Cleavage of RNA by restriction enzymes?

Cleavage of RNA by restriction enzymes?

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Six restriction enzymes discussed in Sequence-specific cleavage of RNA by Type II restriction enzymes (Murray et al.) have the ability to detect and cut RNA strands with that enzyme's recognition sequence.

As you know, restriction enzymes come from a system carried by some bacteria referred to as a restriction-modification system.

With this fact in mind while reading the paper above, I wondered how the bacteria carrying one of these six systems protects it's own RNA from restriction. Do the corresponding methyltranferase in each system methylate DNA as well as RNA? Would this not greatly sacrifice gene expression rates seeing as the methyltranferase surely cannot methylate all the RNA sequences being produced?

Perhaps I misunderstood the paper somehow. Could it be only in some circumstances outside the cell that these restriction enzymes have this unique ability?

This paper found that these enzymes recognize RNA:DNA heteroduplexes. Such duplexes are unlikely to be encountered in vivo. They are present when DNA is primed for replication, but these duplexes are relatively short and thus are less likely to randomly contain a recognition sequence. Furthermore, if the recognition sequence is found in the primer, the gDNA will be methylated specifically to protect it from degradation. RNA:DNA duplexes are also found during transcription, but they are also short and the gDNA would be methylated. Additionally, the transcription bubble where the heteroduplex forms is protected by RNA polymerase, which subsequently displaces the transcript from the gDNA as it leaves the elongating complex.

It's also important to note that this paper found that these enzymes don't cleave RNA:DNA as well as they do DNA:DNA duplexes and these results occurred under experimental conditions with relatively high concentrations of enzyme and substrate. The lower catalytic efficiency is likely due to lower binding affinity, which could be caused by:

  • presence of uracil in RNA (this was explicitly mentioned in the paper)
  • 2'-OH presenting a steric barrier
  • helix shape (RNA tends to adopt an A-form helix)

As to why they don't recognize RNA-DNA heteroduplexes (which are present during transcription, for example), I suspect that the methylation which protects bacterial genomic dsDNA (see the DNA modifying enzyme section of this Columbia University lecture for more info) also protects RNA-DNA hybrids, as the genomic DNA would still be methylated.

Note that most of the enzymes featured in the paper are listed by New England Biolabs as methylation sensitive (for example: AvaII, BanI, HinfI, TaqI).

@canadianer and @user137 provide a host of reasons that these restriction enzymes cannot cleave ds/ssRNA in the comments on canadianer's answer (different helix form, different interaction strengths between strands).

Translocation-coupled DNA cleavage by the Type ISP restriction-modification enzymes

Production of endonucleolytic double-strand DNA breaks requires separate strand cleavage events. Although catalytic mechanisms for simple, dimeric endonucleases are known, there are many complex nuclease machines that are poorly understood. Here we studied the single polypeptide Type ISP restriction-modification (RM) enzymes, which cleave random DNA between distant target sites when two enzymes collide after convergent ATP-driven translocation. We report the 2.7-Å resolution X-ray crystal structure of a Type ISP enzyme−DNA complex, revealing that both the helicase-like ATPase and nuclease are located upstream of the direction of translocation, an observation inconsistent with simple nuclease-domain dimerization. Using single-molecule and biochemical techniques, we demonstrate that each ATPase remodels its DNA-protein complex and translocates along DNA without looping it, leading to a collision complex in which the nuclease domains are distal. Sequencing of the products of single cleavage events suggests a previously undescribed endonuclease model, where multiple, stochastic strand-nicking events combine to produce DNA scission.


Ribonuclease H is a family of endonuclease enzymes with a shared substrate specificity for the RNA strand of RNA-DNA duplexes. By definition, RNases H cleave RNA backbone phosphodiester bonds to leave a 3' hydroxyl and a 5' phosphate group. [7] RNases H have been proposed as members of an evolutionarily related superfamily encompassing other nucleases and nucleic acid processing enzymes such as retroviral integrases, DNA transposases, Holliday junction resolvases, Piwi and Argonaute proteins, various exonucleases, and the spliceosomal protein Prp8. [8] [9]

RNases H can be broadly divided into two subtypes, H1 and H2, which for historical reasons are given Arabic numeral designations in eukaryotes and Roman numeral designations in prokaryotes. Thus the Escherichia coli RNase HI is a homolog of the Homo sapiens RNase H1. [2] [7] In E. coli and many other prokaryotes, the rnhA gene encodes HI and the rnhB gene encodes HII. A third related class, called HIII, occurs in a few bacteria and archaea it is closely related to prokaryotic HII enzymes. [4]

The structure of RNase H commonly consists of a 5-stranded β-sheet surrounded by a distribution of α-helices. [10] All RNases H have an active site centered on a conserved sequence motif composed of aspartate and glutamate residues, often referred to as the DEDD motif. These residues interact with catalytically required magnesium ions. [7] [5]

