CRISPRi screen for enhancing heterologous α-amylase yield in Bacillus subtilis.

Yield improvements in cell factories can potentially be obtained by fine-tuning the regulatory mechanisms for gene candidates. In pursuit of such candidates, we performed RNA-sequencing of two α-amylase producing Bacillus strains and predict hundreds of putative novel non-coding transcribed regions. Surprisingly, we found among hundreds of non-coding and structured RNA candidates that non-coding genomic regions are proportionally undergoing the highest changes in expression during fermentation. Since these classes of RNA are also understudied, we targeted the corresponding genomic regions with CRIPSRi knockdown to test for any potential impact on the yield. From differentially expression analysis, we selected 53 non-coding candidates. Although CRISPRi knockdowns target both the sense and the antisense strand, the CRISPRi experiment cannot link causes for yield changes to the sense or antisense disruption. Nevertheless, we observed on several instances with strong changes in enzyme yield. The knockdown targeting the genomic region for a putative antisense RNA of the 3' UTR of the skfA-skfH operon led to a 21% increase in yield. In contrast, the knockdown targeting the genomic regions of putative antisense RNAs of the cytochrome c oxidase subunit 1 (ctaD), the sigma factor sigH, and the uncharacterized gene yhfT decreased yields by 31 to 43%.

Flagella disruption in Bacillus subtilis increases amylase production yield.

BACKGROUND: Bacillus subtilis is a Gram-positive bacterium used as a cell factory for protein production. Over the last decades, the continued optimization of production strains has increased yields of enzymes, such as amylases, and made commercial applications feasible. However, current yields are still significantly lower than the theoretically possible yield based on the available carbon sources. In its natural environment, B. subtilis can respond to unfavorable growth conditions by differentiating into motile cells that use flagella to swim towards available nutrients. RESULTS: In this study, we analyze existing transcriptome data from a B. subtilis α-amylase production strain at different time points during a 5-day fermentation. We observe that genes of the fla/che operon, essential for flagella assembly and motility, are differentially expressed over time. To investigate whether expression of the flagella operon affects yield, we performed CRISPR-dCas9 based knockdown of the fla/che operon with sgRNA target against the genes flgE, fliR, and flhG, respectively. The knockdown resulted in inhibition of mobility and a striking 2-threefold increase in α-amylase production yield. Moreover, replacing flgE (required for flagella hook assembly) with an erythromycin resistance gene followed by a transcription terminator increased α-amylase yield by about 30%. Transcript levels of the α-amylase were unaltered in the CRISPR-dCas9 knockdowns as well as the flgE deletion strain, but all manipulations disrupted the ability of cells to swim on agar. CONCLUSIONS: We demonstrate that the disruption of flagella in a B. subtilis α-amylase production strain, either by CRISPR-dCas9-based knockdown of the operon or by replacing flgE with an erythromycin resistance gene followed by a transcription terminator, increases the production of α-amylase in small-scale fermentation.

Nuclear retention prevents premature cytoplasmic appearance of mRNA.

In S. cerevisiae cells debilitated in mRNA nuclear export, transcripts are retained in nuclear foci ("dots"). The ultimate fate of dot-mRNA has remained elusive. Here, we use single molecule counting microscopy and (35)S-methionine pulse-labeling assays to quantify cytoplasmic HSP104 RNA levels and estimate HSP104 RNA translation status. HSP104 transcripts, retained in dots as a consequence of the mex67-5 mutation, are slowly released over time for cytoplasmic translation. Thus, dot-mRNA retains function. However, forcing its nuclear export, by overexpressing the Sub2p mRNA export factor, does not elevate Hsp104p protein levels but is instead paralleled by growth deficiency. Nuclear export and growth phenotypes are both counteracted by coexpressing the nuclear RNA quality control factor Rrp6p. Thus, prematurely released dot-mRNA is translationally inactive and possibly toxic. Accordingly, nuclear retention of mRNA may serve a precautionary role during stressful situations such as, e.g., decreased mRNA maturation competence.

Accelerating genome editing in CHO cells using CRISPR Cas9 and CRISPy, a web-based target finding tool.

