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Full Sequence for Plasmid pMG101

Full Sequence for Plasmid pMG101


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I am trying to find the full plasmid sequence for pMG101. I have been looking through other papers that have sequenced this plasmid. The GenBank access numbers I got are the following: AY009372-AY009396, ASRI00000000. However, when I go on genbank it gives me a result saying it is not there… I am stuck and not sure how I am suppose to find this full plasmid sequence. Does anyone know another way to find it?


Looks as if you'll have to do some reconstruction work building the full sequence from the information here and elsewhere in this paper:

Nucleotide sequence accession numbers.The DDBJ/EMBL/GenBank accession numbers for the sequences reported in this paper are AB031076(the SalI-XhoI 3.0-kb fragment of pMG101), D84187(rDNA for strain S55), D86354 (rDNA for USDA 4362), D86355 (rDNA for USDA 4377), AB031077 (plasmid pMG103), and AB031078 (plasmid pMG105).

--Sequence Analysis of the Cryptic Plasmid pMG101 from Rhodopseudomonas palustris and Construction of Stable Cloning Vectors


Here it is: http://www.ncbi.nlm.nih.gov/nuccore/4206623?report=graph

This is the same plasmid as in E. Coli J53. This is the newest sequence.


PWPT-GFP (Plasmid # 12255 )

pWPT or pWPXL can be used for constitutive transgene expression.
pWPXL contains the EF-1alpha promoter + intron that gives you high expression, as RNA loves to be spliced (it goes more efficiently out of the nucleus). pWPT contains only the EF-1alpha promoter

The loxP site in the 3'LTR is duplicated to the 5'LTR during reverse transcription in the target cells. This allows for removal (if necessary) of an integrated provirus by Cre.

Unique restriction sites at key positions will allow you to change promoter and transgene.

Please note that ClaI in this vector is blocked by Dam methylation.
This plasmid needs to be grown in a Dam- bacteria strain if you wish to use ClaI for cloning.

Packaging plasmids for Trono lab lentiviral vectors are also available at Addgene http://www.addgene.org/rnaitools

Please note that the full sequence for this plasmid is approximated and not fully verified. Please visit the Trono lab http://tronolab.epfl.ch for cloning strategies, protocols, publications, and more. See LentiWeb http://www.lentiweb.com for discussions on cloning strategies and protocols.

These plasmids were created by your colleagues. Please acknowledge the Principal Investigator, cite the article in which the plasmids were described, and include Addgene in the Materials and Methods of your future publications.


Molecular basis for resistance to silver cations in Salmonella

Here we report the genetic and proposed molecular basis for silver resistance in pathogenic microorganisms. The silver resistance determinant from a hospital burn ward Salmonella plasmid contains nine open reading frames, arranged in three measured and divergently transcribed RNAs. The resistance determinant encodes a periplasmic silver–specific binding protein (SilE) plus apparently two parallel efflux pumps: one, a P–type ATPase (SilP) the other, a membrane potential–dependent three–polypeptide cation/proton antiporter (SilCBA). The sil determinant is governed by a two–component membrane sensor and transcriptional responder comprising silS and silR, which are co–transcribed. The availability of the sil silver–resistance determinant will be the basis for mechanistic molecular and biochemical studies as well as molecular epidemiology of silver resistance in clinical settings in which silver is used as a biocide.


FUGW (Plasmid # 14883 )

Plasmid pFUGW was constructed by inserting the following into the multicloning site of HR'CS-G: HIV-1 flap sequence PCR-amplified
from the HIV NLA4.3 genome, the human polyubiquitin promoter-C (gift of L. Thiel, Amgen), the EGFP gene, and the WRE (woodchuck
hepatitis virus posttranscriptional regulatory element) (gift of D. Trono, University of Geneva). Lentiviruses can be produced by cotransfecting the HIV-1 packaging vector Delta8.9 and the VSVG envelope glycoprotein into 293 fibroblasts.

Order of elements: CMV LTR PstI flap PacI Ubiquitin promoter SpeI HindIII PstI SalI XbaI BamHI SmaI KpnI GFP NotI EagI XbaI EcoRI EcoRV HindIII ClaI WRE ClaI SalI XhoI KpnI 3'LTR ApaI PmeI.

These plasmids were created by your colleagues. Please acknowledge the Principal Investigator, cite the article in which the plasmids were described, and include Addgene in the Materials and Methods of your future publications.


PSKI015 (Plasmid # 11570 )

The CaMV 35S enhancers in these vectors correspond to nucleotides -417 to -86 relative to the transcription start.

Recommended Agrobacterium strain is GV3101 pMP90RK. Host and helper plasmid markers are kanamycin, gentamycin and rifampicin resistance.

Plasmid selection in E. coli and in A. tumefaciens is ampicillin resistance.

Sites that leave the pBstKS+ sequences intact and cut only on one side between pBstKS+ and either left or right border can be used for plasmid rescue. Note that there are additional BssS1 sites not shown on the map.

The four enhancer repeats in the construct may be unstable in E. coli and Agrobacterium if stored at +4'C for extended time. Check by PCR with T7 (or M13-20) oligo and this one derived from the RB sequence: 5' acc cgc caa tat atc ctg 3'. This should give 1.4 kb band. Note that these oligos will not work in a transgenic plant, as the RB sequence is not transfered. Using this test, we found that after 1-2 weeks in a fridge the Agrobacterium strain loses on average one copy of a repeat, and after a month in a fridge there is only one copy left. It is a good idea to use a freshly streaked colony from a good -70'C glycerol stock for every infiltration.

These plasmids were created by your colleagues. Please acknowledge the Principal Investigator, cite the article in which the plasmids were described, and include Addgene in the Materials and Methods of your future publications.


Complete genome dynamics of a dominant-lineage strain of Xanthomonas oryzae pv. oryzae harbouring a novel plasmid encoding a type IV secretion system

Xanthomonas oryzae pv. oryzae (Xoo) is a serious pathogen causing bacterial blight disease in rice. Population genomic studies have revealed that rampant inter-strain rather than inter-lineage differences are contributing to the evolutionary success of this pathogen. Here, we report the complete genome sequence of BXO1, a strain of Xoo belonging to a dominant lineage from India. A complete genome-based investigation revealed the presence of two plasmids, pBXO1-1 (66.7 kb) and pBXO1-2 (25.6 kb). The pBXO1-1 plasmid encodes 71 genes, 38 of which encode hypothetical proteins of unknown function. However, these hypothetical genes possess atypical GC content, pointing towards their acquisition and movement through horizontal gene transfer. Interestingly, pBXO1-2 encodes a type IV secretion system (T4SS), which is known to play an important role in the conjugative transfer of genetic material, and also provides fitness to pathogenic bacteria for their enhanced survival. Neither plasmid has been reported previously in any other complete Xoo genome published to date. Our analysis also revealed that the pBXO1-2 plasmid is present in Xanthomonas albilineans str. GPE PC73, which is known to cause leaf scald, a lethal disease in sugarcane. Our complete genome sequence analysis of BXO1 has provided us with detailed insights into the two novel strain-specific plasmids, in addition to decoding their functional capabilities, which were not assessable when using the draft genome sequence of the strain. Overall, our study has revealed the mobility of a novel T4SS in two pathogenic species of Xanthomonas that infect the vascular tissues of two economically important monocot plants, i.e. rice and sugarcane.

Keywords: Xanthomonas monocots nanopore sequencing type IV secretion system xylem.

Conflict of interest statement

The authors declare that there are no conflicts of interest.