RNases H2 are larger than H1 and usually have additional helices. The domain organization of the enzymes varies some prokaryotic and most eukaryotic members of the H1 group have an additional small domain at the N-terminus known as the "hybrid binding domain", which facilitates binding to RNA:DNA hybrid duplexes and sometimes confers increased processivity. [2] [7] [11] While all members of the H1 group and the prokaryotic members of the H2 group function as monomers, eukaryotic H2 enzymes are obligate heterotrimers. [2] [7] Prokaryotic HIII enzymes are members of the broader H2 group and share most structural features with H2, with the addition of an N-terminal TATA box binding domain. [7] Retroviral RNase H domains occurring in multidomain reverse transcriptase proteins have structures closely resembling the H1 group. [5]

RNases H1 have been extensively studied to explore the relationships between structure and enzymatic activity. They are also used, especially the E. coli homolog, as model systems to study protein folding. [12] [13] [14] Within the H1 group, a relationship has been identified between higher substrate-binding affinity and the presence of structural elements consisting of a helix and flexible loop providing a larger and more basic substrate-binding surface. The C-helix has a scattered taxonomic distribution it is present in the E. coli and human RNase H1 homologs and absent in the HIV RNase H domain, but examples of retroviral domains with C-helices do exist. [15] [16]

Ribonuclease H enzymes cleave the phosphodiester bonds of RNA in a double-stranded RNA:DNA hybrid, leaving a 3' hydroxyl and a 5' phosphate group on either end of the cut site with a two-metal-ion catalysis mechanism, in which two divalent cations, such as Mg2+ and Mn2+, directly participate in the catalytic function. [17] Depending on the differences in their amino acid sequences, these RNases H are classified into type 1 and type 2 RNases H. [18] [19] Type 1 RNases H have prokaryotic and eukaryotic RNases H1 and retroviral RNase H. Type 2 RNases H have prokaryotic and eukaryotic RNases H2 and bacterial RNase H3. These RNases H exist in a monomeric form , except for eukaryotic RNases H2, which exist in a heterotrimeric form. [20] [21] RNase H1 and H2 have distinct substrate preferences and distinct but overlapping functions in the cell. In prokaryotes and lower eukaryotes, neither enzyme is essential, whereas both are believed to be essential in higher eukaryotes. [2] The combined activity of both H1 and H2 enzymes is associated with maintenance of genome stability due to the enzymes' degradation of the RNA component of R-loops. [22] [23]

Ribonuclease H1 Edit

Ribonuclease H1 enzymes require at least four ribonucleotide-containing base pairs in a substrate and cannot remove a single ribonucleotide from a strand that is otherwise composed of deoxyribonucleotides. For this reason, it is considered unlikely that RNase H1 enzymes are involved in the processing of RNA primers from Okazaki fragments during DNA replication. [2] RNase H1 is not essential in unicellular organisms where it has been investigated in E. coli, RNase H1 knockouts confer a temperature-sensitive phenotype, [7] and in S. cerevisiae, they produce defects in stress response. [24]

In many eukaryotes, including mammals, RNase H1 genes include a mitochondrial targeting sequence, leading to expression of isoforms with and without the MTS present. As a result, RNase H1 is localized to both mitochondria and the nucleus. In knockout mouse models, RNase H1-null mutants are lethal during embryogenesis due to defects in replicating mitochondrial DNA. [2] [25] [26] The defects in mitochondrial DNA replication induced by loss of RNase H1 are likely due to defects in R-loop processing. [23]

Ribonuclease H2 Edit

In prokaryotes, RNase H2 is enzymatically active as a monomeric protein. In eukaryotes, it is an obligate heterotrimer composed of a catalytic subunit A and structural subunits B and C. While the A subunit is closely homologous to the prokaryotic RNase H2, the B and C subunits have no apparent homologs in prokaryotes and are poorly conserved at the sequence level even among eukaryotes. [27] [28] The B subunit mediates protein-protein interactions between the H2 complex and PCNA, which localizes H2 to replication foci. [29]

Both prokaryotic and eukaryotic H2 enzymes can cleave single ribonucleotides in a strand. [2] however, they have slightly different cleavage patterns and substrate preferences: prokaryotic enzymes have lower processivity and hydrolyze successive ribonucleotides more efficiently than ribonucleotides with a 5' deoxyribonucleotide, while eukaryotic enzymes are more processive and hydrolyze both types of substrate with similar efficiency. [2] [30] The substrate specificity of RNase H2 gives it a role in ribonucleotide excision repair, removing misincorporated ribonucleotides from DNA, in addition to R-loop processing. [31] [32] [29] Although both H1 and H2 are present in the mammalian cell nucleus, H2 is the dominant source of RNase H activity there and is important for maintaining genome stability. [29]

Some prokaryotes possess an additional H2-type gene designated RNase HIII in the Roman-numeral nomenclature used for the prokaryotic genes. HIII proteins are more closely related to the H2 group by sequence identity and structural similarity, but have substrate preferences that more closely resemble H1. [7] [33] Unlike HI and HII, which are both widely distributed among prokaryotes, HIII is found in only a few organisms with a scattered taxonomic distribution it is somewhat more common in archaea and is rarely or never found in the same prokaryotic genome as HI. [34]

The active site of nearly all RNases H contains four negatively charged amino acid residues, known as the DEDD motif often a histidine e.g in HIV-1, human or E. coli is also present. [2] [7]

The charged residues bind two metal ions that are required for catalysis under physiological conditions these are magnesium ions, but manganese also usually supports enzymatic activity, [2] [7] while calcium or high concentration of Mg2+ inhibits activity. [11] [35] [36]

Based on experimental evidence and computer simulations the enzyme activates a water molecule bound to one of the metal ions with the conserved histidine. [35] [37] The transition state is associative in nature [38] and forms an intermediate with protonated phosphate and deprotonated alkoxide leaving group. [37] The leaving group is protonated via the glutamate which has an elevated pKa and is likely to be protonated. The mechanism is similar to RNase T and the RuvC subunit in the Cas9 enzyme which both also use a histidine and a two-metal ion mechanism.

