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Molecular Cloning- Blunt end restriction endonucleases

Molecular Cloning- Blunt end restriction endonucleases



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I work in a microbiology lab where we do a lot of cloning. I have always used restriction endonucleases to cleave the DNA to have sticky ends and not blunt ends. I currently am working on a project that suggests using a blunt end restriction enzyme, such as EvoRV, and then to perform a T-tailing of the plasmid, pl4440. My question is why do I not treat my pcr'd insert with a restriction digest just like I would the plasmid as the protocol suggests? How do I know the insert has ends that are able to ligate into the plasmid if I don't design the restriction sites into the primers?


To know if the ends are compatible see compatible cohesive ends (isoschizomers) from NEB website.

If you are unsure then do a blunt end ligation. For blunting a sticky end (From the comments):

Use Klenow or Pfu polymerase (or any proofreading/high fidelity polymerase).

You can use Taq if you intend to to a TA cloning.


Restriction Enzyme Basics

Where would modern-day molecular biology research be without restriction enzymes? These workhorses of the lab have been behind many of the advances in basic biological research and commercial applications for over 40 years. Restriction enzymes (or restriction endonucleases) were first identified in bacteria but have been subsequently found in some archaea. In general, restriction enzymes cleave double-stranded DNA. Each restriction enzyme recognizes specific DNA sequences, and cleavage can occur within the recognition sequence or some distance away, depending on the enzyme. The recognition sequences are generally 4 to 8 base pairs (bp) in length, and cleavage can produce sticky ends (5′ or 3′ protruding ends) or blunt ends (Figure 1).

Figure 1. Sticky or protruding ends (5′ or 3′) or blunt ends produced by specific restriction enzymes.

Today about 4,000 restriction enzymes have been characterized, and over 600 of those are commercially available. REBASE is a useful, browsable resource for comprehensive and up-to-date information about restriction enzymes, including specificity, sensitivity, and commercial sources [1].

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Molecular Cloning- Blunt end restriction endonucleases - Biology

The Polymerase Chain Reaction (PCR) can be used to rapidly generate DNA fragments for cloning, provided that a suitable source of template DNA exists and sufficient sequence information is known to permit design of primers specific for the desired amplicon. Unlike traditional cloning, PCR offers the ability to readily clone DNA fragments that may be of low abundance in a complex sample such as genomic DNA, or cDNAs that correspond to rare mRNA transcripts. PCR products can be digested and ligated by traditional means, ligated directly (blunt or TA ends), or used in ligation independent cloning (LIC) or seamless cloning applications, such as Gibson Assembly ® or NEBuilder HIFI DNA Assembly (NEBuilderHiFi.com).

During a typical PCR, template DNA (containing the region of interest) is mixed with deoxynucleotides (dNTPs), a DNA polymerase and primers. Primers are short segments of complementary DNA that base-pair with the template DNA, upstream of the region of interest, and serve as recruitment sites for the polymerase. PCR involves a series of temperature cycles that are controlled automatically by the use of a thermocycler that precisely controls both the reaction temperature and the duration of each temperature step, ensuring efficient amplification (for more details about PCR, see DNA Amplification).

For routine, robust PCR reactions OneTaq DNA Polymerase is the most common choice of enzyme. This polymerase leaves predominantly template-independent single adenines (A) at the 3&rsquo end of the PCR product. For high-fidelity PCR, a proofreading DNA polymerase should be used. Such enzymes do not create single base overhangs, leaving blunt termini. A consideration of the ends of PCR products, including their phosphorylation status, is important to subsequent cloning strategies (see End Modification). When PCR primers include restriction enzyme sites the PCR products can be digested and ligated by traditional means.

Vector molecules for cloning may also be produced by PCR. Restriction sites included in the primers allow generation of sticky ends (single strand overhangs) to facilitate cloning of restriction fragments. Otherwise, a blunt ended vector can be produced by PCR using a high-fidelity proofreading polymerase or by blunting of the single base 3&rsquo overhang produced by Taq polymerase. Reverse transcription of RNA to first strand complementary DNA (cDNA) followed by PCR (RT-PCR) allows cloning of double-stranded DNA molecules that correspond to the gene transcripts (for mRNA, see the cDNA synthesis).

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Cloning strategies, Part 3: Blunt-end cloning

Blunt-end cloning is one of the easiest and most versatile methods for cloning dsDNA into plasmid vectors. It is easy because the blunt-ended insert requires little to no preparation. Read an overview of blunt-end cloning with tips for making this cloning approach successful.

