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Cutting dna using restriction endonuclease

Cutting dna using restriction endonuclease


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Genomic dna is digested with alu 1 which is a four base pair cutter. What is the frequency with wich it will cut the dna assuming a normal distribution of bases.

(this is not a homework)


Welcome to Biology.SE. I think this question would be better on-topic on math.SE.

Assuming the chromosome is either 4 bases long or infinitely long, then the answer is simply $left(frac{1}{4} ight)^4$. However, chromosomes are not infinitely long so the result should be (negligibly) lower than $left(frac{1}{4} ight)^4$. If chromosomes are only 3 nucleotide long, then of course, the probability will be $0$. If it is 4 bases long the probability is $left(frac{1}{4} ight)^4$. If it is 5 bases long the probability is $frac{2}{4^5} = frac{1}{2}left(frac{1}{4} ight)^4$.

Consider this numerical approximation, to ensure the result is correct with this simple one liner inR

n = 1e7 length(gregexpr('abcd', paste(letters[sample(1:4,n,replace=TRUE)],), perl=TRUE)[[1]])/n [1] 0.0039041

which is really close to $left(frac{1}{4} ight)^4 ≈ 0.00390625$.

So, in a genome of 3 giga bases, that represents an expected $frac{3cdot 10^9}{4^4} = 11,718,750$ (or ~11 millions) such locations where Alu 1 can cut.


DNA Preparation

Molecular cloning involves introducing DNA, as an insert, into a vector molecule. The DNA to be cloned can be obtained by cutting it out of a source DNA by digestion with restriction enzymes, by copying it from a source molecule by either the Polymerase Chain Reaction (PCR) or Reverse Transcription-PCR (RT-PCR), or by assembling it from short DNA pieces (oligonucleotides). These methods all require that the DNA source is sufficiently free of contaminants that could potentially inhibit the enzyme activities (endonucleases, polymerases) involved in processing the DNA for cloning.


Traditional cloning by restriction endonuclease digestion can use any of a number of different source DNA types. Genomic DNA can be digested with a restriction enzyme and cloned into a compatible vector site to produce a library of different inserts, all from the same source DNA. DNA already cloned into one vector can be transferred (subcloned) to a new recipient vector by cutting out the DNA with restriction enzymes and cloning into the corresponding sites of the second vector. This is frequently undertaken to facilitate protein expression or transcription of RNA, for example, which might not be possible from the original vector.

PCR is often used for generating DNA for cloning and frequently restriction sites are incorporated into the primer sites so that the amplified DNA can be digested and cloned into compatible restriction sites of the cloning vector. Any type of DNA containing the desired sequence can serve as the template for PCR. Cloning with two distinct restriction enzymes ensures that non-compatible ends are generated on each molecule, thereby preventing simple vector recircularization and forcing inserts to be cloned directionally. This can be important for ensuring a translational open reading frame for protein expression. Modification of DNA ends following restriction digestion can be helpful in certain situations. For example, in non-directional cloning, where a single restriction enzyme is used, dephosphorylation of the digested vector DNA will prevent recircularization of the vector, thereby increasing the proportion of the desired recombinant DNA molecules.

PCR is increasingly used for preparing DNA for cloning applications. Amplified DNA can either be cloned directly, or following restriction digestion with restriction sites engineered into the primers used for PCR. Alternatively, amplified DNA can be used in Seamless Cloning strategies such as NEBuilder HiFi DNA Assembly or in Ligation Independent Cloning. The vector molecules for these cloning methods may also be produced by PCR. DNA amplified with Taq polymerase has a template-independent single adenosine (A) added at the 3&rsquo ends which allows cloning into complementary T-tailed vectors. High-fidelity proofreading polymerases do not add additional bases allowing cloning of the amplified DNA into blunt-ended restriction sites. Depending on the chosen cloning strategy, the ends of amplified DNA can be modified by A-tailing, blunting, or addition or removal of 5&rsquo-phosphate groups. Complementary DNA (cDNA) generated by reverse transcription of RNA can also be amplified by PCR. This ability to synthesize DNA from RNA templates enables cloning of sequences corresponding to gene transcripts.


Restriction Endonuclease:

Palindromic sequences are sequences that have the same meaning when read from both the sides just like the word ‘MADAM’ & ‘MALAYALAM’.

  • They break the phosphodiester bonds.
  • They are found in bacteria & they protect the bacterium from foreign DNA.
  • Due to methylation of its nucleotide, they don’t act on bacteria’s own DNA.
  • The 1st restriction endonuclease isolated was Hind II.
  • More than 900 R.E have been isolated, from 230 strains of bacteria.
  • There are 3 types of R.E.Type I, II, III.

Type III is used in genetic engineering.

Naming an R.E:

• The 1st letter in a capital: it is the genus of the bacteria from which it is isolated (written in italics).
• The next two letters come from the species of the bacteria (written in italics).
• 3rd is the strain from which it has been isolated.
• 4th is the Roman numeral indicating the order of discovery from that particular strain.

EcoRI →
‘E’ represents Escherichia bacteria,
‘co’ represents colithe species,
‘R’ represents the RY13 strain of the bacteria and
‘I’ it is the 1st to be discovered from that strain.
HindII →
‘H’ represents Haemophilusbacteria,
‘in’ represents influenzae species,
‘d’ represents Rd strain and
‘II’ it is 2nd to be discovered from that strain.

Restricted sites of EcoRI:
5′ GAATTC 3′
3′ CTTAAG 5′

Restricted sites of Hind II:
5′ GTCGAC 3′
3′ CAGCTG 5′

R.E acts in two ways:

Sticky ends: Ends produced when the R.E cuts the strand of DNA a little away from the center of the palindromic sequences. It cuts between the same two bases of the two strands, leaving single-stranded parts at the ends. These single-stranded parts are referred as sticky ends.

Ligase enzyme is used to join DNA fragments with same sticky ends.

Blunt ends: Ends produced by R.E that cuts the two strands of DNA at the center of the restricted site.

