DNA Sequencing and Fragment Analysis

Sanger sequencing methodology requires template DNA to be relatively free from contaminating salts. Template quality is often determined electrophotometrically by loading the template on agarose gel. Agarose gel electrophoresis often shows the faster moving supercoiled DNA and slower moving nicked DNA found in a quality preparation. DNA samples produced by PCR amplification can be similarly tested by agarose gel electrophoresis. A good PCR product is viewed as a single clean band representing the desired length.

Another test is to measure absorbance spectrophotometrically at ultraviolet wavelengths 260 nm and 280 nm. A 260 nm / 280 nm ratio value of 1.8 is an indicator that the DNA is good.

Unfortunately, the result off the capillary sequencer shows double set of peaks. Because one set of peaks is directly over top the other, software cannot determine the correct bases to call as in figure 1. A result showing a double set of peaks is not as uncommon as many might believe.

What are the primary causes of double sequence?

Double Sequence Caused by Template Contamination

One of the early steps in plasmid DNA preparation is to select a desired colony and then inoculate liquid medium. In an overnight culture, the inoculated media would go through rapid bacterial cell division as the cells enter the log growth phase. Because the initial cells came from a single colony, it is believed that every cell is the identical product cloned from a single cell. However, this is not always the case. Colonies do not always come from a single cell. A colony could also be the product of multiple cells that failed to separate when spread on a plate. If the multiple cells are not identical, the sequence result could produce the double sequence shown in figure 1.

The region where double sequence begins is usually a restriction sight where a separate piece of DNA is inserted into a vector. One purpose of bacterial cloning is to produce copies of the inserted DNA. The clean sequence shown in figure 1 is vector sequence before the restriction sight. Despite differences in the two templates, the vector sequence is generally the same. The template contamination becomes apparent at the point where the new DNA has been inserted into the vector.

Often, the reverse or complimentary strand does not produce a double sequence and this is often confusing. If the DNA insert were relatively long, it would require sequencing the whole complimentary strand before double sequence is observed. The reason is simply because double sequence is caused by addition or deletion of an unknown number of bases at one restriction sight only. Therefore, both templates, the desired and contaminant have identical sequence in the complimentary direction.

PCR fragments could also produce double sequence. It is the result of errors in PCR amplification where two products are produced. The PCR fragments could produce two very similar sized products that appear as a single band once tested by agarose gel electrophoresis. Resolution using agarose gel is limited and does not separate DNA samples of very similar size.

Double Priming Results in Dual Amplification

Sanger sequencing is similar to PCR amplification. A primer anneals at the beginning of the region to be sequenced and Taq polymerase adds bases (dNTPs) in extension to produce and identical strand. A primer that matches two regions on either the insert or the plasmid could cause two separate amplifications simultaneously. The result of double priming is shown in figure 2.

Unlike template contamination, two separate products generally do not appear clean in the beginning. It does not require the primer to match identically to both annealing sights as long as the bases of the primer are similar with a match on the 3 prime ends. What happens if one annealing site is downstream from the other annealing site on the same strand of DNA? The downstream primer blocks bases extending from the first primer and the double sequence eventually becomes a single set of peaks.

Single Nucleotide Polymorphisms Could Also Cause Double Sequence

PCR amplification is used to determine potential heterozygous bases as single nucleotide polymorphisms (SNPs). It is one method for detecting mutations that could cause certain genetic diseases. The SNP likely could be observed as a base change in one of two identical alleles amplified together and is generally easy to spot from automated Sanger sequencing. A single base position is represented by two peaks to positively identify a SNP.

However, SNPs could also be the result of a base insertion or deletion (indel) as in figure 3. From the figure it is possible to follow two identical strands of sequence that are different by one base. Both alleles match until the GGCC region. However, one allele is missing the T-base and has shifted upstream by one base. The large clean peaks observed in the double peak region are simply the same base represented by two different products.

Preventing Double Sequencing in Sequencing Results

There are some cases where double sequence cannot be resolved as with PCR fragments that contain indels. However, certain steps could be taken to reduce conditions described in figure 1 and figure 2.

Plasmid preparations that produce double sequence at the point of insert can be prevented using a process called single colony isolation. Once the plasmid cells have been spread on a plate and grown, the select colony is isolated and spread on a second plate. Although the process requires additional time, it is worth the effort in cases when double sequence often occurs.

