An automated small-scale protein expression and purification screening provides beneficial information for protein production

One of the first key steps in structural genomics is high-throughput expression and rapid screening to select highly soluble proteins, the preferred candidates for crystal production. Here we describe the methodology used at the Berkeley Structural Genomics Center (BSGC) for automated parallel expression and small-scale purification of fusion proteins using a 96-well format. Our robotic method includes cell lysis, soluble fraction separation and purification with affinity resins. For detection of His-tagged proteins in the soluble fractions and after affinity resin elution, a dot-blot procedure with an anti-His-antibody is used. The expression level and molecular mass of recombinant proteins are checked by SDS-PAGE. With this approach, we are able to obtain beneficial information to be used for large-scale protein expression and purification.     

High-throughput purification of recombinant proteins using self-cleaving intein tags

High throughput methods for recombinant protein production using E. coli typically involve the use of affinity tags for simple purification of the protein of interest. One drawback of these techniques is the occasional need for tag removal before study, which can be hard to predict. In this work, we demonstrate two high throughput purification methods for untagged protein targets based on simple and cost-effective self-cleaving intein tags. Two model proteins, E. coli beta-galactosidase (βGal) and superfolder green fluorescent protein (sfGFP), were purified using self-cleaving versions of the conventional chitin-binding domain (CBD) affinity tag and the nonchromatographic elastin-like-polypeptide (ELP) precipitation tag in a 96-well filter plate format. Initial tests with shake flask cultures confirmed that the intein purification scheme could be scaled down, with >90% pure product generated in a single step using both methods. The scheme was then validated in a high throughput expression platform using 24-well plate cultures followed by purification in 96-well plates. For both tags and with both target proteins, the purified product was consistently obtained in a single-step, with low well-to-well and plate-to-plate variability. This simple method thus allows the reproducible production of highly pure untagged recombinant proteins in a convenient microtiter plate format.     

Bacterial cell-free system for high-throughput protein expression and a comparative analysis of Escherichia coli cell-free and whole cell expression systems

Sixty-three proteins of Pseudomonas aeruginosa in the size range of 18-159 kDa were tested for expression in a bacterial cell-free system. Fifty-one of the 63 proteins could be expressed and partially purified under denaturing conditions. Most of the expressed proteins showed yields greater than 500 ng after a single affinity purification step from 50 microl in vitro protein synthesis reactions. The in vitro protein expression plus purification in a 96-well format and analysis of the proteins by SDS-PAGE were performed by one person in 4 h. A comparison of in vitro and in vivo expression suggests that despite lower yields and less pure protein preparations, bacterial in vitro protein expression coupled with single-step affinity purification offers a rapid, efficient alternative for the high-throughput screening of clones for protein expression and solubility.     

High-throughput solubility assay for purified recombinant protein immunogens

A high-throughput assay is described for analysis of the solubility of purified recombinant proteins. The assay is based on affinity purification of proteins in the presence of chaotropic agents followed by a dilution and incubation step to investigate the solubility in the absence of high concentrations of such agents. The assay can be performed in a 96-well format, which makes it well suited for high-throughput applications. For 125 recombinant proteins expressed as part of an antibody-based proteomics effort, experimental solubility data were compared to calculated hydrophobicity values based on the amino acid sequence of each protein. This comparison showed only weak correlation between the theoretical and experimental values, which emphasizes the importance of experimental assays to determine the solubility of recombinant proteins.     

High-throughput peptide synthesis and peptide purification strategy at the low micromol-scale using the 96-well format.