Recombinant Vector:

Same R.E to cut the vector and the foreign DNA during the creation of recombinant vector. As it will produce same sticky ends that can be ligated by the ligase enzyme.


Restriction Endonucleases: Molecular Cloning and Beyond

The sequence-specific DNA cleavage activity of restriction endonucleases (REases), combined with other enzymatic activities that amplify and ligate nucleic acids, have enabled modern molecular biology. After more than half a century of research and development, the applications of REases have evolved from the cloning of exogenous DNA and genome mapping to more sophisticated applications, such as the identification and mapping of epigenetic modifications and the high-throughput assembly of combinatorial libraries. Furthermore, the discovery and engineering of nicking endonucleases (NEases) has opened the door to techniques such as isothermal amplification of DNA among others. In this review, we will examine the major breakthroughs of REase research, applications of REases and NEases in various areas of biological research and novel technologies for assembling large DNA molecules.

Siu-Hong Chan, Ph.D., New England Biolabs, Inc.

INTRODUCTION

In the 1950s, a phenomenon known as &ldquohost controlled/induced variation of bacterial viruses&rdquo was reported, in which bacteriophages isolated from one E. coli strain showed a decrease in their ability to reproduce in a different strain, but regained the ability in subsequent infection cycles (1,2). In 1965, Werner Arber&rsquos seminal paper established the theoretical framework of the restriction-modification system, functioning as bacterial defense against invading bacteriophage (3). The first REases discovered recognized specific DNA sequences, but cut at variable distances away from their recognition sequence (Type I) and, thus were of little use in DNA manipulation. Soon after, the discovery and purification of REases that recognized and cut at specific sites (Type II REases) allowed scientists to perform precise manipulations of DNA in vitro, such as the cloning of exogenous genes and creation of efficient cloning vectors. Now, more than 4,000 REases are known, recognizing more than 300 distinct sequences (for a full list, visit REBASE® at rebase.neb.com). With the advent of the Polymerase Chain Reaction (PCR), RT-PCR, and PCR-based mutagenesis methodologies, the traditional cloning workflow transformed biological research in the decades that followed.


DEVELOPMENT OF RESTRICTION ENZYMES AND GENE EDITING TECHNIQUES

ENGINEERING OF RESTRICTION ENZYMES

Traditionally, REases were purified from the native organism. The development of gene cloning vectors and selection methodologies enabled the cloning of REases. Cloning not only allowed the production of large quantities of highly purified enzymes, but also made the engineering of REases possible. Currently, > 250 of the restriction enzymes supplied by New England Biolabs (NEB) are recombinant proteins.

Engineering Improved Performance

Cleavage activity at non-cognate sites (i.e., star activity) had been observed and well-documented for some REases. Of those, some exhibit star activity under sub-optimal reaction conditions, while others have a very narrow range of enzyme units that completely digest a given amount of substrate without exhibiting star activity (4). Through intensive research, scientists at NEB began engineering restriction enzymes that exhibit minimal, if any, star activity with extended reaction times and at high enzyme concentrations. This research enabled the introduction of High Fidelity (HF&trade) REases that have improved performance under a wider range of reaction conditions (for more information, visit www.neb.com/HF).

Engineering New Sequence Specificities

Attempts to alter the sequence specificities of Type IIP REases have been largely unsuccessful, presumably because the sequence specificity determinant is structurally integrated with the active sites of Type IIP REases. MmeI, a Type IIG REase with both methyltransferase (MTase) and REase activities in the same polypeptide, recognizes the target sequence TCCRAC using the target recognition domain (TRD) within its MTase component. This represented an excellent opportunity to engineer altered sequence specificity into the REase. As an added advantage, the sharing of the TRD between the REase and MTase activities resulted in an equivalent change in MTase activity for any change in target sequence cleavage specificity, protecting the new target site from cleavage in recombinant host cells. Through bioinformatics analysis of homologous protein sequences, scientists at NEB identified the amino acid residues that recognized specific bases within the target sequences and created MmeI mutants with altered sequence specificities (5). Rational design of MmeI mutants and homologs unlocked the potential for the creation of REases with hundreds of new sequence specificities.

Figure 1. Nicking Enzyme Engineering
Type IIS REases, such as FokI (light and dark brown) and BstNBI (isoschizomer of BspD6I, light and dark purple), and homing endonuclease I-AniI (cyan), have been engineered to posses nicking enzyme activities.

Engineering Nicking Endonucleases

Basic research involving REases led to surprising findings about the seemingly straightforward mechanism of cleavage. Prototypical Type IIP REases normally act as homodimers, with each of the monomers nicking half of the palindromic site. Type IIS REases, on the other hand, exhibit a broad range of double-stranded cleavage mechanisms, namely heterodimerization, as by BtsI and BbvCI, and sequential cleavage of the dsDNA as monomer, as by FokI. These properties have been exploited to create strand-specific nicking enzymes (NEases) (for more information about nicking enzymes, see review in (6)).

APPLICATIONS UTILIZING RESTRICTION ENZYMES

Traditional Cloning

In combination with DNA ligases, REases facilitated a robust &ldquocut and paste&rdquo workflow where a defined DNA fragment could be moved from one organism to another (Fig. 2). Using this methodology, Stanley Cohen and his colleagues incorporated exogenous DNA into natural plasmids to create the vehicle for cloning-plasmid vectors that self-propagate in E. coli (7). These became the backbone of many present-day vectors, and enabled the cloning of DNA for the study and production of recombinant proteins. Restriction enzymes are also useful as post-cloning confirmatory tools, to ensure that insertions have taken place correctly. The traditional cloning workflow, along with DNA amplification technologies, such as PCR and RT-PCR, has become a mainstream application for REases and facilitated the study of many molecular mechanisms.

Figure 2. Traditional Cloning Workflow
Using PCR, restriction sites are added to both ends of a dsDNA, which is then digested by the corresponding REases. The cleaved DNA can then be ligated to a plasmid vector cleaved by the same or compatible REases with T4 DNA ligase. DNA fragments can also be moved from one vector into another by digesting with REases and ligating to compatible ends of the target vector.

