Duane A. Chung

Tumor cell traffic through the extracellular matrix is controlled by the membrane-anchored collagenase MT1-MMP

As cancer cells traverse collagen-rich extracellular matrix (ECM) barriers and intravasate, they adopt a fibroblast-like phenotype and engage undefined proteolytic cascades that mediate invasive activity. Herein, we find that fibroblasts and cancer cells express an indistinguishable pericellular collagenolytic activity that allows them to traverse the ECM. Using fibroblasts isolated from gene-targeted mice, a matrix metalloproteinase (MMP)–dependent activity is identified that drives invasion independently of plasminogen, the gelatinase A/TIMP-2 axis, gelatinase B, collagenase-3, collagenase-2, or stromelysin-1. In contrast, deleting or suppressing expression of the membrane-tethered MMP, MT1-MMP, in fibroblasts or tumor cells results in a loss of collagenolytic and invasive activity in vitro or in vivo. Thus, MT1-MMP serves as the major cell-associated proteinase necessary to confer normal or neoplastic cells with invasive activity.

Coupling the CRISPR/Cas9 System with Lambda Red Recombineering Enables Simplified Chromosomal Gene Replacement in Escherichia coli.

ABSTRACT To date, most genetic engineering approaches coupling the type II Streptococcus pyogenes clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 system to lambda Red recombineering have involved minor single nucleotide mutations. Here we show that procedures for carrying out more complex chromosomal gene replacements in Escherichia coli can be substantially enhanced through implementation of CRISPR/Cas9 genome editing. We developed a three-plasmid approach that allows not only highly efficient recombination of short single-stranded oligonucleotides but also replacement of multigene chromosomal stretches of DNA with large PCR products. By systematically challenging the proposed system with respect to the magnitude of chromosomal deletion and size of DNA insertion, we demonstrated DNA deletions of up to 19.4 kb, encompassing 19 nonessential chromosomal genes, and insertion of up to 3 kb of heterologous DNA with recombination efficiencies permitting mutant detection by colony PCR screening. Since CRISPR/Cas9-coupled recombineering does not rely on the use of chromosome-encoded antibiotic resistance, or flippase recombination for antibiotic marker recycling, our approach is simpler, less labor-intensive, and allows efficient production of gene replacement mutants that are both markerless and “scar”-less.

Development of an electrotransformation protocol for genetic manipulation of Clostridium pasteurianum.

BackgroundReducing the production cost of, and increasing revenues from, industrial biofuels will greatly facilitate their proliferation and co-integration with fossil fuels. The cost of feedstock is the largest cost in most fermentation bioprocesses and therefore represents an important target for cost reduction. Meanwhile, the biorefinery concept advocates revenue growth through complete utilization of by-products generated during biofuel production. Taken together, the production of biofuels from low-cost crude glycerol, available in oversupply as a by-product of bioethanol production, in the form of thin stillage, and biodiesel production, embodies a remarkable opportunity to advance affordable biofuel development. However, few bacterial species possess the natural capacity to convert glycerol as a sole source of carbon and energy into value-added bioproducts. Of particular interest is the anaerobe Clostridium pasteurianum, the only microorganism known to convert glycerol alone directly into butanol, which currently holds immense promise as a high-energy biofuel and bulk chemical. Unfortunately, genetic and metabolic engineering of C. pasteurianum has been fundamentally impeded due to lack of an efficient method for deoxyribonucleic acid (DNA) transfer.ResultsThis work reports the development of an electrotransformation protocol permitting high-level DNA transfer to C. pasteurianum ATCC 6013 together with accompanying selection markers and vector components. The CpaAI restriction-modification system was found to be a major barrier to DNA delivery into C. pasteurianum which we overcame by in vivo methylation of the recognition site (5’-CGCG-3’) using the M.FnuDII methyltransferase. With proper selection of the replication origin and antibiotic-resistance marker, we initially electroporated methylated DNA into C. pasteurianum at a low efficiency of 2.4 × 101 transformants μg-1 DNA by utilizing conditions common to other clostridial electroporations. Systematic investigation of various parameters involved in the cell growth, washing and pulse delivery, and outgrowth phases of the electrotransformation procedure significantly elevated the electrotransformation efficiency, up to 7.5 × 104 transformants μg-1 DNA, an increase of approximately three order of magnitude. Key factors affecting the electrotransformation efficiency include cell-wall-weakening using glycine, ethanol-mediated membrane solubilization, field strength of the electric pulse, and sucrose osmoprotection.ConclusionsC. pasteurianum ATCC 6013 can be electrotransformed at a high efficiency using appropriately methylated plasmid DNA. The electrotransformation method and tools reported here should promote extensive genetic manipulation and metabolic engineering of this biotechnologically important bacterium.