Chinese hamster ovary (CHO) cells are widely used in the biopharmaceutical industry as a host for the production of complex pharmaceutical proteins. Thus genome engineering of CHO cells for improved product quality and yield is of great interest. Here, we demonstrate for the first time the efficacy of the CRISPR Cas9 technology in CHO cells by generating site-specific gene disruptions in COSMC and FUT8, both of which encode proteins involved in glycosylation. The tested single guide RNAs (sgRNAs) created an indel frequency up to 47.3% in COSMC, while an indel frequency up to 99.7% in FUT8 was achieved by applying lectin selection. All eight sgRNAs examined in this study resulted in relatively high indel frequencies, demonstrating that the Cas9 system is a robust and efficient genome-editing methodology in CHO cells. Deep sequencing revealed that 85% of the indels created by Cas9 resulted in frameshift mutations at the target sites, with a strong preference for single base indels. Finally, we have developed a user-friendly bioinformatics tool, named "CRISPy" for rapid identification of sgRNA target sequences in the CHO-K1 genome. The CRISPy tool identified 1,970,449 CRISPR targets divided into 27,553 genes and lists the number of off-target sites in the genome. In conclusion, the proven functionality of Cas9 to edit CHO genomes combined with our CRISPy database have the potential to accelerate genome editing and synthetic biology efforts in CHO cells.

BSGatlas: a unified Bacillus subtilis genome and transcriptome annotation atlas with enhanced information access.

A large part of our current understanding of gene regulation in Gram-positive bacteria is based on Bacillus subtilis, as it is one of the most well studied bacterial model systems. The rapid growth in data concerning its molecular and genomic biology is distributed across multiple annotation resources. Consequently, the interpretation of data from further B. subtilis experiments becomes increasingly challenging in both low- and large-scale analyses. Additionally, B. subtilis annotation of structured RNA and non-coding RNA (ncRNA), as well as the operon structure, is still lagging behind the annotation of the coding sequences. To address these challenges, we created the B. subtilis genome atlas, BSGatlas, which integrates and unifies multiple existing annotation resources. Compared to any of the individual resources, the BSGatlas contains twice as many ncRNAs, while improving the positional annotation for 70 % of the ncRNAs. Furthermore, we combined known transcription start and termination sites with lists of known co-transcribed gene sets to create a comprehensive transcript map. The combination with transcription start/termination site annotations resulted in 717 new sets of co-transcribed genes and 5335 untranslated regions (UTRs). In comparison to existing resources, the number of 5' and 3' UTRs increased nearly fivefold, and the number of internal UTRs doubled. The transcript map is organized in 2266 operons, which provides transcriptional annotation for 92 % of all genes in the genome compared to the at most 82 % by previous resources. We predicted an off-target-aware genome-wide library of CRISPR-Cas9 guide RNAs, which we also linked to polycistronic operons. We provide the BSGatlas in multiple forms: as a website (https://rth.dk/resources/bsgatlas/), an annotation hub for display in the UCSC genome browser, supplementary tables and standardized GFF3 format, which can be used in large scale -omics studies. By complementing existing resources, the BSGatlas supports analyses of the B. subtilis genome and its molecular biology with respect to not only non-coding genes but also genome-wide transcriptional relationships of all genes.

Ribosome profiling-guided depletion of an mRNA increases cell growth rate and protein secretion.

Recombinant protein production coopts the host cell machinery to provide high protein yields of industrial enzymes or biotherapeutics. However, since protein translation is energetically expensive and tightly controlled, it is unclear if highly expressed recombinant genes are translated as efficiently as host genes. Furthermore, it is unclear how the high expression impacts global translation. Here, we present the first genome-wide view of protein translation in an IgG-producing CHO cell line, measured with ribosome profiling. Through this we found that our recombinant mRNAs were translated as efficiently as the host cell transcriptome, and sequestered up to 15% of the total ribosome occupancy. During cell culture, changes in recombinant mRNA translation were consistent with changes in transcription, demonstrating that transcript levels influence specific productivity. Using this information, we identified the unnecessary resistance marker NeoR to be a highly transcribed and translated gene. Through siRNA knock-down of NeoR, we improved the production- and growth capacity of the host cell. Thus, ribosomal profiling provides valuable insights into translation in CHO cells and can guide efforts to enhance protein production.