Figures

Circular representation of the BXO1…

Circular representation of the BXO1 genome. The rings represent (from outside to inside):…

Circular diagrams showing pBXO1-1 and…

Circular diagrams showing pBXO1-1 and pBXO1-2 plasmids from the BXO1 strain of Xanthomonas…

Comparative analysis of pBXO1-2 with…

Comparative analysis of pBXO1-2 with the X. albilineans str. GPE PC73 plasmid. The…


Identification of IncA/C Plasmid Replication and Maintenance Genes and Development of a Plasmid Multilocus Sequence Typing Scheme

Plasmids of incompatibility group A/C (IncA/C) are becoming increasingly prevalent within pathogenic Enterobacteriaceae They are associated with the dissemination of multiple clinically relevant resistance genes, including blaCMY and blaNDM Current typing methods for IncA/C plasmids offer limited resolution. In this study, we present the complete sequence of a blaNDM-1-positive IncA/C plasmid, pMS6198A, isolated from a multidrug-resistant uropathogenic Escherichia coli strain. Hypersaturated transposon mutagenesis, coupled with transposon-directed insertion site sequencing (TraDIS), was employed to identify conserved genetic elements required for replication and maintenance of pMS6198A. Our analysis of TraDIS data identified roles for the replicon, including repA, a toxin-antitoxin system two putative partitioning genes, parAB and a putative gene, 053 Construction of mini-IncA/C plasmids and examination of their stability within E. coli confirmed that the region encompassing 053 contributes to the stable maintenance of IncA/C plasmids. Subsequently, the four major maintenance genes (repA, parAB, and 053) were used to construct a new plasmid multilocus sequence typing (PMLST) scheme for IncA/C plasmids. Application of this scheme to a database of 82 IncA/C plasmids identified 11 unique sequence types (STs), with two dominant STs. The majority of blaNDM-positive plasmids examined (15/17 88%) fall into ST1, suggesting acquisition and subsequent expansion of this blaNDM-containing plasmid lineage. The IncA/C PMLST scheme represents a standardized tool to identify, track, and analyze the dissemination of important IncA/C plasmid lineages, particularly in the context of epidemiological studies.

Keywords: IncA/C plasmid New Delhi metallo-beta-lactamase functional genomics plasmid multilocus sequence typing uropathogenic E. coli.

Copyright © 2017 American Society for Microbiology.

Figures

Genetic map of pMS6198A. The…

Genetic map of pMS6198A. The outermost rings are CDSs on forward and reverse…

Overview of pMS6198A in vitro…

Overview of pMS6198A in vitro mutagenesis and TraDIS analysis. (1) Tn 5 ::Cm…

Essential gene TraDIS read data.…

Essential gene TraDIS read data. (A) RPKM for each gene listed on the…

Contribution of the partitioning locus…

Contribution of the partitioning locus to IncA/C stability. (A) Plasmid map of mini-IncA/C…

TraDIS-identified essential genes describe IncA/C…

TraDIS-identified essential genes describe IncA/C phylogeny. Shown is a comparison of IncA/C phylogenies…

Minimal spanning tree of IncA/C…

Minimal spanning tree of IncA/C PMLST. The branch lengths indicate the numbers of…

Phylogeny and resistance gene profiles…

Phylogeny and resistance gene profiles of IncA/C plasmids. The tree was built using…


Contents

In the second half of the 19th century, Gregor Mendel's pioneering work on the inheritance of traits in pea plants suggested that specific “factors” (today established as genes) are responsible for transferring organismal traits between generations. [4] Although proteins were initially assumed to serve as the hereditary material, Avery, MacLeod and McCarty established a century later DNA, which had been discovered by Friedrich Miescher, as the carrier of genetic information. [5] These findings paved the way for research uncovering the chemical nature of DNA and the rules for encoding genetic information, and ultimately led to the proposal of the double-helical structure of DNA by Watson and Crick. [6] This three-dimensional model of DNA illuminated potential mechanisms by which the genetic information could be copied in a semiconservative manner prior to cell division, a hypothesis that was later experimentally supported by Meselson and Stahl using isotope incorporation to distinguish parental from newly synthesized DNA. [7] [8] The subsequent isolation of DNA polymerases, the enzymes that catalyze the synthesis of new DNA strands, by Kornberg and colleagues pioneered the identification of many different components of the biological DNA replication machinery, first in the bacterial model organism E. coli, but later also in eukaryotic life forms. [2] [9]

A key prerequisite for DNA replication is that it must occur with extremely high fidelity and efficiency exactly once per cell cycle to prevent the accumulation of genetic alterations with potentially deleterious consequences for cell survival and organismal viability. [10] Incomplete, erroneous, or untimely DNA replication events can give rise to mutations, chromosomal polyploidy or aneuploidy, and gene copy number variations, each of which in turn can lead to diseases, including cancer. [11] [12] To ensure complete and accurate duplication of the entire genome and the correct flow of genetic information to progeny cells, all DNA replication events are not only tightly regulated with cell cycle cues but are also coordinated with other cellular events such as transcription and DNA repair. [2] [13] [14] [15] Additionally, origin sequences commonly have high AT-content across all kingdoms, since repeats of adenine and thymine are easier to separate because their base stacking interactions are not as strong as those of guanine and cytosine. [16]

DNA replication is divided into different stages. During initiation, the replication machineries – termed replisomes – are assembled on DNA in a bidirectional fashion. These assembly loci constitute the start sites of DNA replication or replication origins. In the elongation phase, replisomes travel in opposite directions with the replication forks, unwinding the DNA helix and synthesizing complementary daughter DNA strands using both parental strands as templates. Once replication is complete, specific termination events lead to the disassembly of replisomes. As long as the entire genome is duplicated before cell division, one might assume that the location of replication start sites does not matter yet, it has been shown that many organisms use preferred genomic regions as origins. [17] [18] The necessity to regulate origin location likely arises from the need to coordinate DNA replication with other processes that act on the shared chromatin template to avoid DNA strand breaks and DNA damage. [2] [12] [15] [19] [20] [21] [22] [23]

More than five decades ago, Jacob, Brenner, and Cuzin proposed the replicon hypothesis to explain the regulation of chromosomal DNA synthesis in E. coli. [24] The model postulates that a diffusible, trans-acting factor, a so-called initiator, interacts with a cis-acting DNA element, the replicator, to promote replication onset at a nearby origin. Once bound to replicators, initiators (often with the help of co-loader proteins) deposit replicative helicases onto DNA, which subsequently drive the recruitment of additional replisome components and the assembly of the entire replication machinery. The replicator thereby specifies the location of replication initiation events, and the chromosome region that is replicated from a single origin or initiation event is defined as the replicon. [2]

A fundamental feature of the replicon hypothesis is that it relies on positive regulation to control DNA replication onset, which can explain many experimental observations in bacterial and phage systems. [24] For example, it accounts for the failure of extrachromosomal DNAs without origins to replicate when introduced into host cells. It further rationalizes plasmid incompatibilities in E. coli, where certain plasmids destabilize each other's inheritance due to competition for the same molecular initiation machinery. [25] By contrast, a model of negative regulation (analogous to the replicon-operator model for transcription) fails to explain the above findings. [24] Nonetheless, research subsequent to Jacob's, Brenner's and Cuzin's proposal of the replicon model has discovered many additional layers of replication control in bacteria and eukaryotes that comprise both positive and negative regulatory elements, highlighting both the complexity and the importance of restricting DNA replication temporally and spatially. [2] [26] [27] [28]

The concept of the replicator as a genetic entity has proven very useful in the quest to identify replicator DNA sequences and initiator proteins in prokaryotes, and to some extent also in eukaryotes, although the organization and complexity of replicators differ considerably between the domains of life. [29] [30] While bacterial genomes typically contain a single replicator that is specified by consensus DNA sequence elements and that controls replication of the entire chromosome, most eukaryotic replicators – with the exception of budding yeast – are not defined at the level of DNA sequence instead, they appear to be specified combinatorially by local DNA structural and chromatin cues. [31] [32] [33] [34] [35] [36] [37] [38] [39] [40] Eukaryotic chromosomes are also much larger than their bacterial counterparts, raising the need for initiating DNA synthesis from many origins simultaneously to ensure timely replication of the entire genome. Additionally, many more replicative helicases are loaded than activated to initiate replication in a given cell cycle. The context-driven definition of replicators and selection of origins suggests a relaxed replicon model in eukaryotic systems that allows for flexibility in the DNA replication program. [29] Although replicators and origins can be spaced physically apart on chromosomes, they often co-localize or are located in close proximity for simplicity, we will thus refer to both elements as ‘origins’ throughout this review. Taken together, the discovery and isolation of origin sequences in various organisms represents a significant milestone towards gaining mechanistic understanding of replication initiation. In addition, these accomplishments had profound biotechnological implications for the development of shuttle vectors that can be propagated in bacterial, yeast and mammalian cells. [2] [41] [42] [43]

