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How do I reverse my insert within a plasmid?


I have a plasmid with:

… T7pro. -- RBS -- XBa1 cut site -- ProDpro. -- RBS -- AmilCP -- T -- Spe1 cut site -- T…

How do I design primers to reverse the portion in bold, such that the T7 and ProD promoters are on opposite sides of AmilCP? I am finding it difficult to understand how to design the appropriate primers.


Design your 5' primer upstream of ProDpro. with a Spe1 cut site added to the end (plus a few extra random bases to allow for better cutting). Add an Xba1 cut site to the end of the 3' primer, again with a few extra random bases on the end to allow for better cutting.

Amplify the insert, PCR cleanup, digest, cleanup again, and you should be able to easily clone into the Xba1/Spe1 digested vector that has been gel purified.


Talk:Synthetic Biology:Vectors/Single copy plasmid

I was wondering about the arrangement of the VF/VR sites and the terminators. If the terminators were outside the VF/VR sites we would get somewhat longer sequencing reads. The way they are designed currently I suppose there would be less possibility for non-specific transcription so there are arguments either way --BC

    Currently the VF2 and VR sites lie outside the terminators because typically sequence data take at least


Colony PCR

Colony PCR is a convenient high-throughput method for determining the presence or absence of insert DNA in plasmid constructs. Individual transformants can either be lysed in water with a short heating step or added directly to the PCR reaction and lysed during the initial heating step. This initial heating step causes the release of the plasmid DNA from the cell, so it can serve as template for the amplification reaction. Primers designed to specifically target the insert DNA can be used to determine if the construct contains the DNA fragment of interest. Alternatively, primers targeting vector DNA flanking the insert can be used to determine whether or not the insert is the correct molecular size. Insert specific primers can provide information on both the specificity and size of the insert DNA while the use of vector specific primers allows screening of multiple constructs simultaneously. Colony PCR can also be used to determine insert orientation. PCR amplification of the plasmid using an insert specific primer paired with a vector specific primer can be designed to produce an amplicon of a specific size only if the insert is in the correct orientation. In all experimental designs, presence or absence of a PCR amplicon and size of the product are determined by electrophoresis alongside a DNA size marker on an agarose gel.

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DNA provirus hypothesis

In the mid-20th century there were many advances in molecular biology, including the description of DNA in 1953 by American geneticist and biophysicist James D. Watson and British biophysicists Francis Crick and Maurice Wilkins. By the 1960s it was understood that sarcomas are caused by a mutation that results in uncontrolled cell division. It was also evident that RSV was inherited during the division of cancerous cells. This inheritance occurred in a manner agreeing with the Mendelian laws of genetic inheritance—laws that heretofore had been understood to apply only to DNA molecules (see the articles genetics and heredity).

Scientists hypothesized that, in order for such viral inheritance to occur, a virus would need to transcribe its RNA genome into DNA and then insert this DNA into the host cell genome. Once incorporated into the host genome, the virus would be transcribed as though it were another gene and could produce more RNA virus from its DNA. This hypothesis, called the “DNA provirus hypothesis,” was developed in the late 1950s by American virologist Howard Martin Temin, when he was a postdoctoral fellow in the laboratory of Italian virologist Renato Dulbecco at the California Institute of Technology. Temin’s hypothesis was formally proposed in 1964. The provirus hypothesis came about when experiments demonstrated that an antibiotic called actinomycin D, which is capable of inhibiting DNA and RNA synthesis, inhibited the reproduction of RSV. However, the concept of an RNA molecule’s turning itself into DNA drew very few supporters.


How do you know if it worked?

After transformation, bacteria are grown on a nutrient rich food called agar . Only bacteria containing a plasmid with antibiotic resistance will grow in the presence of antibiotic.

For example, if the bacteria are grown on agar containing the antibiotic ampicillin , only the bacteria that have been transformed with a plasmid containing the resistance gene for ampicillin will survive.

