Restriction Enzymes in Genome Mapping and Analysis

The concept of DNA fingerprinting or profiling arose in the 1980s as a means to genetically identify individuals based on unique patterns of DNA fragment sizes generated from their genomes. In other words, the DNA fragments and their length variations could be used as differentiating markers, or “fingerprints”, for genetic identification (i.e., of alleles) instead of relying solely on phenotypic characteristics.

Today the use of DNA fingerprinting techniques is very common. While much of the general public is aware of its significance in forensics and criminal cases, DNA fingerprinting and mapping have broad applications in other areas such as disease testing and plant breeding. The extensive use of DNA fingerprinting has led to the development of numerous DNA fingerprinting methods, with the choice of method primarily depending on the experimental goals and the study organism(s).

Restriction fragment length polymorphism (RFLP)

Restriction fragment length polymorphism, or RFLP (pronounced “rif-lip”), is the basis for one of the oldest DNA fingerprinting methods. The typical workflow of this method involves restriction digestion, fragment separation, Southern blotting, probe hybridization, and visualization (Figure 1).

Figure 1. Basic workflow for identifying restriction fragment length polymorphisms (RFLPs).

In the first step, purified genomic DNA is digested with one or more restriction enzymes. The choice of restriction enzymes is usually based on the ability to distinguish genetic variability and the cost of the enzymes. The digested fragments are separated by gel electrophoresis and appear as a continuous smear on the gel due to the broad distribution of fragment sizes generated by the enzymes.

To detect the desired fragments, the gel-separated DNA fragments are transferred to a nitrocellulose or PVDF membrane for handling and detection. A labeled single-stranded DNA probe is hybridized to the membrane to identify a subset of fragments. The results are visualized to reveal the unique RFLP fingerprint.    

Probes for RLFPs are based on single- to low-copy number sequences in a genome and usually range between 500 and 2,000 bases. Probes are labeled to detect even low amounts of samples and identify the fragments that will become the basis of the fingerprint. The resulting RFLP markers observed are a result of specific probe and restriction enzyme combinations. For example, Probe A and EcoRI-digested genomic DNA will define one RFLP for a specific genome. Probe A and HindIII-digested DNA will define a different RFLP for that genome (Figure 2A). Knowledge of the template sequence, though not required, allows faster development of useful RFLP probes.

RFLPs and restriction enzymes can also be used to detect DNA differences between two individuals. Figure 2B illustrates probe hybridization and detection on a simplistic level, comparing two individuals for HindIII-based RFLPs of two alleles (labeled “1” and “2”). In this example, probe A detects different restriction fingerprints in the two individuals due to loss or gain of a HindIII restriction site on allele 2. In most cases, however, fragment length variability between individuals is a result of insertion or deletion of DNA sequences outside of the restriction sites, caused by natural recombination and replication. RFLP analysis is also used in applications such as genetic counseling, plant and animal breeding programs, and disease monitoring.

Despite its usefulness, RFLP analysis has some limitations. The analysis requires a large amount of starting sample DNA, and the entire process is slow and cumbersome. With the development of PCR, many of these drawbacks have been addressed—for example, with amplification fragment length polymorphism (AFLP), which requires far less sample and can be performed using more rapid PCR-based protocols.

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Amplification fragment length polymorphism (AFLP)

AFLP is another genetic mapping technique that relies on RFLP followed by selective PCR to generate amplified fragments from genomic DNA of any organism, without prior knowledge of the genomic sequence. In addition, AFLP analysis requires only small amounts of starting template (typically nanograms).

While AFLP was first developed for plant studies [1], it is now used for a variety of applications, such as:

• Creation of genetic maps for new species
• Determination of relatedness among cultivars
• Establishment of linkage groups in crosses
• Studies of genetic diversity and molecular phylogeny

The AFLP procedure involves digestion of genomic DNA to produce a population of restriction fragments, ligation of priming site adaptors, amplification by PCR, separation of fragments by electrophoresis, and finally visualization of PCR amplicons by either autoradiography or fluorescence (Figure 3).

