Restriction Enzymes: A History

By Wil A.M. Loenen, Leiden University Medical Center
April 2019 · 346 pages, illustrated (38 color and 26 B&W)
ISBN 978-1-621821-05-2

<<  Chapter 7   —   Chapter 9  >>

Chapter 8

Chapter doi:10.1101/restrictionenzymes_8

Improved Detection Methods, Single-Molecule Studies, and Whole-Genome Analyses Result in Novel Insights on Structures, Functions, and Applications of Type I, II, III, and IV Restriction Enzymes: ∼2004–2016


As mentioned in Chapter 7, with the arrival of whole-genome sequencing projects it has become clear that the Type I subclasses IA–IE and Type III R-M systems are common in bacteria and archaea (http://rebase.neb.com/rebase/rebase.html). The subdivision of Type II REases in 2003 into 11 subtypes based on behavior and cleavage properties (Roberts et al. 2003; Chapter 7) was helpful, but sometimes puzzling: Some REases would fit in more than one category or in none properly; unrelated proteins could be assigned to one or more of these subtypes; and some REases restricted DNA/RNA hybrids (a useful property to study small regulatory RNAs!) (Roberts et al. 2003; Murray et al. 2010; Loenen et al. 2014b). Although this subdivision remains useful, enzyme structures and/or domain organizations can be used as an alternative for classification (Niv et al. 2007; Pingoud et al. 2014). The notion that REases are evolutionarily related despite the lack of sequence similarity has grown more and more compelling, especially because of the increase in crystal structures with the PD···(D/E)XK fold (called the “PD fold”). As discussed in Chapter 7 (and depicted in Figs. 6 and 7 in that chapter), the structural studies by the group of Virginjuis Šikšnys provided formal evidence for a conserved sequence motif in the active site of REases, called the PD (D/E)XK site. The active site residues of EcoRI (G↓AATTC), NgoMIV (G↓CCGGC), and Cfr10I (R↓CCGGY) appeared to be common to 11 other REases belonging to Type IIP, IIE, and IIF (Kovall and Matthews 1999; Pingoud et al. 2002; Šikšnys et al. 2004). The almost simultaneous appearance of the structures of BamHI, PvuII, and EcoRV elicited much excitement (Winkler et al. 1993; Cheng et al. 1994; Newman et al. 1994), and both the papers on BamHI (Newman et al. 1994) and PvuII (Cheng et al. 1994) discussed the core structural motifs identified in the paper by Venclovas et al. (1994) when the structures of EcoRI and EcoRV were compared.

Early attempts to change specificity had not been very successful (Wolfes et al. 1986; Jeltsch et al. 1996; Lukacs et al. 2000; Pingoud et al. 2014). Substitutions usually resulted in a decrease in activity, but without exception failed to produce substantial changes in specificity. These findings led to the important lesson that recognition did not simply involve amino acids in direct contact with the bases and the backbone but also required water molecules and a complex network of other interactions (Pingoud et al. 2014). Sequence-specific DNA recognition by REases often involved binding to B-DNA in the major groove, with or without DNA distortion, similar to many regulatory proteins (see e.g., Rohs et al. 2010; Pingoud et al. 2014). In contrast, recognition by base flipping was used by enzymes that do chemistry: MTases, DNA repair enzymes (Roberts and Cheng 1998; Cheng and Roberts 2001), but also some REases (Bochtler et al. 2006; Horton et al. 2006; Tamulaitis et al. 2007; Szczepanowski et al. 2008; Miyazono et al. 2014; Manakova et al. 2015).

The new class of Type IV REases, defined in 2003 (Roberts et al. 2003), are modification-dependent enzymes that recognize modified Cs and As (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003). Early findings in the history of modified DNA date back quite a while, long before its importance became known. Modified DNA containing m5C was discovered in 1925 (Johnson and Coghill 1925), followed by m6A (Dunn and Smith 1955a,b), and hm5C in the 1950s (Hershey et al. 1953; Wyatt and Cohen 1953). The analysis of T* mutant phages in 1952 (Chapter 1; Luria and Human 1952) led to the discovery of enzymes that glycosylate hm5C (ghm5C) and of host genes that enable restriction of nonglucosylated phage DNA: rglA and rglB (restricts glucose-less DNA; Luria and Human 1952; Revel and Luria 1970). The rglA and rglB genes were renamed mcrA and mcrBC (modified cytosine restriction) (Noyer-Weidner et al. 1986; Raleigh and Wilson 1986) and were the first designated Type IV REases (Roberts et al. 2003).

Current interest in modified bases is high, as research into the dynamics of DNA modifications (“epigenetic phenomena”) have become of paramount importance for research into all kingdoms (Loenen and Raleigh 2014). Interestingly, hm5C was already discovered in eukaryotic (rat) brain and liver in 1972 (Penn et al. 1972) but did not receive much attention until the discovery of the role of Tet (ten-eleven translocation) proteins, a topic outside the scope of this book (Tet proteins are involved in m5C conversion and hence in control of normal and malignant cell differentiation) (see e.g., Veron and Peters 2011; Pastor et al. 2013; Baumann 2014; Stower 2014; Lu et al. 2015; Hendrickson and Cairns 2016; Jeschke et al. 2016). By 1980, some eight types of modified bases in phage DNA had been described (Warren 1980). A little later the m4C modification was found in Bacillus (Janulaitis et al. 1983), which is often present in thermophilic and mesophilic bacteria (Ehrlich et al. 1985, 1987). For an extensive description of the techniques used to detect and analyze such modified bases, see Weigele and Raleigh (2016), whose review discusses the initial harsh chemical treatments, physiological methods, paper chromatography, anion exchange columns, high-performance liquid chromatography (HPLC), mass spectrometry (MS), and SMRT. SMRT technology analyzes fluorescently labeled nucleotides that are incorporated slightly slower when encountering modified bases in the template strand than unmodified template bases during the sequencing procedure. This method thus allows the analysis of the “methylome” (i.e., the distribution of methylated bases in the DNA of different organisms).