The increasing demand for short- and medium-sized peptides in many fields of biological, medical and pharmaceutical research requires optimized and universally applicable high-throughput synthesis and purification techniques at the low-micromol scale. Here, we describe a continuous peptide synthesis/purification approach using the 96-well format. First, a micromol scale peptide synthesis on resin beads was optimized on a novel miniaturized 96-reaction vessel block employing standard Fmoc/tBu-chemistry. Almost 90% of the synthesized peptides contained the target sequence as the main component, as judged from matrix-assisted laser desorption/ionization (MALDI) mass spectra. Impurities were mostly related to partially protected peptides. Second, we tested the applicability of ion pair reversed-phase solid-phase extraction (IP-RP-SPE) to purify individual peptides. Depending on the length and predicted hydrophobicity of the peptides, elution was performed with 25 or 35% aqueous acetonitrile in the presence of 0.1% trifluoroacetic acid (TFA). Thus, scavengers used during TFA cleavage and partially protected peptides carrying very hydrophobic protecting groups were effectively removed. Using a narrow step gradient, the target peptides were even separated from deleted sequences and protected peptides with similar hydrophobicities. Third, we combined the micromol-scale synthesis in the 96-well format with purification by IP-RP-SPE on a 96-well micro-extraction plate format. This simple, fast and parallel approach was tested on 12-mer and 15-mer peptides to map epitopes of T- and B-cell clones, respectively. Approximately 80% of all peptides were obtained at purities > 90% without purification by RP-HPLC. In summary, this novel approach has several advantages: (i) the micromol-scale reduced the cost of peptide synthesis, (ii) large numbers of peptides were purified faster, (iii) the volumes of eluents and waste were significantly reduced, and (iv) the RP-HPLC column was not contaminated with hydrophobic impurities.     

A rapid screening method for cell lines producing singly-tagged recombinant proteins using the “TARGET tag” system

Recombinant production of extracellular glycoproteins in stable mammalian cell lines is an ideal technique for obtaining a large quantity of high-quality proteins. In most cases, however, current methodologies do not allow for sufficiently rapid cell line development and protein purification. Here, we describe a 21-residue peptide tag (designated as TARGET tag) and its use for rapid stable cell line development and purification. The ability of the anti-tag antibody P20.1 to withstand repetitive regeneration cycles has enabled the development of a sensitive surface plasmon resonance-based screening format that requires only 20 µl of cell culture supernatants. Direct and semi-quantitative screening at the 96-well culture scale eliminated the need for a second screening, re-cloning, or sorting, thereby minimizing culture pre-production time. Using this system, “high producer” cell lines were established in less than a month, and milligram quantities of target proteins could be purified with a standardized protocol.     

Parallel synthesis of peptide libraries using microwave irradiation

The application of microwave irradiation to solid-phase peptide synthesis increases product purity and reduces reaction time. Parallel synthesis in 96-well polypropylene filter plates with microwave irradiation is an efficient method for the rapid generation of combinatorial peptide libraries in sufficient purity to assay the products directly for biological activity without HPLC purification. In this protocol, the solid-phase support is arrayed into each well of a 96-well plate, reagents are delivered using a multichannel pipette and a microwave reactor is used to complete peptide coupling reactions in 6 min and Fmoc-removal reactions in 4 min under temperature-controlled conditions. The microwave-assisted parallel peptide synthesis protocol has been used to generate a library of difficult hexa-beta-peptides in 61% average initial purity (50% yield) and has been applied to the preparation of longer alpha- and beta-peptides. Using this protocol, a library of 96 different hexapeptides can be synthesized in 24 h (excluding characterization).     

Generation and evaluation of mammalian secreted and membrane protein expression libraries for high-throughput target discovery

Expressed protein libraries are becoming a critical tool for new target discovery in the pharmaceutical industry. In order to get the most meaningful and comprehensive results from protein library screens, it is essential to have library proteins in their native conformation with proper post-translation modifications. This goal is achieved by expressing untagged human proteins in a human cell background. We optimized the transfection and cell culture conditions to maximize protein expression in a 96-well format so that the expression levels were comparable with the levels observed in shake flasks. For detection purposes, we engineered a 'tag after stop codon' system. Depending on the expression conditions, it was possible to express either native or tagged proteins from the same expression vector set. We created a human secretion protein library of 1432 candidates and a small plasma membrane protein set of about 500 candidates. Utilizing the optimized expression conditions, we expressed and analyzed both libraries by SDS-PAGE gel electrophoresis and Western blotting. Two thirds of secreted proteins could be detected by Western-blot analyses; almost half of them were visible on Coomassie stained gels. In this paper, we describe protein expression libraries that can be easily produced in mammalian expression systems in a 96-well format, with one protein expressed per well. The libraries and methods described allow for the development of robust, high-throughput functional screens designed to assay for protein specific functions associated with a relevant disease-specific activity.     

Screening for soluble expression of recombinant proteins in a 96-well format

For future structural and functional genomics programs new tools will be required. The implementation of high-throughput (HTP) methods for protein production will be an essential element. Present HTP protein production developments in structural genomics are aimed at obtaining well-expressing and highly soluble proteins, which are preferred candidates for structure-function studies. Here, we describe a cheap and efficient procedure to identify well-expressing soluble proteins in Escherichia coli in a compact 96-well format. Reproducible lysis on filter plates, followed by a filtration step on 96-well filter plates, allows the efficient separation of inclusion bodies from the soluble fraction. In the following step a dot blot procedure using anti-RGS-His(4) antibody (Qiagen) to detect expression of recombinant His-tagged protein is applied allowing direct detection of the target protein in the soluble fraction. The method is well suited for automation and should be applicable to expression screening of most proteins and fusion domains to which specific antibodies are available.     

From genes to proteins: high-throughput expression and purification of the human proteome

The development of high-throughput methods for gene discovery has paved the way for the design of new strategies for genome-scale protein analysis. Lawrence Livermore National Laboratory and Onyx Pharmaceuticals, Inc., have produced an automatable system for the expression and purification of large numbers of proteins encoded by cDNA clones from the IMAGE (Integrated Molecular Analysis of Genomes and Their Expression) collection. This high-throughput protein expression system has been developed for the analysis of the human proteome, the protein equivalent of the human genome, comprising the translated products of all expressed genes. Functional and structural analysis of novel genes identified by EST (Expressed Sequence Tag) sequencing and the Human Genome Project will be greatly advanced by the application of this high-throughput expression system for protein production. A prototype was designed to demonstrate the feasibility of our approach. Using a PCR-based strategy, 72 unique IMAGE cDNA clones have been used to create an array of recombinant baculoviruses in a 96-well microtiter plate format. Forty-two percent of these cDNAs successfully produced soluble, recombinant protein. All of the steps in this process, from PCR to protein production, were performed in 96-well microtiter plates, and are thus amenable to automation. Each recombinant protein was engineered to incorporate an epitope tag at the amino terminal end to allow for immunoaffinity purification. Proteins expressed from this system are currently being analyzed for functional and biochemical properties.     

The RNA polymerase factory: a robotic in vitro assembly platform for high-throughput production of recombinant protein complexes

The in-depth structure/function analysis of large protein complexes, such as RNA polymerases (RNAPs), requires an experimental platform capable of assembling variants of such enzymes in large numbers in a reproducible manner under defined in vitro conditions. Here we describe a streamlined and integrated protocol for assembling recombinant archaeal RNAPs in a high-throughput 96-well format. All aspects of the procedure including construction of redesigned expression plasmids, development of automated protein extraction/in vitro assembly methods and activity assays were specifically adapted for implementation on robotic platforms. The optimized strategy allows the parallel assembly and activity assay of 96 recombinant RNAPs (including wild-type and mutant variants) with little or no human intervention within 24 h. We demonstrate the high-throughput potential of this system by evaluating the side-chain requirements of a single amino acid position of the RNAP Bridge Helix using saturation mutagenesis.     

A colorimetric assay to quantify dehydrogenase activity in crude cell lysates

Nitroblue tetrazolium (NBT) in the presence of phenazine methosulfate (PMS) reacts with the NADPH produced by dehydrogenases to produce an insoluble blue-purple formazan. Endpoint assays taking advantage of this reaction have been successfully used to detect the activity of several dehydrogenases. Here we present a version of this assay suitable for determining the kinetics of 6-phosphogluconate dehydrogenase catalysis in crude lysates of bacterial cells prepared in 96-well plates. Using the assay to screen a small library of variant 6-phosphogluconate dehydrogenases generated by error-prone polymerase chain reaction, we were able to identify three variants with improved activity and thermostability over the parent enzyme. These enzymes were partially purified and shown to be expressed at higher levels than the parent (leading to the increase in activity), and all three variants were indeed more thermostable than the parent (temperature midpoints 4-7 degrees C higher) after purification. Thus the NBT-PMS assay appears suitable for screening libraries of variant dehydrogenases.     

Generation of a fluorescently labeled endogenous protein library in living human cells

We present a protocol to tag proteins expressed from their endogenous chromosomal locations in individual mammalian cells using central dogma tagging. The protocol can be used to build libraries of cell clones, each expressing one endogenous protein tagged with a fluorophore such as the yellow fluorescent protein. Each round of library generation produces 100-200 cell clones and takes about 1 month. The protocol integrates procedures for high-throughput single-cell cloning using flow cytometry, high-throughput cDNA generation and 3' rapid amplification of cDNA ends, semi-automatic protein localization screening using fluorescent microscopy and freezing cells in 96-well format.     

Rapid screening and scale-up of ultracentrifugation-free,membrane-based procedures for purification of His-tagged membrane proteins

This study develops procedures to rapidly screen conditions for purification of membrane proteins (MPs) using 96-well plates containing nickel-functionalized membranes. In addition to their application in the pharmaceutical industry, MPs are important components of new sensors, synthetic membranes, and bioelectronic devices. However, purification of MPs is challenging due to their hydrophobic exterior, which requires stabilization in amphipathic detergent micelles. We examined the extent of extraction of the light-driven sodium transporter, Krokinobacter eikastus rhodopsin 2 (KR2) heterologously expressed in Escherichia coli using different salts and maltoside-based detergents. The extraction was followed by subsequent affinity purification in membranes functionalized with Ni²⁺-nitrilotriacetate complexes that bind the His-tagged KR2. We also employed a hydrophobic chelator to separate detergent micelles from the aqueous phase as an initial isolation step prior to affinity purification. Unlike conventional resin-based capture, which can take a full day or more, the membrane-based screening of purification conditions takes only a few hours, and its scale-up involves changing from a 96-well format to a larger membrane module. The novelty of the method lies in utilizing membrane-based ultracentrifugation-free purification of MPs from cell membrane fragments; the optimized purification conditions from the screening method can potentially be applied to large-scale/conventional resin-based purification of MPs.     

A micro-scale process for high-throughput expression of cDNAs in the yeast Saccharomyces cerevisiae

Methods have been developed aimed at applying at high-throughput technology for expression of cloned cDNAs in yeast. Yeast is a eukaryotic host, which produces soluble recombinant proteins and is capable of introducing post-translational modifications of protein. It is, thus, an appropriate expression system both for the routine expression of various cDNAs or protein domains and for the expression of proteins, which are not correctly expressed in Escherichia coli. Here, we describe a standard system in Saccharomyces cerevisiae, based on a vector for intracellular protein expression, where the gene products are fused to specific peptide sequences (tags). These epitope tags, the N-terminal His(6) tag and the C-terminal StrepII tag, allow subsequent immunological identification and purification of the gene products by a two-step affinity chromatography. This method of dual-tagged recombinant protein purification eliminates contamination by degraded protein products. A miniaturization of the procedures for cloning, expression, and detection was performed to allow all steps to be carried out in 96-well microtiter plates. The system is, thus, suitable for automation. We were able to analyze the simultaneous protein expression of a large number of cDNA clones due to the highly parallel approach of protein production and purification. The microtiter plate technology format was extended to quantitative analysis. An ELISA-based assay was developed that detects StrepII-tagged proteins. The application of this high-throughput expression system for protein production will be a useful tool for functional and structural analyses of novel genes, identified by the Human Genome Project and other large-scale sequencing projects.     

High-throughput screening of soluble recombinant proteins

The aims of high-throughput (HTP) protein production systems are to obtain well-expressed and highly soluble proteins, which are preferred candidates for use in structure-function studies. Here, we describe the development of an efficient and inexpensive method for parallel cloning, induction, and cell lysis to produce multiple fusion proteins in Escherichia coli using a 96-well format. Molecular cloning procedures, used in this HTP system, require no restriction digestion of the PCR products. All target genes can be directionally cloned into eight different fusion protein expression vectors using two universal restriction sites and with high efficiency (>95%). To screen for well-expressed soluble fusion protein, total cell lysates of bacteria culture ( approximately 1.5 mL) were subjected to high-speed centrifugation in a 96-tube format and analyzed by multiwell denaturing SDS-PAGE. Our results thus far show that 80% of the genes screened show high levels of expression of soluble products in at least one of the eight fusion protein constructs. The method is well suited for automation and is applicable for the production of large numbers of proteins for genome-wide analysis.     

High-throughput measurement of protein stability in microtiter plates

The direct determination of protein stability at high throughput has applications in proteomics, directed evolution, and formulation. Each application places different requirements on the accuracy of stability or transition midpoint determination. The measurement of protein stability by chemical denaturation has been previously performed at medium throughput and high accuracy using autotitrating fluorometers, after removal of proteins from the 96-well plate format in which they were expressed and purified. Herein we present a higher-throughput method for measuring and indexing the stability of proteins maintained within the 96-well format using a fluorescence microplate reader. Protein unfolding transitions were monitored by tryptophan fluorescence at 340 nm and assessed using bovine and equine cytochrome c (cyt c), as well as bovine serum albumin (BSA) stabilized with various amounts of palmitic acid. Two different approaches for generating unfolding curves in microtiter plates have been evaluated for their accuracy and applicability. Unfolding curves generated by the serial addition of denaturant into single wells allowed high-throughput stability screens capable of identifying protein variants with unfolding midpoint differences of 0.15 M denaturant concentration or larger. Such a method would be suitable for screening large numbers of proteins, as typically generated for directed evolution. Unfolding curves generated using one well per denaturant concentration allowed for medium-throughput stability screening and generated more accurate and precise stability values (C(1/2) +/- 0.05 M, m(G), and DeltaG(H2O)) for cyt c that are similar to values reported in literature. This method is suitable for screening the smaller numbers of proteins generated in proteomic research programmes. By using BSA stabilized with various palmitate concentrations and simple numerical indexing, it was shown that both experimental methods can successfully rank the order of protein stability.     

High-throughput proteomics: a flexible and efficient pipeline for protein production

Many studies that aim to characterize the proteome require the production of pure protein in a high-throughput format. We have developed a system for high-throughput subcloning, protein expression and purification that is simple, fast, and inexpensive. We utilized ligation-independent cloning with a custom-designed vector and developed an expression screen to test multiple parameters for optimal protein production in E. coli. A 96-well format purification protocol that produced microgram quantities of pure protein was also developed.     

Automated protein precipitation by filtration in the 96-well format

The use of automated protein precipitation by filtration in the 96-well format as a rapid sample preparation technique for high throughput bioanalysis using liquid chromatography tandem mass spectrometry is reported. A robotic sample processor is used to aspirate sequentially a plasma sample and acetonitrile separated by air gaps. These are then mixed by being dispensed into individual channels of a 96-well filter block. The resulting supernatant is separated from the precipitated plasma proteins by the application of gentle vacuum using a custom manifold. The filtered supernatants are collected into a deep well microtitre plate, evaporated to dryness using a heated 96-well dry down station and reconstituted in water prior to analysis. The efficiency of the extraction procedure is measured by the Lowry method for determining protein concentration. This method was used to optimise both the volume and the order of reagent addition, and to compare several prototype 96-well filter blocks. Using the optimised procedure a specific, precise and accurate method was developed for the β-agonist salbutamol in rabbit plasma with a calibration range of 1 to 100 ng/ml from 100 μl of sample.     

Increased efficiency for performing colony formation assays in 96-well plates: novel applications to combination therapies and high-throughput screening

The colony formation assay (CFA) is the gold standard for measuring the effects of cytotoxic agents on cancer cells in vitro; however, in its traditional 6-well format, it is a time-consuming assay, particularly when evaluating combination therapies. In the interest of increased efficiency, the 6-well CFA was converted to a 96-well format using an automated colony counting algorithm. The 96-well CFA was validated using ionizing radiation therapy on the FaDu (human hypopharyngeal squamous cell) and A549 (human lung) cancer cell lines. Its ability to evaluate combination therapies was investigated by the generation of dose-response curves for the combination of cisplatin and radiation therapy on FaDu and A549 cells. The 96-well CFA was then transferred to a robotic platform for evaluating its potential as a high-throughput screening (HTS) readout. The LOPAC1280 library was screened against FaDu cells, and eight putative hits were identified. Using the 96-well CFA to validate the eight putative chemicals, six of the eight were confirmed, resulting in a positive hit rate of 75%. These data indicate that the 96-well CFA can be adopted as an efficient alternative assay to the 6-well CFA in evaluating single and combination therapies in vitro, providing a possible readout that could be used on a HTS platform.