Harnessing heterologous and endogenous CRISPR-Cas machineries for efficient markerless genome editing in Clostridium.

Application of CRISPR-Cas9 systems has revolutionized genome editing across all domains of life. Here we report implementation of the heterologous Type II CRISPR-Cas9 system in Clostridium pasteurianum for markerless genome editing. Since 74% of species harbor CRISPR-Cas loci in Clostridium, we also explored the prospect of co-opting host-encoded CRISPR-Cas machinery for genome editing. Motivation for this work was bolstered from the observation that plasmids expressing heterologous cas9 result in poor transformation of Clostridium. To address this barrier and establish proof-of-concept, we focus on characterization and exploitation of the C. pasteurianum Type I-B CRISPR-Cas system. In silico spacer analysis and in vivo interference assays revealed three protospacer adjacent motif (PAM) sequences required for site-specific nucleolytic attack. Introduction of a synthetic CRISPR array and cpaAIR gene deletion template yielded an editing efficiency of 100%. In contrast, the heterologous Type II CRISPR-Cas9 system generated only 25% of the total yield of edited cells, suggesting that native machinery provides a superior foundation for genome editing by precluding expression of cas9 in trans. To broaden our approach, we also identified putative PAM sequences in three key species of Clostridium. This is the first report of genome editing through harnessing native CRISPR-Cas machinery in Clostridium.

Technical guide for genetic advancement of underdeveloped and intractable Clostridium.

In recent years, the genus Clostridium has risen to the forefront of both medical biotechnology and industrial biotechnology owing to its potential in applications as diverse as anticancer therapy and production of commodity chemicals and biofuels. The prevalence of hyper-virulent strains of C. difficile within medical institutions has also led to a global epidemic that demands a more thorough understanding of clostridial genetics, physiology, and pathogenicity. Unfortunately, Clostridium suffers from a lack of sophisticated genetic tools and techniques which has hindered the biotechnological exploitation of this important bacterial genus. This review provides a comprehensive summary of biotechnological progress made in clostridial genetic tool development, while also aiming to serve as a technical guide for the advancement of underdeveloped clostridial strains, including recalcitrant species, novel environmental samples, and non-type strains. Relevant strain engineering techniques, from genome sequencing and establishment of a gene transfer methodology through to deployment of advanced genome editing procedures, are discussed in detail to provide a blueprint for future clostridial strain construction endeavors. It is expected that a more thorough and rounded-out genetic toolkit available for use in the clostridia will bring about the construction of superior bioprocessing strains and a more complete understanding of clostridial genetics, physiology, and pathogenicity.

Elimination of carbon catabolite repression in Clostridium acetobutylicum--a journey toward simultaneous use of xylose and glucose.

The industrial Gram-positive anaerobe Clostridium acetobutylicum is a valued acetone, butanol, and ethanol (ABE) solvent producer that is able to utilize a vast array of carbon sources in fermentation. When glucose is present in the growth medium, however, C. acetobutylicum, like many Gram-positive organisms, exhibits biphasic growth characteristics in which glucose is used preferentially over secondary carbon sources, a phenomenon known as carbon catabolite repression (CCR). The secondary carbon source is only utilized when the supply of glucose is exhausted, resulting in inefficient use of complex carbon sources. As biofuel production is sought from cheap feedstock, attention has turned to lignocellulosic biomass. Growth of C. acetobutylicum on lignocellulose, however, can be limited by CCR. Here, we present a method to relieve the inhibitory effect of CCR and allow simultaneous utilization of the lignocellulosic sugars of glucose and xylose by C. acetobutylicum. First, we utilized an in vivo gene reporter assay to demonstrate that an identified 14-nucleotide catabolite responsive element (CRE) sequence was sufficient to introduce CCR-mediated transcriptional inhibition, while subsequent mutation of the CRE sequence relieved the inhibitory effect. Next, we demonstrated that C. acetobutylicum harboring a CRE-less plasmid-borne xylose and pentose phosphate pathway operon afforded a 7.5-fold increase in xylose utilization in the presence of glucose as compared to a wild-type CRE plasmid-borne operon, effectively overcoming native CCR effects. The methodology presented here should translate to other members of Clostridium that exhibit CCR to enable simultaneous utilization of a vast array of carbon sources.