This chapter uses the reviews that appeared in 2014 in Nucleic Acids Research as starting material, plus selected talks and posters presented at the 7th NEB meeting in Gdansk in 2015 (Loenen and Raleigh 2014; Loenen et al. 2014a,b; Mruk and Kobayashi 2014; Pingoud et al. 2014; Rao et al. 2014). Groups in Atlanta (Cheng), Bangalore (Rao and Nagaraja), Baltimore (Chandrasegaran), Berlin (Reuter and Kruger), Bristol (Halford and Szczelkun), Delft (Dekker), Edinburgh (Dryden), Gdansk (Mruk and Skowron), Giessen (Pingoud), Moscow (Zavil'gel'skii), New York (Aggarwal), Piscataway (Bogdanova), Pittsburgh (Jen-Jacobson), Portsmouth (Kneale), Seattle (Stoddard), Tokyo (Kobayashi), Tucson (Horton), Vilnius (Lubys and Šikšnys), and Warsaw (Piekarowicz, Bujnicki, and Bochtler) and at NEB (Roberts, Raleigh, Morgan, and Wilson) made important contributions to the field, as discussed throughout this chapter. Control (C) proteins of Type II enzymes were studied by the groups of Bob Blumenthal in Toledo and Geoff Kneale in Portsmouth. Data about the four types from approximately 2004 onward will be discussed, including structures of some Type II REases, as well as the very first structures of the other types, which together reveal many new, unexpected, and amazing details about the mechanisms employed to prevent indiscriminate restriction by the REase (subunit). Other types of control of restriction were elucidated, via transcription regulation, DNA mimics, C proteins, or the cognate MTase. R-M genes and lone MTase genes in pathogenic organisms also became of great interest because they are linked to virulence via “phase variation” (Piekarowicz 2013). Yet another breakthrough was the discovery of the family of the Type II REase MmeI, which would finally allow the generation of the new specificities so long hoped for. As in the previous chapters, this final chapter starts with the Type II REases (Part A), followed by the ATP-dependent Type I (Part B) and III (Part C) R-M systems, and the modification-dependent Type IV REases (Part D). The final section discusses the phenomenon of phase variation used by pathogenic bacteria to combat phage and evade host immunity (Part E).



By 2014, approximately 4000 REases had been identified belonging to more than 350 different prototype Type II REases (i.e., biochemically different) (Roberts et al. 2010; Pingoud et al. 2014). The majority of these prototypes had characterized or putative relatives in sequenced genomes, resulting in more than 8000 publications (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2010; Pingoud et al. 2014). Most Type II REases shared little amino acid sequence similarity, with the exception of, for example, EcoRI and RsrI, an early example of “neutral drift”: EcoRI and RsrI (recognition site G↓AATTC) are identical in places with 50% overall identity (Aiken et al. 1986; Stephenson et al. 1989), allowing the construction of active hybrids (Chuluunbaatar et al. 2007). Also cases of mosaicism occur—for example, EcoRI, MunI (C↓AATTG), and MluCI (↓AATT) (Pingoud et al. 2014). In this large family, “compelling examples” (Pingoud et al. 2014) could be found of convergent (e.g., HaeIII [GG↓CC] and BsuRI [GG↓CC] [Wilson and Murray 1991]) and of divergent evolution (e.g., Bsu36I [CC↓TNAGG], BlpI [GC↓TNAGC], Bpu10I [CCTNAGC], and BbvCI [CCTCAGC] [Heiter et al. 2005]). In addition to the divalent cations Mg2+ and Mn2+ , discussed earlier, some Type II REases use Zn2+ (BslI [CCNNNNN↓NNGG], PacI [TTAAT↓TAA] [Vanamee et al. 2003; Shen et al. 2010; Horton 2015]) and Co2+, Ni2+, and Cu2+ (Pingoud et al. 2014). In the case of the well-known 8-bp cutter NotI (GC↓GGCCGC), the enzyme is dependent on Fe2+, but this Fe2+ is incorporated in a structural Cys4 cluster (Lambert et al. 2008; Pingoud et al. 2014). Despite the high specificity of all enzymes, star activity on noncognate sites does occur, which can be partially inhibited by, for example, spermidine, hydrostatic pressure, mutation, or lowering enzyme concentrations (Pingoud et al. 2014).

The number of crystal structures rose from 16 in 2004 (Chapter 7; summarized in Horton et al. 2004) to 35 by 2014 (Pingoud et al. 2014) and to more than 50 new “de novo” (i.e., the first structure of a particular enzyme) enzyme structures in 2017 (Horton 2015). Figure 1A shows the REase structures in the Protein Data Base (PDB) by February 2, 2018 (http://rebase.neb.com/cgi-bin/cryyearbar 2017). Figure 1B shows a graph displaying the difference between the total number of REase structures and the de novo structures in the PDB (due to follow-up structures with ligands and/or mutations of a particular enzyme that are also deposited in the PDB by April 2017 [Horton 2015]). Most of these enzymes carry the PD fold, but, in addition, structures of REases with PLD, GIY-YIG, HNH, and half-pipe folds have been elucidated (see page 191). The REases that were the first of their (sub)type (adapted from Horton 2015) are indicated in bold in Appendix 1, which lists selected REases studied from approximately 2004 to 2017, including some earlier references, where appropriate. Although these structures would greatly aid modeling studies, even for well-characterized REases the properties that determine specificity and selectivity remain difficult to predict, because the enzyme is fixed in the crystal and changes conformation during the catalysis, and the additional interactions involved in this “transition” state are not evident in the crystal structure (Lanio et al. 2000; Pingoud et al. 2014).


FIGURE 1. (A) The REase structures in the Protein Data Base (PDB) by February 2, 2008 (http://rebase.neb.com/cgi-bin/cryyearbar 2017). (B) Comparison of the total number of REase structures deposited in the PDB with de novo REase structures. (Courtesy of Nancy Horton.)


The picture emerging from all these publications is that (similar to other protein families) the various domains involved in DNA binding, specific recognition, restriction, ATP binding and hydrolysis, and methylation have been fused or separated in all sorts of ways during the course of evolution. As a result, enzymes may have one or two catalytic sites and cleave DNA in one or two steps, with or without sliding and detaching from their DNA and with or without looping (Embleton et al. 2004; Halford and Marko 2004; Halford et al. 2004; Pingoud et al. 2014). Several studies addressed the question of the contribution of 1D and 3D movements of the REases along the DNA in order to find their recognition site (Gowers and Halford 2003; Gowers et al. 2005). Often multimers would bind to two sites rather than acquiring a second catalytic domain, which would be evolutionarily simple. One interesting study by the Bristol group concerns the reaction mechanism of seven REases that recognize GGCGCC and cut at different positions (Gowers et al. 2004). Using plasmids with one or two copies of this sequence revealed five distinct mechanisms, much larger than generally thought at the time (Gowers et al. 2004). Another example includes enzymes specific for the CCNGG sequences (Fig. 1 in Sasnauskas et al. 2015a, adapted in Fig. 3). Nearly 70% of all Type II REases belong to three families; the rest remain “mysteries”: They may be fringe members, or examples of new folds and DNA degradation mechanisms (see Pingoud et al. 2014 for further discussion).

Catalytic Domains of Type II REases

The PD···(D/E)XK Structural Fold

The PD···(D/E)XK fold (called the “PD” domain in this chapter) is present with variations in almost all Type II REases whose structures have been determined and is classified in the SCOP (Structural Classification of Proteins) database (http://scop.mrc-lmb.cam.ac.uk) as the REase-like fold (Niv et al. 2007; Steczkiewicz et al. 2012). The PD motif is often not easy to identify without information from a crystal structure, as the motif may vary and the amino acids involved are often in different locations along the polypeptide chain (Pingoud et al. 2014). Among 289 characterized Type II enzymes, 69% belonged to the PD superfamily (Orlowski and Bujnicki 2008) that includes the four nucleases mentioned in Chapter 7 (lambda exonuclease, MutH, VSR, and TnsA), but also, for example, RecB, Sulfolobus solfataricus Holliday-junction resolvase, and T7 endo I. The mechanism of catalysis continues to be the subject of study and debate. For example, the number of Mg2+ ions needed during catalysis remains uncertain (see Pingoud et al. 2014 for details and discussion).