DNA Mapping

Armed with only a handful of REases in the early 1970s, Daniel Nathans mapped the functional units of SV40 DNA (8), and commenced the era of &ldquorestriction mapping&rdquo and comparison of complex genomes. It has since evolved into sophisticated methodologies that allow the detection of single nucleotide polymorphisms (SNP) and insertions/deletions (Indels) (9), driving applications that include identifying genetic disorder loci, assessing the genetic diversity of populations and parental testing.

Understanding Epigenetic Modifications

REases&rsquo sensitivity to the methylation status of target bases has been exploited to map modified bases within genomes. Restriction Landmark Genome Scanning (RLGS) is a 2-dimensional gel electrophoresis-based mapping technique that employs NotI (GC^GGCCGC), AscI (GG^CGCGCC), EagI (C^GGCCG) or BssHII (G^CGCGC) to interrogate changes in the methylation patterns of the genome during the development of normal and cancer cells. Methylation-Sensitive Amplification Polymorphism (MSAP) takes advantage of the differential sensitivity of MspI and HpaII toward the methylation status of the second C of quadruplet CCGG to identify 5-methylcytosine (5-mC) or 5-hydroxymethylcytosine (5-hmC) (10,11). Scientists at NEB further exploited the property of MspI and HpaII on 5-glucosyl hydroxymethylcytosine (5-ghmC) in the EpiMark® 5-hmC and 5-mC Analysis Kit (NEB #E3317S)(12), which differentiates 5-hmC from 5-mC for more refined epigenetic marker identification and quantitation (for more information, visit EpiMark.com). Additionally, the recently discovered REases that recognize and cleave DNA at 5-mC and 5-hmC sites (e.g., MspJI, FspEI and LpnPI), as well as those that preferentially cleave 5-hmC or 5-ghmC over 5-mC or C (e.g., PvuRts1I, AbaSI) (13), are potential tools for high-throughput mapping of the cytosine-based epigenetic markers in cytosine-methylated genomes (14,15).

In vitro DNA Assembly Technologies

Synthetic biology is a rapidly growing field, in which defined components are used to create biological systems for the study of biological processes and the creation of useful biological devices (16). Novel technologies such as BioBrick&trade originally emerged to facilitate the building of such biological systems. Recently, more robust approaches, such as Golden Gate Assembly and Gibson Assembly&trade, have been widely adopted by the synthetic biology community. Both approaches allow for the parallel and seamless assembly of multiple DNA fragments without resorting to non-standard bases.

BioBrick: The BioBricks community sought to create thousands of &ldquostandardized parts&rdquo of DNAs for rapid gene assembly. With the annual International Genetically Engineered Machines (iGEM) competition (igem.org), the BioBricks community grew and elicited broad interest from many university students in synthetic biology. Based on traditional REase-ligation methodology, BioBrick and its derivative methodologies (BioBrick Assembly Kit, NEB #E0546, and its derivative, BglBricks (17)) are easy to use, but they introduce scar sequences at the junctions. They also require multiple cloning cycles to create a working biological system.

Golden Gate Assembly: Golden Gate Assembly and its derivative methods (19,20) exploit the ability of Type IIS REases to cleave DNA outside of the recognition sequence. The inserts and cloning vectors are designed to place the Type IIS recognition site distal to the cleavage site, such that the Type IIS REase can remove the recognition sequence from the assembly (Fig. 3). The advantages of such an arrangement are three-fold: 1. the overhang sequence created is not dictated by the REase, and therefore no scar sequence is introduced 2. the fragment-specific sequence of the overhangs allows orderly assembly of multiple fragments simultaneously and 3. the restriction site is eliminated from the ligated product, so digestion and ligation can be carried out simultaneously. The net result is the ordered and seamless assembly of DNA fragments in one reaction. The accuracy of the assembly is dependent on the length of the overhang sequences. Therefore, Type IIS REases that create 4-base overhangs (such as BsaI/BsaI-HF, BbsI, BsmBI and Esp3I) are preferred. The downside of these Type IIS REase-based methods is that the small number of overhanging bases can lead to the mis-ligation of fragments with similar overhang sequences (21). It is also necessary to verify that the Type IIS REase sites used are not present in the fragments for the assembly of the expected product. Nonetheless, Golden Gate Assembly is a robust technology that generates multiple site-directed mutations (22) and assembles multiple DNA fragments (23,24). As open source methods and reagents have become increasingly available (see www.addgene.org), Golden Gate Assembly has been widely used in the construction of custom-specific TALENs for in vivo gene editing (25), among other applications.

Figure 3. Golden Gate Assembly Workflow
In its simplest form, Golden Gate Assembly requires a BsaI recognition site (GGTCTC) added to both ends of a dsDNA fragment distal to the cleavage site, such that the BsaI site is eliminated by digestion with BsaI or BsaI-HF (GGTCTC 1/5). Upon cleavage, the overhanging sequences of the adjoining fragments anneal to each other. DNA ligase then seals the nicks to create a new covalently linked DNA molecule. Multiple pieces of DNA can be cleaved and ligated simultaneously. Gibson Assembly: Daniel G. Gibson, of the J. Craig Venter Institute, described a robust exonuclease-based method to assembly DNA seamlessly and in the correct order. The reaction is carried out under isothermal conditions using three enzymatic activities: a 5&rsquo exonuclease generates long overhangs, a polymerase fills in the gaps of the annealed ss regions, and a DNA ligase seals the nicks of the annealed and filled-in gaps (26) (Fig. 4). Applying this methodology, the 16.3 kb mouse mitochondrial genome was assembled from 600 overlapping 60-mers (26). In combination with in vivo assembly in yeast, Gibson Assembly was used to synthesize the 1.1 Mbp Mycoplasma mycoides genome. The synthesized genome was transplanted to a M. capricolum recipient cell creating new self-replicating M. mycoides cells (27).