Most bacterial chromosomes are circular and contain a single origin of chromosomal replication (oriC). Bacterial oriC regions are surprisingly diverse in size (ranging from 250 bp to 2 kbp), sequence, and organization [45] [46] nonetheless, their ability to drive replication onset typically depends on sequence-specific readout of consensus DNA elements by the bacterial initiator, a protein called DnaA. [47] [48] [49] [50] Origins in bacteria are either continuous or bipartite and contain three functional elements that control origin activity: conserved DNA repeats that are specifically recognized by DnaA (called DnaA-boxes), an AT-rich DNA unwinding element (DUE), and binding sites for proteins that help regulate replication initiation. [17] [51] [52] Interactions of DnaA both with the double-stranded (ds) DnaA-box regions and with single-stranded (ss) DNA in the DUE are important for origin activation and are mediated by different domains in the initiator protein: a Helix-turn-helix (HTH) DNA binding element and an ATPase associated with various cellular activities (AAA+) domain, respectively. [53] [54] [55] [56] [57] [58] [59] While the sequence, number, and arrangement of origin-associated DnaA-boxes vary throughout the bacterial kingdom, their specific positioning and spacing in a given species are critical for oriC function and for productive initiation complex formation. [2] [45] [46] [60] [61] [62] [63] [64]

Among bacteria, E. coli is a particularly powerful model system to study the organization, recognition, and activation mechanism of replication origins. E. coli oriC comprises an approximately

260 bp region containing four types of initiator binding elements that differ in their affinities for DnaA and their dependencies on the co-factor ATP. DnaA-boxes R1, R2, and R4 constitute high-affinity sites that are bound by the HTH domain of DnaA irrespective of the nucleotide-binding state of the initiator. [47] [65] [66] [67] [68] [69] By contrast, the I, τ, and C-sites, which are interspersed between the R-sites, are low-affinity DnaA-boxes and associate preferentially with ATP-bound DnaA, although ADP-DnaA can substitute for ATP-DnaA under certain conditions. [70] [71] [72] [63] Binding of the HTH domains to the high- and low-affinity DnaA recognition elements promotes ATP-dependent higher-order oligomerization of DnaA's AAA+ modules into a right-handed filament that wraps duplex DNA around its outer surface, thereby generating superhelical torsion that facilitates melting of the adjacent AT-rich DUE. [53] [73] [74] [75] DNA strand separation is additionally aided by direct interactions of DnaA's AAA+ ATPase domain with triplet repeats, so-called DnaA-trios, in the proximal DUE region. [76] The engagement of single-stranded trinucleotide segments by the initiator filament stretches DNA and stabilizes the initiation bubble by preventing reannealing. [57] The DnaA-trio origin element is conserved in many bacterial species, indicating it is a key element for origin function. [76] After melting, the DUE provides an entry site for the E. coli replicative helicase DnaB, which is deposited onto each of the single DNA strands by its loader protein DnaC. [2]

Although the different DNA binding activities of DnaA have been extensively studied biochemically and various apo, ssDNA-, or dsDNA-bound structures have been determined, [56] [57] [58] [74] the exact architecture of the higher-order DnaA-oriC initiation assembly remains unclear. Two models have been proposed to explain the organization of essential origin elements and DnaA-mediated oriC melting. The two-state model assumes a continuous DnaA filament that switches from a dsDNA binding mode (the organizing complex) to an ssDNA binding mode in the DUE (the melting complex). [74] [77] By contrast, in the loop-back model, the DNA is sharply bent in oriC and folds back onto the initiator filament so that DnaA protomers simultaneously engage double- and single-stranded DNA regions. [78] Elucidating how exactly oriC DNA is organized by DnaA remains thus an important task for future studies. Insights into initiation complex architecture will help explain not only how origin DNA is melted, but also how a replicative helicase is loaded directionally onto each of the exposed single DNA strands in the unwound DUE, and how these events are aided by interactions of the helicase with the initiator and specific loader proteins. [2]

Archaeal replication origins share some but not all of the organizational features of bacterial oriC. Unlike bacteria, Archaea often initiate replication from multiple origins per chromosome (one to four have been reported) [79] [80] [81] [82] [83] [84] [85] [86] [46] yet, archaeal origins also bear specialized sequence regions that control origin function. [87] [88] [89] These elements include both DNA sequence-specific origin recognition boxes (ORBs or miniORBs) and an AT-rich DUE that is flanked by one or several ORB regions. [85] [90] ORB elements display a considerable degree of diversity in terms of their number, arrangement, and sequence, both among different archaeal species and among different origins within in a single species. [80] [85] [91] An additional degree of complexity is introduced by the initiator, Orc1/Cdc6 in archaea, which binds to ORB regions. Archaeal genomes typically encode multiple paralogs of Orc1/Cdc6 that vary substantially in their affinities for distinct ORB elements and that differentially contribute to origin activities. [85] [92] [93] [94] In Sulfolobus solfataricus, for example, three chromosomal origins have been mapped (oriC1, oriC2, and oriC3), and biochemical studies have revealed complex binding patterns of initiators at these sites. [85] [86] [95] [96] The cognate initiator for oriC1 is Orc1-1, which associates with several ORBs at this origin. [85] [93] OriC2 and oriC3 are bound by both Orc1-1 and Orc1-3. [85] [93] [96] Conversely, a third paralog, Orc1-2, footprints at all three origins but has been postulated to negatively regulate replication initiation. [85] [96] Additionally, the WhiP protein, an initiator unrelated to Orc1/Cdc6, has been shown to bind all origins as well and to drive origin activity of oriC3 in the closely related Sulfolobus islandicus. [93] [95] Because archaeal origins often contain several adjacent ORB elements, multiple Orc1/Cdc6 paralogs can be simultaneously recruited to an origin and oligomerize in some instances [94] [97] however, in contrast to bacterial DnaA, formation of a higher-order initiator assembly does not appear to be a general prerequisite for origin function in the archaeal domain. [2]

Structural studies have provided insights into how archaeal Orc1/Cdc6 recognizes ORB elements and remodels origin DNA. [97] [98] Orc1/Cdc6 paralogs are two-domain proteins and are composed of a AAA+ ATPase module fused to a C-terminal winged-helix fold. [99] [100] [101] DNA-complexed structures of Orc1/Cdc6 revealed that ORBs are bound by an Orc1/Cdc6 monomer despite the presence of inverted repeat sequences within ORB elements. [97] [98] Both the ATPase and winged-helix regions interact with the DNA duplex but contact the palindromic ORB repeat sequence asymmetrically, which orients Orc1/Cdc6 in a specific direction on the repeat. [97] [98] Interestingly, the DUE-flanking ORB or miniORB elements often have opposite polarities, [80] [85] [94] [102] [103] which predicts that the AAA+ lid subdomains and the winged-helix domains of Orc1/Cdc6 are positioned on either side of the DUE in a manner where they face each other. [97] [98] Since both regions of Orc1/Cdc6 associate with a minichromosome maintenance (MCM) replicative helicase, [104] [105] this specific arrangement of ORB elements and Orc1/Cdc6 is likely important for loading two MCM complexes symmetrically onto the DUE. [85] Surprisingly, while the ORB DNA sequence determines the directionality of Orc1/Cdc6 binding, the initiator makes relatively few sequence-specific contacts with DNA. [97] [98] However, Orc1/Cdc6 severely underwinds and bends DNA, suggesting that it relies on a mix of both DNA sequence and context-dependent DNA structural features to recognize origins. [97] [98] [106] Notably, base pairing is maintained in the distorted DNA duplex upon Orc1/Cdc6 binding in the crystal structures, [97] [98] whereas biochemical studies have yielded contradictory findings as to whether archaeal initiators can melt DNA similarly to bacterial DnaA. [93] [94] [107] Although the evolutionary kinship of archaeal and eukaryotic initiators and replicative helicases indicates that archaeal MCM is likely loaded onto duplex DNA (see next section), the temporal order of origin melting and helicase loading, as well as the mechanism for origin DNA melting, in archaeal systems remains therefore to be clearly established. Likewise, how exactly the MCM helicase is loaded onto DNA needs to be addressed in future studies. [2]