The mechanism of the release of the cleaved product is still unresolved. Experimental evidence from time-resolved crystallography and similar nucleases points to a role of a third ion in the reaction recruited to the active site. [39] [40]

The human genome contains four genes encoding RNase H:

    , an example of the H1 (monomeric) subtype , the catalytic subunit of the trimeric H2 complex , a structural subunit of the trimeric H2 complex , a structural subunit of the trimeric H2 complex

In addition, genetic material of retroviral origin appears frequently in the genome, reflecting integration of the genomes of human endogenous retroviruses. Such integration events result in the presence of genes encoding retroviral reverse transcriptase, which includes an RNase H domain. An example is ERVK6. [41] Long terminal repeat (LTR) and non-long terminal repeat (non-LTR) retrotransposons are also common in the genome and often include their own RNase H domains, with a complex evolutionary history. [42] [43] [44]

Role in disease Edit

In small studies, mutations in human RNase H1 have been associated with chronic progressive external ophthalmoplegia, a common feature of mitochondrial disease. [26]

Mutations in any of the three RNase H2 subunits are well-established as causes of a rare genetic disorder known as Aicardi–Goutières syndrome (AGS), [3] which manifests as neurological and dermatological symptoms at an early age. [46] The symptoms of AGS closely resemble those of congenital viral infection and are associated with inappropriate upregulation of type I interferon. AGS can also be caused by mutations in other genes: TREX1, SAMHD1, ADAR, and MDA5/IFIH1, all of which are involved in nucleic acid processing. [47] Characterization of mutational distribution in an AGS patient population found 5% of all AGS mutations in RNASEH2A, 36% in 2B, and 12% in 2C. [48] Mutations in 2B have been associated with somewhat milder neurological impairment [49] and with an absence of interferon-induced gene upregulation that can be detected in patients with other AGS-associated genotypes. [47]

Two groups of viruses use reverse transcription as part of their life cycles: retroviruses, which encode their genomes in single-stranded RNA and replicate through a double-stranded DNA intermediate and dsDNA-RT viruses, which replicate their double-stranded DNA genomes through an RNA "pregenome" intermediate. Pathogenic examples include human immunodeficiency virus and hepatitis B virus, respectively. Both encode large multifunctional reverse transcriptase (RT) proteins containing RNase H domains. [51] [52]

Retroviral RT proteins from HIV-1 and murine leukemia virus are the best-studied members of the family. [53] [54] Retroviral RT is responsible for converting the virus' single-stranded RNA genome into double-stranded DNA. This process requires three steps: first, RNA-dependent DNA polymerase activity produces minus-strand DNA from the plus-strand RNA template, generating an RNA:DNA hybrid intermediate second, the RNA strand is destroyed and third, DNA-dependent DNA polymerase activity synthesizes plus-strand DNA, generating double-stranded DNA as the final product. The second step of this process is carried out by an RNase H domain located at the C-terminus of the RT protein. [5] [6] [55] [56]

RNase H performs three types of cleaving actions: non-specific degradation of the plus-strand RNA genome, specific removal of the minus-strand tRNA primer, and removal of the plus-strand purine-rich polypurine tract (PPT) primer. [57] RNase H plays a role in the priming of the plus-strand, but not in the conventional method of synthesizing a new primer sequence. Rather RNase H creates a "primer" from the PPT that is resistant to RNase H cleavage. By removing all bases but the PPT, the PPT is used as a marker for the end of the U3 region of its long terminal repeat. [56]

Because RNase H activity is required for viral proliferation, this domain has been considered a drug target for the development of antiretroviral drugs used in the treatment of HIV/AIDS and other conditions caused by retroviruses. Inhibitors of retroviral RNase H of several different chemotypes have been identified, many of which have a mechanism of action based on chelation of the active-site cations. [58] Reverse-transcriptase inhibitors that specifically inhibit the polymerase function of RT are in widespread clinical use, but not inhibitors of the RNase H function it is the only enzymatic function encoded by HIV that is not yet targeted by drugs in clinical use. [55] [59]

RNases H are widely distributed and occur in all domains of life. The family belongs to a larger superfamily of nuclease enzymes [8] [9] and is considered to be evolutionarily ancient. [60] In prokaryotic genomes, multiple RNase H genes are often present, but there is little correlation between occurrence of HI, HII, and HIII genes and overall phylogenetic relationships, suggesting that horizontal gene transfer may have played a role in establishing the distribution of these enzymes. RNase HI and HIII rarely or never appear in the same prokaryotic genome. When an organism's genome contains more than one RNase H gene, they sometimes have significant differences in activity level. These observations have been suggested to reflect an evolutionary pattern that minimizes functional redundancy among RNase H genes. [7] [34] RNase HIII, which is unique to prokaryotes, has a scattered taxonomic distribution and is found in both bacteria and archaea [34] it is believed to have diverged from HII fairly early. [61]

The evolutionary trajectory of RNase H2 in eukaryotes, especially the mechanism by which eukaryotic homologs became obligate heterotrimers, is unclear the B and C subunits have no apparent homologs in prokaryotes. [2] [28]

Because RNase H specifically degrades only the RNA in double-stranded RNA:DNA hybrids, it is commonly used as a laboratory reagent in molecular biology. Purified preparations of E. coli RNase HI and HII are commercially available. RNase HI is often used to destroy the RNA template after first-strand complementary DNA (cDNA) synthesis by reverse transcription. It can also be used to cleave specific RNA sequences in the presence of short complementary segments of DNA. [62] Highly sensitive techniques such as surface plasmon resonance can be used for detection. [63] [64] RNase HII can be used to degrade the RNA primer component of an Okazaki fragment or to introduce single-stranded nicks at positions containing a ribonucleotide. [62] A variant of hot start PCR, known as RNase H-dependent PCR or rhPCR, has been described using a thermostable RNase HII from the hyperthermophilic archaeon Pyrococcus abyssi. [65] Of note, the ribonuclease inhibitor protein commonly used as a reagent is not effective at inhibiting the activity of either HI or HII. [62]

Ribonucleases H were first discovered in the laboratory of Peter Hausen when researchers found RNA:DNA hybrid endonuclease activity in calf thymus in 1969 and gave it the name "ribonuclease H" to designate its hybrid specificity. [27] [66] [67] RNase H activity was subsequently discovered in E. coli [68] and in a sample of oncoviruses with RNA genomes during early studies of viral reverse transcription. [69] [70] It later became clear that calf thymus extract contained more than one protein with RNase H activity [71] and that E. coli contained two RNase H genes. [72] [73] Originally, the enzyme now known as RNase H2 in eukaryotes was designated H1 and vice versa, but the names of the eukaryotic enzymes were switched to match those in E. coli to facilitate comparative analysis, yielding the modern nomenclature in which the prokaryotic enzymes are designated with Roman numerals and the eukaryotic enzymes with Arabic numerals. [2] [27] [74] [75] The prokaryotic RNase HIII, reported in 1999, was the last RNase H subtype to be identified. [74]

Characterizing eukaryotic RNase H2 was historically a challenge, in part due to its low abundance. [2] Careful efforts at purification of the enzyme suggested that, unlike the E. coli RNase H2, the eukaryotic enzyme had multiple subunits. [76] The S. cerevisiae homolog of the E. coli protein (that is, the H2A subunit) was easily identifiable by bioinformatics when the yeast genome was sequenced, [77] but the corresponding protein was found not to have enzymatic activity in isolation. [2] [24] Eventually, the yeast B and C subunits were isolated by co-purification and found to be required for enzymatic activity. [78] However, the yeast B and C subunits have very low sequence identity to their homologs in other organisms, and the corresponding human proteins were conclusively identified only after mutations in all three were found to cause Aicardi–Goutières syndrome. [2] [3]

USER cloning

USER-friendly DNA engineering and cloning was developed as a restriction enzyme- and ligase-independent DNA assembly method (1-3). The method relies on generating overlapping PCR products suitable for seamless and directional assembly of customized DNA molecules. PCR primers are designed to have 5´-overlapping sequences containing a single deoxyuridine (dU) residue that is incorporated in place of deoxythymidine (dT) at a 6-10 nt distance from the 5´-terminus. The generated PCR fragments are flanked by sequence overlaps and have a single dU residue per each DNA strand. The PCR fragments are then treated with USER/Thermolabile USER II Enzyme, which specifically nicks the dU-containing strands and generates 3´ single-stranded extensions suitable for directional assembly of PCR fragments. NEB sells USER and Thermolabile USER II enzymes for ligase and restriction enzyme independent cloning reactions.


The data presented here indicates that the upstream ATPase of a Type ISP enzyme remodels its downstream MTase-TRD-target complex to allow target release ( Fig. 6 ). Upon binding of ATP, Type ISP domain movement and ensuing rearrangements in the ATPase-DNA interactions, including those involving the β-hairpin loop, would pull the DNA in an upstream direction leading to movement downstream of the complete enzyme without a necessity for DNA loop formation. This model is quite distinct to that of the classical Type I RM enzymes. We suggest that the coupler will play a key role in transferring conformational strain during initiation. The flipped base seen in the MTase active site would also act as an 𠇊nchor” to prevent target release this must be returned to its base-paired location in the DNA before initiation can proceed and this may be accelerated by ATP-induced strain relayed by the DNA and/or coupler.

Model for loop-independent DNA translocation and extensive nucleolytic DNA processing. (1) The pre-initiation complex. The nuclease is in an inactive conformation. (2) The ATPase cycle loosens the MTase-TRD grip on the DNA. (3) dsDNA translocation downstream of the target (red). (4) Convergence of two enzymes initially brings the nucleases approximately 75 bp apart. (5) Example of stochastic 𠇍NA shredding” by a collision complex. Numbers are the order of the nicking events.