The HNH and GIY-YIG Structural Domains

Other endonucleolytic motifs have been identified, including HNH and GIY-YIG motifs, found in homing endonucleases (HEases), Holliday-junction resolvases, exonucleases, nonspecific Serratia nuclease, and colicins (Friedhoff et al. 1999; Galburt et al. 1999; Jurica and Stoddard 1999; Pingoud et al. 2005a; Stoddard 2005; Kleinstiver et al. 2011, 2013). HNH examples are, for example, KpnI (GGTAC↓C) (Saravanan et al. 2004, 2007b; Vasu et al. 2013), Hpy99I (CGWCG↓), and PacI (TTAAT↓TAA), whereas two GIY-YIG REases, Eco29kI (CCGC↓GG) and Hpy188I (TCN↓GA), have been crystallized (Pertzev et al. 1997; Xu et al. 2000b; Bujnicki et al. 2001; Bujnicki 2004; Ibryashkina et al. 2007; Gasiunas et al. 2008; Kaminska et al. 2008; Orlowski and Bujnicki 2008; Mak et al. 2010; Mokrishcheva et al. 2011; Sokolowska et al. 2011). HNH motifs are often difficult to recognize because of the weak connection between the HNH and the residues that form the active site (Sokolowska et al. 2009). HNH enzymes use Mg2+ or Mn2+, but also other ions (Ni2+, Co2+, Zn2+, or Ca2+), sometimes with Cys4-Zn2+ binding elements (called ββα-metal fold), although many Cys4-Zn2+ motifs are not associated with catalytic sites but perform structural roles (Saravanan et al. 2004; Orlowski and Bujnicki 2008; Sokolowska et al. 2009; Shen et al. 2010; Pingoud et al. 2014).

Other Endonuclease Structural Domains

Thought unusual at the time, BfiI (ACTGGG [5/4]) was the first REase found that did not belong to the PD family: It carries the PLD nuclease domain and does not require Mg2+ for restriction (Sapranauskas et al. 2000). BfiI is a homodimer with a carboxy-terminal “B3-like” DNA-binding domain (DBD), which resembles B3 domains of some plant transcription factors. The catalytic site is formed at the interface of the two amino-terminal domains (similar to that of Nuc endonuclease from S. typhimurium), and although it binds to two sites at once, it cleaves only one strand at a time via an unusual covalent enzyme–DNA intermediate. BfiI appears to swivel the catalytic site by 180° and the same residues perform the same reaction on both DNA strands (Lagunavicius et al. 2003; Sasnauskas et al. 2003, 2007, 2010; Gražulis et al. 2005; Golovenko et al. 2014; Pingoud et al. 2014). Using the classification into 11 subtypes, this enzyme may be assigned to six or more of these subtypes (Marshall and Halford 2010; Pingoud et al. 2014). The enzyme AspCNI (GCCGC [9/5]) has a PLD-like domain and cleaves poorly at high concentrations (Heiter et al. 2015). PLD REases are not as rare as previously thought (Sapranauskas et al. 2000), as REBASE BLAST identified more than 40 other putatives (Pingoud et al. 2014). Some ATP-dependent enzymes (e.g., NgoAVII and CglI) contain a B3-like DNA recognition domain and a PLD catalytic domain (Tamulaitienė et al. 2014).

Type II REase Subtypes

This Type II section gives an overview and update with examples of the 11 subtypes, using two reviews (Roberts et al. 2003; Pingoud et al. 2014) as starting material. Figure 2 shows the subunit composition and cleavage mechanism of selected subtypes of Type II REases. Note that Pingoud et al. (2014) do not always follow the REBASE classification (http://rebase.neb.com/rebase/rebase.html). The reason for this is that different subtypes do not necessarily group with the different branches of the REase evolutionary tree, as exemplified by, for example, members of the EcoRII “CCGG family” studied by the Vilnius group (Table 1), which all cut at the same site (in contrast to the site studied by the Bristol group mentioned above [Gowers et al. 2004]): SsoII (↓CCNGG, Type IIP), EcoRII (↓CCWGG, Type IIE), and NgoMIV (G↓CCGGC, Type IIF) have similar DNA-binding sites and catalytic centers (Pingoud et al. 2002; Niv et al. 2007). Specificities for partly related, and even unrelated, sequences can nevertheless depend on the same structural framework: ↓CCNGG (SsoII), ↓CCWGG (PspGI/EcoRII), G↓CCGGC (NgoMIV), R↓CCGGY (Cfr10I), and MboI (↓GATC) (Pingoud et al. 2005c).


FIGURE 2. Subunit composition and cleavage mechanism of selected subtypes of Type II REases. Type IIP enzymes act mainly as homodimers (top) and cleave both DNA strands at once. Some act as dimers of dimers (homotetramers) instead and do the same. Still others act as monomers (bottom) and cleave the DNA strands separately, one after the other. Bright triangles represent catalytic sites. Type IIS enzymes generally bind as monomers but cleave as “transient” homodimers. Type IIB enzymes cleave on both sides of their bipartite recognition sequences. Their subunit/domain stoichiometry and polypeptide chain continuity varies. Three examples of primary forms are shown: BcgI, AloI, and HaeIV. These forms assemble in higher-order oligomers for cleavage. Type IIB enzymes display bilateral symmetry with respect to their methylation and cleavage positions. It is not clear whether they cleave to the left or to the right of the half-sequence bound. Type IIG enzymes (e.g., BcgI) might cleave upstream (left) of their bound recognition half-site. All other Type IIG enzymes (e.g., MmeI) cleave downstream from the site, often with the same geometry. These proteins have very similar amino acid sequences, however, suggesting that somehow the reactions are the same. Type IIT enzymes cleave within or close to asymmetric sequences. Composition varies; they have two different catalytic sites: top-strand-specific and bottom-strand-specific. In some, both subunits/domains interact with the recognition sequence (left cartoons). In others, only the larger subunit/domain recognizes the DNA. (Reprinted from Pingoud et al. 2014.)


Table 1. The CCGG family studied by the Vilnius group of Virgis Šikšnys

Enzyme Recognition site Type Structure PDB IDa Reference(s)





2FQZ, 2GB7

Bochtler et al. 2006






Zhou et al. 2004; Golovenko et al. 2009






Szczepanowski et al. 2008





Ms. in prep.

Manakova et al. 2015





Ms. in prep.