Gibson Assembly can also be used for cloning the assembly of a DNA insert with a restriction-digested vector, followed by transformation, can be completed in a little less than two hours with the Gibson Assembly Cloning Kit (NEB #E5510S, for more information, visit NEBGibson.com). Other applications of Gibson Assembly include the introduction of multiple mutations, assembly of plasmid vectors from chemically synthesized oligonucleotides, and creating combinatorial libraries of genes and pathways.

Figure 4. Gibson Assembly Workflow
Gibson Assembly employs three enzymatic activities in a single-tube reaction: 5´ exonuclease, the 3´ extension activity of a DNA polymerase and DNA ligase activity. The 5´ exonuclease activity chews back the 5´ end sequences and exposes the complementary sequence for annealing. The polymerase activity then fills in the gaps on the annealed regions. A DNA ligase then seals the nick and covalently links the DNA fragments together. The overlapping sequence of adjoining fragments is much longer than those used in Golden Gate Assembly, and therefore results in a higher percentage of correct assemblies. The NEB Gibson Assembly Master Mix (NEB #E2611) and Gibson Assembly Cloning Kit (NEB #E5510S) enable rapid assembly at 50˚C.

Construction of DNA Libraries

SAGE (Serial Analysis of Gene Expression) has allowed the identification and quantification of a large number of mRNA transcripts. It has been widely used in cancer research to identify mutations and study gene expression. REases are key to the SAGE workflow. NlaIII is instrumental as an anchoring enzyme, because of its unique property of recognizing a 4-bp sequence CATG and creating a 4 nucleotide overhang of the same sequence. The use of Type IIS enzymes as tagging enzymes that cleave further and further away from the recognition sequence allows for the higher information content of SAGE analyses (e.g., FokI and BsmFI in SAGE (28), MmeI in LongSAGE (29) and EcoP15I in SuperSAGE (30) and DeepSAGE (31)).

Chromosome conformation capture (3C) and derivative methods allow the mapping of the spatial organizations of genomes in unprecedentedly high resolution and throughput (32). REases plays an indispensible role in creating the compatible ends of the DNA cross-linked to its interacting proteins, such that spatially associated sequences can be ligated and, hence, identified through high-throughput sequencing.

Although REases do not allow for the random fragmentation of DNA that most next-generation DNA sequencing technologies require, they are being used in novel target enrichment methodologies (hairpin adaptor ligation (33) and HaloPlex&trade enrichment (Agilent)). The long-reach REase, AcuI, and USER&trade Enzyme are also used to insert tags into sample DNA, which is then amplified by rolling circle amplification (RCA) to form long, single-stranded DNA &ldquonanoballs&rdquo that serve as template in the high density, chip-based sequencing-by-ligation methodology, developed by Complete Genomics (34). ApeKI was also used to generate the DNA library for a genotyping-by-sequencing technology for the study of sequence diversity of maize (35).

Creation of Nicks in DNA

Before NEases were available, non-hydrolyzable phosphorothioate groups were incorporated into a specific strand of the target DNA such that REases can introduce sequence- and strand-specific nicks into the DNA for applications such as strand displacement amplification (SDA), where a strand-displacing DNA polymerase (e.g., Bst 2.0 DNA Polymerase, NEB #M0537) extends from the newly created 3&rsquo-hydroxyl end, and essentially replicates the complementary sequence (36). Because the nicking site is regenerated, repeated nicking-extension cycles result in amplification of specific single-stranded segments of the sample DNA without the need for thermocycling. NEases greatly streamline the workflow of such applications and open the door to applications that cannot be achieved by REases. Nicking enzyme-based isothermal DNA amplification technologies, such as RCA, NESA, EXPAR and related amplification schemes, have been shown to be capable of detecting very low levels of DNA (37,38). Nicking-based DNA amplification had also been incorporated into molecular beacon technologies to amplify signal (39). The implementation of these sample and/or signal amplification schemes can lead to simple, but sensitive and specific, methods for the detection of target DNA molecules in the field (NEAR, EnviroLogix&trade). By ligating adaptors containing nicking sites to the ends of blunt-ended DNA, the simultaneous actions of the NEase(s) and strand-displacing DNA polymerase can quickly amplify a specific fragment of dsDNA (40). Amplification by nicking-extension cycling is amenable to multiplexing and can potentially achieve a higher fidelity than PCR. The combined activity of NEases and Bst DNA polymerase have also been used to introduce site-specific fluorescent labels into long/chromosomal DNA in vitro for visualization (nanocoding) (41). Innovative applications of nicking enzymes include the generation of reporter plasmids with modified bases or structures (42) and the creation of a DNA motor that transports a DNA cargo without added energy (43). A review of NEases and their applications has been published elsewhere (6).

In vivo Gene Editing

The ability to &ldquocut and paste&rdquo DNA using REases in vitro has naturally led to the quest for performing the art in vivo to correct mutations that cause genetic diseases. Direct use of REases and homing endonucleases in Restriction Enzyme Mediated Integration (REMI) facilitated the generation of transgenic embryos of higher organisms (44,45). There is, however, no control over the integration site. The concept of editing genes through site-specific cleavage has been realized using Zinc Finger Nucleases (ZFNs) and Transcription Activator-like Effector Nucleases (TALENs), due to their ability to create customizable double stranded breaks in complex genomes. With the great success of gene editing in model organisms and livestock (46-50), the therapeutic potential of these gene editing reagents is being put to the first test in the Phase I/II clinical trials of a regime that uses a ZFN to improve CD4+ T-cell counts by knocking out the expression of the CCR5 gene in autologous T-cells from HIV patients (ClinicalTrials.gov indentifier NCT00842634) (51). Recent research on CRISPR, the adaptive defense system of bacteria and archaea, has shown the potential of the Cas9-crRNA complex as programmable RNA-guided DNA endonucleases and strand-specific nicking endonucleases for in vivo gene editing (52,53).