Origin organization, specification, and activation in eukaryotes are more complex than in bacterial or archaeal domains and significantly deviate from the paradigm established for prokaryotic replication initiation. The large genome sizes of eukaryotic cells, which range from 12 Mbp in S. cerevisiae to 3 Gbp in humans, necessitates that DNA replication starts at several hundred (in budding yeast) to tens of thousands (in humans) origins to complete DNA replication of all chromosomes during each cell cycle. [27] [36] With the exception of S. cerevisiae and related Saccharomycotina species, eukaryotic origins do not contain consensus DNA sequence elements but their location is influenced by contextual cues such as local DNA topology, DNA structural features, and chromatin environment. [110] [35] [37] Nonetheless, eukaryotic origin function still relies on a conserved initiator protein complex to load replicative helicases onto DNA during the late M and G1 phases of the cell cycle, a step known as origin licensing. [111] In contrast to their bacterial counterparts, replicative helicases in eukaryotes are loaded onto origin duplex DNA in an inactive, double-hexameric form and only a subset of them (10-20% in mammalian cells) is activated during any given S phase, events that are referred to as origin firing. [112] [113] [114] The location of active eukaryotic origins is therefore determined on at least two different levels, origin licensing to mark all potential origins, and origin firing to select a subset that permits assembly of the replication machinery and initiation of DNA synthesis. The extra licensed origins serve as backup and are activated only upon slowing or stalling of nearby replication forks, ensuring that DNA replication can be completed when cells encounter replication stress. [115] [116] Together, the excess of licensed origins and the tight cell cycle control of origin licensing and firing embody two important strategies to prevent under- and overreplication and to maintain the integrity of eukaryotic genomes. [2]

Early studies in S. cerevisiae indicated that replication origins in eukaryotes might be recognized in a DNA-sequence-specific manner analogously to those in prokaryotes. In budding yeast, the search for genetic replicators lead to the identification of autonomously replicating sequences (ARS) that support efficient DNA replication initiation of extrachromosomal DNA. [117] [118] [119] These ARS regions are approximately 100-200 bp long and exhibit a multipartite organization, containing A, B1, B2, and sometimes B3 elements that together are essential for origin function. [120] [121] The A element encompasses the conserved 11 bp ARS consensus sequence (ACS), [122] [123] which, in conjunction with the B1 element, constitutes the primary binding site for the heterohexameric origin recognition complex (ORC), the eukaryotic replication initiator. [124] [125] [126] [127] Within ORC, five subunits are predicated on conserved AAA+ ATPase and winged-helix folds and co-assemble into a pentameric ring that encircles DNA. [127] [128] [129] In budding yeast ORC, DNA binding elements in the ATPase and winged-helix domains, as well as adjacent basic patch regions in some of the ORC subunits, are positioned in the central pore of the ORC ring such that they aid the DNA-sequence-specific recognition of the ACS in an ATP-dependent manner. [127] [130] By contrast, the roles of the B2 and B3 elements are less clear. The B2 region is similar to the ACS in sequence and has been suggested to function as a second ORC binding site under certain conditions, or as a binding site for the replicative helicase core. [131] [132] [133] [134] [135] Conversely, the B3 element recruits the transcription factor Abf1, albeit B3 is not found at all budding yeast origins and Abf1 binding does not appear to be strictly essential for origin function. [2] [120] [136] [137]

Origin recognition in eukaryotes other than S. cerevisiae or its close relatives does not conform to the sequence-specific read-out of conserved origin DNA elements. Pursuits to isolate specific chromosomal replicator sequences more generally in eukaryotic species, either genetically or by genome-wide mapping of initiator binding or replication start sites, have failed to identify clear consensus sequences at origins. [138] [139] [140] [141] [142] [143] [144] [145] [146] [147] [148] [149] Thus, sequence-specific DNA-initiator interactions in budding yeast signify a specialized mode for origin recognition in this system rather than an archetypal mode for origin specification across the eukaryotic domain. Nonetheless, DNA replication does initiate at discrete sites that are not randomly distributed across eukaryotic genomes, arguing that alternative means determine the chromosomal location of origins in these systems. These mechanisms involve a complex interplay between DNA accessibility, nucleotide sequence skew (both AT-richness and CpG islands have been linked to origins), Nucleosome positioning, epigenetic features, DNA topology and certain DNA structural features (e.g., G4 motifs), as well as regulatory proteins and transcriptional interference. [17] [18] [34] [35] [37] [150] [151] [143] [152] Importantly, origin properties vary not only between different origins in an organism and among species, but some can also change during development and cell differentiation. The chorion locus in Drosophila follicle cells constitutes a well-established example for spatial and developmental control of initiation events. This region undergoes DNA-replication-dependent gene amplification at a defined stage during oogenesis and relies on the timely and specific activation of chorion origins, which in turn is regulated by origin-specific cis-elements and several protein factors, including the Myb complex, E2F1, and E2F2. [153] [154] [155] [156] [157] This combinatorial specification and multifactorial regulation of metazoan origins has complicated the identification of unifying features that determine the location of replication start sites across eukaryotes more generally. [2]

To facilitate replication initiation and origin recognition, ORC assemblies from various species have evolved specialized auxiliary domains that are thought to aid initiator targeting to chromosomal origins or chromosomes in general. For example, the Orc4 subunit in S. pombe ORC contains several AT-hooks that preferentially bind AT-rich DNA, [158] while in metazoan ORC the TFIIB-like domain of Orc6 is thought to perform a similar function. [159] Metazoan Orc1 proteins also harbor a bromo-adjacent homology (BAH) domain that interacts with H4K20me2-nucleosomes. [109] Particularly in mammalian cells, H4K20 methylation has been reported to be required for efficient replication initiation, and the Orc1-BAH domain facilitates ORC association with chromosomes and Epstein-Barr virus origin-dependent replication. [160] [161] [162] [163] [164] Therefore, it is intriguing to speculate that both observations are mechanistically linked at least in a subset of metazoa, but this possibility needs to be further explored in future studies. In addition to the recognition of certain DNA or epigenetic features, ORC also associates directly or indirectly with several partner proteins that could aid initiator recruitment, including LRWD1, PHIP (or DCAF14), HMGA1a, among others. [33] [165] [166] [167] [168] [169] [170] [171] Interestingly, Drosophila ORC, like its budding yeast counterpart, bends DNA and negative supercoiling has been reported to enhance DNA binding of this complex, suggesting that DNA shape and malleability might influence the location of ORC binding sites across metazoan genomes. [31] [127] [172] [173] [174] A molecular understanding for how ORC's DNA binding regions might support the read out of structural properties of the DNA duplex in metazoans rather than of specific DNA sequences as in S. cerevisiae awaits high-resolution structural information of DNA-bound metazoan initiator assemblies. Likewise, whether and how different epigenetic factors contribute to initiator recruitment in metazoan systems is poorly defined and is an important question that needs to be addressed in more detail. [2]

Once recruited to origins, ORC and its co-factors Cdc6 and Cdt1 drive the deposition of the minichromosome maintenance 2-7 (Mcm2-7) complex onto DNA. [111] [175] Like the archaeal replicative helicase core, Mcm2-7 is loaded as a head-to-head double hexamer onto DNA to license origins. [112] [113] [114] In S-phase, Dbf4-dependent kinase (DDK) and Cyclin-dependent kinase (CDK) phosphorylate several Mcm2-7 subunits and additional initiation factors to promote the recruitment of the helicase co-activators Cdc45 and GINS, DNA melting, and ultimately bidirectional replisome assembly at a subset of the licensed origins. [28] [176] In both yeast and metazoans, origins are free or depleted of nucleosomes, a property that is crucial for Mcm2-7 loading, indicating that chromatin state at origins regulates not only initiator recruitment but also helicase loading. [144] [177] [178] [179] [180] [181] A permissive chromatin environment is further important for origin activation and has been implicated in regulating both origin efficiency and the timing of origin firing. Euchromatic origins typically contain active chromatin marks, replicate early, and are more efficient than late-replicating, heterochromatic origins, which conversely are characterized by repressive marks. [27] [179] [182] Not surprisingly, several chromatin remodelers and chromatin-modifying enzymes have been found to associate with origins and certain initiation factors, [183] [184] but how their activities impact different replication initiation events remains largely obscure. Remarkably, cis-acting “early replication control elements” (ECREs) have recently also been identified to help regulate replication timing and to influence 3D genome architecture in mammalian cells. [185] Understanding the molecular and biochemical mechanisms that orchestrate this complex interplay between 3D genome organization, local and higher-order chromatin structure, and replication initiation is an exciting topic for further studies. [2]