Transformed bacteria can then be grown in large amounts. The DNA of interest, or the protein coded for by the DNA, can then be isolated and purified.


General Transfection Protocol

Preparing Cells for Transfection

Removing Adherent Cells Using Trypsin

Trypsinizing cells prior to subculturing or cell counting is an important technique for successful cell culture. The following technique works consistently well when passaging cells.

Materials Required:

  • 1X trypsin-EDTA solution
  • 1X PBS or 1X HBSS
  • adherent cells to be subcultured
  • appropriate growth medium (e.g., DMEM) with serum or growth factors or both added
  • culture dishes, flasks or multiwell plates, as needed
  • hemocytometer
  1. Prepare a sterile trypsin-EDTA solution in a calcium- and magnesium-free salt solution such as 1X PBS or 1X HBSS. The 1X solution can be frozen and thawed for future use, but trypsin activity will decline with each freeze-thaw cycle. The trypsin-EDTA solution may be stored for up to 1 month at 4°C.
  2. Remove medium from the tissue culture dish. Add enough PBS or HBSS to cover the cell monolayer: 2ml for a 150mm flask, 1ml for a 100mm plate. Rock the plates to distribute the solution evenly. Remove and repeat the wash. Remove the final wash. Add enough trypsin solution to cover the cell monolayer.
  3. Place plates in a 37°C incubator until cells just begin to detach (usually 1&ndash2 minutes).
  4. Remove the flask from the incubator. Strike the bottom and sides of the culture vessel sharply with the palm of your hand to help dislodge the remaining adherent cells. View the cells under a microscope to check whether all cells have detached from the growth surface. If necessary, cells may be returned to the incubator for an additional 1&ndash2 minutes.
  5. When all cells have detached, add medium containing serum to cells to inactivate the trypsin. Gently pipet cells to break up cell clumps. Cells may be counted using a hemocytometer, distributed to fresh plates for subculturing, or both.

Typically, cells are subcultured to prepare for transfection the next day. The subculture should bring the cells of interest to the desired confluency for transfection. As a general guideline, plate 5 × 10⁴ cells per well in a 24-well plate or 5.5 × 10⁵ cells for a 60mm culture dish for

80% confluency on the day of transfection. Change cell numbers proportionally for different size plates (see Table 3).

Table 3. Area of Culture Plates for Cell Growth.

Size of Plate Growth Area a (cm²) Relative Area b
24-well 1.88 1X
96-well 0.32 0.2X
12-well 3.83 2X
6-well 9.4 5X
35mm 8.0 4.2X
60mm 21 11X
100mm 55 29X

a This information was calculated for Corning® culture dishes.
b Relative area is expressed as a factor of the total growth area of the 24-well plate recommended for optimization studies. To determine the proper plating density, multiply 5 × 10⁴ cells by this factor.

Preparing DNA for Transfection

High-quality DNA free of nucleases, RNA and chemicals is as important for successful transfection as the transfection reagent chosen. See the Protocols and Applications Guide chapter on DNA purification for information about purifying transfection-quality DNA.

Example Protocol Using ViaFect&trade Reagent

We strongly recommend that you optimize transfection conditions for each cell line. If you have optimized transfection parameters, use the empirically determined conditions for your experimental transfections.

If you choose not to optimize transfection parameters, use the general conditions recommended below.

Materials Required:

  • cell culture medium with serum appropriate for the cell type being transfected
  • serum-free cell culture medium for complex formation (such as Opti-MEM® I reduced-serum medium)
  • 96-well or other culture plates
  • U- or V-bottom dilution plates or microcentrifuge tubes

The total volume of transfection complex (medium, DNA and ViaFect&trade Transfection Reagent) to add per well of a 96-well plate is 5&ndash10&mul. The following protocol is a guideline for transfecting approximately 10&ndash20 wells, depending on the volume of ViaFect&trade Transfection Reagent:DNA mixture used. For additional wells, scale volumes accordingly.

