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Restriction Endonucleases in Genome Mapping and Analysis
Although restriction enzymes are widely used in molecular cloning, their use as molecular tools extends to other common applications in molecular biology. Two important applications are DNA fingerprinting and methylation analysis, which are methods to map sequences and analyze epigenetic patterns in the genome.
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Methylation analysis techniques
Restriction maps are essential tools for genome mapping analysis. Restriction maps provide details of the restriction enzyme cutting sites on in a DNA molecule such as a plasmid. Restriction maps are critical to a wide range of genetic manipulations and analyses.
Restriction enzyme mapping: A bird’s-eye view
Restriction enzyme mapping, also known as restriction mapping, is a method used to determine the locations of restriction enzyme cleavage sites within a DNA strand such as a plasmid. Restriction maps are a blueprint of the locations of restriction enzyme cleavage sites within a DNA molecule. These maps provide a visual representation of the DNA sequence, highlighting where specific restriction enzymes cut the DNA.
Restriction maps are created by digesting DNA with restriction enzymes, analyzing the cut fragments, and a restriction map is constructed, showing the relative positions of the cut sites. Restriction maps have wide ranging applications including gene cloning, mutation detection, plasmid characterization, DNA fingerprinting, and genome mapping. Furthermore, these maps allow comparing DNA sequences among different organisms and are valuable tools in molecular taxonomy.
Figure 1. Restriction map of a pGT232 plasmid [1].
RFLP technique for DNA fingerprinting
Restriction fragment length polymorphism, or RFLP 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. Knowledge of the template sequence, though not required, allows faster development of useful RFLP probes.
Figure 2. Traditional workflow for identifying restriction fragment length polymorphisms (RFLPs).
In the first step of the RFLP workflow, purified genomic DNA is digested with one or more restriction 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.
Simplified RFLP method for molecular taxonomy
16S rRNA gene RFLP analysis is a molecular technique used to identify and classify bacteria based on variations in the 16S ribosomal RNA gene. The 16S rRNA gene is highly conserved among bacteria, making it an ideal target for taxonomic studies.
As with traditional RFLP method genomic DNA from bacterial samples is starting material for this study but instead of immediate digestion the 16S rRNA gene is amplified using specific primers designed to target conserved regions of the gene.
The amplified 16S rRNA gene PCR products are then digested with one or more restriction enzymes, which cut the DNA at specific recognition sites, generating fragments of varying lengths. These DNA fragments are separated by size using agarose gel or PAGE electrophoresis.
The resulting restriction fragment profiles of different bacterial samples are compared. Each bacterial species or strain produces a unique pattern of fragments based on the presence and location of restriction sites within the 16S rRNA gene. The banding patterns are analyzed to identify and classify the bacterial samples and the presence or absence of specific fragments, and the overall pattern can be used to determine genetic relationships and construct phylogenetic trees.
Compared to traditional RFLP analysis this adaptation does not require blotting or probe hybridization and is much simpler and more cost-effective.
Figure 3. 16S rRNA gene RFLP workflow.
RFLPs were widely used in DNA fingerprinting, paternity testing, and analyzing genetic diversity. Figure 2 illustrates how to detect DNA differences between two individuals using restriction digestion and probe hybridization. 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.
Figure 4. Simplified RFLP fingerprinting. (A) Different RFLP markers are obtained from digestion of the same sample with two restriction enzymes separately, EcoRI and HindIII. (B) Two individuals (A and B) are distinguished by HindIII digestions of two alleles (1 and 2).
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, amplification fragment length polymorphism (AFLP) requires far less sample and can be performed using more rapid PCR-based protocols.
Read more on the advantages and limitations of RFLP
Learn more about Thermo Scientific FastDigest HindIII restriction enzyme
DNA fingerprinting for forensic analyses
DNA fingerprinting or profiling arose in the 1980s 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 biomarkers, 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).
DNA fingerprinting for genome mapping
Genome mapping is the process of determining the relative positions of genes and other significant features within a genome. It involves creating a detailed map that shows the locations of genes, regulatory elements, and other sequences of interest along the chromosomes.