Endoplasmic reticulum-directed recombinant mRNA displays subcellular localization equal to endogenous mRNA during transient expression in CHO cells.

When expressing pharmaceutical recombinant proteins in mammalian cells, the protein is commonly directed through the secretory pathway, in a signal peptide-dependent manner, to acquire specific post-translational modifications and to facilitate secretion into the culture medium. One key premise for this is the direction of the mRNA encoding the recombinant protein to the surface of the endoplasmic reticulum (ER) for subsequent protein translocation into the secretory pathway. To evaluate the efficiency of this process in Chinese hamster ovary (CHO) cells, the subcellular localization of recombinant mRNA encoding the therapeutic proteins, erythropoietin (EPO) and Rituximab, was determined. The results show that ER-directed recombinant mRNAs exhibited an efficient recruitment to the ER when compared to an endogenous ER-directed mRNA, with no cytoplasmic translation of ER-directed recombinant proteins observed. These observations indicate that the recombinant mRNA, encoding ER-directed proteins, follows the same distribution pattern as endogenous mRNA directed towards the ER. Furthermore, the previous established fractionation method proves to be an efficient tool to study not only recombinant mRNA localization, but also recombinant protein trafficking between the ER and cytosol in CHO cells.

Development of a VHH-Based Erythropoietin Quantification Assay.

Erythropoietin (EPO) quantification during cell line selection and bioreactor cultivation has traditionally been performed with ELISA or HPLC. As these techniques suffer from several drawbacks, we developed a novel EPO quantification assay. A camelid single-domain antibody fragment directed against human EPO was evaluated as a capturing antibody in a label-free biolayer interferometry-based quantification assay. Human recombinant EPO can be specifically detected in Chinese hamster ovary cell supernatants in a sensitive and pH-dependent manner. This method enables rapid and robust quantification of EPO in a high-throughput setting.

Site-specific integration in CHO cells mediated by CRISPR/Cas9 and homology-directed DNA repair pathway.

Chinese hamster ovary (CHO) cells are the most widely used mammalian hosts for production of therapeutic proteins. However, development of recombinant CHO cell lines has been hampered by unstable and variable transgene expression caused by random integration. Here we demonstrate efficient targeted gene integration into site-specific loci in CHO cells using CRISPR/Cas9 genome editing system and compatible donor plasmid harboring a gene of interest (GOI) and short homology arms. This strategy has enabled precise insertion of a 3.7-kb gene expression cassette at defined loci in CHO cells following a simple drug-selection, resulting in homogeneous transgene expression. Taken together, the results displayed here can help pave the way for the targeting of GOI to specific loci in CHO cells, increasing the likelihood of generating isogenic cell lines with consistent protein production.

One-step generation of triple knockout CHO cell lines using CRISPR/Cas9 and fluorescent enrichment.

The CRISPR/Cas9 genome editing technology has previously been shown to be a highly efficient tool for generating gene disruptions in CHO cells. In this study we further demonstrate the applicability and efficiency of CRISPR/Cas9 genome editing by disrupting FUT8, BAK and BAX simultaneously in a multiplexing setup in CHO cells. To isolate Cas9-expressing cells from transfected cell pools, GFP was linked to the Cas9 nuclease via a 2A peptide. With this method, the average indel frequencies generated at the three genomic loci were increased from 11% before enrichment to 68% after enrichment. Despite the high number of genome editing events in the enriched cell pools, no significant off-target effects were observed from off-target prediction followed by deep sequencing. Single cell sorting of enriched multiplexed cells and deep sequencing of 97 clones revealed the presence of four single, 23 double and 34 triple gene-disrupted cell lines. Further characterization of selected potential triple knockout clones confirmed the removal of Bak and Bax protein and disrupted fucosylation activity as expected. The knockout cell lines showed improved resistance to apoptosis compared to wild-type CHO-S cells. Taken together, multiplexing with CRISPR/Cas9 can accelerate genome engineering efforts in CHO cells even further.