The process for preventing double sequence in PCR fragments is more complicated because PCR conditions need better optimization. It could be something as simple as reducing the amount of primer or number of cycles. Or require small adjustments in PCR conditions including temperature. Guidelines for PCR optimization will be a future topic.

Samples that produce double sequence as the result of double priming can be resolved by extending bases to the custom primer that was selected for sequencing. A better choice is to select a different primer even when double priming is caused from one of the possible universal primers present on a particular vector.

Double sequence result patterns are generally similar for each condition discussed here. Anyone is invited to present results similar to that presented. Questions are also welcome.

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Epigenetics is a relatively new field of research that goes beyond gene expression. It encompasses important areas of research including cell differentiation, cancer and the aging process. Despite complete sequencing of the human genome, there is still much to learn about the complexity of an individual genetic blueprint. Epigenetic research is working to better understand how cells develop into specialized tissue and perform a specified number of functions. The human begins as a single fertilized egg. The egg divides into a growing mass of identical cells. Eventually, individual groups of cells begin to look and act different than other cells. Cell groups develop into tissue and organs. How does this happen when every cell has the same genetic code?

Proximity could play a partial role in cell differentiation. Cells adhering together generally develop into the same tissue. But what boundaries separate each group?

The answers may rest in understanding the epigenome. Epigenetics is a complex field newly emerging in the genetic sciences. Although there is much to learn, there is some basic understanding summarized here.

Histones Play an Important Role in Controlling Gene Expression

DNA is a double stranded molecule that coils around into a helical formation wrapped around histone molecules. Histones are a group of proteins that condense long DNA strands. It is similar to string wrapped around a spool. How tightly DNA is wrapped around the histone molecule determines whether a particular region of DNA is available for transcription and gene expression. Tightly wound DNA is essentially hidden and this region is repressed. Other regions are loosely wound and available for transcription. Transcription is the first step in the protein production process (Figure 1).

Histone molecules are classified into one of five main groups. They are modified in the cell by chemical processes such as methylation, acetylation or other known modifications. Modification is one factor that determines whether DNA is active or repressed. Methylation of certain groups of histones allows genes to be active while others are repressed. However, external factors (like radiation) could effectively influence methylation and activate repressed genes.

Epigenetic Damage is a Cause of Cancer

Let us examine a hypothetical situation in which two individuals have identical genetic content. They are twins. Each individual has identical genes including genes that potentially cause cancer. Why does one individual develop cancer and the other does not?

Research has long understood that cancer could be caused by damage to the DNA making up the genetic blueprint. But external factors can cause damage to DNA leading to cancer as well. Factors that damage the epigenome of an individual can have the same result. An environmental factor like smoking is an example. When cells lose levels of control, they also lose specialization; they de-differentiate. Without such important controls, cells begin to divide uncontrollably into cancer. A better understanding of epigenetic damage has led to new directions in cancer treatment.

Dr. Jean-Pierre Issa explained some general aspects concerning cancer and epigenetic damage during an interview with Nova. See Nova article here.  Epigenetic damage could be related to the aging process. As an individual continues to age, cell division eventually leads to errors in the epigenome. For example, skin constantly exposed to sunlight radiation appears older than normal skin. The aging process is accelerated by stem cells dividing to repair sun damaged skin tissue. The more cells divide, the more the aging processes are accelerated. Dr. Issa utilizes this knowledge in research to determine the apparent age of certain cells.

What Affects the Epigenome?

Current research is attempting to answer this question. There are a number of known environmental conditions that cause damage to the epigenome. Radiation, tobacco and other chemicals have the potential to damage the cellular epigenome. As previously stated, the radiation from excessive sunlight can cause damage to skin tissue. Stem cells divide to repair damaged skin. Damage to skin tissue causes the need for repair and accelerates the aging of stem cells.

Epigenetics is a fascinating new field in the genetic sciences. Understanding factors that affect the epigenome has led medicine into exciting new areas of treatment. This topic is certain to be examined often in the near future.

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DNA synthesis is the production of short, single-stranded DNA molecules (called primers or oligonucleotides) often used in the polymerase chain reaction (PCR) and DNA amplification for Sanger sequencing applications. The region of DNA amplified is determined by an exact match of the primer to its complimentary bases on a given DNA strand. Primer sequences are determined from known sequence since there must be a match to the region of DNA to be amplified.