Why have metazoan replication origins diverged from the DNA sequence-specific recognition paradigm that determines replication start sites in prokaryotes and budding yeast? Observations that metazoan origins often co-localize with promoter regions in Drosophila and mammalian cells and that replication-transcription conflicts due to collisions of the underlying molecular machineries can lead to DNA damage suggest that proper coordination of transcription and replication is important for maintaining genome stability. [139] [141] [143] [146] [186] [20] [187] [188] Recent findings also point to a more direct role of transcription in influencing the location of origins, either by inhibiting Mcm2-7 loading or by repositioning of loaded Mcm2-7 on chromosomes. [189] [152] Sequence-independent (but not necessarily random) initiator binding to DNA additionally allows for flexibility in specifying helicase loading sites and, together with transcriptional interference and the variability in activation efficiencies of licensed origins, likely determines origin location and contributes to the co-regulation of DNA replication and transcriptional programs during development and cell fate transitions. Computational modeling of initiation events in S. pombe, as well as the identification of cell-type specific and developmentally-regulated origins in metazoans, are in agreement with this notion. [140] [148] [190] [191] [192] [193] [194] [152] However, a large degree of flexibility in origin choice also exists among different cells within a single population, [143] [149] [191] albeit the molecular mechanisms that lead to the heterogeneity in origin usage remain ill-defined. Mapping origins in single cells in metazoan systems and correlating these initiation events with single-cell gene expression and chromatin status will be important to elucidate whether origin choice is purely stochastic or controlled in a defined manner. [2]


Applications of GenBrick™

  • Synthetic Genomes: synthetic bacterial and viral genomes are used for a variety of applications, such as gene therapy, vaccine development, and drug delivery.
  • Metabolic Engineering: enables efficient production of biomolecules in high quantities with valuable industrial applications.
  • Industrial Microbiology: bioengineered microbes for waste remediation and generation of alternative forms of fuel.
  • Natural Product Discovery: gene clusters inserted in model organisms to discover novel enzymes for diverse applications.
  • Environmental Microbiology/Bioremediation: generation of bioengineered microorganisms to monitor and break down environmental and to develop sustainable agricultural methods.

Results

Efficient genome editing using all-in-one AAV-sgRNA-hNmeCas9 plasmid in cells and in vivo by hydrodynamic injection

Recently, we have shown that the relatively compact NmeCas9 is active in genome editing in a range of cell types (Amrani et al., manuscript in revision). To exploit the small size of this Cas9 ortholog, we generated an all-in-one AAV construct with human-codon-optimized NmeCas9 under the expression of the mouse U1a promoter and with its sgRNA driven by the U6 promoter (Fig. 1a).

Validation of an all-in-one AAV-sgRNA-hNmeCas9 construct. a Schematic representation of a single rAAV vector expressing human-codon-optimized NmeCas9 and its sgRNA. The backbone is flanked by AAV inverted terminal repeats (ITR). The poly(a) signal is from rabbit beta-globin (BGH). b Schematic diagram of the Pcsk9 (top) and Rosa26 (bottom) mouse genes. Red bars represent exons. Zoomed-in views show the protospacer sequence (red) whereas the NmeCas9 PAM sequence is highlighted in green. Double-stranded break location sites are denoted (black arrowheads). c Stacked histogram showing the percentage distribution of insertions-deletions (indels) obtained by TIDE after AAV-sgRNA-hNmeCas9 plasmid transfections in Hepa1–6 cells targeting Pcsk9 (sgPcsk9) and Rosa26 (sgRosa26) genes. Data are presented as mean values ± SD from three biological replicates. d Stacked histogram showing the percentage distribution of indels at Pcsk9 in the liver of C57Bl/6 mice obtained by TIDE after hydrodynamic injection of AAV-sgRNA-hNmeCas9 plasmid

Two sites in the mouse genome were selected initially to test the nuclease activity of NmeCas9 in vivo: the Rosa26 “safe-harbor” gene (targeted by sgRosa26) and the proprotein convertase subtilisin/kexin type 9 (Pcsk9) gene (targeted by sgPcsk9), a common therapeutic target for lowering circulating cholesterol and reducing the risk of cardiovascular disease (Fig. 1b). Genome-wide off-target predictions for these guides were determined computationally using the Bioconductor package CRISPRseek 1.9.1 [40] with N4GN3 PAMs and up to six mismatches. Many N4GN3 PAMS are inactive, so these search parameters are nearly certain to cast a wider net than the true off-target profile. Despite the expansive nature of the search, our analyses revealed no off-target sites with fewer than four mismatches in the mouse genome (Additional file 1: Figure S1). On-target editing efficiencies at these target sites were evaluated in mouse Hepa1–6 hepatoma cells by plasmid transfections and indel quantification was performed by sequence trace decomposition using the Tracking of Indels by Decomposition (TIDE) web tool [41]. We found > 25% indel values for the selected guides, the majority of which were deletions (Fig. 1c).

To evaluate the preliminary efficacy of the constructed all-in-one AAV-sgRNA-hNmeCas9 vector, endotoxin-free sgPcsk9 plasmid was hydrodynamically administered into the C57Bl/6 mice via tail-vein injection. This method can deliver plasmid DNA to

40% of hepatocytes for transient expression [42]. Indel analyses by TIDE using DNA extracted from liver tissues revealed 5–9% indels 10 days after vector administration (Fig. 1d), comparable to the editing efficiencies obtained with analogous tests of SpyCas9 [43]. These results suggested that NmeCas9 is capable of editing liver cells in vivo.

Knockout of 4-Hydroxyphenylpyruvate dioxygenase rescues the lethal phenotypes of hereditary Tyrosinemia type I mice

Hereditary Tyrosinemia type I (HT-I) is a fatal genetic disease caused by autosomal recessive mutations in the Fah gene, which codes for the fumarylacetoacetate hydroxylase (FAH) enzyme. Patients with diminished FAH have a disrupted tyrosine catabolic pathway, leading to the accumulation of toxic fumarylacetoacetate and succinyl acetoacetate, causing liver and kidney damage [44]. Over the past two decades, the disease has been controlled by 2-(2-Nitro-4-trifluoromethylbenzoyl) -1,3- cyclohexanedione (NTBC), which inhibits 4-hydroxyphenylpyruvate dioxygenase upstream in the tyrosine degradation pathway, thus preventing the accumulation of the toxic metabolites [45]. However, this treatment requires lifelong management of diet and medication and may eventually require liver transplantation [46].

Several gene therapy strategies have been tested to correct the defective Fah gene using site-directed mutagenesis [47] or homology-directed repair by CRISPR-Cas9 [47,48,49]. It has been reported that successful modification of only 1/10,000 of hepatocytes in the liver is sufficient to rescue the phenotypes of Fah mut/mut mice. Recently, a metabolic pathway reprogramming approach has been suggested in which the function of the hydroxyphenylpyruvate dioxygenase (HPD) enzyme was disrupted by the deletion of exons 3 and 4 of the Hpd gene in the liver [35]. This provides us with a context in which to test the efficacy of NmeCas9 editing, by targeting Hpd and assessing rescue of the disease phenotype in Fah mutant mice [50]. For this purpose, we screened and identified two target sites (one each in exon 8 [sgHpd1] and exon 11 [sgHpd2]) within the open reading frame of Hpd (Fig. 2a). These guides induced average indel efficiencies of 10.8% and 9.1%, respectively, by plasmid transfections in Hepa1–6 cells (Additional file 1: Figure S2).