  1. To a sterile tube or U- or V-bottom plate, add 90&ndash99&mul of serum-free medium prewarmed to room temperature so that the final volume after adding the DNA is 100&mul. Add 1&mug of plasmid DNA to the medium, and mix. For a 3:1 ViaFect&trade Transfection Reagent:DNA ratio, add 3&mul of ViaFect&trade Transfection Reagent, and mix immediately.
  2. Incubate the ViaFect&trade Transfection Reagent:DNA mixture for 5&ndash20 minutes at room temperature.
    Optional: Add mixture to cells without an incubation period.
    Note: Longer incubations may adversely affect transfections.
  3. Add 5&ndash10&mul of the ViaFect&trade Transfection Reagent:DNA mixture per well to a 96-well plate containing 100&mul of cells in growth medium. We suggest 10&mul of mixture as a starting point. Mix gently by pipetting or using a plate shaker for 10&ndash30 seconds. Return cells to the incubator for 24&ndash48 hours.
    Note: The total growth medium volume may vary depending on well format and your laboratory&rsquos common practices.
  4. Measure transfection efficiency using an assay appropriate for the reporter gene. For transient transfection, cells are typically assayed 24&ndash48 hours after transfection.

Optimizing Transfection with Lipid Reagents

In previous sections, we discussed factors that influence transfection success. Here we present a method to optimize transfection of a particular cell line with a single transfection reagent. For more modern lipid-based reagents such as the ViaFect&trade Transfection Reagent, we recommend initially testing 50&ndash100ng of DNA per well of a 96-well plate at reagent:DNA ratios of 3:1 or 4:1 for adherent cell lines or 2:1 for suspension cell lines. Figure 4 outlines a typical optimization matrix. When preparing the ViaFect&trade Transfection Reagent:DNA complex, the incubation time may require optimization we recommend 5&ndash20 minutes.

Figure 4. Transfection optimization using the ViaFect&trade Transfection Reagent. TF-1 cells were plated in growth media without antibiotics at 30,000 cells per well in a white 96-well assay plate and transfected with a CMV-luc2 plasmid using various lipid (ViaFect&trade Transfection Reagent):DNA ratios. The DNA concentration was held constant at 1&mug per 100&mul of Opti-MEM® I reduced-serum medium, and the amount of ViaFect&trade Transfection Reagent was varied to obtain the indicated ratios. Either 5 or 10&mul of transfection complex was then added to cells in the 96-well plate. Twenty-four hours after transfection, the ONE-Glo&trade + Tox Luciferase Assay was performed. Note that 0:1 is the negative control with DNA but no lipid. These results show that, for this particular cell line, a 2:1 ViaFect&trade Transfection Reagent:DNA ratio gave optimal results.

For traditional cationic lipid reagents, we recommend testing various amounts of transfected DNA (0.25, 0.5, 0.75 and 1µg per well in a 24-well plate) at two charge ratios of lipid reagent to DNA (2:1 and 4:1 see Figure 5). This brief optimization can be performed using a transfection interval of 1 hour under serum-free conditions. One 24-well culture plate per reagent is required for the brief optimization with adherent cells (three replicates per DNA amount).

Figure 5. Suggested plating format for initial optimization of cationic lipid transfection conditions.

A more thorough optimization can be performed to screen additional charge ratios, time points and effects of serum-containing medium at the DNA amounts found to be optimal during initial optimization studies. One hour or two hours for the transfection interval is optimal for many cell lines. In some cases, however, it may be necessary to test charge ratios and transfection intervals outside of these ranges to achieve optimal gene transfer.

Some transfection methods require removing medium with reagent after incubation others do not. Read the technical literature accompanying the selected transfection reagent to learn which method is appropriate for your system. However, if there is excessive cell death during transfection, consider decreasing time of exposure to the transfection reagent, decreasing the amounts of DNA and reagent added to cells, plating additional cells and removing the reagent after the incubation period and adding complete medium.