MOVING FORWARD

Restriction enzymes have been one of the major forces that enabled the cloning of genes and transformed molecular biology. Novel technologies, such as Golden Gate Assembly and Gibson Assembly, continue to emerge and expand our ability to create new DNA molecules. The potential to generate new recognition specificity in the MmeI family REases, the engineering of more NEases and the discovery of ever more modification-specific REases continues to create new tools for DNA manipulation and epigenome analysis. Innovative applications of these enzymes will take REases&rsquo role beyond molecular cloning by continuing to accelerate the development of biotechnology and presenting us with new opportunities and challenges.


Cutting dna using restriction endonuclease - Biology

BISC411
EXPERIMENTAL MOLECULAR BIOLOGY OF THE CELL

Background Information about Restriction Enzyme Mapping and Pencil Exercise/Questions

Plasmids are extrachromosomal, self-replicating double-stranded DNA molecules. Most plasmids exist as supercoiled molecules (CCC = covalently closed circular DNA). Although small, plasmids encode a number of important gene products. Some may confer selectable phenotypes to their recipient cells such as resistance to certain antibiotics or heavy metals, which indicate that the cell has been transfected with plasmid DNA. Other plasmid genes are essential to maintain the copy number (the number of plasmid DNA molecules per cell) or to provide origin sequences which function in the initiation of plasmid DNA replication. Generally, plasmid replication and gene expression depend entirely on the preexisting host factors required to promote these processes. Many plasmids are said to be cryptic if they do not express selectable phenotypes in cells. Such plasmids are, in general, useless as cloning vehicles since plasmid transfection is not readily assayable.

Although discovered initially in Prokaryotes, several plasmids have been isolated from, or constructed to work in, lower Eukaryotic cells such as yeast and in some plants. In addition, so-called shuttle or bifunctional plasmids have been constructed so that they can replicate in more than one species of bacteria. To be useful as a cloning vehicle for the amplification of inserted foreign genes, plasmids should be low in molecular weight, have selectable phenotypes and a number of unique restriction nuclease cutting sites.

In part I of this experiment, you will isolate two plasmids from cultures of bacterial cells. The plasmids you will be isolating are called expression plasmids. This means that they have been constructed so that, when introduced into bacterial cells under appropriate conditions, the inserted gene can be transcribed and translated by the cells and generate the protein coded by the inserted gene. The protein you will be generating is T4 lyozyme. The expression plasmid has been constructed by inserting the cDNA (complementary DNA, containing only the DNA sequences complementary to the mRNA of the gene) of T4 lysozyme downstream of a promoter in the plasmid that is inducible by a chemical called IPTG. When IPTG is present in the culture, transciption of the cDNA is initiated from the promoter and the mRNA is made and subsequently translated into T4 lysozyme protein. The two types of plasmids you will use and analyze will be discussed thoroughly with you in preparation for the laboratories.

You will do Part I of this experiment in Lab 3 . Part 1 consists of breaking open the bacterial cells and extracting and purifying the plasmid DNAs. During Lab 5 you will cleave the plasmid DNAs with restriction enzymes and separate the DNA fragments produced on agarose gels. By examining the pattern and sizes of these DNA fragments, you will be able to determine which plasmid is which.

Restriction endonuclease digestion of DNA has been extremely useful in the characterization of these molecules since the DNA can be broken down to manageable sizes using them. Because these enzymes recognize a specific nucleotide sequence in DNA, the same enzyme will always produce the same fragments of a certain DNA. Usually, the first step in the analysis of a new DNA is to construct a restriction endonuclease map using one enzyme initially, but eventually using several. In order to construct this map, it is necessary to determine the sizes of all enzyme-generated DNA fragments by agarose gel electrophoresis. A map showing the positions at which the endonuclease cuts the DNA can be created by ordering the fragments on the DNA.

The pH of the electrophoresis buffer used in running the agarose gel is 7.5, therefore, all of the DNA fragments will have a negative charge and will migrate towards the positive electrode. The smaller the size of the DNA fragment, the faster it will move through the pores of the agarose. You will find that the migration during electrophoresis is proportional to the inverse of the log of the molecular weight (i.e. migration distance = K/log MW where K is a constant for each gel condition). The DNA fragments are easily visualized during electrophoresis by including ethidium bromide (EtBr) in the gel. EtBr interchelates between double-stranded DNA and, as a consequence, fluoresces brightly when illuminated with UV light. Although we will not do this, one way to order the fragments is to digest the DNA only partially so that not all potential cleavage sites are used. The "partial" fragments are then isolated, redigested completely with the same enzyme and the resulting fragments are analyzed again by gel electrophoresis. If the "partials" are overlapping, a map will be produced. Another way to order the fragments is to completely digest the DNA separately with two different enzymes. Each limit fragment is then cleaved with the other endonuclease and the resulting fragments are analyzed again. Alternatively, you can include both enzymes in the same digestion and compare the disappearance or appearance of bands to deduce the cutting patterns (we will use this approach). As a pencil exercise, use the following data to order the fragments generated in a virtual experiment and produce a restriction map.

A solution of Virus B DNA (linear double stranded molecule) has been completely digested with EcoRI. An agarose gel of this restriction digest shows the following limit fragments: A limit fragment is what is generated when the DNA is cut at all its EcoRI sites.

Fig 1. Agarose Gel of EcoRI digest of Virus B DNA. Conveniently, these fragments have had their size determined by the use of Lambda Hind III markers-- data not shown.)

Each of these fragments was eluted from the gel and redigested with Taq I.
The following patterns were produced:

____ 7.1 Kb
____ 5.0 Kb
____ 2.9 Kb
____ 1.5 Kb ____ 2.3 Kb

Figure 2 Figure 3 Figure 4

Fig 2. Agarose gel of Taq I fragments from the 8.6 Kb EcoRI fragment of Virus B DNA

Fig 3. Agarose gel of Taq I fragments from the 7.9 Kb EcoRI fragment of Virus B DNA

Fig 4. Agarose gel of Taq I fragments from the 2.3 Kb EcoRI fragment of Virus B DNA

In addition, a Taq I digest was done on the intact viral DNA, giving rise to the following limit fragments:

Fig 5. Agarose gel of Taq I fragments of Virus B DNA

1. Using the data given above, construct a restriction map showing the locations of all Taq I and EcoRI sites and giving the distances (in Kb) between them. Remember that the bands shown represent the sizes of the DNA pieces made after cutting the DNA with a restriction endonuclease.