Remodelling of the MTase-TRD must involve loss of target interactions. However, the remodelled MTase-TRD may remain associated with the DNA to act as a sliding clamp, enhancing processivity. Alternatively, the clamp may open fully, allowing the coupler-MTase-TRD units to swing completely off the DNA. This motion may help explain the

30 bp closest approach of the nuclease domains ( Fig. 5c,d ).

The Type ISP remodelling activity is somewhat similar to the role of the ATPase subunit of the Type III RM enzymes. 12 However, whilst those enzymes use a burst of ATPase activity to break the MTase-target interactions, subsequent bidirectional movement along DNA is thermally-driven without the need for ATP hydrolysis. In contrast, the Type ISP enzymes continue to consume ATP during unidirectional translocation. 18 It remains to be seen whether the Type III enzymes also move towards their cognate MTase subunits, “pushing” them off the target as in the Type ISP scheme ( Fig. 6 ), or move in the opposite direction to “pull” the MTase subunits off the target.

The Type ISP structure also provides a simple, single polypeptide framework to understand other remodelling processes that may involve active disruption of protein-nucleic acid complexes by an ATPase motor, such as the DExH/D RNA helicase, 38 SF1 DNA helicase PcrA, 39,40 the Swi2/Snf2 family ATPase Mot1, 15,17 or nucleosome remodellers. 15,16 The above structural comparison suggests that the ATPase cycle proposed for bona fide helicases, involving domain closure and opening of N- and C-cores upon ATP binding and hydrolysis 26,27,29,30 is conserved in dsDNA translocating ATPases. In turn, the Type ISP ATPase would therefore use an equivalent inchworm mechanism for both remodelling and translocation. Interestingly, the β-hairpin loop is unique to Type ISP ATPases (Supplementary Fig. 7), indicating variation in the translocation mechanisms amongst SF2 ATPases despite conservation of canonical motifs and overall structures.

The wide spacing of the DNA strand breaks produced ( Fig. 5 ) is consistent with the upstream locations of the nuclease domains as predicted in a head-on collision complex (Supplementary Fig. 10a). Since the nucleases cannot interact directly, activation upon collision is most likely due to conformational strain from the ATPases transferred via the couplers. Nonetheless, additional remodelling and movement of the collision complex must be required to further process the DNA to produce a dsDNA break. We propose a 𠇍NA Shredder” model in which cumulative nicking events eventually result in a dsDNA break ( Fig. 6 ). This contrasts with the precise DNA cleavage produced by two closely-spaced nucleases as suggested for Type I and III enzymes (Supplementary Fig. 1) and observed in a great many other, simpler nucleases.

The collision complex would initially result in formation of one or two strand breaks. The location of this initial cleavage will be random, dictated by the preceding stochastic translocation events. Rearrangements of the complex due to interdomain plasticity together with the nuclease’s sequence preference (Supplementary Fig. 13c), can result in varied loci of cleavage and hence a range of initial 3′-3′ spacings. The observed 30 bp minimum could reflect a collision complex where both coupler-MTase-TRD units have swung off the DNA, allowing the closest approach of two helicase-nuclease units that still allows for stress-activation. Continued ATPase activity can then remodel the collision complex, leading to further up- and downstream movement and nicking events, causing compound damage culminating in a dsDNA break ( Fig. 6 ). The increases in the median and skewness values ( Fig. 5 ) are thus due to the longer incubation times during which collision complex motion leads to further cleavage events at other sites. Notably, post-collision mobility can also account for cleavage further upstream than � and for rare 5′ overhangs ( Fig. 5b ), where presumably the collision complex nicks one strand and immediately moves in a 3′ direction before nicking the opposite strand.

The loading of multiple translocating enzymes on the DNA that is a potential consequence of target release could result in a pile-up of enzymes on both sides of the primary collision complex. If these enzymes were activated they could cause a time-dependent increase in 3′-3′ cleavage distance. However, post-collision mobility can more readily account for cleavage events upstream of � and the 5′ overhangs. Nonetheless, rear-end collisions may play a supplementary role in DNA processing. We discounted an alternative model where strand nicking by multiple, separate collision events at random sites generates the widely-spaced overhangs since this would generate 5′ overhangs with equal frequency and would not show a time-dependent increase in distance.