Endpoint Assays

Many transient expression assays use lytic reporter assays like the Dual-Luciferase® Assay System (Cat.# E1910) and Bright-Glo&trade Assay System (Cat.# E2610) 24 hours after transfection. The Nano-Glo® Dual-Luciferase® Reporter Assay System (Cat.# N1610) allows detection in just a few hours after transfection. However, the time frame for most assays can vary (24&ndash72 hours after transfection), depending on protein expression levels. Reporter-protein assays use colorimetric, radioactive or luminescent methods to measure enzyme activity present in a cell lysate. Some assays (e.g., Luciferase Assay System) require that cells are lysed in a buffer after removing the medium, then mixed with a separate assay reagent to determine luciferase activity. Others are homogeneous assays (e.g., Bright-Glo&trade Assay System) that include the lysis reagent and assay reagent in the same solution and can be added directly to cells in medium. Examine the reporter assay results and determine where the greatest expression (highest reading) occurred. These are the conditions to use with your constructs of interest.

Alternative detection methods include histochemical staining of cells (determining the percentage of cells that are stained in the presence of the reporter gene substrate Figure 6), fluorescence microscopy (Figure 7) or cell sorting if using a fluorescent reporter like the Monster Green® Fluorescent Protein phMGFP Vector (Cat.# E6421).

Figure 6. Histochemical staining of RAW 264.7 cells for &beta-galactosidase activity. RAW 264.7 cells were transfected using 0.1µg DNA per well and a 3:1 ratio of FuGENE® HD to DNA. Complexes were formed for 5 minutes prior to applying 5µl of the complex mixture to 50,000 cells/well in a 96-well plate. Twenty-four hours post-transfection, cells were stained for &beta-galactosidase activity using X-gal. Data courtesy of Fugent, LLC.

Figure 7. Fluorescence microscopy of cells transfected with ViaFect&trade Transfection Reagent. iCell® human tissue cells in 96-well plates were transfected with ViaFect&trade Transfection Reagent and a GFP reporter plasmid at the specified reagent:DNA ratio and GFP expression was imaged after transfection. Panel A. iCell® Hepatocytes with a 6:1 reagent:DNA ratio imaged one day after transfection. Panel B. iCell® Cardiomyocytes with a 2:1 reagent:DNA ratio imaged one day after transfection. Panel C. iCell® DopaNeurons with a 4:1 reagent:DNA ratio imaged three days after transfection. Data courtesy of Cellular Dynamics International.

Real-Time Assays

For some types of transfection experiments, especially those examining the changes in gene expression levels associated with pathological mechanisms, monitoring reporter activity in living cells is desirable. Such real-time assays can provide valuable information on the expression of multiple genes in a dynamic fashion. The Nano-Glo® Live Cell detection options (Cat.# N2011) are designed to detect NanoLuc® luminescence from living cells using nonlytic protocols. These assays monitor luminescence at a single time point or continuously for up to 72 hours without compromising cell viability.


Setting up the experiment

The next day I started learning more about GFP, mainly by phoning people. One encouraging fact was that GFP did not need anything added to be fluorescent. i.e., to absorb blue light and emit green light. What I was really hoping to uncover in all my phone calls was a person who was working on obtaining the region of the jellyfish DNA that contained the gene that encoded GFP. We needed the GFP DNA so that we could transfer it and express it in C. elegans. Luckily, I found someone working to identify the GFP gene, Douglas Prasher, a researcher at the Woods Hole Oceanographic Institute. When I reached Douglas by phone, I found that he had very similar ideas about using GFP and at the end of our hour-long conversation, we decided to collaborate.

Douglas said that he would call when he had succeeded in the first step—isolating the jellyfish DNA for GFP. But at that time, this step was not easy. If you want to know more about how he did this, see the “ Dig Deeper 1: cloning jellyfish DNA. ”

Unfortunately, I never got Douglas’ call that he succeeded in cloning GFP (I later learned that he called me when I was away on sabbatical). I subsequently learned that because Doug couldn’t get in touch with me, he thought that I had left science.

