag plant H⁺-ATPase. This versatile method should benefit the optimization of reconstitution conditions and the analysis of many types of membrane proteins.

Graphical abstract:

This protocol describes the recombinant expression of proteins in E. coli containing phosphoserine (pSer) installed at positions guided by TAG codons. The E. coli strains that can be used here are engineered with a ∆serB genomic knockout to produce pSer internally at high levels, so no exogenously added pSer is required, and the addition of pSer to the media will not affect expression yields. For “truncation-free” expression and improved yields with high flexibility of construct design, it is preferred to use the Release Factor-1 (RF1) deficient strain B95(DE3) ∆A ∆fabR ∆serB, though use of the standard RF1-containing BL21(DE3) ∆serB is also described. Both of these strains are serine auxotrophs and will not grow in standard minimal media. This protocol uses rich auto-induction media for streamlined and maximal production of homogeneously modified protein, yielding ~100–200 mg of single pSer-containing sfGFP per liter of culture. Using this genetic code expansion (GCE) approach, in which pSer is installed into proteins during translation, allows researchers to produce milligram quantities of specific phospho-proteins without requiring kinases, which can be purified for downstream in vitro studies relating to phosphorylation-dependent signaling systems, protein regulation by phosphorylation, and protein–protein interactions.

Graphical abstract:

Directed evolution is a powerful technique for identifying beneficial mutations in defined DNA sequences with the goal of improving desired phenotypes. Recent methodological advances have made the evolution of short DNA sequences quick and easy. However, the evolution of DNA sequences >5kb in length, notably gene clusters, is still a challenge for most existing methods. Since many important microbial phenotypes are encoded by multigene pathways, they are usually improved via adaptive laboratory evolution (ALE), which while straightforward to implement can suffer from off-target and hitchhiker mutations that can adversely affect the fitness of the evolved strain. We have therefore developed a new directed evolution method (Inducible Directed Evolution, IDE) that combines the specificity and throughput of recent continuous directed evolution methods with the ease of ALE. Here, we present detailed methods for operating Inducible Directed Evolution (IDE), which enables long (up to 85kb) DNA sequences to be mutated in a high throughput manner via a simple series of incubation steps. In IDE, an intracellular mutagenesis plasmid (MP) tunably mutagenizes the pathway of interest, located on the phagemid (PM). MP contains a mutagenic operon (danQ926, dam, seqA, emrR, ugi, and cda1) that can be expressed via the addition of a chemical inducer. Expression of the mutagenic operon during a cell cycle represses DNA repair mechanisms such as proofreading, translesion synthesis, mismatch repair, and base excision and selection, which leads to a higher mutation rate. Induction of the P1 lytic cycle results in packaging of the mutagenized phagemid, and the pathway-bearing phage particles infect naïve cells, generating a mutant library that can be screened or selected for improved variants. Successive rounds of IDE enable optimization of complex phenotypes encoded by large pathways (as of this writing up to 36 kb), without requiring inefficient transformation steps. Additionally, IDE avoids off-target genomic mutations and enables decoupling of mutagenesis and screening steps, establishing it as a powerful tool for optimizing complex phenotypes in E. coli.

Graphical abstract:

Figure 1. Overview of Inducible Directed Evolution (IDE). Pathways of interest are cloned into a P1 phagemid (PM) backbone and transformed into a strain of E. coli containing MP (diversification strain). The mutagenesis plasmid is induced to generate mutations. Phage lysate is produced and used to infect a strain that expresses the phenotype of interest (screening/selection strain). The resulting strain library is screened to identify those with improved properties. Narrowed-down libraries can then go through another IDE cycle by infecting a fresh diversification strain.

The incorporation of non-standard amino acids (nsAAs) within proteins and peptides through genetic code expansion introduces novel chemical functionalities such as photo-crosslinking and bioconjugation. Given the utility of Bacillus subtilis in fundamental and applied science, we extended existing nsAA incorporation technology from Escherichia coli into B. subtilis, demonstrating incorporation of 20 unique nsAAs. The nsAAs we succeeded in incorporating within proteins conferred properties that included fluorescence, photo-crosslinking, and metal chelation. Here, we describe the reagents, equipment, and protocols to test for nsAA incorporation at a small scale (96-well plate and culture tube scales). We report specific media requirements for certain nsAAs, including two variants for different media conditions. Our protocol provides a consistent and reproducible method for incorporation of a chemically diverse set of nsAAs into a model Gram-positive organism.

Live labelling of active transcription sites is critical to our understanding of transcriptional dynamics. In the most widely used method, RNA sequence MS2 repeats are added to the transcript of interest, on which fluorescently tagged Major Coat Protein binds, and labels transcription sites and transcripts. Here we describe another strategy, using the Argonaute protein NRDE-3, repurposed as an RNA-programmable RNA binding protein. We label active transcription sites in C. elegans embryos and larvae, without editing the gene of interest. NRDE-3 is programmed by feeding nematodes with double-stranded RNA matching the target gene. This method does not require genome editing and is inexpensive and fast to apply to many different genes.

Graphical abstract: