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Chemoproteomic profiling is a powerful approach to define the selectivity of small molecules and endogenous metabolites with the human proteome. In addition to mechanistic studies, proteome specificity profiling also has the potential to identify new scaffolds for biomolecular sensing. Here we report a chemoproteomics-inspired strategy for selective sensing of acetyl-CoA. First, we use chemoproteomic capture experiments to validate the N-terminal acetyltransferase NAA50 as a protein capable of differentiating acetyl-CoA and CoA. A Nanoluc-NAA50 fusion protein retains this specificity and can be used to generate a bioluminescence resonance energy transfer (BRET) signal in the presence of a CoA-linked fluorophore. This enables the development of a ligand displacement assay in which CoA metabolites are detected via their ability to bind the Nanoluc-NAA50 protein ‘host’ and compete binding of the CoA-linked fluorophore ‘guest.’ We demonstrate that the specificity of ligand displacement reflects the molecular recognition of the NAA50 host, while the window of dynamic sensing can be controlled by tuning the binding affinity of the CoA-linked fluorophore guest. Finally, we show the method’s specificity for acetyl-CoA can be harnessed for gain-of-signal optical detection of enzyme activity. Overall, our studies demonstrate the potential of harnessing insights from chemoproteomics for molecular sensing and provide a foundation for future applications in target engagement and selective metabolite detection.

Generating recombinant proteins in insect cells has been made possible via the use of the Baculovirus Expression Vector System (BEVS). Despite the success of many proteins via this platform, some targets remain a challenge due to issues such as cytopathic effects, the unpredictable nature of co-infection and co-expressions, and baculovirus genome instability. Many promoters have been assayed for the purpose of expressing diverse proteins in insect cells, and yet there remains a lack of implementation of those results when reviewing the landscape of commercially available baculovirus vectors. In advancing the platform to produce a greater variety of proteins and complexes, the development of such constructs cannot be avoided. A better understanding of viral gene regulation and promoter options including viral, synthetic, and insect-derived promoters will be beneficial to researchers looking to utilize BEVS by recruiting these intricate mechanisms of gene regulation for heterologous gene expression. Here we summarize some of the developments that could be utilized to improve the expression of recombinant proteins and multi-protein complexes in insect cells.

Protein prenylation is a vital eukaryotic post-translational modification which permits interaction of proteins with cellular membranes. Prenylated proteins are involved in a number of human diseases, and play a major role in cancers driven by the oncogene KRAS, which is normally farnesylated. In cases where the farnesylation machinery is inhibited, however, KRAS eludes inactivation by using an alternative prenylation pathway in which the protein is geranylgeranylated. In order to study this alternative prenylation, large quantities of accurately processed protein are required. We have developed a system to permit high-yield production of geranylgeranylated KRAS which utilizes an engineered baculovirus system. The development of this system helped to elucidate a potential metabolic bottleneck in insect cell production that should enable better production of any geranylgeranylated proteins using this system.

Baculovirus technology has been the most commonly used expression system for insect cells both due to its potential to generate a large amount of recombinant protein as well as the benefit of post-translational modifications. The most commonly used system to generate recombinant baculoviruses is the Tn7 transposition-based technology known as Bac-to-Bac. Although improvements have been made to this system to further improve quality and reproducibility of baculovirus production, recent data suggests that improved strains still have potential issues with contamination of non-recombinant baculovirus caused by improper transposition into a Tn7 site in the E. coli chromosome. Here we describe a new option for alteration of the E. coli genome to completely block the native Tn7 attachment site, leading to far fewer false positive bacmid colonies being selected and eliminating all risk of non-recombinant baculovirus production.

Baculovirus expression vectors are successfully used for the commercial production of complex (glyco)proteins in eukaryotic cells. The genome engineering of single-copy baculovirus infectious clones (bacmids) in E. coli has been valuable in the study of baculovirus biology, but bacmids are not yet widely applied as expression vectors. An important limitation of first-generation bacmids for large-scale protein production is the rapid loss of gene of interest (GOI) expression. The instability is caused by the mini-F replicon in the bacmid backbone, which is non-essential for baculovirus replication in insect cells, and carries the adjacent GOI in between attTn7 transposition sites. In this paper, we test the hypothesis that relocation of the attTn7 transgene insertion site away from the mini-F replicon prevents deletion of the GOI, thereby resulting in higher and prolonged recombinant protein expression levels. We applied lambda red genome engineering combined with SacB counterselection to generate a series of bacmids with relocated attTn7 sites and tested their performance by comparing the relative expression levels of different GOIs. We conclude that GOI expression from the odv-e56 (pif-5) locus results in higher overall expression levels and is more stable over serial passages compared to the original bacmid. Finally, we evaluated this improved next-generation bacmid during a bioreactor scale-up of Sf9 insect cells in suspension to produce enveloped chikungunya virus-like particles as a model vaccine.