Cloning of double-stranded DNA (dsDNA) molecules into plasmid vectors is one of the most commonly employed techniques in molecular biology. The procedure is used for sequencing, building libraries of DNA molecules, expressing coding and non-coding RNA, and many other applications. Past DECODED articles, available online at www.idtdna.com/DECODED, have reviewed the Gibson Assembly&trade method and cohesive-end cloning techniques. There you will also find additional information on plasmid basics in the article: Cloning strategies, Part 2: Cohesive-end cloning, which may be of benefit to readers of this article. Here we will discuss blunt-end cloning as a third method by which DNA fragments can be cloned into a plasmid vector.

Blunt-end cloning overview

Blunt-end cloning involves ligating dsDNA into a plasmid where both the insert and linearized plasmid have no overhanging bases at their termini. It does not benefit from the hydrogen bond stabilization associated with the complementary overhanging bases used in cohesive-end cloning, but the transient associations of the available 5&rsquo phosphate and 3&rsquo hydroxyl groups are sufficient to produce successful clones in the presence of T4 ligase [1]. An illustration of a basic blunt-end cloning experiment is shown in Figure 1.

Blunt-end cloning is also one of the easiest and most versatile methods for cloning dsDNA into plasmid vectors. It is easy because the blunt-ended insert requires little to no preparation&mdashavoiding the enzymatic digestion and subsequent purification needed for cohesive-end cloning. It is versatile because insert and vector have fewer sequence limitations than other methods. This means that blunt-end cloning has some unique advantages, and also disadvantages, over cohesive-end cloning and isothermal assembly methods.

Considerations

There are several factors that make blunt-end cloning challenging. It is 10&ndash100X less efficient than cohesive-end cloning, resulting in fewer colonies and half of these recombinant colonies have inserts incorporated in the wrong orientation. In addition, a lot of plasmid re-circularization occurs. It can be reduced, but is not easily eliminated, resulting in a lot of empty vectors. Therefore, when using this method, you should screen by restriction digestion or PCR prior to sequence-verification.

A major advantage of blunt-end cloning is that the desired insert does not require any restriction sites in the sequence. This makes blunt-end cloning extremely versatile, simplifies planning, and avoids unwanted, artificial sequence additions that might adversely affect some applications. Also, because the insert does not need to be prepared by restriction digestion, blunt-end cloning has the potential to be significantly faster.

Tips for successful blunt-end ligations

Preparing the vector. The vector can be prepared by digestion if the multiple cloning site (MCS) contains a recognition site for a restriction enzyme that produces blunt ends, such as EcoRV (Figure 1). Restrictions sites that generate sequence overhangs can also be used, followed by removal or filling of the overhangs to create blunts ends. However, this approach is not recommended since there is no good method for assessing the success of the blunting reaction, making it hard to troubleshoot unsuccessful reactions.

Alternatively, linearized plasmid can be prepared for blunt-end cloning by amplification with a high-fidelity polymerase and subsequent PCR using primers designed with their 5&rsquo ends at the desired insertion site. The linearized plasmid product appears as a distinct band on an agarose gel, compared to a smear produced by the supercoiled plasmid template, facilitating troubleshooting. The circular template plasmid is eliminated by digesting with DpnI, or a similar restriction enzyme, that cuts the methylated plasmid leaving the unmethylated PCR product

The plasmid is typically dephosphorylated for ligation and amplification methods. However, it is possible to avoid this requirement (see alternative below).

Designing the insert. The blunt-ended insert needs to be phosphorylated. If you plan to anneal oligonucleotides or use IDT gBlocks ® Gene Fragments as your insert, note that these are not synthesized with 5&rsquo phosphate groups unless requested&mdashmake sure to select the 5&rsquo phosphate option when ordering these sequences for blunt-end cloning. Alternatively, phosphates can be added to the 5&rsquo ends of dsDNA with a simple kinase reaction, for example using the commercially available T4 Polynucleotide Kinase. However, this method is less efficient, particularly when bases other than G are kinase targets.

When the ends of the insert are not blunt, a polishing or filling reaction is required. Examples of ends that need polishing or filling include inserts generated by shearing or sonication, or by Taq polymerase, which preferentially leaves a single adenosine overhang at the 3&rsquo ends inserts produced by restriction digests and some inserts produced by annealing multiple oligonucleotides to create longer products. A number of DNA polymerases will remove DNA overhangs and/or can be used to fill in missing bases if there is a 3&rsquo hydroxyl available for priming. Polymerases for such reactions include T4 DNA polymerase, PFU, and the Klenow Fragment of DNA polymerase I. There are many options and kits available so check with the manufacturers to determine which one works best for your application.