In September 1992, the new graduate students joined the department and one of them, Ghia Euskirchen, asked if she could rotate in my lab (many graduate students do short-term projects called “rotations” to try out several labs before deciding where to do their graduate research). I was very happy to have her rotate in my lab, because she had already obtained a master’s degree in chemical engineering at Columbia working on fluorescence. I told Ghia about my idea of using a fluorescent protein as a biological marker. But after waiting 3 years for Douglas’ call, I had given up hope about his experiments. So I suggested that we might have to look for something other than GFP. We then looked up the term “fluorescent protein” on Medline (an early search engine made by the National Institutes of Health) and see what other fluorescent proteins existed. Columbia had just connected the University computers to Medline, another fortunate bit of timing. The first paper that popped up in our search was Douglas’ paper from earlier that year describing the isolation of the GFP DNA. We ran down to the library to find the paper. Then, another stroke of luck: although Douglas had left the Woods Hole Oceanographic Institute, the paper had his new phone number (scientific articles do not often list authors’ telephone numbers). I immediately called Douglas and renewed our collaboration. He sent us the DNA clone a few days later.

With the jellyfish GFP DNA in hand, Ghia’s project was, in one sense, very straightforward: introduce the jellyfish GFP DNA into bacteria ( E. coli ), express the protein, and see whether the bacteria emitted green light when stimulated by blue light.

Most people working on GFP at the time believed that this experiment would never work because GFP is a very unusual protein. Most proteins are linear strings of amino acids. GFP also starts out with the usual linear peptide backbone (the line shown in Figure 3 ), but then it undergoes a chemical rearrangement to make a loop (a chemical ring) in a section of the linear backbone ( Figure 3 ). This chemical ring is essential for the protein to be fluorescent. When we did our experiments, no one knew how this loop was made. People imagined that there was a jellyfish-specific enzyme that produced the loop. Thus, if this enzyme was not present in bacteria (as one would expect), then GFP should not be fluorescent. As a result, our experiment would be considered a fool’s errand the bacteria should not glow green. However, we were not dissuaded and decided to try the experiment anyway.

Figure 3 The Chemistry that Makes GFP fluorescent. Proteins are composed of a linear backbone (traced with the arrow) of connected amino acids with side chains (numbered) branching off the backbone. GFP undergoes an unusual chemical reaction that creates a loop (a chemical ring) in the backbone. This new chemical ring is what allow enables GFP to fluoresce (absorb blue light and emit green light).

There was one additional experimental detail, which proved to be very important and proved fortunate for us. Douglas’ GFP clone had the complete coding sequence for GFP but it also had additional jellyfish DNA on either side. This adjacent DNA did not code for amino acids and we knew nothing about its function. I was leery of this extra DNA and did not want to include it in our experiment and suggested to Ghia that we leave it out and just use the DNA that coded for GFP. We could copy just the GFP coding sequence using the polymerase chain reaction (or commonly known as PCR), a technique that amplifies a precise piece of DNA. See Video 2 for an explanation of how PCR works.

The one problem with PCR at this stage of its development is that it was somewhat error prone (i.e., copying the DNA by PCR could introduce mutations ( Figure 4 ). Thus, researchers who wanted accurate copies of the GFP DNA would have left the jellyfish DNA alone and used the entire DNA fragment obtained by Douglas to express the GFP. In contrast, I reasoned that if some of the PCR DNA products had mutations, this would not be a problem. Certainly some of the DNA would have been copied accurately by PCR, and those bacteria would be easy to spot because they would be glowing green. As you will read later, this seemingly trivial strategic decision allowed us to succeed, while others’ laboratories failed in the same effort using Douglas’ original DNA clone.

Figure 4 Using PCR to Amplify the GFP Gene. The original GFP clone contained the GFP gene as well as extra jellyfish DNA on either side of unknown function. PCR-enabled amplification of just the GFP gene. However, PCR can introduce mutations which could potentially give rise to a non-functional GFP. However, not all of the GFP DNA had to be functional GFP. If some unmutated were present, the bacteria with the normal DNA would allow the bacteria to glow green.

