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Purifying a linear plasmid after restriction digest?

Purifying a linear plasmid after restriction digest?


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I expressed a yeast vector in E.coli and purified about 13µg of it. I then linearized it using a restriction enzyme, and attempted to gel purify it. I attempted this twice. The gel showed a clear bright band at the right spot, but after gel purification I end up with only 0.7µg of linear plasmid. I doubt I made the same mistake twice. What is the best way to purify a linear plasmid after digest?

Edit:

The plasmid is about 3.5 kb. I incubate an overnight culture of E. coli, miniprep the following morning, elute into water. I end up with 13µg of DNA in 200µl of H2O. I add the PmeI (10µl) and the CutSmart buffer (23µl) from NEB, incubate overnight. The following day, I run the 230µl through a 1% gel and get a single band around 3.5 kb. I excise the band, and perform a gel purification using the Biobasic kit, add denaturant (probably guanidine hydrochloride), wait for 10 minutes for the gel to dissolve, run through column, wash 2X with wash buffer with ethanol added, elute in water.

Thanks, Max


Can't exactly say what problem you are facing but you can follow these steps for efficient restriction digestion and elution.

  • Never set up a digestion reaction with more than 1µg DNA per 20µl; if you require more digested DNA, then set up multiple 20µl reactions with maximum of 1µg DNA in each (500-700ng is ideal).
  • 10µl enzyme for a 230µl reaction: bad idea. Set up 10×20µl reactions with not more than 0.5µl enzyme per reaction tube.
  • Avoid overnight incubations; 4 hours should be sufficient (NEB PmeI is 10 units/µl and 1 unit is the amount required to cleave 1µg DNA in 1h in a 50µl reaction. You are adding 5 units in 20µl. Overnight incubation is too much)
  • Deactivate the enzyme after the reaction.
  • Just run a little bit of sample from one tube in the gel to see if digestion has happened. After confirming digestion simply use the PCR purification kit. You can pool your tubes.
  • Optional: Elute in Tris-Cl pH 7.5 instead of water or use warm (60⁰C) water.

Why do a gel purification? you can just use a kit similar to QIAGEN gel extraction kit and do a non gel based plasmid purification and elution, using the buffers used as usual! PCR DNA clean up kits will do as well!


Purifying a linear plasmid after restriction digest? - Biology

Isolated plasmid when mostly in supercoiled form eases most of the downstream processes, but is it essential for restriction digestion also? If there is less supercoiled and more of nicked circular or linear plasmid then it affects transfection efficiency, how does it affect digestion efficiency? Will it be not digested at all? If digested, can't it be used for cloning purposes?

Restriction enzymes are the most effective on the less structured DNA. And contrary to the upper message, there are cases where supercoiled DNA is far less susceptible to digest because DNA structure may hide restriction enzyme cleavage site. NEB has established a table comparing enzyme activity on supercoiled plasmids and on linear lambda DNA:

Digestion with restriction enzymes is not affected if the DNA is supercoiled. For greater efficiency heat the DNA at 94 ° C for 10 minutes to break the supercoiled structure

Restriction enzymes are the most effective on the less structured DNA. And contrary to the upper message, there are cases where supercoiled DNA is far less susceptible to digest because DNA structure may hide restriction enzyme cleavage site. NEB has established a table comparing enzyme activity on supercoiled plasmids and on linear lambda DNA:

Even if you suspect the efficiency of digestion by enzyme, u can linearize DNA and then digest.

True. I agree with the above answers. I would say that during restriction digestion, the concentration of DNA matters more than the form of DNA template taken. Depending on the type of DNA template, the concentration needs to be adjusted to get efficient digestion products. Even if you find some undigested vector on the top in the gel, (In the case of supercoiled DNA as a template), the required digested product (band of your interest) can be gel purified and can very well be used for cloning purposes, It works.

Hi Tias, Dominique is correct regarding restriction digests.

