Therapeutic recombinant protein production relies on industrial scale culture of mammalian cells to produce active proteins in quantities sufficient for clinical use. The combination of stresses from industrial cell culture environment and recombinant protein production can overwhelm the protein synthesis machinery in the endoplasmic reticulum (ER). This leads to a buildup of improperly folded proteins which induces ER stress. Cells respond to ER stress by activating the Unfolded Protein Response (UPR). To restore proteostasis, ER sensor proteins reduce global protein synthesis and increase chaperone protein synthesis, and if that is insufficient the proteins are degraded. If proteostasis is still not restored, apoptosis is initiated. Increasing evidence suggests crosstalk between ER proteostasis and DNA damage repair (DDR) pathways. External factors (e.g., metabolites) from the cellular environment as well as internal factors (e.g., transgene copy number) can impact genome stability. Failure to maintain genome integrity reduces cell viability and in turn protein production. This review focuses on the association between ER stress and processes that affect protein production and secretion. The processes mediated by ER stress, including inhibition of global protein translation, chaperone protein production, degradation of misfolded proteins, DNA repair, and protein secretion, impact recombinant protein production. Recombinant protein production can be reduced by ER stress through increased autophagy and protein degradation, reduced protein secretion, and reduced DDR response.

DNA amplification and reverse transcription enzymes have proven to be invaluable in fast and reliable diagnostics and research applications because of their processivity, specificity, and robustness. Our study focused on the production of mutant Taq DNA polymerase and mutant M-MLV reverse transcriptase in the expression hosts Vibrio natriegens and Escherichia coli under various expression conditions. We also examined nonspecific extracellular production in V. natriegens. Intracellularly, M-MLV was produced in V. natriegens at the level of 11% of the total cell proteins (TCPs) compared with 16% of TCPs in E. coli. We obtained a soluble protein that accounted for 11% of the enzyme produced in V. natriegens and 22% of the enzyme produced in E. coli. Taq pol was produced intracellularly in V. natriegens at the level of 30% of TCPs compared with 26% of TCPs in E. coli. However, Taq pol was almost non-soluble in E. coli, whereas in V. natriegens, we obtained a soluble protein that accounted for 23% of the produced enzyme. We detected substantial extracellular production of Taq pol. Thus, V. natriegens is a suitable alternative host with the potential for production of recombinant proteins.

Plastid engineering offers the potential to carry multigene traits in plants; however, it requires reliable genetic parts to balance expression. The difficulty of chloroplast transformation and slow plant growth makes it challenging to build plants just to characterize genetic parts. To address these limitations, we developed a high-yield cell-free system from Nicotiana tabacum chloroplast extracts for prototyping genetic parts. Our cell-free system uses combined transcription and translation driven by T7 RNA polymerase and works with plasmid or linear template DNA. To develop our system, we optimized lysis, extract preparation procedures (e.g., runoff reaction, centrifugation, and dialysis), and the physiochemical reaction conditions. Our cell-free system can synthesize 34 ± 1 μg/mL luciferase in batch reactions and 60 ± 4 μg/mL in semicontinuous reactions. We apply our batch reaction system to test a library of 103 ribosome binding site (RBS) variants and rank them based on cell-free gene expression. We observe a 1300-fold dynamic range of luciferase expression when normalized by maximum mRNA expression, as assessed by the malachite green aptamer. We also find that the observed normalized gene expression in chloroplast extracts and the predictions made by the RBS Calculator are correlated. We anticipate that chloroplast cell-free systems will increase the speed and reliability of building genetic systems in plant chloroplasts for diverse applications.

BACKGROUND: Research on protein production holds significant importance in the advancement of food technology, agriculture, pharmaceuticals, and bioenergy. Aspergillus niger stands out as an ideal microbial cell factory for the production of food-grade proteins, owing to its robust protein secretion capacity and excellent safety profile. However, the extensive oxidative folding of proteins within the endoplasmic reticulum (ER) triggers ER stress, consequently leading to protein misfolding reactions. This stressful phenomenon results in the accelerated generation of reactive oxygen species (ROS), thereby inducing oxidative stress. The accumulation of ROS can adversely affect intracellular DNA, proteins, and lipids.
RESULTS: In this study, we enhanced the detoxification of ROS in A. niger (SH-1) by integrating multiple modules, including the NADPH regeneration engineering module, the glutaredoxin system, the GSH synthesis engineering module, and the transcription factor module. We assessed the intracellular ROS levels, growth under stress conditions, protein production levels, and intracellular GSH content. Our findings revealed that the overexpression of Glr1 in the glutaredoxin system exhibited significant efficacy across various parameters. Specifically, it reduced the intracellular ROS levels in A. niger by 50%, boosted glucoamylase enzyme activity by 243%, and increased total protein secretion by 88%.
CONCLUSIONS: The results indicate that moderate modulation of intracellular redox conditions can enhance overall protein output. In conclusion, we present a strategy for augmenting protein production in A. niger and propose a potential approach for optimizing microbial protein production system.

Mammalian cell lines are one of the best options when it comes to the production of complex proteins requiring specific glycosylation patterns. Plasmid DNA transfection and stable cell lines are frequently used for recombinant protein production, but they are expensive at large scale or can become time-consuming, respectively. The BacMam baculovirus (BV) is a safe and cost-effective platform to produce recombinant proteins in mammalian cells. The process of generating BacMam BVs is straightforward and similar to the generation of \"insect\" BVs, with different commercially available platforms. Although there are several protocols that describe recombinant protein expression with the BacMam BV in adherent cell lines, limited information is available on suspension cells. Therefore, it is of relevance to define the conditions to produce recombinant proteins in suspension cell cultures with BacMam BVs that facilitate bioprocess transfer to larger volumes. Here, we describe a method to generate a high titer BacMam BV stock and produce recombinant proteins in suspension HEK293 cells.