PCR amplification requires 2 primers that determine the region of sequence amplified in the forward and reverse direction. The forward primer is designed along one strand in the direction toward the reverse primer. Likewise, the reverse primer is designed from the complimentary strand. PCR is exponential amplification in which the newly generated PCR fragment from one cycle also acts as a template for the next cycle (Figure 1A).

Amplification of DNA for Sanger sequencing differs from PCR in that a single primer is used. Amplification is the reproduction of one strand using the compliment from the original strand. Newly generated PCR fragments are single stranded and do not provide a complimentary strand that could act as a template for additional amplification (Figure 1B).

Differences in amplification between PCR and Sanger sequencing were discussed previously (Sanger Sequencing Amplification Compared to Basic PCR).

Simply designing the sequence of the primer from known sequence does not ensure the primer will anneal to the desired region and initiate amplification. The primer should be designed following a set of given standards that improve the chances of success. The guidelines for designing primers used in PCR and sequencing are fairly similar. Primers are designed in the 5’ to 3’ direction to compliment the direction of amplification.

For those utilizing PCR and Sanger sequencing in everyday applications, primer design could seem like yesterday’s news. However, we still believe reviewing good primer design guidelines is helpful to any researcher involved in genetic research.

Guidelines for Primer Design

G1. Primer length should be in the range of 18 to 22 bases. Primers less than 18 bases will have a low melting temperature (Tm values) and might not anneal to the template. There is some flexibility for designing primers longer than 18 bases. Longer primers are frequently designed from template regions that are AT-rich and need additional bases to increase the Tm value.

G2. The primer should have GC content of 50% to 55%.   This is the equivalent of 9 or 10 GC bases included in an 18 base primer. Sometimes there are regions on a template that are AT-rich which prevents meeting this guideline. In those cases it is recommended to design a primer longer than 18 bases.

G3. Primers should have a GC-lock on the 3’ end. A GC-lock is designed when 2 of the final 3 bases is a G or a C. The 3’ base should always be a G or a C.

G4. The melting temperature of any good primer should be in the range of 50OC to 55OC. However, guidelines particularly related to Tm value have some flexibility. Melting temperatures are directly related to the PCR cycle annealing temperature. Tm values that are too low may not anneal well during PCR. High values could be too stringent causing difficulty locating the correct annealing site on the template.

G5. The primer should not include poly base regions. This is when 4 or more bases in a row are the same. This guideline helps prevent potential slippage in which the primer shifts from the annealed position.

G6. Four or more bases that compliment either direction of the primer should be avoided. This prevents the primer from annealing to itself and forming what is referred to as primer-dimer. Primer-dimers have the capability of amplifying the primer itself causing short secondary sequence.

PCR Specific Guidelines

G7. Forward and reverse primers used in PCR amplification should have similar melting temperatures (+/- 2OC). This allows a 4OC difference in total melting temperatures. Researchers involved in using PCR amplification will use primer Tm values in an effort to optimize PCR cycles. Similar Tm values for forward and reverse primers aid optimization efforts. Multiplex PCR applications using multiple primer pairs should all have similar Tm values. A wide range in primer melting temperature complicates PCR optimization.

G8. Forward and reverse primers should not have regions 4 bases or longer that compliment. Just like a primer used in Sanger sequencing, forward and reverse primers used in PCR can anneal to each other and form primer-dimers.

G9. The Tm values for tailed primers should include the tail in calculating melting temperature. Yes, melting temperatures will be greater than 55OC. However, the additional bases in the tail will add to the amplified PCR fragment and become part of the priming site. Tailed primers are often used to add restriction sites to an amplified product.

Primer design is an important aspect relating to many forms of PCR including basic PCR, fragment analysis, quantitative analysis and Sanger sequencing. Below is a link that offers free primer design software…

http://frodo.wi.mit.edu/

We invite any additional guidelines or comments that may be useful to both experienced researchers and those new to primer design.

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Isolation of plasmid DNA in preparation for automated Sanger sequencing has been simplified by the development of commercial kits. The quality of the final sample is generally tested by spectrophotometer measuring absorbance at wavelengths 260 nm and 280 nm. A ratio value of 1.8 calculated for absorbance measurements at 260 nm by 280 nm indicates the DNA sample is good. However, 260 nm and 280 nm absorbance values do not provide a complete profile of whether DNA preparations meet necessary quality standards for Sanger sequencing applications. Samples may still fail. There are a number of reasons why sequencing reactions are not successful. Two well-known techniques eliminate template quality as a reason for failure.