NmeCas9-mediated knockout of Hpd rescues the lethal phenotype in Hereditary Tyrosinemia Type I Mice. a Schematic diagram of the Hpd mouse gene. Red bars represent exons. Zoomed-in views show the protospacer sequences (red) for targeting exon 8 (sgHpd1) and exon 11 (sgHpd2). NmeCas9 PAM sequences are in green and double-stranded break locations are indicated (black arrowheads). b Experimental design. Three groups of Hereditary Tyrosinemia Type I Fah −/− mice are injected with PBS or all-in-one AAV-sgRNA-hNmeCas9 plasmids sgHpd1 or sgHpd2. c Weight of mice hydrodynamically injected with PBS (green), AAV-sgRNA-hNmeCas9 plasmid sgHpd1 targeting Hpd exon 8 (red) or sgHpd2-targeting Hpd exon 11 (blue) were monitored after NTBC withdrawal. Error bars represent three mice for PBS and sgHpd1 groups and two mice for the sgHpd2 group. Data are presented as mean ± SD. d Stacked histogram showing the percentage distribution of indels at Hpd in liver of Fah −/− mice obtained by TIDE after hydrodynamic injection of PBS or sgHpd1 and sgHpd2 plasmids. Livers were harvested at the end of NTBC withdrawal (day 43)

Three groups of mice were treated by hydrodynamic injection with either phosphate-buffered saline (PBS) or with one of the two sgHpd1 and sgHpd2 all-in-one AAV-sgRNA-hNmeCas9 plasmids. One mouse in the sgHpd1 group and two in the sgHpd2 group were excluded from the follow-up study due to failed tail-vein injections. Mice were taken off NTBC-containing water seven days after injections and their weight was monitored for 43 days post injection (Fig. 2b). Mice injected with PBS suffered severe weight loss (a hallmark of HT-I) and were sacrificed after losing 20% of their body weight (Fig. 2c). Overall, all sgHpd1 and sgHpd2 mice successfully maintained their body weight for 43 days overall and for at least 21 days without NTBC (Fig. 2c). NTBC treatment had to be resumed for 2–3 days for two mice that received sgHpd1 and one that received sgHpd2 to allow them to regain body weight during the third week after plasmid injection, perhaps due to low initial editing efficiencies, liver injury due to hydrodynamic injection, or both. Conversely, all other sgHpd1 and sgHpd2 treated mice achieved indels with frequencies in the range of 35–60% (Fig. 2d). This level of gene inactivation likely reflects not only the initial editing events but also the competitive expansion of edited cell lineages (after NTBC withdrawal) at the expense of their unedited counterparts [46, 47, 49]. Liver histology revealed that liver damage is substantially less severe in the sgHpd1- and sgHpd2-treated mice compared to Fah mut/mut mice injected with PBS, as indicated by the smaller numbers of multinucleated hepatocytes compared to PBS-injected mice (Additional file 1: Figure S3).

In vivo genome editing by NmeCas9 delivered by a rAAV vector

Although plasmid hydrodynamic injections can generate indels, therapeutic development will require less invasive delivery strategies, such as rAAV. To this end, all-in-one AAV-sgRNA-hNmeCas9 plasmids were packaged in hepatocyte-tropic AAV8 capsids to target Pcsk9 (sgPcsk9) and Rosa26 (sgRosa26) (Fig. 1b) [51, 52]. Pcsk9 and Rosa26 were used in part to enable NmeCas9 AAV delivery to be benchmarked with that of other Cas9 orthologs delivered similarly and targeted to the same loci [18]. Vectors were administered into C57BL/6 mice via tail vein (Fig. 3a). We monitored cholesterol level in the serum and measured PCSK9 protein and indel frequencies in the liver tissues 25 and 50 days post injection.

AAV-delivery of NmeCas9 for in vivo genome editing. a Experimental outline of AAV8-sgRNA-hNmeCas9 vector tail-vein injections to target Pcsk9 (sgPcsk9) and Rosa26 (sgRosa26) in C57Bl/6 mice. Mice were sacrificed at 14 (n = 1) or 50 days (n = 5) post injection and liver tissues were harvested. Blood sera were collected at days 0, 25, and 50 post injection for cholesterol level measurement. b Serum cholesterol levels. p values are calculated by unpaired t test. c Stacked histogram showing the percentage distribution of indels at Pcsk9 or Rosa26 in livers of mice, as measured by targeted deep-sequencing analyses. Data are presented as mean ± SD from five mice per cohort. d A representative anti-PCSK9 western blot using total protein collected from day 50 mouse liver homogenates. A total of 2 ng of recombinant mouse PCSK9 (r-PCSK9) was included as a mobility standard. The asterisk indicates a cross-reacting protein that is larger than the control recombinant protein

Using a colorimetric endpoint assay, we determined that the circulating serum cholesterol level in the sgPcsk9 mice decreased significantly (p < 0.001) compared to the PBS and sgRosa26 mice at 25 and 50 days post injection (Fig. 3b). Targeted deep-sequencing analyses at Pcsk9 and Rosa26 target sites revealed very efficient indels of 35% and 55%, respectively, at 50 days post vector administration (Fig. 3c). Additionally, one mouse of each group was euthanized at 14 days post injection and revealed on-target indel efficiencies of 37% and 46% at Pcsk9 and Rosa26, respectively (Fig. 3c). As expected, PCSK9 protein levels in the livers of sgPcsk9 mice were substantially reduced compared to the mice injected with PBS and sgRosa26 (Fig. 3d). The efficient editing, PCSK9 reduction, and diminished serum cholesterol indicate the successful delivery and activity of NmeCas9 at the Pcsk9 locus.

SpyCas9 delivered by viral vectors is known to elicit host immune responses [19, 53]. To investigate if the mice injected with AAV8-sgRNA-hNmeCas9 generate anti-NmeCas9 antibodies, we used sera from the treated animals to perform IgG1 ELISA. Our results show that NmeCas9 elicits a humoral response in these animals (Additional file 1: Figure S4). Despite the presence of an immune response, NmeCas9 delivered by rAAV is highly functional in vivo, with no apparent signs of abnormalities or liver damage (Additional file 1: Figure S5).

NmeCas9 is highly specific in vivo

A significant concern in therapeutic CRISPR/Cas9 genome editing is the possibility of off-target edits. We and others have found that wild-type NmeCas9 is a naturally high-accuracy genome editing platform in cultured mammalian cells [32] (Amrani et al., manuscript in revision). To determine if NmeCas9 maintains its minimal off-targeting profile in mouse cells and in vivo, we screened for off-target sites in the mouse genome using genome-wide, unbiased identification of DSBs enabled by sequencing (GUIDE-seq) [22]. Hepa1–6 cells were transfected with sgPcsk9, sgRosa26, sgHpd1, and sgHpd2 all-in-one AAV-sgRNA-hNmeCas9 plasmids and the resulting genomic DNA was subjected to GUIDE-seq analysis. Consistent with our previous observations in human cells (Amrani et al., manuscript in revision), GUIDE-seq revealed very few off-target (OT) sites in the mouse genome. Four potential OT sites were identified for sgPcsk9 and another six for sgRosa26. We were unable to detect off-target edits with sgHpd1 and sgHpd2 (Fig. 4a), thus reinforcing our previous observation that NmeCas9 is often intrinsically hyper-accurate (Amrani et al., manuscript in revision).