As with other cloning methods, shorter inserts will usually ligate more efficiently than longer ones. Manufacturers typically include size recommendations for plasmid inserts in the plasmid documentation as a useful guide in planning your cloning experiment.

Finally, unlike cohesive-end cloning, blunt-end cloning does not automatically recreate a restriction site following ligation of the insert unless bases are added to the insert sequence to complete the missing site (Figure 1).

Ligation conditions. In blunt-end ligations, the association of 5&rsquo phosphate groups and 3&rsquo hydroxyl groups is more transient than in cohesive-end ligations. Because they lack the hydrogen bond stabilization of cohesive ends, blunt-end ligations are more sensitive to reaction conditions, especially to the concentrations of the reaction components.

The likelihood of an insert associating with a linearized plasmid is increased by having a high concentration of available insert blunt ends. However, the intramolecular circularization of the plasmid, after one end of the insert has been joined, works best at lower concentrations. Insert concentrations that are too high, or overall DNA concentrations that are too high, can result in plasmids with multiple inserts, or concatemers. Although it is not always necessary, some researchers perform a short 1hr incubation with high concentrations of insert and ligase, then dilute the reaction 20X with ligase buffer and allow the ligation to proceed for 4 more hours in order to facilitate the second step in the ligation reaction [1].

T4 ligase quality and concentration are also important. Blunt-end ligations typically take place in the presence of higher concentrations of ligase than cohesive-end ligations. For example, whereas a cohesive-end ligation may use 1 unit T4 ligase/20 &muL reaction, a blunt reaction may use up to 3 units/20 &muL reaction. Commercially available T4 ligases typically state whether they are optimized for blunt-end ligations or not. Follow the manufacturer&rsquos guidelines to determine which ligase is appropriate for your cloning experiment and what concentration to use. Note that Taq ligase as well as several other ligases do not ligate blunt ends.

An alternative blunt-end method. As was mentioned, unless the insert is designed with the necessary bases to recreate the restriction site, the blunt restriction site used to linearize the vector is not normally recreated by the ligation of the insert (Figure 1). This allows for an alternative, less common, blunt-end cloning method that does not require the vector to be dephosphorylated. Instead, it relies on competing digestion and ligation reactions to decrease empty vector background.

In this method, the circularized plasmid and insert are placed in a reaction mixture containing the blunt-end&minusproducing restriction enzyme, as well as the T4 ligase. The circular plasmid is cut and the insert ligated in a single tube reaction. Empty plasmid that is produced by T4 ligation is subsequently re-cut by the restriction enzyme. As long as the insert is not designed to produce that specific restriction site, all circularized plasmids should contain the desired insert. Specifics for this method&mdashalso described with a polishing enzyme component&mdashcan be found in Sambrook and Russell [1]. One reason why researchers may want to avoid this technique is that it involves mixing multiple optimized enzyme buffers, which may adversely affect the activity of individual enzymes. Check with the enzyme manufacturer(s) to determine if your chosen enzymes are compatible with this method.


Molecular Cloning: A Laboratory Manual (Fourth Edition)

Molecular Cloning has served as the foundation of technical expertise in labs worldwide for 30 years. No other manual has been so popular, or so influential. Molecular Cloning, Fourth Edition, by the celebrated founding author Joe Sambrook and new co-author, the distinguished HHMI investigator Michael Green, preserves the highly praised detail and clarity of previous editions and includes specific chapters and protocols commissioned for the book from expert practitioners at Yale, U Mass, Rockefeller University, Texas Tech, Cold Spring Harbor Laboratory, Washington University, and other leading institutions. The theoretical and historical underpinnings of techniques are prominent features of the presentation throughout, information that does much to help trouble-shoot experimental problems.

For the fourth edition of this classic work, the content has been entirely recast to include nucleic-acid based methods selected as the most widely used and valuable in molecular and cellular biology laboratories.

Core chapters from the third edition have been revised to feature current strategies and approaches to the preparation and cloning of nucleic acids, gene transfer, and expression analysis. They are augmented by 12 new chapters which show how DNA, RNA, and proteins should be prepared, evaluated, and manipulated, and how data generation and analysis can be handled.

The new content includes methods for studying interactions between cellular components, such as microarrays, next-generation sequencing technologies, RNA interference, and epigenetic analysis using DNA methylation techniques and chromatin immunoprecipitation. To make sense of the wealth of data produced by these techniques, a bioinformatics chapter describes the use of analytical tools for comparing sequences of genes and proteins and identifying common expression patterns among sets of genes.

Building on thirty years of trust, reliability, and authority, the fourth edition of Molecular Cloning is the new gold standard—the one indispensable molecular biology laboratory manual and reference source.