In the first step of AFLP, genomic DNA samples are enzymatically digested, typically with EcoRI and MseI. The GC content of the genomic DNA is relevant to the effectiveness of digestion and the resulting fingerprint. For example, optimal digestion with MseI is obtained when the GC content is <50%. High GC content (e.g., >65%) may hinder MseI digestion and result in a low quantity of fragments. Similarly, low GC content favors more complete digestion by EcoRI.

After digestion is complete, two adaptors are ligated to the ends of the fragments. The design of these adaptors includes an additional 19–22 bp to allow subsequent primer binding, and their ligation results in the loss of the original MseI and EcoRI recognition sites (Figure 3B).

Next, two rounds of PCR amplification are carried out, one that is pre-selective followed by a more selective reaction. During the pre-selective PCR, DNA fragments with ligated EcoRI and MseI adaptors are amplified (Figure 3C). After this step, a more selective PCR is performed with primer sets that carry up to three additional bases at the 3′ end of the primer (Figure 3D). These primers include selective nucleotides that, in combination with stringent PCR conditions, help ensure that amplification occurs only for those DNA fragments that share sequence homology with the 3′ ends of the selective primers. In the selective amplification step, the primers to the EcoRI adaptors are either radioactively or fluorescently labeled for downstream detection of fragments, whereas MseI primers are unlabeled.

The combination of selective primers and selective labeling refines the subsequent map to reveal a unique fingerprint by gel or capillary electrophoresis(Figure 3E). The optimal range for a fingerprint after amplification is between 50 and 200 fragments, ranging from 45 to 500 nucleotides in length. Of the visualization methods, fluorescent labeling in combination with capillary gel electrophoresis is preferred, since the process can be automated from fragment separation to data collection for efficiency and robustness.

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Methylation is an endogenous DNA modification and regulatory strategy occurring in the genomes of both eukaryotes and prokaryotes. While it is part of the restriction-modification system in prokaryotes (see Restriction enzyme basics), DNA methylation plays an important role in regulation of gene expression in higher eukaryotes [2]. DNA methylation has been found to be involved in the regulation of many cellular processes and disease states such as embryonic development, X-chromosome gene silencing, cell cycle regulation, and oncogenesis.

In many plants and animals, methylation normally occurs at 5′-CpG-3′, with a methyl group added to the fifth carbon of the nitrogenous ring of cytosine to produce 5-methylcytosine [3]. In plants, cytosine methylation may also occur at 5′-CpNpG-3′ and 5′-CpNpN-3′, where N represents any nucleotide but guanine. Also of importance in methylation analysis are CpG islands, 500–2,000 base pair long, GC-rich DNA segments typically found in promoters and the first exons of the genes. CpG islands are intensively studied because of the potential role of their methylation state(s) in the inactivation and activation of gene transcription.

Among methods available to investigate DNA methylation state, restriction enzymes are a popular tool due to their availability and ease of use. Some restriction enzymes’ recognition sites contain CpG sequences, and the enzymes may display varying sensitivity towards methylated substrates (e.g., unaffected, impaired, blocked, or dependent; see also Restriction enzyme key considerations). Taking advantage of this property, researchers widely use restriction endonucleases as a basis for locus-specific methylation analyses. Some well-known approaches include combined bisulfite restriction analysis (COBRA) and restriction digestion followed by qPCR (also known as quantitative analysis of DNA methylation using real-time PCR, or qAMP).

Combined bisulfite restriction analysis (COBRA)

The COBRA assay is one of the most popular methods for locus-specific CpG methylation analysis. Its workflow involves chemical conversion of cytosine by bisulfite treatment, amplification of the bisulfite-treated DNA by PCR, restriction digestion of the PCR products, and finally, examination of the restriction pattern to investigate cytosine methylation at the locus of interest [4,5].