Regarding transfections, supercoiled DNA transforms

100x more efficiently than linear (relaxed/nicked is intermediate). This is due to the increased space that the relaxed and linear forms take, making it more difficult to a charged molecule through the lipid membrane. However, linear DNA integrates


Using plasmids for cloning

One of the more common techniques available to scientists working in molecular biology is cloning. In this technique, sequences of DNA containing genes of interest are inserted into vectors which are then used to introduce these genes into cells or organisms to study the effects of the expression of the genes.

Vectors are based on bacterial plasmids – short circular pieces of DNA separate to the main bacterial chromosome which may be transferred between bacteria. Scientists source plasmid vectors from biological supply companies, which create them by ligating together pre-existing genes and sequences of DNA built from scratch using sequencing technology. A map of an example of a commercial vector (the pGEM-T Easy system) is presented in the figure below.

Note that this map shows a number of regions contained within the vector. The numbers refer to how many base pairs along the sequence (out of a total of 3015) a particular region is found.

In our experiment, the pGEM-T Easy vector has had a short fragment of the gene for the protein polo-like kinase 1 (PLK1) cloned in at the insertion site (located at around “3 o’clock” if you imagine the picture of the vector to be a clock face). This portion of the gene codes for a region of the protein called the polobox domain, which assists in the localization of the protein at various point during the cell cycle.

Prior to today’s project, this cloned vector was used to transform E. coli cells. The cells were used to inoculate an agar plate containing ampicillin and the plates incubated overnight. Because of the presence of the ampicillin resistance gene in the pGEM-T vector, the only cells to grow into colonies were those that had been transformed by the plasmid. These colonies were then used to inoculate a culture broth, which has been provided to you.

The alkaline lysis mini-plasmid preparation

Using transformed bacteria is a very efficient means to generate DNA needed for research. Bacteria have minimal requirements for nutrition and so the production of large quantities of DNA can be done quickly and at a minimal cost. Once a culture of transformed cells is established, this culture can be used to seed new cultures, so transformation need only be performed once. However once we have our culture, we need a way of recovering the DNA in a relatively pure form.

DNA (including plasmid DNA) is not generally secreted by cells. In order to recover it, we need to disrupt the cells and then purify the DNA we need from the other cellular contents. This is the purpose of the alkaline lysis mini-plasmid preparation (or mini-prep).

The first stage of the mini-prep involves bursting the cells using an alkaline solution. This releases their contents into the surrounding liquid. An acidic solution is then added, which neutralizes the alkaline solution and denatures the proteins, causing them to become insoluble. They can be removed from the cell lysate through centrifugation (see figure below).

The second stage of the mini-prep involves passing the cell lysate through a column. These columns bind onto plasmid DNA, and allow chromosomal DNA and other cell products to pass through. After washing the column several times, we can make the column release the plasmid DNA by passing through an elution solution (see figure below).

Restriction digests

By the end of the mini-prep procedure, you should have approximately one drop of a colourless liquid. To demonstrate that you have recovered the plasmid DNA you will need to run the sample on an electrophoresis gel. However, before your sample is ready to run, you must first prepare it using a restriction digest.

Plasmid DNA is circular. For DNA to be demonstrated on a gel, it needs to be linearised. This is done by using enzymes to cut the DNA (think of cutting a rubber band once to obtain a straight strip of rubber). Restriction endonucleases are enzymes which cut the DNA strand at very specific locations, normally given by sequences of half a dozen or so base pairs called restriction sites. If we know the sequence of a length of DNA, we can select enzymes which cut the DNA once (ie. the restriction site sequence occurs once in the entire DNA sequence) or even twice. If a plasmid is cut twice, you should end up with DNA fragments of two different sizes. Molecular biologists often use this double cutting to “drop out” an insert they have cloned into a vector.

The pGEM-T Easy vector has been created with a number of restriction sites on either side of the insertion point. Some of these restriction sites are only found on one side of the insertion point, and so can be used to linearise the vector to make it ready for electrophoresis. Others are found on both sides, and so may be used to drop out the insert to check its size. In this exercise, we will be using the enzyme EcoRI which has restriction sites just upstream (before) and downstream (after) the insertion site. This will allow us to drop out the polobox insert, resulting in DNA of two different sizes : around 3015 base pairs long for the vector, and 800 base pairs long for the insert. The two linearised fragments are now ready to be demonstrated using electrophoresis.

Further information on restriction digests can be found here.

Agarose gel electrophoresis

If we wanted to sort sand from gravel from larger rocks, we would use a series of sieves of different sizes. Each sized sieve lets smaller particles pass through but retains the larger fragments. Electrophoresis can be thought of as a sieve for large molecules like DNA or protein.

In agarose gel electrophoresis, a DNA sample is loaded towards one end of a block of a jelly-like substance called agarose. When an electrical current is passed through the gel, the DNA molecules are pushed through the gel away from the negative electrode (DNA has an overall negative charge and like charges repel). Smaller fragments of DNA can move more easily through the gel than larger fragments, so in a given period of time, DNA of different sizes accumulates in regions of the gel. If we include a dye which binds to the DNA, , these regions are visible as bands – the further towards the positive electrode a band is located, the smaller the fragments of DNA are found in that band.

To get an idea of the size of a band seen on a gel, we always run a sample consisting of a mixture of DNA fragments of known sizes alongside our test samples. This is called a marker, or a “ladder”, as the multiple bands of DNA seen on the gel resembles the rungs on a ladder. By matching the position of a band in our test sample to those representing DNA of known size in the ladder, we can estimate the size of DNA fragments in our test. The part of the PLK1 gene which codes for the polobox domain is 800 base pairs (bp) long. Therefore, if our digest has been a success, we should see a band corresponding to our DNA markers which is 800 base pairs long representing the polo-box insert, and another representing the pGEM-T vector at 3000 base pairs long.


Restriction Cloning Tips

For many applications, conventional restriction cloning is still the best method. A few simple tricks will help to ensure that your cloning goes smoothly.

  1. First and foremost, be careful at each step of a procedure. This approach saves time in the long run.
  2. Make sure the DNA is very clean. Start with about 2 μg of DNA when preparing a vector or excising a fragment to be inserted.
  3. After each enzymatic reaction, purify the DNA with a spin-column. Elute the DNA in 40 – 45 μl of 10 mM Tris (pH 8.5), and then perform the next reaction. (There is no need to use a spin-column before purifying a DNA fragment with a gel.)
  4. To maximize efficiency, minimize the number of steps in a procedure. For example, it works better to clone a blunt-ended fragment into a blunt vector site, such as a SmaI site, than into a site that has been blunted with Klenow or T4 DNA polymerase.
  5. When in doubt, purify a DNA fragment with a gel. Cleaner starting material will yield a better outcome.

The most common problem with restriction cloning is that the starting vector is recovered after the procedure. This problem has two causes: incomplete digestion of the vector, and re‑ligation of the cut vector with itself. The tricks described below will minimize these effects.

    Ensure that digestion of the vector goes to completion. Use an excess of restriction enzyme (

20 U for 2 μg DNA), and digest for

With the vector, ensure that the digestion and phosphatase reactions go to completion. Vortex the mixtures thoroughly and repeatedly, and do not allow any droplets of liquid to escape enzyme treatment. It is a good idea to spin briefly after vortexing to ensure that no droplets are left on the side of the tube.

Preparing the Inserted Fragment

  1. For preparation of an inserted fragment, incomplete digestion is not a serious concern because partial recovery of the fragment is acceptable. You can use relatively short digestion times, and for a double digest, you can use a reaction buffer that is suboptimal for one or both enzymes.
  2. Purify the inserted fragment with a gel. In addition to removing unwanted or incompletely digested DNA, this procedure removes enzymes and small molecules. When visualizing DNA bands with a transilluminator, do not use short wavelength UV light of 312 nm or below because the DNA will be severely damaged. Use 360-365 nm UV light instead.
  3. If the inserted fragment was generated by PCR, purify it with a gel or spin-column before doing any restriction digests. If you plan to clone a blunt-ended PCR fragment (generated with a polymerase such as Pfu) into a phosphatase-treated vector, ensure that your PCR primers have 5′-phosphates.

