Biological Engineering

Pathological conditions of the cervix ranging from cervical cancer to structural dysfunction associated with preterm labor all have limited treatment options. Thus, there is a need for physiologically relevant preclinical models that recapitulate the structure and function of this human organ. Here, we describe a protocol for engineering and studying a highly functional in vitro model of the human cervix that is composed of a commercially available, dual-channel, microfluidic, organ-on-a-chip (Organ Chip) device lined by primary cervical epithelial (CE) cells interfaced across a porous membrane with cervical stromal cells. The provision of dynamic and customized media flow through both the epithelial and stromal compartments results in cell growth and differentiation, including the accumulation of a thick mucus layer overlying the epithelium. The resulting model closely mimics the structure, epithelial barrier, mucus composition and structure, and biochemical properties of the in vivo human cervix, as well as its responsiveness to female hormones, pH, and microbiome. This Cervix Chip protocol also includes noninvasive techniques for longitudinal monitoring of the live 3D tissue model. The Cervix Chip offers a powerful preclinical platform for replicating in vivo cervical physiology, studying disease mechanisms, and facilitating the development of new therapeutics and diagnostics.

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

The sesquiterpene lactone compound artemisinin is a natural medicinal product of commercial importance. This Artemisia annua–derived secondary metabolite is well known for its antimalarial activity and has been studied in several other biological assays. However, the major shortcoming in its production and commercialization is its low accumulation in the native plant. Moreover, the chemical synthesis of artemisinin is difficult and expensive due to its complex structure. Hence, an alternative and sustainable production system of artemisinin in a heterologous host is required. Previously, heterologous production of artemisinin was achieved by Agrobacterium-mediated transformation. However, this requires extensive bioengineering of modified Nicotiana plants. Recently, a technique involving direct in vivo assembly of multiple DNA fragments in the moss, P. patens, has been successfully established. We utilized this technique to engineer artemisinin biosynthetic pathway genes into the moss, and artemisinin was obtained without further modifications with high initial production. Here, we provide protocols for establishing moss culture accumulating artemisinin, including culture preparation, transformation method, and compound detection via HS-SPME, UPLC-MRM-MS, and LC-QTOF-MS. The bioengineering of moss opens up a more sustainable, cost effective, and scalable platform not only in artemisinin production but also other high-value specialized metabolites in the future.

Regulated cell death plays a key role in immunity, development, and homeostasis, but is also associated with a number of pathologies such as autoinflammatory and neurodegenerative diseases and cancer. However, despite the extensive mechanistic research of different cell death modalities, the direct comparison of different forms of cell death and their consequences on the cellular and tissue level remain poorly characterized. Comparative studies are hindered by the mechanistic and kinetic differences between cell death modalities, as well as the inability to selectively induce different cell death programs in an individual cell within cell populations or tissues. In this method, we present a protocol for rapid and specific optogenetic activation of three major types of programmed cell death: apoptosis, necroptosis, and pyroptosis, using light-induced forced oligomerization of their major effector proteins (caspases or kinases).

Engineered aptamers for new compounds are typically produced by using in vitro selection methods. However, aptamers that are developed in vitro might not function as expected when introduced into complex cellular environments. One approach that addresses this concern is the design of initial RNA pools for selection that contain structural scaffolds from naturally occurring riboswitch aptamers. Here, we provide guidance on design and experimental principles for developing riboswitch-inspired aptamers for new ligands. The in vitro selection protocol (based on Capture-SELEX) is generalizable to diverse RNA scaffold types and amenable to multiplexing of ligand candidates. We discuss strategies to avoid propagation of selfish sequences that can easily dominate the selection. We also detail the identification of aptamer candidates using next-generation sequencing and bioinformatics, and subsequent biochemical validation of aptamer candidates. Finally, we describe functional testing of aptamer candidates in bacterial cell culture.

Key features

• Develop riboswitch-inspired aptamers for new ligands using in vitro selection.

• Ligand candidates can be multiplexed to conserve time and resources.

• Test aptamer candidates in bacterial cells by grafting the aptamer back onto its expression platform.

Graphical overview

The easyPACId (easy Promoter Activation and Compound Identification) approach is focused on the targeted activation of natural product biosynthetic gene clusters (BGCs) encoding non-ribosomal peptide synthetases (NRPS), polyketide synthases (PKS), NRPS-PKS hybrids, or other BGC classes. It was applied to entomopathogenic bacteria of the genera Xenorhabdus and Photorhabdus by exchanging the natural promoter of desired BGCs against the L-arabinose inducible P_BAD promoter in ∆hfq mutants of the respective strains. The crude (culture) extracts of the cultivated easyPACId mutants are enriched with the single compound or compound class and can be tested directly against various target organisms without further purification of the produced natural products. Furthermore, isolation and identification of compounds from these mutants is simplified due to the reduced background in the ∆hfq strains. The approach avoids problems often encountered in heterologous expression hosts, chemical synthesis, or tedious extraction of desired compounds from wild-type crude extracts. This protocol describes easyPACId for Xenorhabdus and Photorhabdus, but it was also successfully adapted to Pseudomonas entomophila and might be suitable for other proteobacteria that carry hfq.