We speculate that, depending on the cellular host, broken DNA with variable 3′-overhangs produced by Type ISP enzymes would not be a substrate for the corresponding dsDNA break repair enzymes (RecBCD or AddAB), which have a preference for blunt ends. 41,42,43 Consequently, even if the foreign DNA were to have the regulatory sequences required for homologous recombination, they would not enter that repair pathway. dsDNA break formation by multiple nicking events would additionally prevent simple religation of the DNA ends. Where the cell also encodes a CRISPR-Cas system, the small DNA fragments generated during cleavage by Type ISP enzymes (or during post-cleavage processing by classical Type I enzymes 44 ) may feed into the spacer acquisition pathway (adaptation), as suggested for the DNA fragments generated by RecBCD. 45

In conclusion, the distinct nucleolytic activity of the Type ISP enzymes contrasts with mechanisms employed by dimeric nucleases and further illustrates yet another strategy evolved towards resistance in the perpetual bacteriophage-host arms race. 2

Transcription of bacteriophage φX174 in vitro: Analysis with restriction enzymes

The three phage φX174 promotors (PA′, PA and PG) defined and mapped relative to one another by Axelrod (1976) have been mapped onto the bacteriophage genome. RNA chains were specifically initiated in vitro at each promotor with oligonucleotide primers, pulse-labeled with (α- 32 P)-labeled ribonucleoside triphosphates, chased into full-length molecules with an excess of unlabeled ribonucleoside triphosphates, and purified on polyacrylamide gels. The isolated RNA species were hybridized to a series of purified, unlabeled DNA fragments generated by cleavage of φX174 RFI † DNA with restriction endonucleases from Hemophilus influenzae (endonuclease R) or H. aegyptius (endonuclease Z). The 32 P-labeled RNA sequences (approx. 100 to 400 bases) originating from promotors A′, A or G hybridized preferentially to DNA fragments R-4, R-8 and R-6b/Z-3, respectively. The DNA fragments have previously been aligned with the bacteriophage genetic map by independent methods in other laboratories (Chen et al., 1973 Hutchison et al., 1973 Lee and Sinsheimer, 1974a, Lee and Sinsheimer, 1974b Borrias et al., 1976). Comparison of the above hybridization results to that fragment map shows that promotors A′, A and G lie near the starts of cistrons A, B and D.

Abbreviation used: RFI, the double-stranded closed circular supercoiled replicative form DNA of φX174.

Restriction Enzyme Cleavage: &lsquosingle-site&rsquo enzymes and &lsquomulti-site&rsquo enzymes

Restriction enzymes are proteins used to fragment and clone DNA, but their biological function is to protect bacteria and archaea against viral infections. All bind to double-stranded (ds) DNA at specific sequences of base pairs (the &lsquorecognition sequence&rsquo) and cleave the DNA strands. Given their catalytic similarities, we might expect all restriction enzymes to be similar to each other, but they are in fact extremely diverse, and vary widely in amino acid sequence, three-dimensional structure, subunit composition, and modes of action. Restriction enzymes of identical specificity (&lsquoisoschizomers&rsquo) are sometimes similar, and represent diverged versions of the same ancestral protein, but those of different specificity are often unique, and display no more similarity to one another than to unrelated proteins chosen at random. It seems likely that restriction enzymes arose independently many times during microbial evolution and under varied circumstances.

Accompanying this diversity are subtle differences in the ways that restriction enzymes behave. In one comparison, seven enzymes that cleave the sequence GGCGCC at four different positions were examined, and five distinct reaction pathways were discerned (1). Perhaps the most important mechanistic difference when using restriction enzymes as molecular biology reagents concerns the number of recognition sites a restriction enzyme must bind to in order to cleave. Most restriction enzymes act as simple monomers (one protein chain, e.g. MspI (NEB #R0106) (2)) or homodimers (two identical protein chains, e.g. BamHI (NEB #R3136)), which bind and cleave one recognition site at a time. These enzymes cut substrates with one site as efficiently as they cut substrates with several sites. Others are more complex, and undergo allosteric activation, or form &lsquotransient&rsquo dimers (e.g. FokI (NEB #R0109) (3-5)), tetramers (e.g. NgoMIV (NEB #R0564) (6)), or even larger assemblages (e.g. BcgI (NEB #R0545) (7,8)), and these cut only when bound to two, and sometimes more (9), sites at once. In some cases, this &lsquomulti-site&rsquo, behavior might be an adaptation against accidental cleavage of the bacterium&rsquos own DNA. Structurally, it likely stems from the subunit organizations of the enzymes and how their recognition and catalytic domains fit together (8). Irrespective of the why and how, the need to bind to more than one site in order to cleave can make substrates with only one site difficult to cut in vitro.

Restriction enzymes that bind several sites in order to cleave exhibit several characteristics:

    Cleavage kinetics. Substrates with single sites are cleaved slowly and in some cases incompletely because enzymes must interact with (&lsquobridge&rsquo) two or more DNA molecules at once. The probability of doing so declines precipitously at low DNA concentrations. Adding more enzyme to try to improve cleavage in this situation can do the opposite and make matters even worse, because increasing enzyme concentration in effect reduces the relative substrate concentration (refer to #2 below). If the contacts between the subunits are fragile, as they are thought to be for transient dimers such as FokI (NEB #R0109) (10), then enzymes bridging sites in trans, in different molecules, can be unstable and ineffective. In contrast, when multiple sites are present in the same DNA molecule, the local concentration of sites is higher, and enzymes bridging sites in cis are more stable, both of which lead to faster, more complete, cleavage. Multi-site enzymes vary in the degree to which they are affected by these and related factors. Some, such as BspMI/BfuAI (NEB #R0502) (11) and NmeAIII (NEB #R0711) barely cleave 1-site substrates at all. Others (e.g. SacII (NEB #R0157) cut partially, and yet others (e.g. PluTI (NEB #R0713)) can cleave to near-completion.