GUIDE-seq genome-wide specificities of NmeCas9. a Number of GUIDE-seq reads for the on-target (OnT) and off-target (OT) sites. b Targeted deep sequencing to measure the lesion rates at each of the OT sites in Hepa1–6 cells. The mismatches of each OT site with the OnT protospacers is highlighted (blue). Data are presented as mean ± SD from three biological replicates. c Targeted deep sequencing to measure the lesion rates at each of the OT sites using genomic DNA obtained from mice injected with all-in-one AAV8-sgRNA-hNmeCas9 sgPcsk9 and sgRosa26 and sacrificed at day 14 (D14) or day 50 (D50) post injection. Data are presented as mean ± SD

Several of the putative OT sites for sgPcsk9 and sgRosa26 lack the NmeCas9 PAM preferences (N4GATT, N4GCTT, N4GTTT, N4GACT, N4GATA, N4GTCT, and N4GACA) (Fig. 4b) and may therefore represent background. To validate these OT sites, we performed targeted deep sequencing using genomic DNA from Hepa1–6 cells. By this more sensitive readout, indels were undetectable above background at all these OT sites except OT1 of Pcsk9, which had an indel frequency < 2% (Fig. 4b). To validate NmeCas9’s high fidelity in vivo, we measured indel formation at these OT sites in liver genomic DNA from the AAV8-NmeCas9-treated, sgPcsk9-targeted, and sgRosa26-targeted mice. We found little or no detectable off-target editing in mice liver sacrificed at 14 days at all sites except sgPcsk9 OT1, which exhibited < 2% lesion efficiency (Fig. 4c). More importantly, this level of OT editing stayed below < 2% even after 50 days and also remained either undetectable or very low for all other candidate OT sites. These results suggested that extended (50 days) expression of NmeCas9 in vivo does not compromise its targeting fidelity (Fig. 4c).


Contents

The Ti plasmid is a member of the RepABC plasmid family found in Alphaproteobacteria. [3] These plasmids are often relatively large in size, ranging from 100kbp to 2Mbp. They are also often termed replicons, as their replication begins at a single site. Members of this family have a characteristic repABC gene cassette. [4] Another notable member of this family is the root inducing (Ri) plasmid carried by A. rhizogenes, which causes another plant disease known as hairy root disease. [1]

A key feature of Ti plasmids is their ability to drive the production of opines, which are derivatives of various amino acids or sugar phosphates, in host plant cells. These opines can then be used as a nutrient for the infecting bacteria, which catabolizes the respective opines using genes encoded in the Ti plasmid.

Accordingly, Ti plasmids have been classified based on the type of opine they catabolize, namely: nopaline-, octopine- or mannityl-types, which are amino acid derivatives, or agrocinopine-type, which are sugar phosphate derivatives. [1]

The identification of A. tumefaciens as the cause of gall tumours in plants paved the way for insights into the molecular basis of crown gall disease. [5]

The first indication of a genetic effect on host plant cells came in 1942-1943, where plant cells of secondary tumours were found to lack any bacterial cells within. However, these tumour cells did possess the ability to produce opines metabolized by the infecting bacterial strain. [6] Crucially, the production of the respective opines occurred regardless of the plant species and occasionally only within crown gall tissues, indicating that the bacteria had transferred some genetic material to the host plant cells in order to allow opine synthesis. [5]

However, how and to what extend did DNA transfer occur remained an open question. Adding A. tumefaciens DNA alone did not cause tumors in plants, [7] while very little A. tumefaciens DNA was found to be integrated into the host plant cell genome. [8] The addition of deoxyribonucleases (DNases) to degrade DNA also failed to prevent the formation and growth of the plant tumors. [9] These suggested that little, if any, of the A. tumefaciens DNA is transferred to the host plant cell to cause disease and, if DNA is indeed transferred from the bacteria to the plant, it must occur in a protected manner.

Subsequently, oncogenic bacterial strains were found to be able to convert non-pathogenic bacteria into pathogens via the process of conjugation, where the genes responsible for virulence were transferred to the non-pathogenic cells. [10] The role of a plasmid in this pathogenic ability was further supported when large plasmids were found only in pathogenic bacteria but not avirulent bacteria. [11] Eventually, the detection of parts of bacterial plasmids in host plant cells was established, confirming that this was the genetic material responsible for the genetic effect of infection. [12]

With the identification of the Ti plasmid, many studies were carried out to determine the characteristics of the Ti plasmid and how the genetic material is transferred from the Agrobacterium to the plant host. Some notable early milestones in the studies of Ti plasmids include the mapping of a Ti plasmid in 1978 and the studying of sequence similarity between different Ti plasmids in 1981. [13] [14]

Between 1980–2000, the characterization of the T-DNA region and the 'vir' region was also pursued. Studies into the T-DNA region determined their process of transfer and identified genes allowing the synthesis of plant hormones and opines. [15] Separately, early work aimed to determine the functions of the genes encoded in the 'vir' region - these were broadly categorized into those that allowed bacterial-host interactions and those that enabled T-DNA delivery. [2]

The replication, partitioning and maintenance of the Ti plasmid depends on the repABC gene cassette, which is mainly made up of three genes: repA, repB and repC. repA and repB each encode for proteins involved in plasmid partitioning, while repC encodes a replication initiator. [1] These genes are expressed from 4 different promoters located upstream of repA. repE encodes for a small antisense RNA and is located between repB and repC. [4] Additionally, there is a partitioning site (parS) and an origin of replication (oriV) present within the repABC cassette. [1]

Replication of the Ti plasmid Edit

The replication of the Ti plasmid is driven by the RepC initiator protein ( P05684 ), which possesses two protein domains: an N-terminal domain (NTD) that binds to DNA and a C-terminal domain (CTD). Mutational analyses have shown that without a functional RepC protein, the Ti plasmid is unable to replicate. [4] Meanwhile, the oriV sequence is around 150 nucleotides in length and is found within the repC gene. [3] Laboratory experiments have shown that the RepC protein binds to this region, suggesting its role as the origin of replication. [16] Therefore, while the complete process behind the replication of the Ti plasmid has not been fully described, the initial step of replication would likely depend on the expression of RepC and its binding to oriV. Of note, the RepC protein only acts in cis, where it only drives the replication of the plasmid it is encoded in and not any other plasmid also present in the bacterial cell. [16]

Partitioning of the Ti plasmid Edit

Components involved in the RepA/RepB partitioning system of Ti plasmids [1]
Component Function
RepA (ParA), P05682 A weak ATPase that negatively autoregulates the expression of the repABC cassette and can form filaments to aid in the partitioning of the plasmid during cell division
RepB (ParB), P05683 A DNA binding protein that serves as an adaptor between RepA and the parS site
parS The palindromic binding site for the ParB protein consensus GTTNNCNGCNGNNAAC

The partitioning system of the Ti plasmid is similar to the ParA/ParB system used in other plasmids and bacterial chromosomes and is thought to act in the same way. [17] Mutations in either of the proteins RepA or RepB have resulted in a decrease in plasmid stability, indicating their role and importance in plasmid partitioning. [4] The ability of RepA to form filaments allows it to create a physical bridge along which DNA can be pulled to opposite poles of a dividing cell. Meanwhile, the RepB protein can bind specifically to the parS sequence, forming a complex with DNA that can be recognized by RepA. [1] [4] This system is particularly important for the proper partitioning of the Ti plasmid, as the plasmid is only present in few copy numbers in the bacterial cell.

Maintenance of the Ti plasmid Edit

The Ti plasmid is maintained at low copy numbers within a bacterial cell. This is partly achieved by influencing the expression of the replication initiator RepC. [1] When bound to ADP, RepA is activated to work with RepB, acting as a negative regulator of the repABC cassette. [3] The levels of RepC is therefore kept low within a cell, preventing too many rounds of replication from occurring during each cell division cycle. Furthermore, there is a small RNA known as RepE encoded between repB and repC that lowers the expression of repC. [18] RepE is complementary to RepC and will bind with the repC mRNA to form a double-stranded molecule. This can then block the translational production of the RepC protein. [18]

Separately, the expression of the repABC cassette and hence the copy number of the Ti plasmid is also influenced via a quorum sensing system in Agrobacterium. [4] Quorum sensing systems respond to bacterial population densities by sensing a molecule, known as an autoinducer, that is produced by the bacterial cells at low levels and would build up to a threshold level when there is a high density of bacteria present. [18] In this case, the autoinducer is the N-3-oxooctanoyl-L-homoserine lactone (3-O-C8-AHL) molecule, which is sensed by a regulator known as TraR. [4] When activated, TraR will bind to regions known as tra boxes in the repABC gene cassette's promoter regions to drive expression. Therefore, a high level of population density increases the number of plasmids present within each bacterial cell, likely to support pathogenesis in the plant host. [4]

Virulence operon Edit

The expression of the vir region is usually repressed under normal conditions, and only becomes activated when the bacteria senses plant-derived signals from wound sites. This activation is necessary for the production of Vir proteins and the transfer of DNA and proteins into host plant cells. [1]