Ligation and Transformation

  1. With a phosphatase-treated vector, perform a control ligation in which the inserted fragment is omitted. This control ligation should yield very few transformants because only circular DNA molecules transform E. coli efficiently, and re‑circularization of the vector cannot occur without ligation to a phosphorylated fragment.
  2. If the DNA has only sticky ends, ligate at room temperature for


Molecular cloning of PCR products: Restriction digestion guide

Cloning is a ubiquitous multi-step technique in molecular biology labs, and involves inserting a target gene (insert) into a circular, double-stranded, self-replicating plasmid vector backbone that is carrying an antibiotic resistance gene (most often ampicillin or kanamycin). This is made possible by restriction digestion, a process that ensures that the target gene and the receiving vector have compatible ends. Following ligation of the gene into the vector, the resulting recombinant DNA molecule is introduced into Escherichia coli cells (transformation), and antibiotic pressure is applied to select those bacteria that are carrying the plasmid. These transformed bacteria can then be used to replicate the recombinant plasmid DNA (plasmid preparation). In other cases, the bacterial cellular machinery can be directed to express the target gene and synthesize its corresponding protein (protein expression).

Cloning is a ubiquitous multi-step technique in molecular biology labs, and involves inserting a target gene (insert) into a circular, double-stranded, self-replicating plasmid vector backbone that is carrying an antibiotic resistance gene (most often ampicillin or kanamycin). This is made possible by restriction digestion, a process that ensures that the target gene and the receiving vector have compatible ends. Following ligation of the gene into the vector, the resulting recombinant DNA molecule is introduced into Escherichia coli cells (transformation), and antibiotic pressure is applied to select those bacteria that are carrying the plasmid. These transformed bacteria can then be used to replicate the recombinant plasmid DNA (plasmid preparation). In other cases, the bacterial cellular machinery can be directed to express the target gene and synthesize its corresponding protein (protein expression).

In the following series of posts, I will explore each step (beginning with restriction digestion) and discuss tips to ensure success and minimal troubleshooting.

Restriction digestion

1. Restriction endonucleases: REs are bacterial enzymes that recognize short DNA sequences (6-8 base pairs) called restriction sites, and cleave (digest) double-stranded DNA at these sites. Most REs are palindromic: much like language palindromes (Madam I’m Adam), these enzymes read the same sequences on both DNA strands in the 5’ to 3’ direction. Some enzymes create single-stranded overhangs, also called sticky or cohesive ends. Other enzymes cut straight down the middle of a target sequence and leave no overhangs, but blunt or flush ends. For cloning purposes, REs that create sticky ends are more often used because they result in a much higher cloning efficiency.

2. Methodology: To prepare the insert (e.g. a PCR product) for cloning, it is most often cut with two different REs, and these same REs are used to digest the vector. This guarantees the production of non-compatible ends within the same molecule, forces the insert to be cloned in one direction (directional cloning), and prevents self-ligation of the vector. On the other hand, because the insert and the vector are digested with the same enzymes, insert-vector compatible tails are created. These ends can base pair, and can later be joined by DNA ligase, thus forming a chimeric molecule. If the insert is digested with only one enzyme, the same ends will be created on both ends, and the target can be inserted in either direction. Any obtained clones will need to be screened to determine that the gene orientation is correct. A vector digested with only one enzyme will need to be de-phosphorylated to prevent self-ligation. A vector that ligates upon itself will get transformed more efficiently into E. coli and will thus decrease the efficiency of the cloning reaction.

3. Tips for a successful restriction digest:

a. If your insert is a PCR product, you will probably add the restriction sites to the 5’ end of both PCR primers. To ensure efficient binding and digestion, make sure to include six bases between the recognition site and the 5’ end of the primer. Remember to confirm that the restriction sites you select do not occur within your insert!

b. Make sure the DNA is free of contaminants, such as phenol, chloroform, ethanol, and salt. If using a PCR product, you can use a PCR clean-up kit. If your PCR reaction generates a nonspecific band, you will have to cut the target band out of the gel and purify the product using a DNA gel extraction kit.

c. The amount of enzyme used should not exceed 10% of the reaction volume, because glycerol carried over from the enzyme storage buffer may inhibit the reaction.