Lysine acetyltransferases (KATs) play a critical role in the regulation of transcription and other genomic functions. However, a persistent challenge is the development of assays capable of defining KAT activity directly in living cells. Towards this goal, here we report the application of a previously reported dCas9-p300 fusion as a transcriptional reporter of KAT activity. First we benchmark the activity of dCas9-p300 relative to other dCas9-based transcriptional activators, and demonstrate its compatibility with second generation short guide RNA architectures. Next, we repurpose this technology to rapidly identify small molecule inhibitors of acetylation-dependent gene expression. These studies validate a recently reported p300 inhibitor chemotype, and reveal a role for p300’s bromodomain in dCas9-p300-mediated transcriptional activation. Comparison with other CRISPR-Cas9 transcriptional activators highlights the inherent ligand tuneable nature of dCas9-p300 fusions, suggesting new opportunities for orthogonal gene expression control. Overall, our studies highlight dCas9-p300 as a powerful tool for studying gene expression mechanisms in which acetylation plays a causal role, and provide a foundation for future applications requiring spatiotemporal control over acetylation at specific genomic loci.

Protein prenylation is a common posttranslational modification that enhances the ability of proteins to interact with membrane components within the cell. In many cases, these prenylated proteins are involved in important human diseases, including aging-related disorders and cancer. To effectively study these proteins or develop therapeutics, large quantities of properly modified proteins are required. Historically, production of fully modified farnesylated and methylated proteins at high yield has been challenging. Recently, an engineered insect cell system which is capable of producing authentically modified KRAS protein was used to generate material for structural studies and assay development. Here we describe protocols for extending this work to other farnesylated and methylated substrates.

Background: Trichoplusia ni derived cell lines are commonly used to enable recombinant protein expression via baculovirus infection to generate materials approved for clinical use and in clinical trials. In order to develop systems biology and genome engineering tools to improve protein expression in this host, we performed de novo genome assembly of the Trichoplusia ni-derived cell line Tni-FNL. Methods: By integration of PacBio single-molecule sequencing, Bionano optical mapping, and 10X Genomics linked-reads data, we have produced a draft genome assembly of Tni-FNL. Results: Our assembly contains 280 scaffolds, with a N50 scaffold size of 2.3 Mb and a total length of 359 Mb. Annotation of the Tni-FNL genome resulted in 14,101 predicted genes and 93.2% of the predicted proteome contained recognizable protein domains. Ortholog searches within the superorder Holometabola provided further evidence of high accuracy and completeness of the Tni-FNL genome assembly. Conclusions: This first draft Tni-FNL genome assembly was enabled by complementary long-read technologies and represents a high-quality, well-annotated genome that provides novel insight into the complexity of this insect cell line and can serve as a reference for future large-scale genome engineering work in this and other similar recombinant protein production hosts.

Protein prenylation is a common posttranslational modification that enhances the ability of proteins to interact with membrane components within the cell. In many cases, these prenylated proteins are involved in important human diseases, including aging-related disorders and cancer. To effectively study these proteins or develop therapeutics, large quantities of properly modified proteins are required. Historically, production of fully modified farnesylated and methylated proteins at high yield has been challenging. Recently, an engineered insect cell system which is capable of producing authentically modified KRAS protein was used to generate material for structural studies and assay development. Here we describe protocols for extending this work to other farnesylated and methylated substrates.

Lysine acetyltransferases (KATs) play a critical role in the regulation of transcription and other genomic functions. However, a persistent challenge is the development of assays capable of defining KAT activity directly in living cells. Towards this goal, here we report the application of a previously reported dCas9-p300 fusion as a transcriptional reporter of KAT activity. First we benchmark the activity of dCas9-p300 relative to other dCas9-based transcriptional activators, and demonstrate its compatibility with second generation short guide RNA architectures. Next, we repurpose this technology to rapidly identify small molecule inhibitors of acetylation-dependent gene expression. These studies validate a recently reported p300 inhibitor chemotype, and reveal a role for p300’s bromodomain in dCas9-p300-mediated transcriptional activation. Comparison with other CRISPR-Cas9 transcriptional activators highlights the inherent ligand tuneable nature of dCas9-p300 fusions, suggesting new opportunities for orthogonal gene expression control. Overall, our studies highlight dCas9-p300 as a powerful tool for studying gene expression mechanisms in which acetylation plays a causal role, and provide a foundation for future applications requiring spatiotemporal control over acetylation at specific genomic loci.