Manakova et al. 2015






Bozic et al. 1996





3V1Z, 3V20, 3V21, 1KNV

Gražulis et al. 2002; Manakova et al. 2012





4ABT (cited in Manakova et al. 2012)

Deibert et al. 2000






Tamulaitis et al. 2015






Manakova et al. 2015; Tamulaitienė et al. 2017





4C3G cryoEM
3MQY, 3N78, 3N7B, 3MQ6, 3DVO, 3DW9, 3DPG

Lyumkis et al. 2013; Little et al. 2011; Park et al. 2010; Dunten et al. 2008






Sasnauskas et al. 2015, 2017

Updated by Elena Manak, Gintautas Tamulaitis, and Giedrius Sasnauskas (November, 2017).

a PDB ID is the identification number in the Protein Data Bank.


Type IIA

Type IIA enzymes usually have separate R and S domains, recognize asymmetric sequences, and cleave within or at a defined position in or close to this site (Roberts et al. 2003; Pingoud et al. 2014). Many have two MTases each modifying one strand of the recognition sequence, rather than a single MTase. Others are combined R-M enzymes, some with separate MTases. Kinetic studies indicate that Type IIA enzymes transiently dimerize for cooperative cleavage. Examples are BbvCI, which uses two different catalytic sites from different subunits (Bellamy et al. 2005; Heiter et al. 2005), and Mva1269I (GAATGC [1/−1], IIA/IIS), which uses two sites from different domains within the same protein (Armalyte et al. 2005).

Type IIB

Type IIB enzymes cleave on both sides of a bipartite site releasing ∼34 bp (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003; Marshall et al. 2007; Pingoud et al. 2014). Some enzymes comprise a large single RMS polypeptide with features in common with Type I enzymes. Sometimes SAM acts as the cofactor for R as well as for S. Most IIB enzymes can only restrict when bound to two sites, preferably in cis, or in trans on concatenates. The first IIB R-M system, BcgI (cloned in 1994 [Kong et al. 1994] and extensively studied by the Halford group), concertedly cleaves two double-strand bonds [(10/12) CGANNNNNNTGC (12/10)] (Kong et al. 1993; Marshall and Halford 2010; Sasnauskas et al. 2010; Marshall et al. 2011; Smith et al. 2013a,b; Pingoud et al. 2014). It can be considered IIB/G/H/S (Kong and Smith 1998; Jurenaite-Urbanaviciene et al. 2007; Marshall and Halford 2010; Marshall et al. 2011; Smith et al. 2013a,b; Pingoud et al. 2014), like some other Type IIB enzymes (http://rebase.neb.com/rebase/rebase.html). BcgI comprises two subunits, RM and S, for cleavage and methylation with a stoichiometry of (RM)2S1, comparable to the Type I pentamer R2M2S1, but cutting at fixed positions (Kong et al. 1994; Kong and Smith 1997; Kong 1998). Other enzymes include, for example, BaeI [(10/15) ACNNNNGTAYC (12/7)], BsaXI [(9/12) ACNNNNCTCC (10/7)], and NgoAVIII [(12/14) GACNNNNNTGA (13/11)] (Sears et al. 1996; Marshall and Halford 2010). The BcgI-like enzymes modify both strands of their recognition sequences without additional MTases, and cleavage requires multiple (RM)2S1 complexes for double-strand cleavage on both sides of the recognition site (Marshall et al. 2007, 2011). The exact mechanism requires further investigation (see Marshall and Halford 2010 for discussion). Other IIB enzymes are, for example, AloI ([7/12] GAACNNNNNNTC [12/7]), PpiI ([7/12] GAACNNNNNCTC [13/8]), CjeI ([8/14] CCANNNNNNGT [15/9]), and TstI ([8/13] CACNNNNNNTCC [12/7]) (Jurenaite-Urbanaviciene et al. 2007; Smith et al. 2014). Domain-swapping experiments suggest that, like Type I enzymes, TRD swapping may also be used to generate hybrid specificities of Type II enzymes (Jurenaite-Urbanaviciene et al. 2007). Domain swapping and circular permutation of subdomains of BsaXI ([9/12] ACNNNNNCTCC [10/7]), or deletion, resulted in either active protein with altered specificity, poor protein yields, or inactive enzymes, which allowed mapping of critical amino acids for the interaction between the RM subunit and the TRD of the S subunit (Xu et al. 2015).

Type IIC

Type IIC are combined RM enzymes (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003; Pingoud et al. 2014). Most IIC bind as monomers to continuous and asymmetric sequences and cleave on one side of the recognition site at 1 turn, 1½ turn, or 2 turns away, whereas others cleave on both sides (i.e., IIB). Cleavage via transient dimerization is likely and is more efficient on DNA with multiple recognition sites or on addition of oligonucleotides. Examples are Eco57I (CTGAAG [16/14]), MmeI (TCCRAC [20/18]), and BpuSI (also called RM·BpuSI, GGGAC [10/14]), which are also considered to be IIC as well as IIE or IIG, respectively (see pages 202–203).

Type IIE

The prototype Type IIE enzymes are EcoRII and NaeI with separate domains for cleavage and allosteric activation (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003; Pingoud et al. 2014), as discussed in Chapter 7 and, for example, Reuter et al. (2004). The Type IIE enzymes prove to be diverse in structure (Fig. 3). Figure 3 shows a comparison of the structures of NaeI (Fig. 3A), EcoRII (Fig. 3B), and a new Type IIE enzyme from an unknown bacterium, named UbaLAI (Sasnauskas et al. 2017) by the CCGG group in Vilnius (Fig. 4).


FIGURE 3. Diversity of Type IIE REases. In all panels PD · · · (D/E)XK subunits are colored in different shades of green, monomeric MvaI-like PD · · · (D/E)XK domains are red, catabolite activator protein (CAP)-like domains are orange and light brown, and B3-like domains are blue. Yellow diamonds in the cartoon representations denote the catalytic center(s) present in each enzyme. (A) NaeI (GCC↓GGC) is a Type II homodimer that simultaneously binds two recognition sites. One is cleaved by the EcoRV-like dimer of the catalytic N domains (Endo domains), whereas the second one bound to the CAP DNA-binding motif in the carboxy-terminal domain (Topo domain) stimulates cleavage of the first site (PDB ID: 1IAW) (Embleton et al. 2001; Huai et al. 2001). (B) EcoRII (↓CCWGG) is a Type IIE homodimer capable of simultaneous binding of three recognition sites. One is cleaved by the PspGI-like dimer of the catalytic C domains, whereas two others, one per EcoRII-N effector domain, stimulate cleavage of the first site (PDB IDs: 3hqf and 3hqg) (Tamulaitis et al. 2006a,b; Golovenko et al. 2009). (C) UbaLAI (CC↓WGG) is a novel monomeric REase consisting of an MvaI-like catalytic domain (red) and an EcoRII-N-like effector domain (blue; PDB ID to be published). UbaLAI requires two recognition sites for optimal activity, and, like NaeI and EcoRII, uses one copy of a recognition site to stimulate cleavage of a second copy. UbaLAI-N acts as a handle that tethers the monomeric UbaLAI-C domain to the DNA, thereby helping UbaLAI-C to perform two sequential DNA nicking reactions on the second recognition site during a single DNA-binding event (Sasnauskas et al. 2017). The structure of the UbaLAI-C domain is a model built using Modeller (Webb and Sali 2016). (Portions reprinted from Sasnauskas et al. 2017, courtesy of Gintautas Tamulaitis.)