Highlights of the new edition:

  • Extensive new content: 12 entirely new chapters are devoted to the most exciting current research strategies, including epigenetic analysis, RNA interference, genome sequencing, and bioinformatics
  • Expanded scope: the nucleic-acid-based techniques selected for inclusion have promoted recent advances in gene transfer, protein expression, RNA analysis, and expression of cloned genes
  • Classic content: 10 original core chapters have been updated to reflect developments and innovations in standard techniques and to introduce new cutting-edge protocols
  • Easy-to-follow format: the previous editions’ renowned attention to detail and accuracy are fully retained
  • Essential appendices: an up-to-date collection of reagents, vectors, media, detection systems, and commonly used techniques are included
  • Expanded authorship: chapters and protocols have been specifically commissioned from renowned experts at leading institutions

Praise for the previous edition:

“Any basic research laboratory using molecular biology techniques will benefit from having a copy on hand of the newly published Third Edition of Molecular Cloning: A Laboratory Manual. the first two editions of this book have been staples of molecular biology with a proven reputation for accuracy and thoroughness.” —The Scientist

“In every kitchen there is at least one indispensable cookbook. Molecular Cloning: A Laboratory Manual fills the same niche in the laboratory (with) information to help both the inexperienced and the advanced user. (It) has once again established its primacy as the molecular laboratory manual and is likely to be found on lab benches. around the world.” ——Trends in Neurosciences

“Molecular Cloning: A Laboratory Manual has always been the laboratory mainstay for protocols and techniques. It has a pure-bred ancestry, and the new edition does not disappoint. (It) includes information panels at the end of each chapter that describe the principles behind the protocols. The addition of this information extends Molecular Cloning from an essential laboratory resource into a new realm, one merging the previous prototype with a modern molecular monograph. the next generation of Molecular Cloning not only carries on the proud heritage of the first two editions but also admirably expands on that tradition to provide a truly essential laboratory manual.” —Trends in Microbiology


Joining DNA in vitro to form recombinant molecules

Restriction endonucleasescut at defined sequences of (usually) 4 or 6 bp. This allows the DNA of interest to be cut at specific locations. The physiological function of restriction endonucleases is to serve as part of system to protect bacteria from invasion by viruses or other organisms. (See Chapter 7)

Table 3.1. List of restriction endonucleases and their cleavage sites. A ' means that the nuclease cuts between these 2 nucleotides to generate a 3' hydroxyl and a 5' phosphate.
Enzyme Site Enzyme Site
AluI AG'CT NotI GC'GGCCGC
BamHI G'GATCC PstI CTGCA'G
BglII A'GATCT PvuII CAG'CTG
EcoRI G'AATTC SalI G'TCGAC
HaeIII GG'CC Sau3AI 'GATC
HhaI GCG'C SmaI CCC'GGG
HincII GTY'RAC SpeI A'CTAGT
HindIII A'AGCTT TaqI T'CGA
HinfI G'ANTC XbaI T'CTAGA
HpaII C'CGG XhoI C'TCGAG
KpnI GGTAC'C XmaI C'CCGGG
MboI 'GATC

a. Sticky ends

(1) Since the recognition sequences for restriction endonucleases are pseudopalindromes, an off-center cleavage in the recognition site will generate either a 5' overhang or a 3' overhang with self-complementary (or "sticky") ends.

e.g. 5' overhang EcoRI G'AATTC

(2) When the ends of the restriction fragments are complementary,

the ends can anneal to each other. Any two fragments, regardless of their origin (animal, plant, fungal, bacterial) can be joined in vitro to form recombinant molecules (Figure 3.3).

b. Blunt ends

(1) The restriction endonuclease cleaves in the center of the pseudopalindromic recognition site to generate blunt (or flush) ends.

T4 DNA ligase is used to tie together fragments of DNA (Figure 3.4). Note that the annealed "sticky" ends of restriction fragments have nicks(usually 4 bp apart). Nicks are breaks in the phosphodiester backbone, but all nucleotides are present. Gapsin one strand are missing a string of nucleotides.

T4 DNA ligase uses ATP as source of adenylyl group attached to 5' end of the nick, which is a good leaving group after attack by the 3' OH. (See Chapter 5 on Replication).

At high concentration of DNA ends and of ligase, the enzyme can also ligate together blunt‑ended DNA fragments. Thus any two blunt‑ended fragments can be ligated together. Note: Any fragment with a 5' overhang can be readily converted to a blunt‑ended molecule by fill‑in synthesis catalyzed by a DNA polymerase (often the Klenow fragment of DNA polymerase I). Then it can be ligated to another blunt‑ended fragment.