d. Add the reaction components in this order: restriction enzyme buffer, water (the enzyme may get denatured if added directly after the concentrated buffer), DNA, enzyme.

e. Keep the enzyme on ice or in a cooling block while you set up the reaction.

f. Mix the reaction components by gently pipetting the reaction mixture up and down. Avoid generating air bubbles because the enzyme may get trapped at the liquid-air interface and become denatured. Follow with a quick spin-down in a microcentrifuge. Do not vortex the reaction.

g. Check the success of your digestion reaction by running digested vs. undigested product side-by-side on an agarose gel. NEBcutter is a helpful tool from New England BioLabs that can help you predict the outcome of a digestion reaction.

h. Finally, and before proceeding to ligation, the RE present in the reaction needs to be inactivated. Otherwise, any remaining active RE will digest any ligated products. Many REs can be heat-inactivated. For those that can’t, a gel purification step is recommended.

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Experiment 1:  Restriction Digestion

Restriction Digests begin by mixing the DNA and the RE, but it's unfortunately not quite as simple as that. Restriction Enzymes are delicate and need to be treated carefully. Because enzymes are proteins and proteins denature as the temperature is increased, RE's are always stored in a freezer until they are used. In fact, all of the ingredients in a Restriction Digest are kept on ice until it's time for the reaction to begin. The actual reaction conditions vary from one enzyme to the next, and include temperature, NaCl and/or MgCl 2 concentration, pH, etc. All of these variables except temperature are optimized by mixing the enzyme and DNA with a buffer specific for the enzyme of choice. Once all the ingredients are mixed in the reaction tube, the tube is incubated at the Restriction Enzyme's optimal temperature for 1 hour or longer. Then finally when the digest has run for the appropriate amount of time, the reaction tube is put back on ice to prevent nonspecific degradation of your DNA. Once the Restriction Digest is completed, Agarose Gel Electrophoresis is performed to separate the digest fragments by size and visualize the fragments and perhaps purify them for further experiments.


Plasmid DNA Isolation and Restriction Mapping - PowerPoint PPT Presentation

Plasmid DNA Isolation and Restriction Mapping Plasmid DNA isolation Agarose gel electrophoresis Determine DNA fragment size Restriction . &ndash PowerPoint PPT presentation

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Integrating after CEN Excision (ICE) Plasmids: Combining the ease of yeast recombination cloning with the stability of genomic integration

Randolph Y. Hampton, Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, 9500 Gilman Dr., La Jolla, CA 92093-0347.

Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California

Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California

Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, La Jolla, California

Randolph Y. Hampton, Division of Biological Sciences, Section of Cell and Developmental Biology, University of California San Diego, 9500 Gilman Dr., La Jolla, CA 92093-0347.

Abstract

Yeast recombination cloning is a straightforward and powerful method for recombining a plasmid backbone with a specific DNA fragment. However, the utility of yeast recombination cloning is limited by the requirement for the backbone to contain an CEN/ARS element, which allows for the recombined plasmids to propagate. Although yeast CEN/ARS plasmids are often suitable for further studies, we demonstrate here that they can vary considerably in copy number from cell to cell and from colony to colony. Variation in plasmid copy number can pose an unacceptable and often unacknowledged source of phenotypic variation. If expression levels are critical to experimentation, then constructs generated with yeast recombination cloning must be subcloned into integrating plasmids, a step that often abrogates the utility of recombination cloning. Accordingly, we have designed a vector that can be used for yeast recombination cloning but can be converted into the integrating version of the resulting vector without an additional subcloning. We call these “ICE” vectors, for “Integrating after CEN Excision.” The ICE series was created by introducing a “rare-cutter” NotI-flanked CEN/ARS element into the multiple cloning sites of the pRS series yeast integration plasmids. Upon recovery from yeast, the CEN/ARS is excised by NotI digest and subsequently religated without need for purification or transfer to new conditions. Excision by this approach takes

3 hr, allowing this refinement in the same time frame as standard recombination cloning.

