Duchenne muscular dystrophy (DMD) is a devastating disease affecting about 1 out of 5000 male births and caused by mutations in the dystrophin gene. Genome editing has the potential to restore expression of a modified dystrophin gene from the native locus to modulate disease progression. In this study, adeno-associated virus was used to deliver the CRISPR/Cas9 system to the mdx mouse model of DMD to remove the mutated exon 23 from the dystrophin gene. This includes local and systemic delivery to adult mice and systemic delivery to neonatal mice. Exon 23 deletion by CRISPR/Cas9 resulted in expression of the modified dystrophin gene, partial recovery of functional dystrophin protein in skeletal myofibers and cardiac muscle, improvement of muscle biochemistry, and significant enhancement of muscle force. This work establishes CRISPR/Cas9-based genome editing as a potential therapy to treat DMD.

We report the improved production of recombinant proteins in E. coli , reliant on tightly controlled autoinduction, triggered by phosphate depletion in stationary phase. The method, reliant on engineered strains and plasmids, enables improved protein expression across scales.Expression levels using this approach have reached as high as 55% of total cellular protein.Initial use of the method in instrumented fed batch fermentations enables cell densities of~30 grams dry cell weight (gCDW) per liter and protein titers up to 8.1+/-0.7 g/L (~270 mg/gCDW).The process has also been adapted to an optimized autoinduction media, enabling routine batch production at culture volumes of 20 L (384 well plates), 100 L (96 well plates), 20 mL and 100 mL. In batch cultures, cells densities routinely reach~5-7 gCDW per liter, offering protein titers above 2 g/L. The methodology has been validated with a set of diverse heterologous proteins and is of general use for the facile optimization of routine protein expression from high throughput screens to fed-batch fermentation.

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Phage-derived "recombineering" methods are utilized for bacterial genome editing. Recombineering results in a heterogeneous population of modified and unmodified chromosomes, and therefore selection methods, such as CRISPR-Cas9, are required to select for edited clones. Cells can evade CRISPR-Cas-induced cell death through recA-mediated induction of the SOS response. The SOS response increases RecA dependent repair as well as mutation rates through induction of the umuDC error prone polymerase. As a result, CRISPR-Cas selection is more efficient in recA mutants. We report an approach to inhibiting the SOS response and RecA activity through the expression of a mutant dominant negative form of RecA, which incorporates into wild type RecA filaments and inhibits activity. Using a plasmid-based system in which Cas9 and recA mutants are coexpressed, we can achieve increased efficiency and consistency of CRISPR-Cas9-mediated selection and recombineering in E. coli, while reducing the induction of the SOS response. To date, this approach has been shown to be independent of recA genotype and host strain lineage. Using this system, we demonstrate increased CRISPR-Cas selection efficacy with over 10 000 guides covering the E. coli chromosome. The use of dominant negative RecA or homologues may be of broad use in bacterial CRISPR-Cas-based genome editing where the SOS pathways are present.

A key challenge in synthetic biology is the successful utilization of characterized parts, such as promoters, in different biological contexts. We report the robustness testing of a small library of E. coli PhoB regulated promoters that enable heterologous protein production in two-stage cultures. Expression levels were measured both in a rich Autoinduction Broth as well as a minimal mineral salts media. Media dependent differences were promoter dependent. 4 out of 16 promoters tested were identified to have tightly controlled expression which was also robust to media formulation. Improved promoter robustness led to more predictable scale up and consistent expression in instrumented bioreactors. This subset of PhoB activated promoters, useful for two-stage autoinduction, highlight the impact of the environment on the performance of biological parts, and the importance of robustness testing in synthetic biology.

We report improved release of recombinant proteins in Escherichia coli, which relies on combined cellular autolysis and DNA/RNA autohydrolysis, conferred by the tightly controlled autoinduction of both phage lysozyme and the nonspecific DNA/RNA endonuclease from Serratia marcescens. Autoinduction occurs in a two‐stage process wherein heterologous protein expression and autolysis enzymes are induced upon entry into stationary phase by phosphate depletion. Cytoplasmic lysozyme and periplasmic endonuclease are kept from inducing lysis until membrane integrity is disrupted. After cell harvest, the addition of detergent (0.1% Triton X‐100) and a single 30 min freeze‐thaw cycle results in >90% release of protein, green fluorescent protein. This cellular lysis is accompanied by complete oligonucleotide hydrolysis. The approach has been validated for shake flask cultures, high‐throughput cultivation in microtiter plates, and larger scale stirred‐tank bioreactors. This tightly controlled system enables robust growth and resistance to lysis in routine media when cells are propagated and autolysis/hydrolysis genes are only induced upon phosphate depletion.

CRISPR/Cas systems have become ubiquitous for genome editing in eukaryotic as well as bacterial systems. Cas9 associated with a guide RNA (gRNA) searches DNA for a matching sequence (target site) next to a protospacer adjacent motif (PAM) and once found, cuts the DNA.The number of PAM sites in the genome are effectively a non-target pool of inhibitory substrates, competing with the target site for the Cas9/gRNA complex. We demonstrate that increasing the number of non-target sites for a given gRNA reduces on-target activity in a dose dependent manner. Furthermore, we show that the use of Cas9 mutants with increased PAM specificity towards a smaller subset of PAMs (or smaller pool of competitive substrates) improves cutting rates. Decreasing the non-target pool by increasing PAM specificity provides a path towards improving on-target activity for slower high fidelity Cas9 variants. These results demonstrate the importance of competitive non-target sites on Cas9 activity and, in part, may help to explain sequence and context dependent activities of gRNAs. Engineering improved PAM specificity to reduce the competitive non-target pool offers an alternative strategy to engineer Cas9 variants with increased specificity and maintained on-target activity.

We report the improved production of recombinant proteins in E. coli , reliant on tightly controlled autoinduction, triggered by phosphate depletion in stationary phase. The method, reliant on engineered strains and plasmids, enables improved protein expression across scales.Expression levels using this approach have reached as high as 55% of total cellular protein.Initial use of the method in instrumented fed batch fermentations enables cell densities of 10 grams dry cell weight (gCDW) per liter and protein titers up to 2.7+/-0.2 g/L (270 mg/gCDW).The process has also been adapted to an optimized autoinduction media, enabling routine batch production at culture volumes of 20 L (384 well plates), 100 L (96 well plates), 20 mL and 100 mL. In batch cultures, cells densities routinely reach~5-7 gCDW per liter, offering protein titers above 2 g/L. The methodology has been validated with a set of diverse heterologous proteins and is of general use for the facile optimization of routine protein expression from high throughput screens to fed-batch fermentation Highlights :• Stationary phase protein expression results in high titers.• Autoinduction by phosphate depletion enables protein titers from 2-3 g/L.• Autoinduction has been validated from 384 well plates to instrumented bioreactors.