FIGURE 4. (A) CCGG group photo. From left to right: Inga Songailiene, Gintautas Tamulaitis, Elena Mankova, Saulius Gražulis, Virgis Šikšnys, Giedrė Tamulaitienė, Giedrius Sasnauskas, and Mindaugas Zaremba. (B) Graduate students and postdoctoral colleagues from 1977 through 2011 at Steve Halford’s retirement party (2011). His group, from left to right: Stuart Bellamy, Dave Scott, Rachel Smith, Kelly Sanders, Niall Gormley, Tim Nobbs, Mark Watson, Panos Soultanos, Geoff Baldwin, Steve Halford (in his DNA jumper), Darren Gowers, Mark Szczelkun, Neil Stanford, Jacqui Marshall, Barry Vipond, Yana Kovacheva, Katie Wood, Tony Maxwell, Isobel Kingston, John Taylor, Sophie Castell, Michelle Embleton, Christian Vermote, Alistair Jacklin, Alison Ackroyd, Fiona Preece, Susan Retter, Lucy Catto, Shelley Williams. Christian Parker and Denzil Bilcock were at the party but not in the photo. (Absent: Pete Luke, Paul Bennett, Samantha Hall, Lois Wenztell, Symon Erskine, Mark Oram, Abigail Bath, David Rusling, and Sumita Ganguly). (C) Werner Arber, Noreen Murray, and D.N. Rao at the 6th NEB meeting in Bremen (2010). (D) Participants of the CSHL meeting in 2013: History of Restriction Enzymes. (D, Courtesy Cold Spring Harbor Laboratory Archives.)


Nearly a dozen papers were published on EcoRII in collaborations between experts in the field of crystallography, AFM, and single-molecule studies (Zhou et al. 2002, 2003, 2004; Kruger and Reuter 2005; Tamulaitis et al. 2006a,b, 2008; Shlyakhtenko et al. 2007; Gilmore et al. 2009; Golovenko et al. 2009; Szczepek et al. 2009). A high-resolution crystal structure of the dimeric EcoRII was published in 2004, which revealed a hinge loop connecting the catalytic and allosteric activation domains (Zhou et al. 2002, 2003, 2004). The catalytic domain (comprised of two copies of the carboxy-terminal domain) had the PD fold, whereas the two amino-terminal regulatory/effector domains had a different DNA recognition fold with a large cleft. This fold was novel at the time, but is in fact the B3-like fold mentioned above and present in BfiI and NgoAVII (more specifically, it is a SCOP double-split β-barrel fold, of the DNA-binding pseudobarrel domain superfamily). The structure explained the mechanism of autoinhibition/activation of EcoRII, which was novel in REases, but similar to that described for various transcription factors (Zhou et al. 2004). This structure contained three possible DNA-binding regions, and in line with this, only a plasmid with three recognition sites yielded linear DNA during a single turnover, whereas the same plasmid with only one or two sites did not (Tamulaitis et al. 2006b). AFM studies showed two-loop structures with an EcoRII dimer at the core of the three-site synaptosome (Shlyakhtenko et al. 2007). A variant of AFM (called high-speed AFM) allowed single-molecule imaging of the EcoRII protein (Gilmore et al. 2009). In this way, binding, translocation, and dissociation could be monitored, and they indicated that EcoRII can translocate along the DNA to search for a second binding site, after finding the first site. Dissociation from the loop structure resulted in either two monomers bound to the two sites or one dimer to one site (Gilmore et al. 2009). Further experiments showed the very different ways in which the enzyme interacted with the effector and substrate DNA. The carboxy-terminal domain flipped the central T:A base pair out, and interacted with the CC:GG half-sites, whereas the effector domain bound asymmetrically without pushing out the T:A base pair (Golovenko et al. 2009). Interestingly, the 7-bp cutter PfoI (T↓CCNGGA) also uses base flipping as part of its DNA recognition mechanism. But in this case the extrahelical bases are captured in binding pockets that are quite different from those in the related structurally characterized enzymes Ecl18kI, PspGI, and EcoRII-C (Manakova et al. 2015). PspGI (↓CCWGG) and Ecl18kI/SsoII (↓CCNGG) flip the central A and T (W) bases out of the helix, compressing the recognition sequence in effect to just CC-GG (Bochtler et al. 2006; Tamulaitis et al. 2007; Szczepanowski et al. 2008). Repression of catalysis by the amino-terminal domain was further analyzed by site-directed mutagenesis and addition of soluble peptides in trans, which revealed the structural elements essential for autoinhibition (Szczepek et al. 2009). The crystal structure of MvaI identified MvaI as a monomer that recognizes its pseudosymmetric target sequence (CC↓WGG) asymmetrically (Kaus-Drobek et al. 2007). The enzyme has two lobes: a catalytic one that contacts the bases from the minor groove side, and the other that contacts those from the major groove. MvaI resembles BcnI (CC↓SGG), and also MutH, which nicks DNA rather than cutting both strands. The reason for this is clear: MvaI, BcnI, and MutH have a single catalytic site and just nick their substrates upon binding. Because the substrates of MvaI and BcnI are symmetric, these two enzymes can then bind in the opposite orientation and nick the other strand resulting in double-strand cleavage. The substrate of MutH (hemimethylated GATC) is asymmetric, and so MutH can only bind in one orientation and thus cannot cut the second strand. Different responses to slight substrate asymmetries, which could be altered by protein engineering, determine whether these monomeric REases make single-strand nicks or double-strand breaks (Sokolowska et al. 2007a; see Kaus-Drobek et al. 2007 for further details). For some other studies on the EcoRII and CCGG family, see Kubareva et al. (1992, 2000); Šikšnys et al. (2004); Pingoud et al. (2005b); Sud'ina et al. (2005); Zaremba et al. (2006); Fedotova et al. (2009); and Abrosimova et al. (2013), and the Type IIF section.