2. For this experiment, you will be doing a several double digests. Think about how this differs from the technique described above. Diagram a gel of the restriction fragments that would arise from a double digest using TaqI/EcoRI of the Virus B DNA. What do you perceive to be the advantages or disadvantages of each technique?

3. How do these approaches (double digest, limit digest/elution/redigestion with a different enzyme) compare to partial digests?

4. Many viral and bacterial DNA molecules are circular rather than linear. How would this influence the degree of difficulty experienced in construction of a restriction map? For purposes of illustration, change our linear Virus B DNA molecule into a circular molecule by ligating the ends. What happens to the various patterns of fragments generated? Show using gel patterns and a restriction map. (Assume ligation does not generate a Taq I or EcoRI site.)


Restriction Enzyme Digestion

Read about Type II restriction enzymes and the distinguishing properties of the four principle subtypes.

High throughput sequencing methods have revolutionized genomic analysis by producing millions of sequence reads from an organism’s DNA at an ever decreasing cost.

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Videos

What is a Type II Restriction Enzyme?

Type II restriction enzymes are most commonly used for molecular biology applications, as they recognize stereotypical sequences and produce a predictable cleavage pattern. Learn more about how Type II REs work.

What is a Type I Restriction Enzyme?

Type I restriction enzymes are a group of endonucleases that recognize a bipartite sequence, but do not produce a predictable cleavage pattern. Learn more about how Type I REs work.

What is a Type III Restriction Enzyme?

Type III restriction enzymes are a group of endonucleases that recognize a non-pallindromic sequence, comprising two inversely oriented sites. Learn more about these poorly understood enzymes.

Cloning With Restriction Enzymes

Restriction enzymes are an integral part of the cloning workflow, for generating compatible ends on fragments and vectors. This animation discusses three guidelines for determining which restriction enzymes to use in your cloning experiment.

Standard Protocol for Restriction Enzyme Digests

Let one of NEB's restriction enzyme experts help you improve your technique and avoid common mistakes in digest setup.

Why is My Restriction Enzyme Not Cutting DNA?

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Restriction Enzyme Digest Problem: Too Many DNA Bands

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What is Restriction Enzyme Star Activity?

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Reduce Star Activity with High-Fidelity Restriction Enzymes

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NEB ® Restriction Enzyme Double Digest Protocol

Double digestions can save you time, and this video can offer tips for how to achieve the best results, no matter which of NEB's restriction enzymes you're using.

Restriction Enzyme Digest Protocol: Cutting Close to DNA End

When cutting close to the end of a DNA molecule, make sure you know how many bases to add to the ends of your PCR primers.

Restriction Enzyme Digestion Problem: DNA Smear on Agarose Gel

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RESTRICTION ENDONUCLEASES: MOLECULAR SCISSORS FOR SPECIFICALLY CUTTING DNA

Today, in the age of molecular biology, the study of an organism’s genome (its complete DNA) is a central component driving our understanding of biology. When scientists first considered studying genomes they were faced with a problem: how to reproducibly cut a genome’s DNA into fragments that were small enough to handle? It was a significant problem. Genomes are composed of large DNA chunks on the order of millions of units, while a scientist could reasonable only handle pieces of DNA a few thousand units long. A discrepancy far too large to bridge, thus a method for reproducibly cutting DNA into manageable pieces was required to move the genomic studies forward. When this question was first posed in the 1970s it was becoming a relatively simple exercise to isolate DNA and then randomly cut it up using chemical or mechanical means. Unfortunately, this random cutting was not a satisfactory way to obtain smaller pieces of DNA, since it was impossible to tell what the original order of the DNA fragments were, an important point since the specific order of DNA is essential for its function. The biologists were stuck. A breakthrough was needed.

As with many of the important discoveries in biology, it was the study of bacteria that yielded this breakthrough. It was discovered that a type of bacterial enzyme was found to have the ability to cut DNA in a test tube [1-2]. These restriction endonucleases, so named because they cut double stranded DNA at restricted sites, were discovered as a natural part of the bacterial machinery. In a bacterial cell, restriction endonucleases (often referred to as restriction enzymes) act as a kind of immune system, protecting the cell from the invasion of foreign DNA, as would occur when a virus attempted to infect a bacterial cell. These restriction endonucleases provided biologists with a tool to study and manipulate DNA by enabling the generation of consistently sized DNA fragments. They are now used for a wide range of applications, including cloning, Southern hybridization analysis, DNA sequencing and global gene expression analysis (SAGE). Their importance to biotechnology is apparent when looking through the catalogue of any company that supplies reagents for molecular biological work-there are pages and pages devoted to listing different restriction enzymes. Lists that continue to grow as more enzymes are discovered. Truth be told, many recombinant DNA technologies, which the field of biotechnology heavily relies on, are unlikely to have been developed without the discovery of restriction enzymes.

The Discovery of Restriction Enzymes

Restriction endonucleases were discovered during experiments to determine the ability of a bacteriophage (the name given to viruses that infect bacteria) to infect two different laboratory strains of Escherichia coli called strain B and strain K [2]. In these experiments, bacteriophages were incubated with the different strains of E. coli and the ability of the bacteriophage to kill the E. coli cells was monitored. A bacteriophage will normally cause an infected bacterial cell to break open, killing the cell, while releasing millions of bacteriophage that had replicated within the cell. When bacteriophage spilling out of E. coli strain B cells were isolated, they were found to be very successful at re-infecting E. coli strain B cells. But when these bacteriophages were incubated with E. coli strain K, it was found that only a few of the bacteriophage could manage to replicate. This was a curious observation. Even more curious was the fact that after a series of incubations of the bacteriophage with strain K, the ability of the bacteriophage to infect and kill strain K increased. With a switch back to strain B, these bacteriophage that were now able to infect strain K to a high degree, showed a reduction of their ability to infect strain B (see Figure 1). It appeared that these bacteriophages, which were previously able to infect strain B to a high degree, had switched to become strain K-infective over the course of growing within strain K cells.


