Microbiology

Recombinant proteins of high quality are crucial starting materials for all downstream applications, but the inherent complexities of proteins and their expression and purification create significant challenges. The Pichia pastoris yeast is a highly useful eukaryotic protein expression system. Pichia’s low cost, genetic tractability, rapid gene expression, and scalability make it an ideal expression system for foreign proteins. Here, we developed a protocol that has optimized the expression and isolation of a non-mammalian secreted metalloprotease, where we can routinely generate recombinant proteins that are pure and proteolytically active. We maximized growth and protein production by altering the feeding regime, through implementation of a non-fermentable and non-repressing carbon source during the methanol-induction phase. This approach increased biomass production and yielded milligrams of recombinant protein. Downstream applications involving active, recombinant fungal proteases, such as conjugation to nanoparticles and structure-related studies, are greatly facilitated with this improved, standardized approach.

Graphical abstract

Genome-wide screens using yeast or phage displays are powerful tools for identifying protein–ligand interactions, including drug or vaccine targets, ligand receptors, or protein–protein interactions. However, assembling libraries for genome-wide screens can be challenging and often requires unbiased cloning of 10⁵–10⁷ DNA fragments for a complete representation of a eukaryote genome. A sub-optimal genomic library can miss key genomic sequences and thus result in biased screens. Here, we describe an efficient method to generate genome-wide libraries for yeast surface display using Gibson assembly. The protocol entails genome fragmentation, ligation of adapters, library cloning using Gibson assembly, library transformation, library DNA recovery, and a streamlined Oxford nanopore library sequencing procedure that covers the length of the cloned DNA fragments. We also describe a computational pipeline to analyze the library coverage of the genome and predict the proportion of expressed proteins. The method allows seamless library transfer among multiple vectors and can be easily adapted to any expression system.

Based on previous in-depth characterisation, aldehyde dehydrogenases (ALDH) are a diverse superfamily of enzymes, in terms of both structure and function, present in all kingdoms of life. They catalyse the oxidation of an aldehyde to carboxylic acid using the cofactor nicotinamide adenine dinucleotide (phosphate) (NAD(P)⁺), and are often not substrate-specific, but rather have a broad range of associated biological functions, including detoxification and biosynthesis. We studied the structure of ALDH_Tt from Thermus thermophilus, as well as performed its biochemical characterisation. This allowed for insight into its potential substrates and biological roles.

In this protocol, we describe ALDH_Tt heterologous expression in E. coli, purification, and activity assay (based on Shortall et al., 2021). ALDH_Tt was first copurified as a contaminant during caa₃-type cytochrome oxidase isolation from T. thermophilus. This recombinant production system was employed for structural and biochemical analysis of wild-type and mutants, and proved efficient, yielding approximately 15–20 mg/L ALDH_Tt. For purification of the thermophilic his-tagged ALDH_Tt, heat treatment, immobilized metal affinity chromatography (IMAC), and gel filtration chromatography were used. The enzyme activity assay was performed via UV-Vis spectrophotometry, monitoring the production of reduced nicotinamide adenine dinucleotide (NADH).

Graphical abstract:

Flow chart outlining the steps in ALDH_Tt expression and purification, highlighting the approximate time required for each step.

Cell lysis, a process that releases host oligonucleotides, is required in many biotechnological applications. However, intact oligonucleotides in crude cellular lysates increase the viscosity of lysates, which complicates downstream processes and routine laboratory workflows. To address this, nucleases that hydrolyze the intact oligonucleotides are commonly added, either as purified enzymes or co-expressed in genetically engineered bacterial strains. To measure oligonucleotide hydrolysis, common DNA quantification methods, such as qPCR or fluorescence-based, require expensive reagents and equipment, and cannot distinguish different-sized DNA fragments. Here, we outline a simple alternative method for measuring DNA/RNA hydrolysis in cellular lysates, by measuring their viscosity. This method only requires common laboratory supplies and a cell phone camera.

Recombinant protein expression is extensively used in biological research. Despite this, current protein expression and extraction methods are not readily scalable or amenable for high-throughput applications. Optimization of protein expression conditions using traditional methods, reliant on growth-associated induction, is non-trivial. Similarly, protein extraction methods are predominantly restricted to chemical methods, and mechanical methods reliant on expensive specialized equipment more tuned for large-scale applications. In this article, we outline detailed protocols for the use of an engineered autolysis/autohydrolysis E. coli strain, in two-stage fermentations in shake-flasks. This two-stage fermentation protocol does not require optimization of expression conditions and results in high protein titers. Cell lysis in an engineered strain is tightly controlled and only triggered post-culture by addition of a 0.1% detergent solution. Upon cell lysis, a nuclease digests contaminating host oligonucleotides, which facilitates sample handling. This method has been validated for use at different scales, from microtiter plates to instrumented bioreactors.