Type IIF

Type IIF bind two recognition sites and cleave all four strands at once as pairs of back-to-back dimers (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003; Šikšnys et al. 2004; Zaremba et al. 2005; Pingoud et al. 2014). The structures of Cfr10I (R↓CCGGY), Bse634I (R↓CCGGY), and NgoMIV (G↓CCGGY) (Chapter 7) and the observed transient tetramerization of Ecl18kI (↓CCNGG) indicated that the boundaries between IIE and IIF are not strict (Šikšnys et al. 2004; Zaremba et al. 2005, 2010; Pingoud et al. 2014). Work on Bse634I continued in Vilnius (Zaremba et al. 2005, 2006, 2012; Manakova et al. 2012). The tetramer could be converted to a dimeric enzyme by mutation, and kinetic studies indicated two types of communication signals via the dimer–dimer interface in the tetramer: an inhibitory and an activating signal, which somehow control the catalytic and regulatory properties of the Bse634I and mutant proteins (Zaremba et al. 2005, 2006). Contrary to expectation, dimeric enzymes have the same fidelity toward their recognition site as the tetramer, because they act concertedly at two sites, thus providing a safety catch against cleavage at a single unmodified site (Zaremba et al. 2012). The structures of the SfiI (GGCCNNNN↓NGGCC) tetramer in complex with cognate DNA provided details on how SfiI recognized and cleaved its target DNA sites (Viadiu et al. 2003; Vanamee et al. 2005). Some other Type IIF enzymes are PluTI (GGCGC↓C) (Khan et al. 2010; Pingoud et al. 2014) and SgrAI (CR↓CCGGYG). SgrAI (Laue et al. 1990; Tautz et al. 1990; Capoluongo et al. 2000; Bitinaite and Schildkraut 2002; Daniels et al. 2003; Hingorani-Varma and Bitinaite 2003; Wood et al. 2005; Dunten et al. 2008, 2009; Park et al. 2010; Little et al. 2011; Lyumkis et al. 2013; Ma et al. 2013b; Horton 2015) is also a member of the CCGG family and preferentially cleaves concertedly at two sites. Interestingly, SgrAI assembles into homotetramers, and then other molecules join to generate helical structures with one DNA-bound homodimer after another. Adjacent homodimers are not back-to-back (i.e., 180°), but at ∼90°, and four homodimers form almost one turn of a left-hand spiral of 18 homodimers or perhaps even more. These SgrAI filaments have some star activity, probably as a result of asymmetry generated by the multimerization process (Fig. 5). Another interesting enzyme is the Type IIF homotetrameric GIY-YIG Cfr42I enzyme that is rather similar to the monomeric/dimeric Eco29kI enzyme, which supports the notion of convergent evolution of REases belonging to unrelated nuclease families toward homotetramers with a “safety catch” (Gasiunas et al. 2008).


FIGURE 5. CryoEM structure of SgrAI bound to DNA. Each SgrAI dimer is colored uniquely. This picture was made using PDB coordinates and surface rendering. (The original figure in the paper by Lyumkis et al. [2013] was the actual cryoEM envelope, carved up into different subunits.) (Adapted from Lyumkis et al. 2013, with permission from Elsevier.)


Type IIG

Type IIG are Type I-like combined RM systems, with an amino-terminal PD domain, and a γ-class MTase domain in a single protein (http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003; Niv et al. 2007; Loenen et al. 2014a; Pingoud et al. 2014). The S specificity subunit may be present as a separate subunit or as a domain attached to the carboxyl terminus of RM. IIG are stimulated by SAM or are SAM-dependent. This definition includes most IIB and IIC REases (Loenen et al. 2014a; Pingoud et al. 2014). Only one catalytic site is present in these domains, and cleavage of duplex DNA is thought to occur by the transient dimerization of neighboring enzyme molecules. Examples are Eco57I, MmeI, and BpuSI.

Eco57I was the first member of a new class of monomeric enzymes, initially called Type IV (like BspLU11III [GGGAC (10/14)] from Bacillus sp. LU11 [Lepikhov et al. 2001]), but renamed Type IIG enzymes (Janulaitis et al. 1992a,b), although it can also be considered IIE as it is accompanied by one additional MTase. It is a large RMS protein, cuts one and one-half turns away, and is useful for engineering (Janulaitis et al. 1992b; Rimseliene et al. 2003; Pingoud et al. 2014). It methylates the top strand of its asymmetric recognition site (CTGAAG [16/14]), whereas a separate MTase, M·Eco57I, methylates the adenine in the bottom strand (Janulaitis et al. 1992a). M·Eco57I can also methylate the same adenine in the top strand as Eco57I (Janulaitis et al. 1992a; Loenen et al. 2014a). Some other monomeric Type IIG have accompanying MTases that methylate m5C (BpuSI) or m4C (BseRI, GAGGAG [10/8]) (Loenen et al. 2014a).

MmeI is IIE/IIG/IIC and cuts two turns away (TCCRAC [20/18]). It was the first Type IIG enzyme to be purified and belongs to a large family of closely related enzymes with many different specificities (Boyd et al. 1986; Morgan et al. 2008; Loenen et al. 2014a). Based on in vitro studies, MmeI has also been named Type IIL, for lone-strand DNA modification (Morgan et al. 2009). As the enzyme does not require a head-to-head approach in vitro, there is disagreement on its mode of action: Does in vivo MmeI act on two inverted (head-to-head) recognition sequences like Type III enzymes (Dryden et al. 2011; Schwarz et al. 2011; Loenen et al. 2014a), using sliding or DNA looping between adjacent sites (Halford et al. 1999; Halford 2001), or perhaps bind DNA as a monomer and then form dimers or multimers before methylation or cleavage, similar to Type I enzymes (Loenen et al. 2014a)? But why would MmeI slide along the DNA, as the adenine that will eventually be methylated is likely to be flipped into the binding pocket on specific site recognition (Cooper et al. 2017; Bogdanove et al. 2018)? MmeI requires at least two bound specificity sites for cutting. Unlike FokI, adding excess enzyme in solution, without a specific site, does not stimulate cutting. Richard (Rick) Morgan suggests a model that includes the requirement for enzyme bound at two (or possibly four) sites to come together for cutting (Cooper et al. 2017; Bogdanove et al. 2018). As methylation is effective at single sites, this process does not require dimerization of the enzyme.

MmeI has been well characterized (Boyd et al. 1986; Tucholski et al. 1995, 1998; Nakonieczna et al. 2007, 2009; Morgan et al. 2008, 2009; Callahan et al. 2011, 2016), and rational engineering based on sequence alignments and mutational analysis led to altered specificities that could be predicted (Morgan and Luyten 2009; Morgan et al. 2009). Changes in the S domain alter the recognition site for both R and M (like Type I enzymes), and hence members of the MmeI family have been able to diverge widely in the course of evolution (Morgan et al. 2008, 2009; Morgan and Luyten 2009). Certain different pairs of amino acids are specific for alternative base pairs in the recognition sequence: for example, Glu806···Arg808 in MmeI (TCCRAC) specifies the third C, whereas Lys806···Asp808 specifies G at that position (TCCRAG). The crystal structure has been solved (Callahan et al. 2011, 2016). Together with the structure of MmeI in complex with DNA (and SAM-analog sinefungin) (Callahan et al. 2011), these data on the MmeI family allowed the construction of REases with novel predictable DNA recognition and restriction properties, which had “long been a goal of modern biology” (Callahan et al. 2016) and previously denied for EcoRI and EcoRV. With this in mind, Geoff Wilson pondered whether one could predict and design new specificities of other enzymes (e.g., Type I HsdS) or even predict those of putative HsdS subunits in REBASE based on sequence data alone (Loenen et al. 2014a). The answer to this is yes, as in recent times Rick Morgan has predicted and made specificity changes in Type I HsdS systems (R Morgan, in prep.).