Figure 1. B and K-infective bacteriophages.

For the researchers, the results of this experiment suggested two things. The first was that strain K and strain B each had some mechanism that “restricted” the infective ability of bacteriophage isolated from the opposite strain. This was supported by the reduction of infectivity of the bacteriophage when it was initially switched from one strain to the other. The second was that the restriction of the bacteriophage’s infectivity was lost as the bacteriophage’s DNA was replicated inside the bacterial cell. This was evidenced by the return of the bacteriophage’s infectivity after several passages inside of the new host strain of bacteria.

What was this restrictive mechanism? It turns out that a bacterium “labels” its own DNA by modifying it in special ways. For example, in some bacteria the cytosine nucleotide has an extra single carbon group (referred to as a methyl group) added to it. This modification is referred to as methylation [3]. Each time a bacterium replicates its DNA these modifications are added to the newly synthesized DNA. When DNA comes from outside of the cell, as when a bacteriophage injects its DNA into a bacterium, it doesn’t have these modifications and thus is recognized as a foreign object to the cell. As you can imagine, a bacterium doesn’t want foreign and potentially harmful DNA floating around inside it and accordingly makes every attempt to degrade it when detected. To perform this function a series of enzymes evolved to chop up DNA that was not methylated-enzymes that were later named restriction endonucleases [2]. The existence of these enzymes explains the differences in the infection rate between E. coli B and K strains above. If bacteriophage DNA did manage to escape destruction, the replication of this foreign DNA by the bacterium over time would mark the bacteriophage DNA with the same modifications as the host, it would then no longer be recognized as foreign and be able to infect a bacterium at a much greater frequency. Discovery of this relatively simple system for detecting the difference between host and foreign DNA provided biologist with a new molecular tool: bacterial enzymes that could cut DNA, just like scissors cut paper.

How Restriction Enzymes Work and are Used

It has already been mentioned that restriction endonucleases recognize foreign DNA by their lack of methylation3, but the way these enzymes cut DNA is varied. Some cut the foreign DNA randomly. While others recognize a particular DNA recognition sequence and then either cut within this DNA sequence or several nucleotides away from it. Where a restriction endonuclease cuts within a DNA molecule is one of the primary characteristics by which it is classified [2]. For example, type II restriction endonucleases recognize a particular sequence and cut inside that sequence, while type III restriction endonucleases cut outside of their recognition site, the cut site being potentially a dozen nucleotide base pairs away or several thousand bases away.

Type II restriction endonucleases in particular have become an exceptional tool for the molecular biologist to specifically cut DNA. These enzymes bind to DNA at any position and then travel along the strand of DNA until they reach a recognition sequence [2]. Tight binding of the enzyme at the recognition site causes its structure to change, bringing the parts of the enzyme necessary for DNA cleavage into close proximity of the DNA strand. Once accomplished the “backbone” of the DNA molecule can be cut to produce two DNA fragments from one (see Figure 2). It is the recognition sites that provide the key to how one DNA fragment can be cut into two in a specific manner. These sites can vary in size with some restriction endonucleases recognizing sequence motifs in DNA that are four nucleotides long, while others recognizing sequences that are twenty nucleotides long. The nucleotide pattern that is recognized by different restriction enzymes is quite variable, although it is frequently palindromic. A palindromic sequence is the same when read in 5′ to 3′ direction on either complementary strand of DNA, an example being the palindromic sequence recognized by the restriction enzyme known as EcoRI (see Figure 2).


Figure 2. An EcoRI restriction enzyme.

EcoRI recognizes a six-nucleotide pattern that reads GAATTC from the 5′ to the 3′ end of the DNA molecule. The complement of this sequence (on opposite DNA strand) also reads GAATTC when read from 5′ to 3′. In figure 2 you can see an illustration of an EcoRI molecule binding to and cleaving a strand of DNA, here you can see the palindromic recognition for the molecule. In this scenario the cleavage of the DNA molecule by EcoRI is symmetrical-with the enzyme cutting at the same point inside the sequence (between the G and the A when reading 5′ to 3′) on both strands. An important feature of this cleavage is the “overhanging” ends of the DNA molecule that are produced. These overhangs are often referred to as “sticky ends, ” since they can bind, or stick, to a complementary sequence of DNA (in this case 5′-AATT-3′). This stickiness is often utilized in the process of DNA cloning to help the adhesion of two DNA fragments. Overall, restriction endonucleases are quite variable in the DNA ends that they produce upon cleavage, for example leaving overhangs at the 5′ (as in EcoRI in figure 2), a 3′ overhang, or no overhang at all (referred to as “blunt” ends).

The frequency with which a given restriction endonucleases cuts DNA depends on the recognition site of the enzyme. As mentioned before, some enzymes recognize sites that are four nucleotides long (simply referred to as “four cutters”). With a quick calculation and a couple of basic assumptions, one can use this knowledge to estimate how frequently it should cut a piece of DNA. For example, with the four nucleotide bases that make up DNA the probability of any one nucleotide occurring at a given location is ¼. In the case of a “four cutter” a specific sequence of four nucleotides must be present and assuming that each nucleotide has an equal chance (i.e. ¼) of occurring at any particular site, then ¼ x ¼ x ¼ x ¼ = 1/256, a four-cutter should on average cut once every 256 base pairs.

A similar calculation can be applied to any restriction enzyme as long as the size of its recognition site is known, making it possible to predict the size and number of a DNA fragments that would be obtained by cutting a DNA molecule of known size. This fact gave molecular biologists the method they required to produce DNA fragments of known size for their experiments, as in gene mapping and genetic engineering.