VirA and VirG form a two-component regulatory system within Agrobacterium. [19] This is a type of sensing and signalling system found commonly in bacteria in this case, they act to sense plant-derived signals to drive the expression of the vir region. During the sensing, VirA, a histidine sensor kinase, will become phosphorylated before passing on this phosphate group to the response regulator VirG. [20] The activated response regulator VirG can then bind to a region of DNA known as the vir box, located upstream of each vir promoter, to activate the expression of the vir region. [1] [19] One possible downstream functions of the sensing mediated by VirA and VirG is the directional movement, or chemotaxis, of the bacteria towards plant-derived signals this allows the Agrobacterium to move towards the wound site in plants. [21] Furthermore, with the induction of the vir region, the transfer of T-DNA can be mediated by the Vir proteins. [22]

The virB operon is the largest operon in the vir region, encoding for 11 VirB proteins involved in the transfer process of T-DNA and bacterial proteins into host plant cells (see transfer apparatus below). [23] [24]

The virC operon encodes for two proteins: VirC1 and VirC2. These proteins influence the pathogenesis of the Agrobacterium towards different plant hosts, and mutations can reduce but not remove the virulence of the bacteria. [25] Both the virC and virD operons can be repressed by a chromosomally encoded protein known as Ros. [26] [27] Ros binds to a region of DNA that overlaps with the binding site of the VirG regulator, and therefore competes with VirG to control their expression levels. [26] [27] Functionally, VirC1 and VirC2 promote the assembly of a relaxosome complex during the conjugative transfer of T-DNA from the bacteria to the host plant cell. [28] This is an energy-dependent process mediated via their NTPase activity, and occurs as they bind to a region of DNA known as overdrive. [28] As a result, they act to increase the amount of T-DNA strands produced. Following the production of the DNA strand to be transferred (transfer strand, T-strand), the VirC proteins can also help to direct the transfer strand to the transfer apparatus. [28]

The virD operon encodes for 4 proteins: VirD1-D4. [29] VirD1 and VirD2 are involved in the processing of T-DNA during conjugation to produce the T-strand this is the single-stranded DNA molecule that is transported to the host plant cell (see transfer apparatus below). [30] During the processing, VirD1 will act as a topoisomerase to unwind the DNA strands. [30] VirD2, a relaxase, will then nick one of the DNA strands and remain bound to the DNA as it is transferred to the recipient cell. [31] [32] Within the recipient cell, VirD2 will also work together with VirE2 to direct the transferred DNA to the recipient cell's nucleus. There are suggestions that VirD2 may be phosphorylated and dephosphorylated by different proteins, affecting its ability to deliver DNA. [33] Conversely, little is known about VirD3, and mutational analyses have not provided any support for its role in the virulence of Agrobacterium. [34] Finally, VirD4 is a crucial part of the conjugation process, serving as a coupling factor that recognizes and transfers the T-strand to the transport channel. [35]

The virE operon encodes for 2 proteins: VirE1 and VirE2. [36] VirE2 is an effector protein translocated together with the T-strand into host plant cells. There, it binds to the T-strand to direct its delivery to the nucleus of the host plant cell. [37] [38] Part of this activity involves the presence of nuclear localization sequences within the protein, which marks the protein and the associated DNA for entry into the nucleus. It also protects the T-strand from nuclease attack. [39] There is some speculation regarding the role of VirE2 as a protein channel, allowing DNA to move through the plant cytoplasmic membrane. [40] On the other hand, VirE1 may be involved in promoting the transfer of the VirE2 protein into the host plant cell. [41] It binds to the ssDNA-binding domain of VirE2, therefore preventing the VirE2 protein from prematurely binding to the T-strand within the bacterial cell. [42]

virF is a host specificity factor found in some but not all types of Ti plasmids for example, octopine-type Ti plasmids possess virF but nopaline-types do not. [43] [44] The ability of A. tumefaciens to induce crown gall tumours in certain plant species but not others has been attributed to the presence or absence of this virF gene. [43] [44]

The virH operon encodes for 2 proteins: VirH1 and VirH2. [45] A bioinformatics study of the amino acid sequences of the VirH protein showed similarities between them and a superfamily of proteins known as cytochrome P450 enzymes. [46] VirH2 was then discovered to metabolize certain phenolic compounds detected by VirA. [45]

Transfer DNA (T-DNA) Edit

The T-DNA of Agrobacterium is approximately 15-20 kbp in length and will become integrated into the host plant genome upon its transfer via a process known as recombination. This process utilizes preexisting gaps in the host plant cell's genome to allow the T-DNA to pair with short sequences in the genome, priming the process of DNA ligation, where the T-DNA is permanently joint to the plant genome. [37] The T-DNA region is flanked at both ends by 24bp sequences.

Within the host plant cell's genome, the T-DNA of Agrobacterium is expressed to produced two main groups of proteins. [1] One group is responsible for the production of plant growth hormones. As these hormones are produced, there will be an increase in the rate of cell division and therefore the formation of crown gall tumors. [47] The second group of proteins are responsible for driving the synthesis of opines in the host plant cells. The specific opines produced depends on the type of the Ti plasmid but not on the plant host. These opines cannot be utilized by the plant host, and will instead be exported out of the plant cell where it can be taken up by the Agrobacterium cells. The bacteria possess genes in other regions of the Ti plasmid that allows the catabolism of opines. [1]

Transfer apparatus Edit

Transfer apparatuses encoded within the Ti plasmid have to achieve two objectives: allow the conjugative transfer of the Ti plasmid between bacteria and allow the delivery of the T-DNA and certain effector proteins into host plant cells. These are achieved by the Tra/Trb system and the VirB/VirD4 system respectively, which are members of the type IV secretion system (T4SS). [47]

For the Ti plasmid and T-DNA to be transferred via conjugation, they must first be processed by different proteins, such as the relaxase enzyme (TraA/VirD2) and the DNA transfer and replication (Dtr) proteins. Together, these proteins will recognize and bind to a region known as the origin of transfer (oriT) in the Ti plasmid to form the relaxosome complex. For the T-DNA, a nick will be created at the T-DNA's border sequence, and the nicked T-strand will be transported to the cell membrane, where the rest of the transfer machinery is present. [31]

Within the VirB/VirD4 system, the VirD2 relaxase is aided by the accessory factors VirD1, VirC1 and VirC2 while it processes the DNA substrate. [48] Furthermore, the VirD2 relaxase and the VirC proteins will contribute to the delivery of the DNA strand to the VirD4 receptor at the cell membrane. [28] This receptor is an essential component of T4SSs, and is thought to energize and mediate the transfer of the DNA into the translocation channel between two cells. [49] The table below summarizes the proteins encoded in the virB operon that makes up the translocation channel of the VirB/VirD4 system. [1]

Protein(s) Function
VirB4, VirB11 ATPases that provide the energy for DNA transfer [50] [51]
VirB3, VirB6, VirB8 Subunits of a putative inner membrane translocase [50] [52] [53]
VirB7, VirB9, VirB10 Forms a core complex that stabilizes the channel subunits [50] [54]
VirB2 The major pilin subunit of the conjugative pilus [50]
VirB1, VirB5 Minor components of the conjugative pilus [55] [56]

The ability of Agrobacterium to deliver DNA into plant cells opened new doors for plant genome engineering, allowing the production of genetically modified plants (transgenic plants). [57] Proteins involved in mediating the transfer of T-DNA will first recognize the border sequences of the T-DNA region. Therefore, it is possible for scientists to use T-DNA border sequences to flank any desired sequence of interest - such a product can then be inserted into a plasmid and introduced into Agrobacterium cells. [58] There, the border sequences will be recognized by the transfer apparatus of A. tumefaciens and delivered in a standard manner into the target plant cell. [1] Moreover, by leaving behind only the border sequences of the T-DNA, the resulting product will edit the plant genome without causing any tumours in plants. [59] This method has been used to modify several crop plants, including rice, [60] barley [61] and wheat. [62] Further work have since extended the targets of A. tumefaciens to include fungi and human cell lines. [63] [64]


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