Linkers are short duplex oligonucleotides that contain a restriction endonuclease cleavage site. They can be ligated onto any blunt‑ended molecule, thereby generating a new restriction cleavage site on the ends of the molecule. Ligation of a linker on a restriction fragment followed by cleavage with the restriction endonuclease is one of several ways to generate an end that is easy to ligate to another DNA fragment.

Annealing of homopolymer tails are another way to joint two different DNA molecules.

The enzyme terminal deoxynucleotidyl transferasewill catalyze the addition of a string of nucleotides to the 3' end of a DNA fragment. Thus by incubating each DNA fragment with the appropriate dNTP and terminal deoxynucleotidyl transferase, one can add complementary homopolymers to the ends of the DNAs that one wants to combine. E.g., one can add a string of G's to the 3' ends of one fragment and a string of C's to the 3' ends of the other fragment. Now the two fragments will join together via the homopolymer tails.

Figure 3.5. Use of linkers (left) and homopolymer tails (right) to make recombinant DNA molecules.


APPLICATIONS AND IMPORTANCE OF ENZYMES USED IN MOLECULAR BIOLOGY EXPERIMENTS

RESTRICTION ENDONUCLEASE: Restriction endonucleases are enzymes that cut nucleic acids (inclusive of DNA and RNA molecules) at specific sites. With restriction enzymes or endonucleases, molecular biologists can cut or nick DNA in a precise and reproducible manner which is crucial for gene cloning techniques and other molecular biology experimentation. Restriction endonucleases are DNA cutting enzymes specifically found and isolated from bacteria and which nick specific sites on a nucleotide sequence known as restriction sites. Restriction sites are the different sites on a DNA molecule that is nicked by a particular restriction enzyme. Several types of restriction enzymes exist, and they are primarily sourced from microorganisms (particularly bacteria) but they can also be chemically synthesized. Restriction enzymes or endonucleases are usually divided into three (3) groups viz: Type I, Type II, and Type III restriction enzymes (REs). Type I and Type III REs bind to the nucleic acid molecule at their recognition sequences but they specifically nick the DNA molecule at a considerable distance away from the restriction sites. However, Type II REs are much more applicable for a variety of molecular biology manipulations because they cut DNA molecules at exactly their restriction sites.

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Some examples of restriction enzymes are EcoRI (from E. coli), HaeIII (from Haemophilus aegyptius), BamHI (from Bacillus amyloliquefaciens), PvuI (from Proteus vulgaris), HindII (from Haemophilus influenzae)and Sau3A (from S. aureus) amongst others. The nomenclature of restriction endonucleases is pretty simple. Restriction enzymes are generally named after the bacteria they were actually isolated from. The first three alphabets in the name of a restriction enzyme represent the generic and species name of the bacteria while the remaining alphabets or numbers attached to the name represent the order of discovery of the enzymes in that particular organism. These later alphabets or numbers (usually in Roman numerals) may also represent the strain designate of the restriction endonucleases and the type. For example, EcoRI is E. coli restriction enzyme I. Restriction endonucleases are applied in many molecular biology techniques including but not limited to gene cloning, gene amplification, DNA sequencing and in blotting techniques.

ALKALINE PHOSPHATASE: Alkaline phosphatase is an enzyme produced by organisms such as E. coli and the intestinal tissue of calf, and which removes the phosphate group that is present at the 5′ end of a DNA molecule. It specifically removes phosphate groups from the 5′ terminus of DNA to give the 5′-OH terminus that can join to the 3′ end of the vector that transports an exogenous DNA into a recipient host cell. Thus, preventing unwanted re-ligation or re-joining of already nicked or cut DNA molecules.

DNA LIGASE: DNA ligase is an enzyme that specifically joins cut (nicked) DNA molecules together. They repair and join two individual single-stranded DNA fragments (i.e. the cloning vector and the DNA molecule to be cloned) cut by the same restriction endonuclease. Ligation carried out by DNA ligase enzyme is usually the last step in the gene cloning technique prior to the transformation of the recipient bacterial cell. The joining of the DNA molecule to be cloned with the vector leads to the formation of a new molecule known as the recombinant DNA (rDNA) molecule. Recombinant DNA molecule is a DNA molecule that is created by the ligation of a vector with a cut DNA molecule, and this normally occurs in a test tube where the cut DNA, vector and the DNA ligase enzymes are mixed together for the reaction to occur. In gene cloning technique for example, the same restriction enzyme used to cut the DNA of interest should be used to cut the vehicle or vector meant to carry the gene of interest into the recipient host cell. Using the same restriction enzyme to do the nicking or cutting will ensure that one part is not abnormally cut, but are equally nicked so that they can be properly ligated or joined by the DNA ligase enzyme, which is mainly sourced from a genetically modified E. coli.