Prenylated proteins play key roles in several human diseases including cancer, atherosclerosis and Alzheimer’s disease. KRAS4b, which is frequently mutated in pancreatic, colon and lung cancers, is processed by farnesylation, proteolytic cleavage and carboxymethylation at the C-terminus. Plasma membrane localization of KRAS4b requires this processing as does KRAS4b-dependent RAF kinase activation. Previous attempts to produce modified KRAS have relied on protein engineering approaches or in vitro farnesylation of bacterially expressed KRAS protein. The proteins produced by these methods do not accurately replicate the mature KRAS protein found in mammalian cells and the protein yield is typically low. We describe a protocol that yields 5–10 mg/L highly purified, farnesylated and methylated KRAS4b from insect cells. Farnesylated and methylated KRAS4b is fully active in hydrolyzing GTP, binds RAF-RBD on lipid Nanodiscs and interacts with the known farnesyl-binding protein PDEδ.

The RAS Reference Reagents program within the NCI RAS Initiative is produces and distributes qualified DNA vectors, cell lines, and viruses to the external community to facilitate research into RAS biology and development of RAS cancer therapeutics. DNA vectors representing a variety of RAS mutations, as well as a complete collection of 180 genes in the RAS pathway are available as Gateway-compatible Entry clones. These clones represent the most commonly expressed transcript forms of the 180 genes, many of which were not previously available in full-length, wildtype forms. The vectors can be used with the FNLCR combinatorial cloning library to easily generate a large number of different types of expression clones with various promoters, fusion tags, and plasmid backbones. Additional reagents under construction include validated plasmid and lentiviral reagents for shRNA and CRISPR delivery, qualified cell lines for use in research into the RAS pathway, and systems for the generation of highly purified and characterized Ras and Ras-related proteins. A highlight of these reagents are materials for the high yield production of properly processed, prenylated KRAS protein using an engineered insect cell expression system. This system could be used to produce other prenylated small GTPases at levels not previously attainable. All of the resources of the RAS Reference Reagents program are readily available to the academic community through simple mechanisms, and can be licensed to industry scientists as well. The program is also interested in identifying the needs of the community for additional RAS-related resources, and is developing methods for vetting requests from the outside to provide additional reagents. Citation Format: Dominic Esposito, Jennifer Mehalko, Vanessa Wall, Carissa Grose, William Gillette, Robert Stephens. Development of reference reagents to accelerate research on the RAS pathway. [abstract]. In: Proceedings of the 107th Annual Meeting of the American Association for Cancer Research; 2016 Apr 16-20; New Orleans, LA. Philadelphia (PA): AACR; Cancer Res 2016;76(14 Suppl):Abstract nr 1872.

Background Uncharacterized proteases naturally expressed by bacterial pathogens represents important topic in infectious disease research, because these enzymes may have critical roles in pathogenicity and cell physiology. It has been observed that cloning, expression and purification of proteases often fail due to their catalytic functions which, in turn, cause toxicity in the E. coli heterologous host. Results In order to address this problem systematically, a modified pipeline of our high-throughput protein expression and purification platform was developed. This included the use of a specific E. coli strain, BL21(DE3) pLysS to tightly control the expression of recombinant proteins and various expression vectors encoding fusion proteins to enhance recombinant protein solubility. Proteases fused to large fusion protein domains, maltosebinding protein (MBP), SP-MBP which contains signal peptide at the N-terminus of MBP, disulfide oxidoreductase (DsbA) and Glutathione S-transferase (GST) improved expression and solubility of proteases. Overall, 86.1% of selected protease genes including hypothetical proteins were expressed and purified using a combination of five different expression vectors. To detect novel proteolytic activities, zymography and fluorescence-based assays were performed and the protease activities of more than 46% of purified proteases and 40% of hypothetical proteins that were predicted to be proteases were confirmed. Conclusions Multiple expression vectors, employing distinct fusion tags in a high throughput pipeline increased overall success rates in expression, solubility and purification of proteases. The combinatorial functional analysis of the purified proteases using fluorescence assays and zymography confirmed their function.