Scanning Spectrophotometry Provides Important Information about Template Quality

Nanodrop technology has the advantage of highly sensitive spectrophotometric scanning of DNA samples using just 1 ul of volume. Most spectrophotometers available today are capable of performing a wavelength absorbance scan in the ultra violet spectrum. DNA is scanned between wavelengths ranging from 220 nm to 320 nm. The scan profile for quality DNA is shown in figure 1.

A typical DNA profile is a Gaussian (bell-shaped) curve with the maximum peak height absorbing at 260 nm. A secondary peak will also begin to form around 220 nm. Deviations in the shape of either curve could indicate the presence of salts or other contaminants.

Figure 2 provides a profile of plasmid DNA containing salts compared to clean DNA. The Gaussian curve should drop almost to the baseline when DNA is free of contaminating salts in the range 230 nm to 240 nm. Salts present in the sample will absorb wavelengths in this range. Excessive amounts of salt in DNA samples may show little or no dip in the curve between 220 nm and 260 nm.

Figure 3 represents a profile of DNA that contains either protein (DNA from tissue) or phenol (plasmid DNA). Phenol residue in DNA can result from phenol/ chloroform extraction (a method of extraction that was widely used prior to the development of commercial kits). Despite the availability of kit-based methods, many researchers opt to use phenol/ chloroform instead. It is important to remove phenol residues with ethanol precipitation in the final purification step. Phenol residue can negatively affect the PCR amplification in Sanger sequencing.

Scanning spectrophotometry successfully identifies salt contaminants in plasmid DNA and DNA samples amplified in the Polymerase Chain Reaction (PCR). PCR fragments will have a lower concentration. But purified PCR products should still show a scan similar to plasmid preparations.

Agarose Gel Electrophoresis Identifies Nicked DNA

Once plasmid DNA has been determined to be free from salt contaminants it should work well for Sanger sequencing. However, this is not always the end result. Scanning spectrophotometry does not reveal whether DNA is supercoiled or nicked as described previously (Nicked Plasmid DNA Prevents Automated Sanger Sequencing). Nicked DNA has been damaged mechanically. Aggressive vortexing during DNA purification is one cause of DNA damage. Nicked DNA does not amplify in Sanger sequencing applications because the double stranded helix does not maintain a tight formation. Taq polymerase is unable to fulfill a lock-key attachment to the DNA and catalyze extension. For automated sequencing to work, plasmid DNA must maintain a supercoiled structure.

Agarose gel electrophoresis successfully separates nicked DNA (slow migrating) and supercoiled (fast migrating) as shown in figure 4. Nicked DNA moves more slowly through the gel because it is a larger molecule than supercoiled DNA. It should be noted that plasmid preparations typically have a mixture of nicked and supercoiled DNA. Presence of some nicked DNA should not cause concern. It is the lack of supercoiled DNA that causes sequencing failures.

Scanning spectrophotometry and agarose gel electrophoresis combined provide a good assessment of whether template DNA purification has been successful. However, it does not guarantee Sanger sequencing will be successful every time. Primer selection, GC content and presence of poly base regions and hairpins could also affect sequencing. Problems of this nature are related more to base content and not preparation of the sample.

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Cesium chloride and phenyl chloroform were two early methods used for isolating a sample of DNA from a bacterial vector. Cesium chloride produced a sample that was very clean. However, the method was labor intensive. Phenyl chloroform was less labor intensive, but phenyl contamination was difficult to remove even using the final alcohol precipitation step. Fortunately, commercial kits were developed around the same time Sanger sequencing became automated. Presently there are numerous commercial kits from which a researcher could choose.

However, using a commercial kit does not guarantee that a resulting DNA sample will be clean and free from contaminants. The final step of the sample preparation often includes Tris or another buffer (salt) for elution. Other times the DNA may incur physical damage and become nicked. Preparing a sample for automated sequencing includes following the recommended guidelines provided for the commercial kit. However, low yields can require additional steps to concentrate the sample. Simple drying methods concentrate everything in the sample including buffer salts.

Steps Involved in Commercial DNA Preparation Kits

The general protocol for commercial kits used to isolate purified DNA from plasmids is fairly similar regardless of the kit. After plasmid bacteria have been grown overnight in liquid media, the cells are pelleted by centrifugation into a solid mass. The media is then discarded before beginning the kit provided procedure (Figure 1).