A plasmid that supports lysine prototrophy is not subject to positive selection in medium lacking lysine. (A) Cells bearing a pTDH3::LYS1-GFP YCp were subjected to flow cytometry after outgrowth in medium lacking leucine (red histogram) or lysine (blue histogram). (B) Mean fluorescence after outgrowth in -LEU or -LYS medium was recorded using flow cytometry. N = 3. Bars show the mean of three experiments. Error bars represent standard deviation.

Simplified schematic of a LEU2 ICE plasmid. (A) The key features of an ICE plasmid include a CEN/ARS flanked by NotI sites, defined sequences (dashed line) upstream and downstream to the MCS (denoted by the unique cutter SpeI and XhoI) that can be added to an expression cassette by overhang primers, and a yeast selection cassette (LEU2) with a unique cut site (AgeI) that can be used for plasmid linearization and genomic integration.

Graphical protocol of ICE plasmid workflow. (A) In preparation for YRC, an ICE plasmid is linearized by digestion. Any restriction enzyme with a unique cut site in the MCS is suitable. We recommend using two restriction enzymes, as this can lower the frequency of plasmid closure by non-homologous end joining. (B) To prepare an insert for recombination, amplify Your Favorite Gene (YFG) using PCR. Primers should include overhangs with homology to the ICE plasmid backbone. (C) PCR product and cut backbone are co-transformed into yeast to achieve recombination (dotted lines). We recommend between a 1:9 and 1:18 vector to insert ratio. It should also be noted that recombination often outcompetes non-homologous end joining, and that a lack of enrichment between a cut-vector-only control and a vector-plus-insert transformation should not always be interpreted as an unsuccessful attempt at cloning. (D) Isolated colonies are subjected to a yeast miniprep, and the new plasmid is recovered. At this stage, we recommend performing a diagnostic PCR with yeast miniprep to determine if cloning has been successful. Desired plasmids are then subjected to digestion by NotI, religation by T4 DNA ligase, and transformation into Escherichia coli. We suggest using a no-ligase control as an indicator of successful CEN excision and plasmid religation. A successful product of the ICE plasmid protocol has a single, reconstituted NotI site, which can be used in a diagnostic digest to demonstrate presence or absence of the

500 bp CEN/ARS fragment. (E) The YIp generated by CEN excision is prepared for integration by plasmid linearization. Any restriction enzyme with a unique cut site in the yeast selectable marker is suitable (E.g. AgeI). Upon transformation, the plasmid integrates at the homologous genomic locus by recombination and supports prototrophy or drug resistance. Notably, this method of plasmid integration can lead to tandem plasmid integration. Users should identify strains with a single plasmid integration by PCR, Western blot, or flow cytometry. Our preferred PCR strategy detects a novel junction between multiple plasmid integrations. This novel junction is produced where the plasmid is juxtaposed to an additional plasmid rather than the genome.

Yeast Strains used in this study.

Plasmids used in this study.

Oligonucleotides used in this study.

Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing content) should be directed to the corresponding author for the article.


Midiprep

Maxiprep

ZymoPURE Plasmid Purification Kits Product Description

The ZymoPURE plasmid purification kits are the best method for rapid isolation of transfection ready plasmid DNA. Plasmid purification for Mini preps, Midi preps and Maxi preps is performed in less than 20 minutes and Giga preps in 50 minutes. This family of plasmid purification kits feature a patented binding chemistry that enables simple purification of highly-concentrated (up to 3 mg/ml) endotoxin-free plasmid DNA on a spin-column. The streamlined workflow eliminates slow gravity flow anion-exchange columns and isopropanol precipitation steps found in other kits. ZymoPURE plasmid purification kits reduce processing time by up to 7x using a vacuum manifold or centrifugation. Simply bind, wash and elute transfection ready, endo-free plasmid in minutes.

The ZymoPURE plasmid purification kits are optimized to ensure the eluted DNA is free of endotoxins, salt, protein, and RNA, resulting in plasmids that are suitable for use in sensitive applications. ZymoPURE II plasmid purification kits include the EndoZero columns that enable rapid isolation of endotoxin-free plasmid DNA. The resulting endo-free plasmid DNA is ideal for transfection 1 (including sensitive and primary cells), CRISPR-Cas9 and gene editing 2 , lentiviral vectors 3 , adenovirus vectors 3 and AAV vectors 3 , gene therapy, chimeric antigen receptor (CAR) generation 4 , recombinant antibody generation 5 , in vitro transcription 6 , synthetic biology 7 , PCR 8 , transgenic organism generation 9 and microinjections 10 , molecular cloning 11 , restriction endonuclease digestions, site-directed mutagenesis 12 , plasmid transformation of competent cells, Sanger sequencing 13 , and other sensitive downstream applications.