BackgroundIn the postgenomic era, high throughput protein expression and protein microarray technologies have progressed markedly permitting screening of therapeutic reagents and discovery of novel protein functions. Hexa-histidine is one of the most commonly used fusion tags for protein expression due to its small size and convenient purification via immobilized metal ion affinity chromatography (IMAC). This purification process has been adapted to the protein microarray format, but the quality ofin situHis-tagged protein purification on slides has not been systematically evaluated. We established methods to determine the level of purification of such proteins on metal chelate-modified slide surfaces. Optimizedin situpurification of His-tagged recombinant proteins has the potential to become the new gold standard for cost-effective generation of high-quality and high-density protein microarrays.ResultsTwo slide surfaces were examined, chelated Cu2+slides suspended on a polyethylene glycol (PEG) coating and chelated Ni2+slides immobilized on a support without PEG coating. Using PEG-coated chelated Cu2+slides, consistently higher purities of recombinant proteins were measured. An optimized wash buffer (PBST) composed of 10 mM phosphate buffer, 2.7 mM KCl, 140 mM NaCl and 0.05% Tween 20, pH 7.4, further improved protein purity levels. UsingEscherichia colicell lysates expressing 90 recombinantStreptococcus pneumoniaeproteins, 73 proteins were successfully immobilized, and 66 proteins werein situpurified with greater than 90% purity. We identified several antigens among thein situ-purified proteins via assays with anti-S. pneumoniaerabbit antibodies and a human patient antiserum, as a demonstration project of large scale microarray-based immunoproteomics profiling. The methodology is compatible with higher throughput formats ofin vivoprotein expression, eliminates the need for resin-based purification and circumvents protein solubility and denaturation problems caused by buffer exchange steps and freeze-thaw cycles, which are associated with resin-based purification, intermittent protein storage and deposition on microarrays.ConclusionAn optimized platform forin situprotein purification on microarray slides using His-tagged recombinant proteins is a desirable tool for the screening of novel protein functions and protein-protein interactions. In the context of immunoproteomics, such protein microarrays are complimentary to approaches using non-recombinant methods to discover and characterize bacterial antigens.

We have developed and evaluated a highly parallel protein expression and purification system using ORFs derived from the pathogenic bacterium Streptococcus pneumoniae as a representative test case in conjunction with the Gateway cloning technology. Establishing high throughput protein production capability is essential for genome-wide characterization of protein function. In this study, we focused on protein expression and purification outcomes generated from an expression vector which encodes an NH2-terminal hexa-histidine tag and a COOH-terminal S-tag. Purified recombinant proteins were validated by SDS–PAGE, followed by in-gel digestion and identification by MALDI-TOF/TOF analysis. Starting with 1360 sequence-validated destination clones we examined correlation analyses of expression and solubility of a wide variety of recombinant proteins. In total, 428 purified proteins (31%) were recovered in soluble form. We describe a semi-quantitative scoring method using an S-tag assay to improve the throughput and efficiency of expression and solubility studies for recombinant proteins. Given a relatively large dataset derived from proteins representing all functional groups in a microbial genome we correlated various protein characteristics as they relate to protein expression outcomes.

KRAS4b, which is frequently mutated in pancreatic, colon and lung cancers, is processed in several steps including farnesylation by farnesyl-transferase (FNT); proteolytic cleavage at the C-terminus by ras converting enzyme 1 (RCE1) and carboxymethylation by isoprenylcysteine carboxymethyl transferase (ICMT). Plasma membrane localization of KRAS4b requires this processing as does KRAS4b-dependent RAF kinase activation. Previous attempts to produce modified KRAS have relied on protein engineering approaches or in vitro farnesylation of bacterially expressed KRAS protein. The proteins produced by these methods do not completely replicate the final KRAS protein found in mammalian cells and the yield and quality of the protein are typically low. Using an engineered baculovirus containing the two subunits of farnesyl-transferase (FNTA/B) and a maltose-binding protein (MBP)-KRAS4b fusion we are able to produce highly purified, farnesylated, and methylated KRAS4b (FMe-KRAS4b) in insect cells at yields of approximately 10mg/L. This protein has been extensively characterized by electrospray-ionization mass spectrometry, analytical ultracentrifugation, and circular dichroism. In addition, we demonstrate that FMe-KRAS4b is fully active in hydrolyzing GTP, binding to RAF-RBD, and interacting with lipid nanodiscs. Citation Format: Andrew G. Stephen, William K. Gillette, Carissa Grose, Dominic Esposito. Production and biochemical characterization of farnesylated and methylated KRAS4b using engineered baculovirus. [abstract]. In: Proceedings of the 106th Annual Meeting of the American Association for Cancer Research; 2015 Apr 18-22; Philadelphia, PA. Philadelphia (PA): AACR; Cancer Res 2015;75(15 Suppl):Abstract nr 3925. doi:10.1158/1538-7445.AM2015-3925

Toward achieving rapid and large scale genome modification directly in a target organism, we have developed a new genome engineering strategy that uses a combination of bioinformatics aided design, large synthetic DNA and site-specific recombinases. Using Cre recombinase we swapped a target 126-kb segment of the Escherichia coli genome with a 72-kb synthetic DNA cassette, thereby effectively eliminating over 54 kb of genomic DNA from three non-contiguous regions in a single recombination event. We observed complete replacement of the native sequence with the modified synthetic sequence through the action of the Cre recombinase and no competition from homologous recombination. Because of the versatility and high-efficiency of the Cre-lox system, this method can be used in any organism where this system is functional as well as adapted to use with other highly precise genome engineering systems. Compared to present-day iterative approaches in genome engineering, we anticipate this method will greatly speed up the creation of reduced, modularized and optimized genomes through the integration of deletion analyses data, transcriptomics, synthetic biology and site-specific recombination.