Microbiology

Eukaryotic cells rely on actin to support cellular structure, motility, transport, and a wide variety of other cytoplasmic functions and nuclear activities. Humans and other mammals express six closely related isoforms of actin, four of which are found primarily in skeletal, cardiac, and smooth muscle tissues. The final two isoforms, β and γ, are found in non-muscle cells. Due to the ease of purification, many biochemical studies surveying the functions of actin and its regulators have been carried out with protein purified from skeletal muscle. However, it has become increasingly clear that some activities are isoform specific, necessitating more accessible sources of non-muscle actin isoforms. Recent innovations permit the purification of non-muscle actins from human cell culture and heterologous systems, such as insect cell culture and the yeast Pichia pastoris. However, these systems generate mixtures of actin types or require additional steps to remove purification-related tags. We have developed strains of Saccharomyces cerevisiae (budding yeast) that express single untagged isoforms of either human non-muscle actin (β or γ) as their sole actin, allowing the purification of individual homogeneous actin isoforms by conventional purification techniques.

Key features

• Easy growth of yeast as a source of human cytoplasmic actin isoforms.

• Uses well-established actin purification methods.

• The tag-free system requires no post-purification processing.

Graphical overview

Isolating human cytoplasmic actins from yeast

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.

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.

To determine boron quantity in soil, water and biological samples, several protocols are available. Colorimetric assays are the simplest and cheapest methods which can be used to determine boron concentration. However, published protocols do not give straightforward guidance for beginners to adopt these protocols for routine use in the laboratory. Based on a previously published available procedure, we present a detailed and modified version of a curcumin based colorimetric protocol to determine boron concentration extracted from any sample. Our modified protocol is able to determine up to 0.2 nmole of Boron in a sample volume of 300 µl.

This protocol describes the methods used to engineer and deploy genetically encoded fluorescence activity reporters for nitrate and peptide transporter activity in yeast cells. Fusion of the dual-affinity nitrate transceptor CHL1/AtNRT1.1/AtNPF6.3 or four different peptide transporters (AtPTR1, 2, 4, and 5) from Arabidopsis to a pair of fluorescent proteins with different spectral properties, enabled us to engineer the NiTracs (nitrate transporter activity tracking sensors) and the PepTracs (peptide transporter activity tracking sensors), ratiometric fluorescence activity sensors that monitor the activity of the plasma membrane nitrate transceptor or the peptide transporters in vivo (Ho et al., 2014). The NiTrac1 sensor responds specifically and reversibly to the addition of nitrate, while the PepTracs respond to addition of dipeptides, either by a reduction in donor and acceptor emission, while acceptor-excited emission remains unaltered, or by a change in ratio of the fluorophore emission. All sensors are suitable for ratiometric imaging. The similarity of the biphasic kinetics of the NiTrac1 sensor response [from µM to mM (Liu and Tsay, 2003)] and the nitrate transport kinetics of the native nitrate transceptor, intimates that NiTrac1 provides information on conformational rearrangements during the transport cycle, thereby reporting transporter activity over a wide range of external nitrate concentrations. Several variants of NiTrac have been engineered, which differ with respect to their affinity for nitrate (NiTrac1: CHL1; NiTracT101A: CHL1T101A). NiTrac also recognizes chlorate. Here we describe a simple method for the design, implementation, and detection of nitrate transceptor activity in yeast cells using a spectrofluorimeter.

Heterologous expression of genes in budding yeast Saccharomyces cerevisiae (S. cerevisiae) is especially suitable to functionally study the corresponding encoded protein at the cellular level (Bonneaud et al., 1991). This is mainly because many strains defective in specific activities are available and could be complemented by homologous genes existing across the eukaryotic kingdom (http://www.yeastgenome.org/). However, the protocol we describe here is not a complementation but a “gain-of-function” assay. It is based on a drop-test assay that we have set up to assess the cellular zinc tolerance conferred by the expression of heterologous genes in the wild-type S. cerevisiae. Different dilutions of a yeast culture expressing the heterologous gene of interest are grown on a range of zinc-enriched plates, and are then compared to the control yeast expressing the empty vector. Working with different concentrations of both yeast and zinc are essential to succeed in describing zinc tolerance phenotype upon yeast transformation (Mirouze et al., 2006). This test has also proven to be valuable to differentiate among related members of gene families as exemplified for Arabidopsis Plant Defensin type1 (Shahzad et al., 2013).