Expansion of the genetic toolkit for metabolic engineering of Clostridium pasteurianum: chromosomal gene disruption of the endogenous CpaAI restriction enzyme.

BackgroundClostridium pasteurianum is one of the most promising biofuel producers within the genus Clostridium owing to its unique metabolic ability to ferment glycerol into butanol. Although an efficient means is available for introducing foreign DNA to C. pasteurianum, major genetic tools, such as gene knockout, knockdown, or genome editing, are lacking, preventing metabolic engineering of C. pasteurianum.ResultsHere we present a methodology for performing chromosomal gene disruption in C. pasteurianum using the programmable lactococcus Ll.ltrB group II intron. Gene disruption was initially found to be impeded by inefficient electrotransformation of Escherichia coli-C. pasteurianum shuttle vectors, presumably due to host restriction. By assessing the ability of various vector deletion derivatives to electrotransform C. pasteurianum and probing the microorganism’s methylome using next-generation sequence data, we identified a new C. pasteurianum Type I restriction-methylation system, CpaAII, with a predicted recognition sequence of 5′-AAGNNNNNCTCC-3′ (N = A, C, G, or T). Following rescue of high-level electrotransformation via mutation of the sole CpaAII site within the shuttle vectors, we retargeted the intron to the cpaAIR gene encoding the CpaAI Type II restriction endonuclease (recognition site of 5′-CGCG-3′). Intron insertion was potentially hindered by low retrohoming efficiency, yet this limitation could be overcome by a procedure for enrichment of the intron insertion. The resulting ΔcpaAIR mutant strain was efficiently electrotransformed with M.FnuDII-unmethylated plasmid DNA.ConclusionsThe markerless and plasmidless ΔcpaAIR mutant strain of C. pasteurianum developed in this study can serve as a general host strain for future genetic and metabolic manipulation. Further, the associated gene disruption protocol should not only serve as a guide for chromosomal gene inactivation studies involving mobile group II introns, but also prove invaluable for applying metabolic engineering strategies to C. pasteurianum.

Extending CRISPR-Cas9 Technology from Genome Editing to Transcriptional Engineering in the Genus Clostridium

ABSTRACT The discovery and exploitation of the prokaryotic adaptive immunity system based on clustered regularly interspaced short palindromic repeats (CRISPRs) and CRISPR-associated (Cas) proteins have revolutionized genetic engineering. CRISPR-Cas tools have enabled extensive genome editing as well as efficient modulation of the transcriptional program in a multitude of organisms. Progress in the development of genetic engineering tools for the genus Clostridium has lagged behind that of many other prokaryotes, presenting the CRISPR-Cas technology an opportunity to resolve a long-existing issue. Here, we applied the Streptococcus pyogenes type II CRISPR-Cas9 (SpCRISPR-Cas9) system for genome editing in Clostridium acetobutylicum DSM792. We further explored the utility of the SpCRISPR-Cas9 machinery for gene-specific transcriptional repression. For proof-of-concept demonstration, a plasmid-encoded fluorescent protein gene was used for transcriptional repression in C. acetobutylicum. Subsequently, we targeted the carbon catabolite repression (CCR) system of C. acetobutylicum through transcriptional repression of the hprK gene encoding HPr kinase/phosphorylase, leading to the coutilization of glucose and xylose, which are two abundant carbon sources from lignocellulosic feedstocks. Similar approaches based on SpCRISPR-Cas9 for genome editing and transcriptional repression were also demonstrated in Clostridium pasteurianum ATCC 6013. As such, this work lays a foundation for the derivation of clostridial strains for industrial purposes. IMPORTANCE After recognizing the industrial potential of Clostridium for decades, methods for the genetic manipulation of these anaerobic bacteria are still underdeveloped. This study reports the implementation of CRISPR-Cas technology for genome editing and transcriptional regulation in Clostridium acetobutylicum, which is arguably the most common industrial clostridial strain. The developed genetic tools enable simpler, more reliable, and more extensive derivation of C. acetobutylicum mutant strains for industrial purposes. Similar approaches were also demonstrated in Clostridium pasteurianum, another clostridial strain that is capable of utilizing glycerol as the carbon source for butanol fermentation, and therefore can be arguably applied in other clostridial strains.