The CRISPR-Cas9 system is a revolutionary tool in genetic engineering, offering unprecedented precision and efficiency in genome editing. Cas9, an enzyme derived from bacteria, is guided by RNA to edit DNA sequences within cells precisely. However, while CRISPR-Cas9 presents notable benefits and encouraging outcomes as a molecular tool and a potential therapeutic agent, the process of producing and purifying recombinant Cas9 protein remains a formidable hurdle. In this study, we systematically investigated the expression of recombinant SpCas9-His in four distinct Escherichia coli (E. coli) strains (Rosetta2, BL21(DE3), BL21(DE3)-pLysS, and BL21(DE3)-Star). Through optimization of culture conditions, including temperature and post-induction time, the BL21(DE3)-pLysS strain demonstrated efficient SpCas9 protein expression. This study also presents a detailed protocol for the purification of recombinant SpCas9, along with detailed troubleshooting tips. Results indicate successful SpCas9 protein expression using E. coli BL21(DE3)-pLysS at 0.5 mM IPTG concentration. Furthermore, the findings suggest potential avenues for further enhancements, paving the way for large-scale Cas9 production. This research contributes valuable insights into optimizing E. coli strains and culture conditions for enhanced Cas9 expression, offering a step forward in the development of efficient genome editing tools and therapeutic proteins.

With an increasing number of blockbuster drugs being recombinant mammalian proteins, protein production platforms that focus on mammalian proteins have had a profound impact in many areas of basic and applied research. Many groups, both academic and industrial, have been focusing on developing cost-effective methods to improve the production of mammalian proteins that would support potential therapeutic applications. As it stands, while a wide range of platforms have been successfully developed for laboratory use, the majority of biologicals are still produced in mammalian cell lines due to the requirement for posttranslational modification and the biosynthetic complexity of target proteins. An unbiased high-throughput RNAi screening approach can be an efficient tool to identify target genes involved in recombinant protein production. Here, we describe the process of optimizing the transfection conditions, performing the genome-wide siRNA screen, the activity and cell viability assays, and the validation transfection to identify genes involved with protein expression.

The production of recombinant proteins has helped in understanding of their function and developing new therapies. However, one of the major bottlenecks for protein production is the establishment of reliable mammalian cell lines with high expression levels. In this chapter, we describe a simple and robust system that allows for the quick establishment of stable transgenic 293 cell lines with reproducible and high protein expression levels. This methodology is based on the piggyBac transposon system and enables the inducible production of the protein of interest. Finally, this methodology can easily be used in conventional laboratory cell culture settings without requiring specialized devices.

BACKGROUND: The process of producing proteins in bacterial systems and secreting them through ATP-binding cassette (ABC) transporters is an area that has been actively researched and used due to its high protein production capacity and efficiency. However, some proteins are unable to pass through the ABC transporter after synthesis, a phenomenon we previously determined to be caused by an excessive positive charge in certain regions of their amino acid sequence. If such an excessive charge is removed, the secretion of any protein through ABC transporters becomes possible.
RESULTS: In this study, we introduce \'linear charge density\' as the criteria for possibility of protein secretion through ABC transporters and confirm that this criterion can be applied to various non-secretable proteins, such as SARS-CoV-2 spike proteins, botulinum toxin light chain, and human growth factors. Additionally, we develop a new algorithm, PySupercharge, that enables the secretion of proteins containing regions with high linear charge density. It selectively converts positively charged amino acids into negatively charged or neutral amino acids after linear charge density analysis to enable protein secretion through ABC transporters.
CONCLUSIONS: PySupercharge, which also minimizes functional/structural stability loss of the pre-mutation proteins through the use of sequence conservation data, is currently being operated on an accessible web server. We verified the efficacy of PySupercharge-driven protein supercharging by secreting various previously non-secretable proteins commonly used in research, and so suggest this tool for use in future research requiring effective protein production.

(1) Recombinant protein production in mammalian cells is either based on transient transfection processes, often inefficient and underlying high batch-to-batch variability, or on laborious generation of stable cell lines. Alternatively, BacMam, a transduction process using the baculovirus, can be employed. (2) Six transfecting agents were compared to baculovirus transduction in terms of transient and stable protein expression characteristics of the model protein ACE2-eGFP using HEK293-6E, CHO-K1, and Vero cell lines. Furthermore, process optimization such as expression enhancement using sodium butyrate and TSA or baculovirus purification was assessed. (3) Baculovirus transduction efficiency was superior to all transfection agents for all cell lines. Transduced protein expression was moderate, but an 18-fold expression increase was achieved using the enhancer sodium butyrate. Ultracentrifugation of baculovirus from a 3.5 L bioreactor significantly improved the transduction efficiency and protein expression. Stable cell lines were obtained with each baculovirus transduction, yet stable cell line generation after transfection was highly unreliable. (4) This study demonstrated the superiority of the BacMam platform to standard transfections. The baculovirus efficiently transduced an array of cell lines both transiently and stably and achieved the highest efficiency for all tested cell lines. The feasibility of the scale-up of baculovirus production was demonstrated and the possibility of baculovirus purification was successfully explored.