Taq POLYMERASE: Taq polymerase enzyme is a heat-stable DNA polymerase enzyme that is isolated from a thermostable bacteria (particularly Thermus aquaticus), and which is used in PCR techniques to extend primers along the single stranded DNA molecule in the 5′-3′ direction.

DNA POLYMERASE I: DNA polymerase I is an enzyme that synthesizes DNA molecules complementary to a DNA template in the 5′-3′ direction. It generally synthesizes DNA on a DNA or RNA template. The DNA polymerase I usually start from an oligonucleotide primer with a 3′ OH terminus and it is used to extend the oligonucleotide primers along the single stranded DNA molecules. DNA polymerases are synthesized by E. coli, and they perform similar activity like the Taq polymerase enzyme. They base-pair and polymerizes the growing DNA strands until the termination site is reached.

RNase ENZYME: RNase enzyme is a nuclease enzyme which digests RNA.

DNase ENZYME: DNase enzyme is a nuclease enzyme which digests DNA.

NUCLEASES: Nucleases are degradative enzymes that cut, shorten and degrade nucleic acids (inclusive of DNA and RNA). They are usually used in PCR reaction and other molecular biology experimentations to digest nucleic acid molecules.

Alberts B, Bray D, Lewis J, Raff M, Roberts K and Watson J.D (2002). The molecular Biology of the Cell. Fourth edition. New York, Garland, USA.

Cooper G.M and Hausman R.E (2004). The cell: A Molecular Approach. Third edition. ASM Press.

Dale J (2003). Molecular genetics of bacteria. Jeremy W. Dale and Simon Park (4 th eds.). John Wiley & Sons Ltd, West Sussex, UK. Pp. 312-313.

Lewis R (2004). Human Genetics: Concepts and Applications. Sixth edition. McGraw Hill Publishers, USA.

Robert L. Nussbaum, Roderick R. McInnes and Huntington F. Willard (2001). Genetics in Medicine. Philadelphia, USA. Saunders publishers.

Sambrook, J., Russell, D.W. (2001). Molecular Cloning: a Laboratory Manual, 3rd edn. Cold Spring Harbor Laboratory Press, New York.

Tamarin Robert H (2002). Principles of Genetics. Seventh edition. Tata McGraw-Hill Publishing Co Ltd, Delhi.

Twyman R.M (1998). Advanced Molecular Biology: A Concise Reference. Bios Scientific Publishers. Oxford, UK.


What is the difference between sticky ends and blunt ends?

Restriction enzymes cut double-stranded DNA in half. Depending on the restriction enzyme, the cut can result in either a sticky end or a blunt end. Sticky ends are more useful in molecular cloning because they ensure that the human DNA fragment is inserted into the plasmid in the right direction.

Beside above, what are blunt ends used for? Blunt ends have no overhang. They cannot match up as specifically as DNA with sticky ends however, they can be useful when sticky ends can't be used. SmaI is a restriction enzyme that makes blunt ends.

Just so, what is the difference between blunt ends and sticky ends quizlet?

Sticky ends are when the enzymes make staggered cuts in the two strands. Cuts that are not directly opposite of each other. Sticky ends are most useful in rDNA because they can be used to join two different pieces of DNA that were cut by the same restriction enzyme.

How do you convert blunt ends to sticky ends?

Blunt and sticky ends can be inter converted by either adding restriction cognitive sites or by removing some of them. Blunt end are converted into sticky end prior to cloning.


Methods

Cell lines and media

The E. coli HB101 was used for the preparation of plasmid DNA. The bacteria were cultured in Luria-Bertani (LB) media. Human embryonic kidney (HEK) 293 T cells were cultured in Dulbecco’s Modified Eagle medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, 2 mM L-glutamine, 100 mg/ml streptomycin, and 100 units/ml penicillin. A myeloid leukemia cell line C1498 [52], was cultured in Roswell Park Memorial Institute (RPMI) 1640 medium supplemented with the same reagents used for DMEM. Cells were split every other day to keep them on log phase.

Plasmids, primers, PCR and sequencing

A plasmid containing the coding sequence of the tdTomato gene, plasmid containing an alpha-retroviral vector, and plasmids containing codon-optimized alpharetroviral gag/pol [53] were kindly provided by Axel Schambach (MHH Hannover, Germany). A forward (5′- ACCGGTGCCACCATGGCCACAACCATGGTG-3′) and a reverse (5′-GTCGACTTACTTGTACAGCTCGTCCATGCC-3′) primer used for the amplification of the tdTomato gene were synthesized by Eurofins Genomics (Ebersberg, Germany).