Improved Draft Genome Sequence of Clostridium pasteurianum Strain ATCC 6013 (DSM 525) Using a Hybrid Next-Generation Sequencing Approach

ABSTRACT We present an improved draft genome sequence for Clostridium pasteurianum strain ATCC 6013 (DSM 525), the type strain of the species and an important solventogenic bacterium with industrial potential. Availability of a near-complete genome sequence will enable strain engineering of this promising bacterium.

Disruption of the Reductive 1,3-Propanediol Pathway Triggers Production of 1,2-Propanediol for Sustained Glycerol Fermentation by Clostridium pasteurianum

ABSTRACT Crude glycerol, the major by-product of biodiesel production, is an attractive bioprocessing feedstock owing to its abundance, low cost, and high degree of reduction. In line with the advent of the biodiesel industry, Clostridium pasteurianum has gained prominence as a result of its unique capacity to convert waste glycerol into n-butanol, a high-energy biofuel. However, no efforts have been directed at abolishing the production of 1,3-propanediol (1,3-PDO), the chief competing product of C. pasteurianum glycerol fermentation. Here, we report rational metabolic engineering of C. pasteurianum for enhanced n-butanol production through inactivation of the gene encoding 1,3-PDO dehydrogenase (dhaT). In spite of current models of anaerobic glycerol dissimilation, culture growth and glycerol utilization were unaffected in the dhaT disruption mutant (dhaT::Ll.LtrB). Metabolite characterization of the dhaT::Ll.LtrB mutant revealed an 83% decrease in 1,3-PDO production, encompassing the lowest C. pasteurianum 1,3-PDO titer reported to date (0.58 g liter−1). With 1,3-PDO formation nearly abolished, glycerol was converted almost exclusively to n-butanol (8.6 g liter−1), yielding a high n-butanol selectivity of 0.83 g n-butanol g−1 of solvents compared to 0.51 g n-butanol g−1 of solvents for the wild-type strain. Unexpectedly, high-performance liquid chromatography (HPLC) analysis of dhaT::Ll.LtrB mutant culture supernatants identified a metabolite peak consistent with 1,2-propanediol (1,2-PDO), which was confirmed by nuclear magnetic resonance (NMR). Based on these findings, we propose a new model for glycerol dissimilation by C. pasteurianum, whereby the production of 1,3-PDO by the wild-type strain and low levels of both 1,3-PDO and 1,2-PDO by the engineered mutant balance the reducing equivalents generated during cell mass synthesis from glycerol. IMPORTANCE Organisms from the genus Clostridium are perhaps the most notable native cellular factories, owing to their vast substrate utilization range and equally diverse variety of metabolites produced. The ability of C. pasteurianum to sustain redox balance and glycerol fermentation despite inactivation of the 1,3-PDO pathway is a testament to the exceptional metabolic flexibility exhibited by clostridia. Moreover, identification of a previously unknown 1,2-PDO-formation pathway, as detailed herein, provides a deeper understanding of fermentative glycerol utilization in clostridia and will inform future metabolic engineering endeavors involving C. pasteurianum. To our knowledge, the C. pasteurianum dhaT disruption mutant derived in this study is the only organism that produces both 1,2- and 1,3-PDOs. Most importantly, the engineered strain provides an excellent platform for highly selective production of n-butanol from waste glycerol.