Genome mapping vs genome sequencing
Genome mapping determines the relative positions of genes or genetic markers on a chromosome. It provides a framework that shows the order and approximate distances between genes or markers. Genome mapping helps identify the location of genes associated with specific traits or diseases. In contrast, genome sequencing determines the order of nucleotides in a DNA fragment. It provides a detailed readout of the genetic code. Sequencing is used to identify the complete genetic makeup of an organism.
Amplification fragment length polymorphism (AFLP) for genome mapping
AFLP is a genome 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, it is now used for a variety of applications including creation of genetic maps for new species, determination of relatedness among cultivars, establishment of linkage groups in crosses, and studies on genetic diversity and molecular phylogeny [2].
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.
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Figure 5. Illustration of the basic amplified fragment length polymorphism (AFLP) workflow.
In the first step of AFLP, genomic DNA samples are enzymatically digested, typically with EcoRI and MseI. The GC content of the genomic DNA correlates 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 over >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 5B).
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 5D). 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 5E). 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.
How does fragment analysis work?
Fragment analysis refers to a genetic analysis technique used for a wide variety of applications such as mutation detection, genotyping, DNA profiling, genetic mapping, and linkage analysis.
Both restriction fragment length polymorphism (RFLP) and amplified fragment length polymorphism (AFLP) analyses rely on various fragment analysis techniques to separate, visualize, and interpret DNA fragments.
Methylation analysis with restriction enzymes
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, DNA methylation plays an important role in regulation of gene expression in higher eukaryotes [3]. 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 [4]. 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 extensively 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 including unaffected, impaired, blocked, or dependent. 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
The combined bisulfite restriction analysis (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 [5,6].
Figure 6 illustrates the concept and steps of the COBRA assay. Bisulfite treatment of DNA samples convert 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.
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Figure 6. Illustration of the combined bisulfite restriction analysis (COBRA) workflow with a 5’-CGCG-3’ locus sequence and a restriction enzyme (Bsh1236I) that recognizes a CG-containing sequence.
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, for example, 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.
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 [7,8].
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 7A).
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 7B).
Figure 7. Methylation analysis by restriction digestion followed by gel electrophoresis and qPCR.
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 7A. 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, which is reflected in the lower CT values. Using this principle, the methylation status of the locus can be determined from the qPCR analysis (Figure 7C).
References
- Heng NCK, Bateup JM, Loach DM, et al. (1999) Influence of different functional elements of plasmid pGT232 on maintenance of recombinant plasmids in Lactobacillus reuteri populations in vitro and in vivo. Appl Environ Microbiol 65:5378–5385. doi: 10.1128/AEM.65.12.5378-5385.
- Vos P, Hogers R, Bleeker M, et al. (1995) AFLP: a new technique for DNA fingerprinting. Nucleic Acids Res 23(21):4407–4414. doi: 10.1093/nar/23.21.4407.
- Costello JF, Plass C (2001) Methylation matters. J Med Genet 38(5):285–303. doi: 10.1136/jmg.38.5.285.
- Adams RL (1995) Eukaryotic DNA methyltransferases – structure and function. Bioessays 17: 139–145. doi: 10.1002/bies.950170209.
- Xiong Z, Laird PW (1997) COBRA: a sensitive and quantitative DNA methylation assay. Nucleic Acids Res 25(12):2532–2534. doi: 10.1093/nar/25.12.2532.
- Boyko A, Kovalchuk I (2010) Analysis of locus-specific changes in methylation patterns using a COBRA (combined bisulfite restriction analysis) assay. Methods Mol Biol 631:23–31. doi: 10.1007/978-1-60761-646-7_3.
- Oakes CC, La Salle S, Trasler JM et al. (2009) Restriction digestion and real-time PCR (qAMP). Methods Mol Biol 507:271–280. doi: 10.1007/978-1-59745-522-0_20.
- Mao R, Chou LS (2010) Methylation analysis by restriction endonuclease digestion and real-time PCR. Clin Chem 56(7):1050–1052. doi: 10.1373/clinchem.2010.146654.
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