Applications of Use

  1. Transfection experiments (stable transfection and transient transfection) require highly-concentrated plasmid DNA that is free of salt contaminants and endotoxins otherwise transfection efficiency is reduced. An example of ZymoPURE used in transfections:

Koblan, LW, et al., Improving cytidine and adenine base editors by expression optimization and ancestral reconstruction. Nature Biotechnology, 843-846 (2018).

Findlay, GM, et al. Accurate classification of BRCA1 variants with saturation genome editing. Nature 562, 217-222 (2018).

Hu, JH, et al. Evolved Cas9 variants with broad PAM compatibility and high DNA specificity. Nature, 57-63 (2018).

Salasova, A, et al. A proteomic analysis of LRRK2 binding partners reveals interactions with multiple signaling components of the WNT/PCP pathway. Molecular Neurodegeneration, 12(54) (2017).

Jaitin, D.A, et al. Dissecting immune circuits by linking CRISPR-Pooled Screens with Single-Cell RNA-Seq. Cell, 1883-1896 (2016).

Salasova, A, et al. A proteomic analysis of LRRK2 binding partners reveals interactions with multiple signaling components of the WNT/PCP pathway. Molecular Neurodegeneration, 12(54) (2017).

Chamberlain, K, et al. A Calsequestrin Cis-Regulatory Motif Coupled to a Cardiac Troponin T Promoter Improves Cardiac Adeno-Associated Virus Serotype 9 Transduction Specificity. Human Gene Therapy, 29(8) (2018).

Takemasa, T, et al. Rapid Construction of Antitumor T-Cell Receptor Vectors from Frozen Tumors for Engineered T-Cell Therapy. Cancer Immunology Research 6(5) (2018).

Vazquez-Lombardi, R, et al. Transient expression of human antibodies in mammalian cells. Nature Protocols, 13(1), 99–117 (2017).

Marshall, R, et al. Rapid and Scalable Characterization of CRISPR Technologies Using an E. coli Cell-Free Transcription-Translation System. Molecular Cell, 146-157 (2017).

Peñalber-Johnstone, C, et. al. Optimizing cell-free protein expression in CHO: Assessing small molecule mass transfer effects in various reactor configurations. Biotechnology and Bioengineering, 114(7), 1478–1486 (2017).

Hunter, DJB, et al. Unexpected instabilities explain batch-to-batch variability in cell-free protein expression systems. Biotechnology and Bioengineering, 115(8), 1904–1914 (2018).

Lillacci, G, et al. Synthetic control systems for high performance gene expression in mammalian cells. Nucleic Acids Research 46(18), 9855-9863 (2018).

Champer J, et al. Novel CRISPR/Cas9 gene drive constructs reveal insights into mechanisms of resistance allele formation and drive efficiency in genetically diverse populations. PLoS Genet 13(7), (2017).

Aram, R, et al. Tools for Mos1-mediated single copy insertion (mosSCI) with excisable unc-119(+) or NeoR (G418) selection cassettes. microPublication Biology (2019).

An example of ZymoPURE purified plasmid after Gibson assembly:

Lillacci, G, et al. Synthetic control systems for high performance gene expression in mammalian cells. Nucleic Acids Research, 46 (18), 9855-9863 (2018).

An example of ZymoPURE purified plasmid after Gateway cloning:

Xiong, J, et al. Development of a Time-Resolved Fluorescence Resonance Energy Transfer Ultrahigh-Throughput Screening Assay for Targeting the NSD3 and MYC Interaction. Assay and Drug Development Technologies 16(2), 96-106 (2018).