BpuSI (GGGAC [10/14]) is IIG or IIS and has two MTases (Shen et al. 2011; Sarrade-Loucheur et al. 2013; Pingoud et al. 2014). The crystal structure indicates that it resembles the well-characterized carboxy-terminal cleavage domain of FokI (GGATG [9/13]) and produces 5′ sticky ends (Wah et al. 1997, 1998; Shen et al. 2011). This is unusual because most Type IIG enzymes create 3′ overhangs, indicating that their catalytic sites cleave across the minor groove of DNA rather than across the major groove. BpuSI was crystallized without DNA and evidently must undergo significant structural rearrangements to bind DNA and carry out catalysis (Shen et al. 2011). This means that the carboxy-terminal S domain must rotate with respect to the R and M domains and reorganize in order to bind DNA (also seen with other REases) (Shen et al. 2011; Sarrade-Loucheur et al. 2013; Pingoud et al. 2014).

Type IIH

Type IIH are hybrid Type IIP-like (e.g., GACNNN↓NNGTC) REases with an m6A MTase (Pingoud et al. 2014). M·AhdI is a tetramer of M and S subunits, suggestive of the ancestral form of Type I MTases. As such, they have been called “Type 1½” RM systems, and a “missing link” between Type I and II IIH enzymes, but as they have proved rather common, this distinction may no longer be relevant (Marks et al. 2003; Pingoud et al. 2014).

Type IIM

Type IIM enzymes recognize methylated DNA. The well-known DpnI (Lacks and Greenberg 1975; Pingoud et al. 2014) cuts Gm6A↓TC as a monomer, one strand at a time. The complementary specificities of DpnI and DpnII have been useful for site-directed mutagenesis, as DpnII cuts unmethylated ↓GATC) sites (Lacks and Greenberg 1977). DpnI has the amino-terminal PD domain and a carboxy-terminal winged-helix (wH) allosteric activator domain. Both domains bind methylated DNA with sequence specificity (Lacks and Greenberg 1975; Siwek et al. 2012; Mierzejewska et al. 2014; Pingoud et al. 2014). A new addition to this subtype is the BisI (Gm5CNGC) enzyme and its relatives (Xu et al. 2016). Some enzymes that recognize methylated DNA and are classified as Type IV enzymes would also fit into the IIM subtype, if they cut at specific sites (see the section Part D: Type IV Enzymes).

Type IIP

The best-known orthodox Type IIP palindromic REases are, of course, EcoRI and EcoRV. Type IIP cleave symmetric recognition sequences and have a single domain in which recognition and cleavage functions are integrated (Pingoud et al. 2014). They tend to have a single cognate MTase, although some have two MTases. The IIP REases can be monomeric but most are homodimers or homotetramers. Multimers usually cleave both DNA strands in one binding event, whereas monomers need to cleave sequentially first one strand, then the other, because of the opposite 5′ to 3′ polarity of the DNA strands (Gowers et al. 2004; Pingoud et al. 2014). In line with this prediction, the BcnI (CC↓SGG) monomer first localizes the recognition site by 1D and 3D diffusion, and nicks one DNA strand; it then diffuses from the nicked site, turns 180°, diffuses back, and cleaves the other (unnicked) strand (Sokolowska et al. 2007b; Kostiuk et al. 2011, 2015, 2017; Sasnauskas et al. 2011).

Single-molecule studies with EcoRV provided evidence for fast 3D sliding and jumping of EcoRV on nonspecific DNA following a slow initial 1D diffusion (Bonnet et al. 2008). Using optical tweezers with fluorescence tracking, it became clear that the enzyme stays in close contact with the DNA during sliding (Bonnet et al. 2008; Biebricher et al. 2009). Aneel Aggarwal and coworkers analyzed the structure of BstYI, a thermophilic REase that cleaves 5′-Pu/GATCPy-3′, a degenerate version of the BamHI (G↓GATCC) and BglII (A↓GATCT) recognition sites. A comparison of free BstYI with BamHI and BglII revealed a strong structural likeness between these enzymes, but in addition, BstYI also contained an extra “arm” domain possibly related to the thermostability of BstYI (Townson et al. 2004). The cocrystal structure with DNA revealed a mechanism of degenerate DNA recognition, which will stimulate thoughts about the possibilities and limitations in altering specificities of closely related REases (Townson et al. 2005). Interestingly, an isoschizomer of BamHI, OkrAI (G↓GATCC), is a much smaller version of BamHI, which recognizes the DNA in a similar manner, “a rare opportunity to compare two REases that work on exactly the same DNA substrate” (Vanamee et al. 2011).

The group of Ichizo Kobayashi studied regulation of the EcoRI operon (see page 214) (Liu and Kobayashi 2007; Liu et al. 2007), whereas the group of Linda Jen-Jacobsen in Pittsburgh continued studies on EcoRI with respect to the mechanism of coupling between DNA recognition specificity and catalysis (Kurpiewski et al. 2004), the inhibition by Cu2+ ions of Mg2+-catalyzed DNA cleavage (Ji et al. 2014), and the relaxed specificity and structure of promiscuous mutants of EcoRI that cleave at EcoRI* sites (Sapienza et al. 2005, 2007, 2014). As EcoRI* sites are not protected by M·EcoRI, promiscuous mutants are deleterious to the host. They encountered “unanticipated and counterintuitive observations” that three EcoRI mutants with such relaxed specificity in vivo nevertheless bound more tightly than wild-type EcoRI to the cognate site (GAATTC) in vitro and even preferred that site to EcoRI* sites (Sapienza et al. 2005). How could this be? Using structural and thermodynamic analyses, this question was addressed further (Sapienza et al. 2007, 2014). The crystal structure of the promiscuous mutant A138T homodimer in complex with the cognate site was nearly identical to that of the wild-type complex, except that the threonine138 side chains interacted with bases 5′ to the GAATTC site. This would enable A138T to form complexes with EcoRI* sites that structurally resembled the specific wild-type complex with GAATTC (Sapienza et al. 2007). The importance of these flanking bases was also confirmed by the finding that AAATTC sites with an adjacent 5′-purine-pyrimidine (5′-RY) were cleaved much faster (up to 170× faster!). This and further thermodynamic analyses supported the notion that specificity relied on a series of cooperative events that were “uniquely associated with specific recognition” (Sapienza et al. 2014).

