Summary It is generally assumed that the allocation and synthesis of total cellular resources in microorganisms are uniquely determined by the growth conditions. Adaptation to a new physiological state leads to a change in cell size via reallocation of cellular resources. However, it has not been understood how cell size is coordinated with biosynthesis and robustly adapts to physiological states. We show that cell size in Escherichia coli can be predicted for any steady-state condition by projecting all biosynthesis into three measurable variables representing replication initiation, replication-division cycle, and the global biosynthesis rate. These variables can be decoupled by selectively controlling their respective core biosynthesis using CRISPR interference and antibiotics, verifying our predictions that different physiological states can result in the same cell size. We performed extensive growth inhibition experiments, and discovered that cell size at replication initiation per origin, namely the initiation mass or “unit cell,” is remarkably invariant under perturbations targeting transcription, translation, ribosome content, replication kinetics, fatty acid and cell-wall synthesis, cell division, and cell shape. Based on this invariance and balanced resource allocation, we explain why the total cell size is the sum of all unit cells. These results provide an overarching framework with quantitative predictive power over cell size in bacteria.

Recombination with single-strand DNA oligonucleotides (oligos) in E. coli is an efficient and rapid way to modify replicons in vivo. The generation of a nucleotide alteration by oligo recombination provides novel assays for studying cellular processes. Single-strand exonucleases inhibit oligo recombination, and by mutating all four known exonucleases recombination is increased. Increasing the oligo concentration or addition of non-specific carrier oligo titrates out the exonucleases. In a model for oligo recombination, λ Beta protein anneals the oligo to complementary single-strand DNA at the replication fork. Mismatches are created and the methyldirected mismatch repair (MMR) system acts to eliminate the mismatches inhibiting recombination. Three ways to evade MMR through oligo design include, in addition to the desired change 1) a C~C mismatch six bp from that change, 2) four or more adjacent mismatches, or 3) mismatches at four or more consecutive wobble positions. The latter proves useful for making high frequency changes that alter only the target amino-acid sequence and even allows modification of essential genes. Efficient uptake of DNA is important for oligo-mediated recombination. Uptake of oligos or plasmids is growth media-dependent and is 10,000-fold reduced for cells grown in minimal vs rich medium. Genome-wide engineering technologies utilizing recombineering will benefit from both optimized recombination frequencies and a greater understanding of how biological processes such as DNA replication and cell division impact recombinants formed at multiple chromosomal loci. Recombination events at multiple loci in individual cells are described here.

The two-step process of selection and counter-selection is a standard way to enable genetic modification and engineering of bacterial genomes using homologous recombination methods. The tetA and sacB genes are contained in a DNA cassette and confer a novel dual counter-selection system. Expression of tetA confers bacterial resistance to tetracycline (TcR) and also causes sensitivity to the lipophillic chelator fusaric acid; sacB causes sensitivity to sucrose. These two genes are introduced as a joint DNA cassette into Escherichia coli by selection for TcR. A medium containing both fusaric acid and sucrose has been developed, in which, coexpression of tetA-sacB is orders of magnitude more sensitive as a counter-selection agent than either gene alone. In conjunction with the homologous recombination methods of recombineering and P1 transduction, this powerful system has been used to select changes in the bacterial genome that cannot be directly detected by other counter-selection systems.

The bacterial chromosome and bacterial plasmids can be engineered in vivo by homologous recombination using PCR products and synthetic oligonucleotides as substrates. This is possible because bacteriophage-encoded recombination proteins efficiently recombine sequences with homologies as short as 35 to 50 bases. Recombineering allows DNA sequences to be inserted or deleted without regard to location of restriction sites. This unit first describes preparation of electrocompetent cells expressing the recombineering functions and their transformation with dsDNA or ssDNA. It then presents support protocols that describe several two-step selection/counter-selection methods of making genetic alterations without leaving any unwanted changes in the targeted DNA, and a method for retrieving onto a plasmid a genetic marker (cloning by retrieval) from the Escherichia coli chromosome or a co-electroporated DNA fragment. Additional protocols describe methods to screen for unselected mutations, removal of the defective prophage from recombineering strains, and other useful techniques.