The optimal buffers for enzymes or other reagents were provided by the manufacturers along with the corresponding enzymes or inside the kits. If available by the manufacturers, the pH and ingredients of buffers are mentioned. Primers were dissolved in ultrapure water at a stock concentration of 20 pmol/μl. The template plasmid was diluted in water at a stock concentration of 50 ng/μl. For PCR, the following reagents were mixed and filled up with water to a total volume of 50 μl: 1 μl plasmid DNA (1 ng/μl final concentration), 1.25 μl of each primer (0.5 pmol/μl final concentration for each primer), 1 μL dNTP (10 mM each), 10 μl of 5X Phusion HF buffer (1X buffer provides 1.5 mM MgCl2), and 0.5 μl Phusion DNA polymerase (2U/μl, Thermo Scientific).

PCR was performed using a peqSTAR thermocycler (PEQLAB Biotechnologie) at: 98°C for 3 minutes 25 cycles at 98°C for 10 seconds, 66°C for 30 seconds, 72°C for 30 seconds and 72°C for 10 minutes. To prepare a 0.8% agarose gel, 0.96 g agarose (CARL ROTH) was dissolved in 120 ml 1X TAE buffer (40 mM Tris, 20 mM acetic acid, 1 mM EDTA, pH of 50X TAE: 8.4) and boiled for 4 minutes. Then 3 μl SafeView nucleic acid stain (NBS Biologicals) was added to the solution and the mixture was poured into a gel-casting tray.

DNA was mixed with 10 μl loading dye (6X) (Thermo Scientific) and loaded on the agarose gel (CARL ROTH) using 80 V for one hour in TAE buffer. The separated DNA fragments were visualized using an UV transilluminator (365 nm) and quickly cut to minimize the UV exposure. DNA was extracted from the gel slice using Zymoclean™ Gel DNA Recovery Kit (Zymo Research). The concentration of DNA was determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific).

For sequence validation, the PCR product was subcloned using CloneJET PCR cloning kit (Thermo Scientific). 1 μl of blunt vector (50 ng/μl), 50 ng/μl of the PCR product, and 10 μl of 2X reaction buffer (provided in the kit) were mixed and filled with water to a total volume of 20 μl. 1 μl of T4 DNA ligase (5 U/μl) was added to the mixture, mixed and incubated at room temperature for 30 minutes. For bacterial transfection, 10 μl of the mixture was mixed with 100 μl of HB101 E. coli competent cells and incubated on ice for 45 minutes. Then the mixture was heat-shocked (42°C/2 minutes), put on ice again (5 minutes), filled up with 1 ml LB medium and incubated in a thermomixer (Eppendorf) for 45 minutes/37°C/450RPM. Then the bacteria were spun down for 4 minutes. The pellet was cultured overnight at 37°C on an agarose Petri dish containing 100 μg/mL of Ampicillin. The day after, colonies were picked and cultured overnight in 3 ml LB containing 100 μg/mL of ampicillin.

After 16 hours (overnight), the plasmid was isolated from the cultured bacteria using the QIAprep spin miniprep kit (QIAGEN) according to the manufacturer’s instructions. 720 to 1200 ng of plasmid DNA in a total of 12 μl water were sent for sequencing (Seqlab) in Eppendorf tubes. The sequencing primers pJET1.2-forward (5′-CGACTCACTATAGGGAG-3′), and pJET1.2-reverse (5′-ATCGATTTTCCATGGCAG-3′), were generated by the Seqlab Company (Göttingen, Germany). An ABI 3730XL DNA analyzer was used by the Seqlab Company to sequence the plasmids applying the Sanger method. Sequence results were analyzed using NCBI Blast as explained in the Results and discussion section.

Manipulation of DNA fragments

For viewing plasmid maps, Clone Manager suite 6 software (SciEd) was used. Restriction endonuclease enzymes (Thermo Scientific) were used to cut plasmid DNA. 5 μg plasmid DNA, 2 μl buffer O (50 mM Tris–HCl (pH 7.5 at 37°C), 10 mM MgCl2, 100 mM NaCl, 0.1 mg/mL BSA, Thermo Scientific), 1 μl SalI (10 U), and 1 μl AgeI (10 U) were mixed in a total of 20 μl water and incubated (37°C) overnight in an incubator to prevent evaporation and condensation of water under the tube lid. The next day, DNA was mixed with 4 μl loading dye (6X) (Thermo Scientific) and run on a 0.8% agarose gel at 80 V for one hour in TAE buffer. The agarose gel (120 ml) contained 3 μl SafeView nucleic acid stain (NBS Biologicals). The bands were visualized on a UV transilluminator (PEQLAB), using a wavelength of 365 nm, and quickly cut to minimize the UV damage. DNA was extracted from the gel slices using the Zymoclean™ gel DNA recovery kit (Zymo Research). The concentration of DNA was determined using a NanoDrop 2000 spectrophotometer (Thermo Scientific).