Now I will turn the narrative over to Ghia who can tell you what she did.


Restriction enzyme

A restriction enzyme, restriction endonuclease, or restrictase is an enzyme that cleaves DNA into fragments at or near…

Since REs cut at specific sequences, you can see how APE is useful for us. APE comes with a library of restriction enzymes and the sequence they recognize and cut at. So when you import a new plasmid into APE, you can quickly and easily determine which RE will cut your plasmid, and where.

We keep a stock of RE enzymes, and they can be purchased from companies like New England Biolabs:

The NEB website is useful since they have many free online tools for figuring out which enzymes can be used, what buffers, and other reaction conditions are necessary, etc. NEB sells much more than REs, but that is how they got their start.

Other companies who also sell REs include the following:


CDNAs and Gene Analysis

Plasmid vectors are limited in the size of the cloned DNA that can be incorporated and successfully reintroduced into the bacterium, typically holding a maximum of about 15 kb (kilobases [1 kb equals 1,000 bases]) of foreign DNA. One common use for plasmid vectors is to make cDNA (complementary DNA) libraries cDNA molecules are DNA copies of messenger ribonucleic acid (mRNA) molecules, produced in vitro by action of the enzyme reverse transcriptase . Because cDNAs represent only the portions of eukaryotic genes that are transcribed into the mRNA, cDNA clones are particularly useful for analysis of gene expression and cell specialization. The existence of a cDNA is also evidence that the gene is active, or transcribed, in the cells or tissues from which the mRNA was isolated. Such information can be used to compare gene activities in healthy versus diseased cells, for instance.

Frequently the simpler sequence of a cDNA is easier to analyze than the corresponding genomic sequence since it will not contain noncoding, or intervening, sequences (introns). Another advantage of cDNA is that generally the sequence does not include enhancers or regulatory sequences to direct their transcription. As a result, they can be combined with other regulatory systems in the clone to direct their expression.

Genome sequencing projects typically generate sequence information from many different cDNA clones. The cDNA cloned sequence is termed an Ȯxpressed sequence tag" (EST), and, when correlated with the whole genomic DNA sequence, EST information can help determine the locations and sizes of genes.

In order to obtain the cDNA for a specific gene, it is first necessary to construct a cDNA "library." This is a collection of bacteria that contain all the cDNAs from the cell or tissue type of interest. To make a library, the thousands of different mRNAs are first harvested from the cell of interest, and cDNA is made using reverse transcriptase. The cDNA is then cloned into plasmids, and introduced into bacteria. Under the right conditions, each bacterium will take up only one cDNA. The bacteria are then grown in Petri dishes on a solid medium. A library therefore consists of a mixed population of bacteria, each carrying one type of cDNA. To find the bacterium containing a particular type of cDNA, one can either search for the gene itself with a nucleotide probe or for its protein product with an antibody .

Screening a library depends either on having a probe bearing part of the nucleotide sequence or an antibody or other way of recognizing the protein coded by the gene. Screening by nucleotide probes (labeled with radioactive or chemical tags for detection) depends on base pair complementarity between the single-stranded target DNA and the probe DNA this allows the label to mark the cell with the desired cDNA. Screening by labeled antibody depends on binding of the antibody to the protein encoded by the gene. Literally thousands of cloned genes have been isolated this way from libraries of many different species. One of the most powerful observations in biology is that the same or similar gene sequences can be isolated from different species, ranging from bacteria to humans.