Given the diversity of restriction enzymes, many exceptions occur, but single-site and multi-site enzymes partition fairly well into two distinguishable groups based on positions of cleavage.

    Single-site enzymes. The majority of restriction enzymes that cleave within or very close to their recognition sequence are active at single-sites. These enzymes are classified as &lsquoType IIP&rsquo if their sequences are palindromic (symmetric), and &lsquoType IIT&rsquo if their sequences are asymmetric and two different catalytic sites are used to cleave the DNA strands. They cleave to completion DNA substrates with only one site as efficiently as they cleave substrates with several sites. Their cleavage ability is not inhibited at high enzyme concentration due to site-saturation, and is not enhanced by the addition of specific oligos.

Type IIT enzymes AciI (NEB #R0551) and EarI (NEB #R0528) are possible exceptions to this generalization, as too are several Type IIP enzymes. These enzymes cleave 1-site substrates slowly, and in some cases, incompletely cleavage is inhibited at high enzyme concentrations due to site-saturation and cleavage can be enhanced by the addition of specific oligos. These Type IIP enzyme exceptions include: AluI (NEB #R0137) BsaWI (NEB #R0567) (25) BsrFI/Cfr10I (NEB #R0562) (14,19,20) Ecl18kI (12,18) EcoRII (9,17,19,20) HpaII (NEB #R0171) (19,20) NaeI (NEB #R0190) (13,24,26,27) NarI/Mly113I (NEB #R0191) (1,19,20,24,28) NgoMIV ((NEB #R0564) 6) PluTI/BbeI (NEB #R0713) (1) RsrII (NEB #R0501) SacII/Cfr42I (NEB #R0157) (19,29) Sau3AI (NEB #R0169) (19,20) SfiI (NEB #R0123) (16,30-35) SgrAI (NEB #R0603) (19,20,36-38) SmlI and XmaI/Cfr9I (NEB #R0180) (19,24), all of which act to one degree or another as multi-site enzymes.


If you are using an enzyme that may require more than one recognition site on the substrate to cleave optimally, we suggest two possible optimization methods: 1) Titrate the units of enzyme used in the reaction to determine the optimal enzyme to substrate ratio. As a starting point, we recommend using 1-2 &mul of restriction enzyme (at the supplied units/&mul) per microgram of substrate and performing 2-fold serial dilutions of the enzyme, keeping the DNA concentration constant. 2) If that is not possible, add duplex oligonucleotides that contain the recognition site. The oligos provide the additional recognition site needed to activate substrate-bound, but dormant enzyme monomers. Addition of duplex DNA containing a recognition site can compete for binding and reduce the effective enzyme concentration. To use oligos effectively, the stoichiometry of the reaction needs to be established beforehand by performing a series of titrations and identifying the optimum range for the concentration of oligos. We recommend keeping the enzyme concentration constant at 2-4 fold above the optimum established in step 1, above, while performing 2-fold serial dilutions of the oligo. As a starting point, we recommend a ratio of 4:1 (oligo sites:substrate sites) and performing a 2-fold serial dilution of the oligo, keeping the enzyme and substrate concentrations constant.

These authors contributed equally: Heng Zhang, Zhuang Li, Renjian Xiao.


Department of Biological Sciences, Purdue University, West Lafayette, IN, USA

Heng Zhang, Zhuang Li, Renjian Xiao & Leifu Chang

Purdue University Center for Cancer Research, Purdue University, West Lafayette, IN, USA

You can also search for this author in PubMed Google Scholar

You can also search for this author in PubMed Google Scholar

You can also search for this author in PubMed Google Scholar

You can also search for this author in PubMed Google Scholar


H.Z., R.X. and Z.L. prepared samples. Z.L., H.Z. and L.C. collected and processed cryo-EM data. H.Z. and R.X. performed biochemical analysis. Z.L., H.Z. and L.C. prepared figures. All authors analyzed the data. H.Z., Z.L. and L.C. prepared the manuscript with input from R.X.

Corresponding authors

Restriction Enzyme Digestion

Read about Type II restriction enzymes and the distinguishing properties of the four principle subtypes.

High throughput sequencing methods have revolutionized genomic analysis by producing millions of sequence reads from an organism’s DNA at an ever decreasing cost.


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What is a Type II Restriction Enzyme?

Type II restriction enzymes are most commonly used for molecular biology applications, as they recognize stereotypical sequences and produce a predictable cleavage pattern. Learn more about how Type II REs work.

What is a Type I Restriction Enzyme?

Type I restriction enzymes are a group of endonucleases that recognize a bipartite sequence, but do not produce a predictable cleavage pattern. Learn more about how Type I REs work.

What is a Type III Restriction Enzyme?

Type III restriction enzymes are a group of endonucleases that recognize a non-pallindromic sequence, comprising two inversely oriented sites. Learn more about these poorly understood enzymes.

Cloning With Restriction Enzymes

Restriction enzymes are an integral part of the cloning workflow, for generating compatible ends on fragments and vectors. This animation discusses three guidelines for determining which restriction enzymes to use in your cloning experiment.