For the ligation of vector and insert fragments, a ligation calculator was designed (the Excel file available in the Additional file 1) for easy calculation of the required insert and vector volumes. The mathematical basis of the calculator is inserted into the excel spreadsheet. The size and concentration of the vector and insert fragments and the molar ratio of vector/insert (normally 1:3) must be provided for the calculation. Calculated amounts of insert (tdTomato) and vector (alpha-retroviral backbone) were mixed with 2 μl of 10X T4 ligase buffer (400 mM Tris–HCl, 100 mM MgCl2, 100 mM DTT, 5 mM ATP (pH 7.8 at 25°C), Thermo Scientific), 1 μl of T4 ligase (5 U/μl, Thermo Scientific), filled up to 20 μl using ultrapure water and incubated overnight at 16°C. The day after, HB101 E. coli was transfected with the ligation mixture as mentioned above. The clones were picked and consecutively cultured for one day in LB medium containing ampicillin. Plasmid DNA was isolated using Zyppy™ plasmid miniprep kit (Zymo Research) and digested with proper restriction enzymes for screening. Digested plasmids were mixed with the loading dye and run on an agarose gel as mentioned above. The separated DNA fragments were visualized using a Gel Doc™ XR+ System (BIO-RAD) and analyzed by the Image Lab™ software (BIO-RAD). The positive clone was cultured overnight in 450 ml LB medium containing ampicillin. Plasmid DNA was isolated using QIAGEN plasmid maxi kit (QIAGEN), diluted in ultrapure water and stored at −20°C for later use.

Production of viral supernatant and transduction of cells

HEK293T cells were thawed, split every other day for one week and grown in log phase. The day before transfection, 3.5 × 10 6 cells were seeded into tissue culture dishes (60.1 cm 2 growth surface, TPP). The day after, the cells use to reach about 80% confluence. If over confluent, transfection efficiency decreases. The following plasmids were mixed in a total volume of 450 μl ultrapure water: codon-optimized alpharetroviral gag/pol (2.5 μg), VSVG envelope (1.5 μg), and the alpharetroviral vector containing the tdTomato gene (5 μg). Transfection was performed using calcium phosphate transfection kit (Sigma-Aldrich). 50 μl of 2.5 M CaCl2 was added to the plasmid DNA and the mixture was briefly vortexed. Then, 0.5 ml of 2X HEPES buffered saline (provided in the kit) was added to a 15 ml conical tube and the calcium-DNA mixture was added dropwise via air bubbling and incubated for 20 minutes at room temperature. The medium of the HEK293T cells was first replaced with 8 ml fresh medium (DMEM containing FCS and supplement as mentioned above) containing 25 μM chloroquine. Consecutively the transfection mixture was added. Plates were gently swirled and incubated at 37°C. After 12 hours, the medium was replaced with 6 ml of fresh RPMI containing 10% FCS and supplements. Virus was harvested 36 hours after transfection, passed through a Millex-GP filter with 0.22 μm pore size (Millipore), and used freshly to transduce C1498 cells. Before transduction, 24 well plates were coated with retronectin (Takara, 280 μl/well) for 2 hours at room temperature. Then, retronectin was removed and frozen for later use (it can be re-used at least five times) and 300 μl of PBS containing 2.5% bovine serum albumin (BSA) was added to the wells for 30 minutes at room temperature. To transduce C1498 cells, 5 × 10 4 of cells were spun down and resuspended with 1 ml of fresh virus supernatant containing 4 μg/ml protamine sulfate. The BSA solution was removed from the prepared plates and plates were washed two times with 0.5 ml PBS. Then cells were added to the wells. Plates were centrifuged at 2000RPM/32°C/90 minutes. Fresh medium was added to the cells the day after.

Flow cytometry and fluorescence microscope

For flow cytometry assessment, cells were resuspended in PBS containing 0.5% BSA and 2 mM EDTA and were acquired by a BD FACSCanto™ (BD Biosciences) flow cytometer. Flow cytometry data were analyzed using FlowJo software (Tree Star). Imaging was performed with an Olympus IX71 fluorescent microscope equipped with a DP71 camera (Olympus). Images were analyzed with AxioVision software (Zeiss). Fluorescent images were superimposed on bright-field images using adobe Photoshop CS4 software (Adobe).


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