Human insulin was the first medicine to be created through recombinant DNA technology. Insulin is a protein hormone produced by the pancreas that is vital for regulation of blood sugar. In the disease insulin-dependent diabetes mellitus (IDDM), the immune system attacks and destroys the insulin-producing cells. A person with IDDM requires daily injections of insulin to control blood sugar. Before 1980, insulin was isolated from pigs or other animals. Animal insulin has a slightly different amino acid sequence from the human form. In the early 1980s, recombinant DNA technology was used to splice the human insulin gene into bacteria, which were grown in vats to make large amounts of the human protein. Recombinant human insulin was the first recombinant drug approved for human use. Since then more than two dozen other drugs have been created in this way, including growth hormone, blood clotting factors, and tissue plasminogen activator, used to break up blood clots following a stroke. Gene sequence similarities indicate that all living organisms have descended from shared common ancestors, back to the beginning of life.


NZY Reverse Transcriptase ( MB124 )

Description: NZY Reverse Transcriptase is a modified recombinant form of the Moloney Murine Leukemia Virus (M-MuLV) Reverse Transcriptase purified from Escherichia coli. The enzyme has been modified in order to promote stability. NZY Reverse Transcriptase synthesizes the complementary DNA strand in the presence of a primer using either RNA (cDNA synthesis) or single-stranded DNA as a template in a wide range of temperatures (37-50°C). This is useful when using templates with high GC-content or with high degree of secondary structure. The enzyme lacks 3´→5´ exonuclease activity and has no RNase H activity, enabling improved synthesis of full-length cDNA even for long mRNA, using random priming.

Features:
– Synthesizes of cDNA from RNA or ssDNA in only 30 min
– High yields of first-strand cDNA
– No intrinsic RNase H activity
– Thermostable – working temperature range 37-50 ºC

Applications:
– First-strand cDNA synthesis for RT-PCR and RT-qPCR
– Reverse transcription at elevated temperatures (ideal for high GC-content templates)
– Synthesis of cDNA for cloning and expression
– Analysis of RNA by primer extension

High sensitivity of NZY Reverse Transcriptase:
The high sensitivity of NZY Reverse Transcriptase is demonstrated using decreasing amounts of total RNA isolated from mouse liver (1 μg – 10 pg) as starting material in a 20 μL first-strand cDNA synthesis reaction with 200 units of the enzyme.

After the reverse transcription, 1 μL of the resultant cDNA was used as a template to amplify the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) gene in a 25 μL end-point PCR reaction with Supreme NZYTaq II DNA polymerase. NC: No template control. Lane M: NZYDNA Ladder III

Specifications:

Product length: up to 7 kb
Sensitivity: high
Optimal reaction temperature: 50 ºC
Available as Kits: NZY First-Strand cDNA Synthesis Kit (MB125) NZY First-Strand cDNA Synthesis Kit, separate oligos (MB170)
Speed: 30 min
Denaturing conditions: NZY Reverse Transcriptase is inhibited in the presence of metal chelators (e.g. EDTA), inorganic phosphate, pyrophosphate and polyamines. The enzyme is inactivated at 85 °C for 5 min
Lyo format available: Lyo NZY Reverse Transcriptase (MB409)
Storage conditions: Store at -20 ºC
Shipping conditions: Shipped with dry ice

Components:
– NZY Reverse Transcriptase (200 U/μL)
– Reaction Buffer (10x)

NZYTech Reverse Transcriptases: Selection Guide

NZY Reverse Transcriptase vs the competition. A 10-fold serial dilution of total RNA of mouse liver (1 μg to 1 ng) was reverse transcribed using 200 units of NZY Reverse Transcriptase or a popular reverse transcriptase. The resultant cDNA was then used as template in a PCR using specific primers for amplification of the mouse GAPDH gene with Supreme NZYTaq II DNA polymerase. NC: No template control. Lane M: NZYDNA Ladder III.

1. What primers are used for Reverse Transcription?
There are three different approaches for priming cDNA reactions: oligo(dT) primers, random primers, or sequence-specific primers. These primers differently bind to the template RNA strand, by providing a starting point for the cDNA synthesis, and each one has advantages and disadvantages. The choice between these three priming methods will depend on the size and type of RNA, on the reverse transcription temperature, or on the intended downstream applications.