The pellet of cells is resuspended in buffer and transferred into a 1.5 or 2 ml tube (Step 1). Lysis reagent is added (Step 2). The lysis reagent disrupts the bacterial cell membranes freeing internal components. The lysis buffer is neutralized with neutralizing solution and cell components form a precipitate with the DNA still in solution (Step 3). The resulting solution is centrifuged to pellet the cell waste leaving the DNA in solution.

DNA is captured on a filter or resin by transferring the DNA solution to a collection vessel (Step 4). Excess liquid is pulled through the filter by centrifugation or vacuum filtration. The remaining DNA on the filter is then washed removing any cell waste left (step 5). The DNA is then collected in a clean tube using water or elution buffer (step 6). Elution buffer is provided in the commercial kits. It is comprised of Tris buffer in water. Fortunately, manufacturers no longer include EDTA in elution buffer. EDTA interferes with magnesium chloride, a necessary component in any PCR amplification. Water is the preferred media for DNA to be used for Sanger sequencing as no salts are being introduced to the sample.

The process may take as little as 15 minutes after bacterial cells are grown overnight. The final DNA sample is relatively pure and clean from other cellular debris that could interfere with Sanger sequencing.

Quality Testing of the DNA Sample

DNA sample quality can be determined using two simple methods. The methods will be discussed in detail in the next article, but are summarized below.

Scanning spectrophotometry using ultraviolet wavelengths between 220 nm and 310 nm provides a general profile of the overall quality of the DNA (Figure 2). DNA absorbs light in the range of 240 nm to 300 nm with the maximum peak at 260 nm. Absorbance at 260 nm is also used to calculate the concentration of the DNA. Another factor often used when measuring DNA quality is calculating the 260 nm / 280 nm absorbance ratio. A good value for this ratio is 1.8 to 1.9. Ratio values below 1.6 indicate the DNA sample contains contaminants.

A good scanning profile does not always prove the DNA is good. Nicked DNA cannot be identified by scanning and may inhibit quality Sanger sequencing. (https://agctsequencing.wordpress.com/2012/02/16/nicked-plasmid-dna-prevents-automated-sanger-sequencing/)

Agarose gel electrophoresis will separate DNA into bands indicating whether the DNA has remained supercoiled.  This supercoiled DNA is required for sequencing. It will also show if the DNA has been damaged and the supercoil has loosened when the DNA is nicked. Nicked DNA migrates more slowly through an agarose gel and will separate from the supercoiled DNA.

Both quality tests provide necessary data to show whether the final DNA sample is clean and of high quality.

Additional Concentration Required

Once the DNA sample has been isolated and appears clean using test procedures, it may be necessary to adjust the concentration to meet submission guidelines. Samples with a concentration higher than required are diluted in water. Although some buffer is still present, the buffer is diluted along with the DNA. Generally dilute buffers (without EDTA) do not interfere with amplification for Sanger sequencing. What if the sample needs to be more concentrated?

Drying a sample in elution buffer is not an effective means for concentrating a sample. Ethanol precipitation is a preferred method because salts are removed along with excess water. Scanning spectrophotometry is an effective means of quantifying the DNA.

What is most important for researchers to remember when isolating plasmid DNA with a commercial kit is to thoroughly read the directions. The kits use enzymes to disrupt the bacterial cell membrane and remove components such as RNA. Enzymes are fragile. Excess shaking or vortexing could damage the enzymes. Commercial kits often include precautions to use care when working with these important enzymes.

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Automated capillary sequencers are also capable of performing fragment analysis applications. The process is similar to sequencing because fluorescent dyes are used to label products amplified by PCR. Fragment analysis is often performed manually loading amplified products on agarose gel. It is a relatively simple and cost effective method for conducting fragment analysis. However, there are also limitations. First, there is no method for accurately calculating fragment sizes even with a size ladder. Second, multiplexing is not possible when product sizes from multiple primer pairs (markers) are similar in size. Finally, agarose gels does not have the resolution capabilities that capillary electrophoresis has. Capillary electrophoresis, used in forensics fingerprinting, provides remarkable accuracy to fragment analysis applications.

Automated Fragment Analysis Uses Color Fluorescent Dyes

The Polymerase Chain Reaction (PCR) first amplifies samples that will be compared by fragment analysis. For manual applications, the forward and reverse primer are simply unlabeled oligonucleotides. For automated capillary equipment the forward primer contains a fluorescent label on the 5’end. The amplified product will also be labeled following PCR because the product includes both primers (figure 1). The forward and reverse primers (markers) determine the region and base pair size of the resulting amplification.