Standard Protocol for Restriction Enzyme Digests

Let one of NEB's restriction enzyme experts help you improve your technique and avoid common mistakes in digest setup.

Why is My Restriction Enzyme Not Cutting DNA?

Not getting the cleavage you expected? Let an NEB scientist help you troubleshoot your reaction.

Restriction Enzyme Digest Problem: Too Many DNA Bands

Are you finding unexpected bands in your digestion reaction? Here are some tips to help you determine the cause.

What is Restriction Enzyme Star Activity?

Learn what Star Activity is, why it is detrimental to accurate restriction enzyme digestion, and how NEB's HF enzymes are engineered to avoid it.

Reduce Star Activity with High-Fidelity Restriction Enzymes

NEB has engineered HF® enzymes to eliminate star activity. Learn how, and what this means for your digests.

NEB ® Restriction Enzyme Double Digest Protocol

Double digestions can save you time, and this video can offer tips for how to achieve the best results, no matter which of NEB's restriction enzymes you're using.

Restriction Enzyme Digest Protocol: Cutting Close to DNA End

When cutting close to the end of a DNA molecule, make sure you know how many bases to add to the ends of your PCR primers.

Restriction Enzyme Digestion Problem: DNA Smear on Agarose Gel

Learn more about what causes this common problem, and how NEB's enzymes are QC'd to avoid DNA smearing.

Restriction Enzymes in Isothermal Amplification

Isothermal amplification generates many copies of a target sequence in a short period of time, at a constant temperature. Learn more about isothermal amplification.

Restriction Enzymes in Optical Mapping

Optical mapping is a method that allows production of restriction maps of whole chromosomes or genomes. Learn more about optical mapping.

Dicer-like enzymes with sequence cleavage preferences

Researchers keep discovering new functions of small RNAs. For instance, they can be used as a defense mechanism against viruses or self-replicating genome invaders. These tiny pieces of RNA are often produced by a cleavage of long precursors by so called Dicer proteins. To their surprise, researchers from the University of Bern have found that some Dicers acquired a unique and as yet unknown feature that allow them to cleave the RNA precursors in a very specific way, resulting in small RNAs that work much more efficiently.

In the human cell as in society: It is all about information. Genes are read, information molecules (called messenger RNAs or short mRNA) are produced and proteins are synthesized according to the instructions. And there are a whole lot more of these messenger substances, the cellular apparatus is a huge information hub, a meticulously regulated business. And as if all these information exchanges were not complicated enough: in the past years RNA-related research has uncovered more and more evidence that regulation works on a higher level too. The information management itself is highly dynamic – information substances are being held back, they are changed, at times even shredded right away.

In eukaryotes especially, the shredded information molecules, called small RNAs (sRNAs), have critical roles in development, gene expression, and genome stability. They are produced by a range of different proteins. One of the best known is called Dicer and cuts them from long double-stranded RNA templates. Up to now, Dicer was thought to be a bit of a lumberjack when it comes to tearing down RNA – just hacking it into short fragments of roughly equal size, paying no special attention to the form and content of the fragments. The proteins have never been shown to have sequence cleavage preferences, but recently a research group from the Institute of Cell Biology of the University of Bern headed by Mariusz Nowacki has found Dicer-like enzymes with truly surprising qualities. The research could lead to new insights into the regulation of cellular communication and might open up as yet uncharted opportunities for future protein engineering.

The research group, which is part of the NCCR "RNA and disease," has found that in the microorganism Paramecium, which uses small RNAs to guide the elimination of invading transposable DNA elements, some Dicer-like proteins are cutting up RNA with much greater diligence than expected. As published in Cell, Cristina Hoehener, Iris Hug, and Mariusz Nowacki report that their Dicer-like enzymes have strict size and pronounced sequence preferences. These preferences result in the production of sRNAs precisely matching the ends of their target DNA elements, which leads the researchers to propose an as yet unknown biological role for these newly characterized enzymes and their cleavage products.

Small RNAs repair the DNA

They think that these Dicer-like proteins facilitate the elimination of unwanted DNA elements by accumulating precisely at their ends. These accumulations then trigger the elimination of the "highlighted" parts of the DNA from the genetic material. Transposons are alien parts of the genome, infiltrated into the DNA, with the ability to jump from one part of the DNA to another. This makes them especially dangerous as they can trigger all kinds of unwanted genetic effects. So an organism has every reason to get rid of these DNA segments quickly and efficiently.

Especially intriguing for the researchers is the level of precision with which these Dicer enzymes do their job. In contrast to restriction enzymes that cut only when they recognize very specific sequences, the Paramecium Dicer proteins function somewhat sloppily—Nowacki calls it a "relaxed preference." The researchers believe that a too strict preference would not be appropriate here: for the DNA repair to be effective it is important that a lot of sRNA fragments can assemble at the right segment. If Dicer would be too picky, it would not be able to produce suitable snippets for all the dangerous foreign DNA. "Fuzziness is not necessarily a bad thing," says Nowacki. Over the course of the evolution many biological processes have evened out between precision and chance—one of the reasons why cells can react with surprisingly tailor-made solutions to all kinds of different threats.

Watch the video: Exonucleases (May 2022).