An overview of different priming methods for reverse transcription

Oligo(dT) primers specifically bind to the poly(A)-tail found at the 3´-end of most eukaryotic mRNAs. The capacity to generate many different cDNAs from the same starting RNA pool, makes these primers the preferred choice for two-step RT-PCR reactions. Different types of oligo(dT)s are available. Oligo (dT)18 primer mix, a homogenous mixture of 18-mer thymidines, is available at NZYTech for the synthesis of full-length cDNA from poly(A)-tailed mRNA. In contrast to the standard oligo (dT), which randomly bind within the poly(A) tail of the eukaryotic mRNA, the anchored oligo(dT) primers bind at the beginning of the tail. This avoids an unnecessary reverse transcription of this often long region as well as erroneous products synthesized by mispriming.

Random hexamers are preferred for long transcripts or if they contain significant secondary structures. In addition, random hexamers are used for non-polyadenylated target templates (as prokaryotic mRNA). They will perform random priming throughout the entire length of the RNA to generate a cDNA pool containing various lengths of cDNA. With this method, all RNAs present in a population constitute templates for cDNA synthesis experiment. NZYTech provides a Random hexamer mix that includes oligonucleotides representing all possible hexamer sequences.

A mixture of both oligo(dT) and random hexamer primers is usually used to improve the efficiency of cDNA synthesis and qPCR sensitivity.

Gene-specific primers (GSPs) enhance sensitivity by allowing the reverse transcription of a specific RNA sequence. This priming method is chosen to perform one-step RT-PCR reactions once the same primer is used in both the RT and PCR steps. However, GSPs offer less flexibility than oligo(dT) and random primers, since each cDNA synthesis is limited to one target gene.

2. Is the NZYReverse Transcriptase RNase H minus?
The enzyme has no intrinsic RNase H activity.

3. The reaction buffer in my NZYReverse Transcriptase has precipitated. Can I still use?
The NZYReverse Transcriptase 10x Reaction Buffer can on occasions form a white precipitate after repeated freeze/thaws. If this happens, mix thoroughly to resuspend the precipitate. If you verify non-conformity of results, then the buffer needs to be replaced.

4. Should I treat the synthesized cDNA with RNase H before PCR?
Addition of RNase H after first-strand synthesis will degrade the RNA used as template, by removing it from the cDNA:RNA hybrid molecule. The RNA is still present when using RNase H – versions of reverse transcriptase, as is the case of the NZYReverse Transcriptase. Presence of RNA during PCR could inhibit annealing of the primers to the cDNA and then affect the amplification reaction, especially for long fragments. However, the 95°C denaturing step could cause RNA degradation of the RNA-cDNA hybrids and therefore RNase H treatment may not be necessary. We recommend performing RNase H digestion before PCR when using lower levels of template or when amplifying long fragments.

5. How much synthesized cDNA should be used in a PCR reaction?
Do not exceed 10% of the final PCR reaction volume. The volume of cDNA used will depend on the amount of RNA used as template for first-strand synthesis, as well as the abundance of the target gene.

6. Little or no RT-PCR/RT-qPCR amplification product is observed. What should I do?
This may result from several factors, such as:
a) RNA damage or degradation. Analyze RNA by denaturing gel electrophoresis to verify nucleic acid integrity. Use aseptic conditions while working with RNA to prevent RNase contamination. Ensure the use of NZY Ribonuclease Inhibitor: addition of this inhibitor is essential when using less than 50 ng of RNA in order to safeguard the template against degradation due to ribonuclease contamination. Replace RNA if necessary.
b) Presence of RT inhibitors. Some inhibitors of RT enzymes include: SDS, EDTA, glycerol, sodium phosphate, spermidine, formamide and guanidine salts. They can be problematic if present in smaller reaction volumes. If necessary, remove inhibitors by ethanol precipitation of the RNA preparation before use wash the pellet with 70% (v/v) ethanol.
c) Not enough starting RNA. Increase the concentration of starting RNA.