Figure 4 illustrates the concept and steps of the COBRA assay. Bisulfite treatment of DNA samples converts unmethylated cytosines to uracils but has no effect on methylated cytosines (5-methylcytosines). In subsequent PCR, primers are designed to amplify CpG islands in the selected locus. During formation of new strands in PCR, uracils are replicated as thymines while 5-methylcytosines are replicated as cytosines. The PCR products are then digested using restriction enzyme(s) with CpG-containing recognition site(s) in the amplified region. If the original locus sequence is unmethylated, no cleavage occurs since cytosines in the restriction site are now substituted with thymines after PCR. If the original sequence contains 5-methylcytosines in the restriction site, the enzyme(s) will cleave the DNA as their recognition site(s) are unmodified after PCR. Using the cleavage pattern of the PCR products, the methylation state of the original locus sequence can be determined.

There are several considerations in each step of the COBRA assay. In the bisulfite treatment, the efficiency of cytosine conversion is critical since any remaining unmethylated cytosines would yield false positive results. However, bisulfite treatment is harsh on the DNA and should be carried out no longer than necessary, to avoid DNA damage. Follow the kit manufacturer’s recommended time for bisulfite conversion, or determine the conversion efficiency empirically after treatment. As the DNA sequence would contain uracils in place of unmethylated cytosines after bisulfite treatment, the DNA polymerase must be able to read through uracil-containing DNA templates during PCR (e.g., Taq-based DNA polymerase). Finally, in the digestion step, the selected restriction enzyme(s) must be able to cleave the PCR products efficiently to obtain reliable cleavage patterns for methylation analysis.  Once these aspects are appropriately addressed, COBRA represents an elegant method for methylation analysis.

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Quantitative DNA methylation analysis using restriction enzymes and real-time PCR

Another popular way to analyze locus-specific DNA methylation involves direct restriction digestion of DNA templates, followed by real-time or quantitative PCR (qPCR). This method requires a set of isoschizomers or neoschizomers with different sensitivity to methylation, to investigate cytosine methylation at a specific locus. By circumventing bisulfite conversion of DNA (as with the COBRA assay), this approach requires less DNA input, avoids potential DNA damage from bisulfite treatment, and simplifies the workflow. In addition, qPCR provides a more quantitative examination of sample methylation than the COBRA assay [6,7].

The criteria in choosing isoschizomers or neoschizomers with differing methylation sensitivity include the locus of interest (e.g., CpG islands or gene bodies) and availability of the enzymes. A commonly used enzyme pair for CpG methylation studies is MspI and HpaII, which share the recognition site 5′-CCGG-3′. Cytosine methylation at the target site completely blocks HpaII cleavage, while MspI activity is unaffected. Alternatively, to study methylation at gene bodies (defined as the transcribed sequence of a gene) or low-GC genomic regions, one may instead choose isoschizomers like HpyF30I and TaqI, which share the recognition sequence 5′-TCGA-3′. When the cytosine in the sequence is methylated, cleavage with HpyF30I is blocked but with TaqI it is not affected. In addition to methylation-sensitive iso/neoschizomers, a methylation-dependent restriction enzyme that only cleaves methylcytosine-containing sequences may be included for a more thorough characterization of locus-specific methylation (Figure 5A).

After restriction digestion, qualitative analysis of the methylation can be achieved by running the DNA samples directly on a gel. Please note that digested DNA may appear as smears or multiple smaller bands. With gel electrophoresis, higher DNA loading may be necessary to visualize the fragments, depending on the sensitivity of the detection method (Figure 5B).

For quantitative methylation analysis, the digested DNA samples are subjected to qPCR. The PCR primers are designed to flank and amplify the locus of interest such that DNA substrates that remain undigested serve as the PCR templates. Levels of remaining uncleaved DNA depend upon the methylation sensitivities of the restriction enzymes used on the loci, as illustrated in Figure 5A. The higher the level of intact DNA, the more DNA template available for PCR amplification, and the sooner DNA amplification is detected in real-time PCR (i.e., lower C_t values). Using this principle, the methylation status of the locus can be determined from the qPCR analysis (Figure 5C).

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