The environmental toxin TCDD (2,3,7,8-tetrachlorodibenzop-dioxin, dioxin) produces diverse toxic effects including a lethal wasting syndrome whose hallmark is suppressed hepatic gluconeogenesis. All TCDD toxicities require activation of the aryl hydrocarbon receptor (AHR), a ligand-activated transcription factor. Whereas the mechanism for AHR induction of target genes is well understood, it is not known how AHR activation produces any TCDD toxicity. This report identifies for the first time an AHR target gene, TiPARP (TCDD-inducible poly(ADPribose) polymerase, PARP7) that can mediate a TCDD toxicity, i.e. suppression of hepatic gluconeogenesis. TCDD suppressed hepatic glucose production, expression of key gluconeogenic genes, phosphoenolpyruvate carboxykinase (PEPCK), and glucose-6-phosphatase (G6Pase), and NAD ؉ levels, and increased PARP activity and

The ability to control the level of gene expression is a major quest in biology. A widely used approach employs deletion of a nonessential gene of interest (knockout), or multi-step recombineering to move a gene of interest under a repressible promoter (knockdown). However, these genetic methods are laborious, and limited for quantitative study. Here, we report a tunable CRISPR-cas system, “tCRISPRi”, for precise and continuous titration of gene expression by more than 30-fold. Our tCRISPRi system employs various previous advancements into a single strain: (1) We constructed a new strain containing a tunable arabinose operon promoter PBAD to quantitatively control the expression of CRISPR-(d)Cas protein over two orders of magnitude in a plasmid-free system. (2) tCRISPRi is reversible, and gene expression is repressed under knockdown conditions. (3) tCRISPRi shows significantly less than 10% leaky expression. (4) Most important from a practical perspective, construction of tCRISPRi to target a new gene requires only one-step of oligo recombineering. Our results show that tCRISPRi, in combination with recombineering, provides a simple and easy-to-implement tool for gene expression control, and is ideally suited for construction of both individual strains and high-throughput tunable knockdown libraries.

Targeted gene repair mediated by single-stranded oligonucleotides (SSOs) has great potential for use in functional genomic studies and gene therapy. Genetic changes have been created using this approach in a number of prokaryotic and eukaryotic systems, including mouse embryonic stem cells. However, the underlying mechanisms remain to be fully established. In one of the current models, the ‘annealing-integration’ model, the SSO anneals to its target locus at the replication fork, serving as a primer for subsequent DNA synthesis mediated by the host replication machinery. Using a λ-Red recombination-based system in the bacterium Escherichia coli, we systematically examined several fundamental premises that form the mechanistic basis of this model. Our results provide direct evidence strongly suggesting that SSO-mediated gene repair is mechanistically linked to the process of DNA replication, and most likely involves a replication intermediate. These findings will help guide future experiments involving SSO-mediated gene repair in mammalian and prokaryotic cells, and suggest several mechanisms by which the efficiencies may be reliably and substantially increased.

SummarySynthetic single-strand oligonucleotides (oligos) with homology to genomic DNA have proved to be highly effective for constructing designed mutations in targeted genomes, a process referred to as recombineering. The cellular functions important for this type of homologous recombination have yet to be determined. Towards this end, we have identified Escherichia coli functions that process the recombining oligo and affect bacteriophage l Red-mediated oligo recombination. To determine the nature of oligo processing during recombination, each oligo contained multiple nucleotide changes: a single base change allowing recombinant selection, and silent changes serving as genetic markers to determine the extent of oligo processing during the recombination. Such oligos were often not incorporated into the host chromosome intact; many were partially degraded in the process of recombination. The position and number of these silent nucleotide changes within the oligo strongly affect both oligo processing and recombination frequency. Exonucleases, especially those associated with DNA Polymerases I and III, affect inheritance of the silent nucleotide changes in the oligos. We demonstrate for the first time that the major DNA polymerases (Pol I and Pol III) and DNA ligase are directly involved with oligo recombination.