Biological Engineering


Protocols in Current Issue
Protocols in Past Issues
0 Q&A 404 Views Apr 20, 2023

Plant protoplasts are useful to study both transcriptional regulation and protein subcellular localization in rapid screens. Protoplast transformation can be used in automated platforms for design-build-test cycles of plant promoters, including synthetic promoters. A notable application of protoplasts comes from recent successes in dissecting synthetic promoter activity with poplar mesophyll protoplasts. For this purpose, we constructed plasmids with TurboGFP driven by a synthetic promoter together with TurboRFP constitutively controlled by a 35S promoter, to monitor transformation efficiency, allowing versatile screening of high numbers of cells by monitoring green fluorescent protein expression in transformed protoplasts. Herein, we introduce a protocol for poplar mesophyll protoplast isolation followed by protoplast transformation and image analysis for the selection of valuable synthetic promoters.

Graphical overview

0 Q&A 645 Views Nov 5, 2022

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.

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0 Q&A 919 Views Nov 5, 2022

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) ∆AfabRserB, 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.

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0 Q&A 1424 Views Oct 20, 2022

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.

0 Q&A 724 Views Sep 5, 2022

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.

0 Q&A 1125 Views Jun 5, 2022

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.

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0 Q&A 1642 Views Jun 5, 2022

A multitude of membrane-localized receptors are utilized by cells to integrate both biochemical and physical signals from their microenvironment. The clustering of membrane receptors is widely presumed to have functional consequences for subsequent signal transduction. However, it is experimentally challenging to selectively manipulate receptor clustering without altering other biochemical aspects of the cellular system. Here, we describe a method to fabricate multicomponent, ligand-functionalized microarrays, for spatially segregated and simultaneous monitoring of receptor activation and signaling in individual living cells. While existing micropatterning techniques allow for the display of fixed ligands, this protocol uniquely allows for functionalization of both mobile membrane corrals and immobile polymers with selective ligands, as well as microscopic monitoring of cognate receptor activation at the cell membrane interface. This protocol has been developed to study the effects of clustering on EphA2 signaling transduction. It is potentially applicable to multiple cell signaling systems, or microbe/host interactions.

Graphical abstract:

A side-by-side comparison of clustered or non-clustered EphA2 receptor signaling in a single cell.

0 Q&A 1403 Views Mar 20, 2022

Transbilayer movement of phospholipids in biological membranes is mediated by a diverse set of lipid transporters. Among them are scramblases that facilitate rapid bi-directional movement of lipids without metabolic energy input. In this protocol, we describe the incorporation of phospholipid scramblases into giant unilamellar vesicles (GUVs) formed from scramblase-containing large unilamellar vesicles by electroformation. We also describe how to analyze their activity using membrane-impermeant sodium dithionite, to bleach symmetrically incorporated fluorescent ATTO488-conjugated phospholipids. The fluorescence-based readout allows single vesicle tracking for a large number of settled/immobilized GUVs, and provides a well-defined experimental setup to directly characterize these lipid transporters at the molecular level.

Graphic abstract:

Giant unilamellar vesicles (GUVs) are formed by electroformation from large unilamellar vesicles (LUVs) containing phospholipid scramblases (purple) and trace amounts of a fluorescent lipid reporter (green).
The scramblase activity is analyzed by a fluorescence-based assay of single GUVs, using the membrane-impermeant quencher dithionite. Sizes not to scale. Modified from Mathiassen et al. (2021).

0 Q&A 2241 Views Mar 5, 2022

Double-strand breaks (DSBs) are lesions in DNA that, if not properly repaired, can cause genomic instability, oncogenesis, and cell death. Multiple chromatin posttranslational modifications (PTMs) play a role in the DNA damage response to DSBs. Among these, RNF168-mediated ubiquitination of lysines 13 or 15 at the N-terminal tail of histone H2A (H2AK13/15Ub) is essential for the recruitment of effectors of both the non-homologous end joining (NHEJ) and the homologous recombination (HR) repair pathways. Thus, tools and techniques to track the spatiotemporal dynamics of H2AK13/15 ubiquitination at DNA DSBs are important to facilitate studies of DNA repair. Previous work from other groups used the minimal focus-forming region (FFR) of the NHEJ effector 53BP1 to detect H2AK15Ub generated upon damage induced by gamma or laser irradiation in live cells. However, 53BP1-FFR only binds nucleosomes modified with both H2AK15Ub and dimethylation of lysine 20 on histone H4 (H4K20me2); thus, 53BP1-FFR does not recognize H2AK13Ub–nucleosomes or nucleosomes that contain H2AK15Ub but lack methylation of H4K20 (H4K20me0). To overcome this limitation, we developed an avidity-based sensor that binds H2AK13/15Ub without dependence on the methylation status of histone H4K20. This sensor, called Reader1.0, detects DNA damage-associated H2AK13/15Ub in live cells with high sensitivity and selectivity. Here, we present a protocol to detect the formation of H2AK13/15Ub at laser-induced DSBs using Reader1.0 as a live-cell reporter for this histone PTM.

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0 Q&A 1837 Views Feb 20, 2022

At the end of about 80% of the operon in Escherichia coli, translation termination decouples transcription, leading to Rho-dependent transcription termination (RDT). However, no in vitro or in vivo assay system has proven to be good enough to see the 3’ end of the mRNA generated by RDT. Here, we present a cell-free assay system that could provide detailed information on the 3’ end of a transcript RNA generated by RDT. Our protocol shows how to extract transcript RNA generated by transcription reactions from a cell-free extract, followed by an RNA oligomer ligation to the 3’ end of a transcript RNA of interest. The 3' end of the RNA is amplified using RT-PCR. Its genetic location can be determined using a gene-specific primer extension reaction. The 3’ ends of mRNA can be visualized and quantified by polyacrylamide gel electrophoresis. One significant advantage of a cell-free assay system is that factors involved in the generation of the 3' end, such as proteins and sRNA, can be directly assayed by exogenously adding factor(s) to the reaction.

Graphic abstract:

An illustration of the experimental methodology.

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