Any catalogue of molecular biological reagents provides a sample of the great variety of restriction endonucleases available for research, and their names are quite strange: EcoRI, BamHI, BglII (pronounced “Bagel two”) and the racy SexAI. Their discovery has pushed the frontiers of molecular biology to greater things, allowing the development of techniques to create genetically modified proteins, to localize genes and to detect mutations that cause disease. Who would have thought that a pair of very small scissors could be so handy?

Text Consulted and Additional Reading

1. Old R.W., Primrose S.B., eds. (1994). Principles of Gene Manipulation: An Introduction to Genetic Engineering, 5th Ed. Blackwell Scientific Publications, U.S.A.
2. Burrell M.M., ed. (1993). Enzymes of Molecular Biology. Humana Press Inc., New York.
3. Kendrew J., ed. (1994). The Encyclopedia of Molecular Biology. Blackwell Scientific Publications, Oxford.

(Art by Jane Wang – note that high res versions of image files available here)


Case 1: Screening for the sickle-cell gene

The only difference between the two genes is the substitution of a T for an A in the middle position of codon 6.

  • converts a GAG codon (for Glu) to a GTG codon for Val and
  • abolishes a sequence (CTGAGG, which spans codons 5, 6, and 7) recognized and cut by one of the restriction enzymes.

When the normal gene (beta A ) is digested with the enzyme and the fragments separated by electrophoresis, the probe binds to a short fragment (between the red arrows).

However, the enzyme cannot cut the sickle-cell gene at this site, so the probe attaches to a much larger fragment (between the blue arrows).

The figure (from data provided by S. E. Antonarakis) shows the pedigree of a family whose only son has sickle-cell disease. Both his father and mother were heterozygous (semifilled box and circle respectively) as they had to be to produce an afflicted child (solid box). The electrophoresis patterns for each member of the family are placed directly beneath them. Note that the two homozygous children (1 and 3) have only a single band, but these are more intense because there is twice as much DNA in them.

In this example, a change of a single nucleotide produced the RFLP. This is a very common cause of RFLPs and now such polymorphisms are often referred to as single nucleotide polymorphisms or SNPs. (However, not all RFLPs arise from SNPs. Link to an example of one that didn't.)

How can these tools be used?

By testing the DNA of prospective parents, their genotype can be determined and their odds of producing an afflicted child can be determined. In the case of sickle-cell disease, if both parents are heterozygous for the genes, there is a 1 in 4 chance that they will produce a child with the disease. Amniocentesis and chorionic villus sampling make it possible to apply the same techniques to the DNA of a fetus early in pregnancy. The parents can learn whether the unborn child will be free of the disease or not. They may choose to have an abortion rather than bring an afflicted child into the world.

  • The mutations that cause most human genetic diseases are more varied than the single mutation associated with sickle-cell disease. Over a thousand different mutations in the cystic fibrosis gene can cause the disease. A probe for one will probably fail to identify a second. A mixture of probes, one for each of the more common mutations, can be used. But there remains the problem of "false negatives": people who are falsely told they do not carry a mutant gene.
  • There are many diseases which result from several mutant genes working together to produce the disease phenotype.
  • There are still genetic diseases for which no gene has yet been discovered. Until the gene can be located, cloned, and sequenced, no probe can be made to detect it directly. However, it is sometimes possible to find a genetic "marker" that can serve as a surrogate for the gene itself. Let's see how.

Biology

Restriction enzymes are produced naturally by bacteria and cut the DNA at specific sequences. The sequences that are recognized are usually inverted repeats (palindromic) and the cuts that are made on the DNA are double stranded symmetrical cuts. Cleavage produces either cohesive or blunt ends. The mode of action of these enzymes is that they hydrolyze the phosphodiester bonds at specific cleavage sites present in the recognition sequences in each sugar-phosphate backbone of the DNA strand (Black, 2005 Pierce, 2003 Barnum, 1998 Stryer, 2002).

In partial digestion, the restriction enzymes are allowed to act on the DNA sequences for only a limited time. The objective is to prevent the enzyme from cleaving all the available restriction sites in the DNA. This generates large overlapping fragments or contigs that can be cloned using suitable vectors. Separation of the DNA fragments on the basis of their size differences can thereafter be carried out using gel electrophoresis (Pierce, 2003).

This experiment sought to partially digest the pRcCMV plasmid using BamH1 by varying the reaction times. BamH1 is an example of a type II restriction enzyme and it is isolated from Bacillus amyloliquefaciens. The recognition sequence for this enzyme is 3-CGATCC-5 and it produces cohesive ends. The pRcCMV plasmid is 5542 bp long and has 3 BamH1 restriction sites and these are situated at 909 bp, 1306 bp and 3350 bp respectively (Pierce, 2003). Since the DNA is circular, cleavage of the plasmid using BamH1 was expected to yield 3 fragments for each of the test reactions. The sizes of the fragments were expected to be 397 bp (1306-909), 2044 bp (3350-1306) and 2192 bp (5542-3350) respectively. The control was expected to yield one fragment since no cleavage occurs as the plasmid DNA was not exposed to the enzyme. The band representing the fragment produced was expected to be located close to the loading point due to its relatively big size.

Materials and methods
The BamH1 restriction enzyme was used to cleave the plasmid DNA in reactions that were performed at 37c. The reactions were carried out in 4 different tubes, one of which was the control tube. The reaction buffer, the plasmid DNA and enzyme were added to all the test tubes. No enzyme was added to the control tube. The reactions in the 3 test tubes were timed and were allowed to proceed for 2, 10 and 30 minutes respectively. GlycerolEDTA was used to stop the reactions. The loading dye was added into each of the 4 tubes, the gel prepared and the samples loaded into separate wells. The gel was then placed into the tank and run for 5-7 minutes at 275V. The bands obtained were then photographed.

Difficulties encountered during the experiment included leakage of the gel mixture and that the gel had speckles after destaining had been done. Leakage of the gel mixture was stopped by correctly aligning the spacers and the bottom of the glass plates. To get rid of the speckles, the stain was filtered and this removed the precipitates present which were the cause of the speckles.


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Watch the video: RESTRICTION ENZYMES (September 2022).


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