Graphic abstract:

Two-stage protein expression, cell autolysis and DNA/RNA autohydrolysis.

Reprinted with permission from Menacho-Melgar et al. (2020a). Copyright 2020 John Wiley and Sons.

We have previously described the development of two specialized Escherichia coli strains for high-level recombinant membrane protein (MP) production. These engineered strains, termed SuptoxD and SuptoxR, are capable of suppressing the cytotoxicity caused by MP overexpression and of producing greatly enhanced MP yields. Here, we present a Bio-protocol that describes gene overexpression and culturing conditions that maximize the accumulation of membrane-integrated and well-folded recombinant MPs in these strains.

Microbial production of alkanes employing synthetic biology tools has gained tremendous attention owing to the high energy density and similarity of alkanes to existing petroleum fuels. One of the most commonly studied pathways includes the production of alkanes by AAR (acyl-ACP (acyl carrier protein) reductase)-ADO (aldehyde deformylating oxygenase) pathway. Here, the intermediates of fatty acid synthesis pathway are used as substrate by the AAR enzyme to make fatty aldehyde, which is then deformylated by ADO to make linear chain alkane. However, the variation in substrate availability to the first enzyme of the pathway, i.e., AAR, via fatty acid synthesis pathway and low turnover of the ADO enzyme make calculation of yields and titers under in vivo conditions extremely difficult. In vivo assay employing external addition of defined substrates for ADO enzyme into the medium helps to monitor the influx of substrate hence providing a more accurate measurement of the product yields. In this protocol, we include a detailed guide for implementing the in vivo assay for monitoring alkane production in E. coli.

Roseburia and Eubacterium species of the human gut microbiota play an important role in the maintaince of human health, partly by producing butyrate, the main energy source of our colonic epithelial cells. However, our knowledge of the biochemistry and physiology of these bacteria has been limited by a lack of genetic manipulation techniques. Conjugative transposons previously introduced into Roseburia species could not be easily modified, greatly limiting their applicability as genetic modification platforms. Modular plasmid shuttle vectors have previously been developed for Clostridium species, which share a taxonomic order with Roseburia and Eubacterium, raising the possibility that these vectors could be used in these organisms. Here, we describe an optimized conjugation protocol enabling the transfer of autonomously replicating plasmids from an E. coli donor strain into Roseburia inulinivorans and Eubacterium rectale. The modular nature of the plasmids and their ability to be maintained in the recipient bacterium by autonomous replication makes them ideal for investigating heterologous gene expression, and as a platform for other genetic tools including antisense RNA silencing or mobile group II interon gene disruption strategies.

Yeast are an ideal system to study Heat Shock Protein 70 (Hsp70) function in a cellular context. This protocol was generated to analyze the function of non-native Hsp70 proteins by expressing them as the sole cytosolic Hsp70 in yeast. As an initial step, Hsp70 variants (such as Ssa1 point mutants and non-yeast versions such as Nematostella vectensis NvHsp70A, B and D) are cloned into an appropriate expression plasmid. Next, these plasmids are transformed into ssa1-4∆ yeast [expressing native Ssa1 from an uracil-based (URA3) plasmid] which are subsequently cured of the original yeast on 5-Fluroorotic Acid (5-FOA). The resulting cells can be screened for a variety of phenotypes which match to the activity of well-studied cellular pathways.

Human pancreatic lipase (HPL) is the main lipolytic enzyme involved in the digestion of dietary fat. An active recombinant human pancreatic lipase (recHPL) was successfully prepared for the first time in an Escherichia coli (E. coli) expression system using a short Strep-tag II (ST II). The recHPL-ST II was solubilized with 8 M urea from the E. coli lysate and purified on a Strep-Tactin-Sepharose column. After refolding by stepwise dialyses against decreasing concentrations of urea in the presence of glycerol and Ca²⁺ for two days followed by gel filtration FPLC, 1.8-6 mg of active recHPL-ST II was obtained from 1 L of culture. Here we report the expression, purification, and optimized refolding procedures for active recHPL from E. coli, thus establishing it as a suitable system for the production of recHPL of high purity and scaling up.