There are advantages to using fluorescent labels in the amplification process. Capillary analyzers recognize fluorescent emission wavelengths (different colors). This adds the capacity to multiplex more than one set of primers in an amplification reaction. For example, forensic science currently runs 16 separate markers in one multiplexed reaction. Markers are labeled with four different fluorescent dyes. Fragment sizes add a second parameter because fragments with the same dye, but amplified with different markers, never overlap.

A Standard is Added to Every Sample

Another parameter that increases accuracy and resolution in automated fragment analysis applications is the addition of a given standard to every sample. The base pair sizes of test samples are calculated using this standard. Motility variance that could occur from one capillary to the next is eliminated because a standard curve is determined for each capillary. Each fragment in the standard would need to be labeled with fluorescent dye. However, the standard uses a unique label not used for any of the samples, typically ROX or TAMRA.

Resolution of Automated Capillary Fragment Analysis is 1 Base Pair

The resolution capability of capillary fragment analysis is 1 base pair. The same technology that separates each base in sequencing is also applied to fragment analysis. A single base pair separation is nearly impossible using an agarose gel (figure 2). Fragment size accuracy is also limited by manual methods because there is no way to view the standard and sample fragments loaded in the same lane.

The introduction of slab gel sequencers improved capacity to perform fragment analysis over manual agarose methods. Development of capillary analyzers has brought greater sensitivity and speed to this well-established process. Very little genomic DNA is now required to perform automated fragment analysis. Although science has developed new methods for sequencing, fragment analysis still remains an efficient and cost effective method to analyze genomic variability; particularly related to genetic disease and forensics.

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Development of simple-to-use purification kits from a number of commercial providers has simplified preparations of DNA samples used for automated DNA sequencing. The isolated DNA is generally clean with good yield. The template can then be quantified by spectrophotometry in preparations to submit to a sequence service provider. In addition, a spectrophotometric scan (220 nm to 310 nm) will indicate whether the plasmid DNA contains salts that could interfere with a sequencing reaction. Even though the template appears clean with good concentration, some samples fail. What could cause this? One possibility is that the DNA could be nicked.

What is Nicked DNA?

Plasmid DNA is characteristically a double-stranded supercoiled molecule. A restriction enzyme is often used to cut both strands linearizing the DNA molecule. A nick is an isolated break in one of the two strands keeping the supercoiled form intact (figure 1).

How Does DNA Become Nicked?

DNA can be enzymatically nicked for certain applications. However, nicked DNA is undesirable for automated sequencing. It is likely the DNA was damaged physically by shearing during purification. Causes of damage include excessive vortexing or pipetting that physically break the DNA. Over-drying can also damage supercoiled DNA. Most commercial kits warn against over-drying a DNA preparation.

How to Detect Nicked DNA

The best method for determining whether the DNA has become nicked is using agarose gel electrophoresis. Nicked DNA cannot be identified using spectrophotometry. A typical plasmid preparation is shown in figure 2. Plasmid preparations almost always have some nicked DNA. However, supercoiled DNA should have the darker band in the resulting gel. Lane 3 provides the best quality DNA for automated DNA sequencing. Samples loaded in lane 1 and lane 2 would most likely fail.

Why Does Nicked DNA Fail to Sequence?

The enzyme lock-key model for enzymes provides a simple explanation why nicked DNA does not sequence. The enzyme, Taq polymerase, needs to sit down on the DNA to catalyze the addition of the bases during extension. A nick in the DNA loosens the strands of the supercoiled DNA strands and the enzyme no longer fits the DNA molecule.

How to Prevent DNA from Becoming Nicked

As previously stated, plasmid preparation kits generally isolate both nicked and supercoiled DNA. It is possible to reduce the amount of damage during the purification procedure. One recommendation is to thoroughly read the directions provided for the kit. There are specific steps where shaking DNA in solution should be minimal. Often it could also be recommended to mix reagents and DNA gently. Excessive vortexing and pipetting should be avoided. And the final DNA isolate should not be overly dried.

DNA is relatively stable and could be useful for years when stored under the proper conditions. But DNA has some fragile characteristics as well. Special care in preparation could reduce damage that inhibits successful automated sequencing applications.