Conway, JM, et al. Novel multidomain, multifunctional glycoside hydrolases from highly lignocellulolytic Caldicellulosiruptor species. AIChE Journal, 64(12) (2018).

McCullock, TW, et al. Comparing the performance of mScarlet-I, mRuby3, and mCherry as FRET acceptors for mNeonGreen. PLoS ONE, 15(2) (2020).


Purifying a linear plasmid after restriction digest? - Biology

Neoschizomers: BfuCI, Bsp143I, BstENII, BstKTI,BstMBI, DpnII, Kzo9I, NdeII

JBSpeed Restriction Enzyme

For in vitro use only!

Unit Definition: One unit is the amount of enzyme required to completely digest 1 &mug of pBR322 (22 sites) in 1 hour in a total reaction volume of 50 &mul. Enzyme activity was determined in the recommended reaction buffer.

Shipping: shipped on blue ice

Storage Conditions: store at -20 °C
avoid freeze/thaw cycles

Shelf Life: 12 months

Form: liquid (Supplied in 10 mM Tris-HCl pH 7.4, 400 mM KCl, 0.1 mM EDTA, 1 mM DTT, 200 &mug/ml BSA and 50 % [v/v] glycerol)

Concentration: 10 units/&mul

Source: Diplococcus pneumoniae G41, recombinant, E. coli

Supplied with: 10x Universal Buffer (UB)

Recommended 50 &mul assay

5 &mul10x Universal Buffer (UB)
1 &mugpure DNA 1 or PCR product 2
10 unitsenzyme
fill up to 50 &mulPCR grade water
  • The enzyme should not exceed 10 % of total reaction volume.
  • Add enzyme as last component. Mix components well before adding enzyme. After enzyme addition, mix gently by pipetting. Do not vortex.
  • Incubate 5 to 10 min. at 37 °C.
  • Stop reaction by alternatively:
    - Addition of 2.1 &mul EDTA pH 8.0 [0.5 M], final 20 mM
    - Heat Inactivation (20 min. at 80 °C)
    - Spin Column DNA Purification (e.g. PCR Purification Kit, Cat.-No. PP-201S/L)
    - Gel Electrophoresis and Single Band Excision (e.g. Agarose Gel Extraction Kit, Cat.-No. PP-202S/L)
    - Phenol-Chloroform Extraction or Ethanol Precipitation.

Double Digestion - Buffer Compatibility:
B1 - 75-100 % Relative Activity
B2 - 75-100 % Relative Activity
B3 - 50-75 % Relative Activity
B4 - 10 % Relative Activity
B5 - 75-100 % Relative Activity
1x UB - 100 % Relative Activity (recommended)

Please note that the optimum digestion condition for this enzyme is 1x UB. Within the Universal Buffer (UB) system, the most majority of our enzymes display 100% Relative Activity in 1x UB and only few either in 0.5x or 2x UB. If optimum condition for second enzyme is different than the recommended for the first enzyme, we suggest carrying out first the restriction at the higher recommended concentration of UB and dilute the reaction volume to the adequate UB concentration for further proceeding with the second restriction.


Reaction Enzymes Buffer Guide:

Reaction Buffer Compatibility:
Our restriction enzymes are fully compatible to restrictases and buffer systems from other manufacturers and can be used along in double digestions. To obtain best results, consult the corresponding manuals of all involved products.

Ligation and recutting:
After 50-fold overdigestion with DpnI, >70 % of the DNA fragments can be ligated and >95 % of these can be recut.

Methylation dependency:
DpnI is specific for methylated and hemimethylated DNA. Since DNA isolated from most E. coli strains is dam methylated, it is susceptible to DpnI digestion. Hence, DpnI is frequently used after a PCR reaction to digest the methylated parental DNA template and select for the newly synthesized DNA containing mutations.

Quality Control:
All preparations are assayed for contaminating endonuclease, 3'-exonuclease, 5' exonuclease/ 5' phosphatase, as well as nonspecific single- and doublestranded DNase activities.


Watch the video: ΚΑΘΑΡΙΣΜΟΣ ΠΟΤΑΜΩΝ ΣΤΗΝ ΑΡΓΟΛΙΔΑ 2522020 (February 2023).