SwaI (ATTT↓AAAT) (Dedkov and Degtyarev 1998) and PacI (TTAAT↓ TAA) both recognize AT-rich DNA sequences, but their protein structures are completely different (Shen et al. 2010). In the case of PacI, the normal base-pairing is completely disrupted in the bound structure: “two bases on each strand are unpaired, four are engaged in noncanonical A:A and T:T base pairs, and the remaining two bases are matched with new Watson–Crick partners.” This suggests that PacI is an unusual REase that recognizes its target site via contacts not visualized in the DNA-bound cocrystal structure (Shen et al. 2010). Whereas PacI is elongated and follows the track of the DNA helix (Shen et al. 2010), SwaI is flattened and horseshoe-shaped (Shen et al. 2015). SwaI has an open conformation with the DNA-binding surface accessible from the outside. When bound to DNA, the enzyme is closed and completely encircles the DNA. Like PacI, SwaI profoundly distorts the DNA on binding, but in a different way (Shen et al. 2010). In SwaI, the central T:A and A:T bases are unpaired, and the two adenines switch positions and stack on each other in the reverse order. This is accompanied by a ∼50° bend in the helix and severe compression of the major groove, much as is seen in EcoRV (GAT↓ATC) (Winkler et al. 1993). The authors had no idea “how this surprising reversal in base order takes place” (Shen et al. 2015, 2017). Like EcoP15I, which has been used to count Huntington's disease CAG repeats, TseI (G↓CWGC) is also useful for the analysis of A:A and T:T mismatches in CAG and CTG repeats in this dreadful disease (Moncke-Buchner et al. 2002; Ma et al. 2013a).

Type IIS

Type IIS cut at a fixed distance from the recognition site (http://rebase.neb.com/rebase/rebase.html; Szybalski et al. 1991; Roberts et al. 2003; Welsh et al. 2004; Niv et al. 2007; Pingoud et al. 2014). The recognition and cleavage domains are separated by a linker region allowing fusion of the cleavage domain to other recognition modules, thus generating novel specificities. They usually have two MTases, each methylating one of the two strands (m6A or m5C). The name Type IIS (for “shifted”) enzymes was first coined by Wacław Szybalski and coworkers at the University of Wisconsin, who devised “ingenious applications” in the 1980s (Hasan et al. 1986; Kim et al. 1988; Pingoud et al. 2014). All Type IIB, IIC, and IIG REases can be considered IIS (cut outside their recognition sites), but all share the integral γ-class MTase as described above.

FokI (GGATG [9/13]) is one of the earliest, most-studied, Type IIS enzymes with a DNA recognition domain and a separate cleavage domain, which has been used extensively for genome engineering (Sugisaki and Kanazawa 1981; Nwankwo and Wilson 1987; Mandecki and Bolling 1988; Kaczorowski et al. 1989; Kita et al. 1989a,b; Landry et al. 1989; Looney et al. 1989; Sugisaki et al. 1989; Goszczynski and McGhee 1991; Szybalski et al. 1991; Li et al. 1993; Skowron et al. 1993; Kim et al. 1994; Waugh and Sauer 1994; Yonezawa and Sugiura 1994; Kim et al. 1996a, 1997, 1998; Skowron et al. 1996; Hirsch et al. 1997; Wah et al. 1997, 1998; Bitinaite et al. 1998; Leismann et al. 1998; Chandrasegaran and Smith 1999; Friedrich et al. 2000; Vanamee et al. 2001; Catto et al. 2006; Laurens et al. 2012; Pernstich and Halford 2012; Rusling et al. 2012; Guilinger et al. 2014b; Mino et al. 2014; Pingoud et al. 2014). The accompanying two MTases are fused into a single protein. Single-particle EM studies provided new insights into the activation mechanism of FokI and avoidance of aspecific cleavage (Vanamee et al. 2007). FokI crystals show the catalytic domain to be hidden behind the DNA recognition domain, which will require a substantial conformational change before cutting can take place after dimerization of two catalytic domains (Bitinaite et al. 1998; Pingoud et al. 2014). Details on the cleavage mechanism still need to be sorted out, but the need for two enzyme molecules for catalysis appears to be quite common (Embleton et al. 2001; Welsh et al. 2004; Catto et al. 2006, 2008; Gemmen et al. 2006; Sanders et al. 2009; Pingoud et al. 2014). Other enzymes also have a FokI-like domain—for example, StsI (GGATG [10/14]) (Kita et al. 1992a,b) and Mva1269I (GAATGC [1/−1]) (Armalyte et al. 2005).

Type IIT

Type IIT enzymes are heterodimers with two subunits (e.g., Bpu10I, BbvCI) or heterotetramers (e.g., BslI [CCNNNNN↓NNGG] [http://rebase.neb.com/rebase/rebase.html; Roberts et al. 2003; Pingoud et al. 2014]). IIT use two different catalytic sites for cleavage. Some enzymes are single chain (e.g., Mva1269I uses an EcoRI-like domain and a FokI-like domain) (Armalyte et al. 2005; Pingoud et al. 2014). Type IIT systems usually have two MTases (either separate proteins or fused as a single protein) that each modify one strand. They are useful after conversion to strand-specific nicking enzymes (see Chan et al. 2011 for a review, and page 209)—for example, BbvCI has two catalytic sites from different subunits, each cleaving its own strand (Bellamy et al. 2005; Heiter et al. 2005).

The “Half-Pipe”

PabI of Pyrococcus abyssi (GTA↓C) was thought to be a bona fide REase, as it was found near a MTase gene (Pingoud et al. 2014). However, it is a homodimeric DNA glycosylase with a unique structure and flips all four purines out of the helix, leaving the pyrimidines as intrahelical “orphans” (Ishikawa et al. 2005; Watanabe et al. 2006; Miyazono et al. 2007, 2014; Pingoud et al. 2014; Kojima and Kobayashi 2015). Close isoschizomers of PabI are ubiquitous in Helicobacter pylori strains. Whether PabI is involved in genetic rearrangements remains to be investigated (Pingoud et al. 2014).

Type II Enzymes as Tools for Gene Targeting


As briefly discussed in Chapter 7, Srinivasan Chandrasegaran at Johns Hopkins School of Medicine pioneered what is now termed gene targeting by fusing the REase endonuclease domain of FokI to a zinc-finger protein to create a novel engineered zinc-finger nuclease (ZFN) (Li and Chandrasegaran 1993). ZFNs usually contain three to six Zn fingers (each ∼30 aa) with a ββα fold that binds one Zn2+ via 2Cys + 2His (Miller et al. 1985; Klug 2010a,b). Each finger recognizes a 3-bp target sequence via four amino acids that project from the α-helix into the major DNA groove (Durai et al. 2005; Wu et al. 2007). Two different three-finger ZFNs will recognize an 18-bp sequence, sufficient to be unique in the human genome. Such constructs have been used with considerable success, although they tend to be less specific than expected (Urnov et al. 2005, 2010; Carroll 2011a,b; Gabriel et al. 2011; Handel and Cathomen 2011; Pattanayak et al. 2011; Perez-Pinera et al. 2012a; see also Carroll 2014; Carroll and Beumer 2014; Hendel et al. 2015).

ZNF-based engineered highly specific REases can be used for gene targeting by introducing a dsDNA break into a complex genome and thereby stimulating homologous recombination (Yanik et al. 2013; Carroll 2014). With the exception of engineered homing endonucleases (“meganucleases”) with integrated DBD and catalytic domains (Galetto et al. 2009), the other engineered nucleases have distinct DBD and catalytic domains. ZFNs usually have the cleavage domain of FokI (Li et al. 1992, 1993; Li and Chandrasegaran 1993; Waugh and Sauer 1993; Kim et al. 1994, 1996b; Chandrasegaran and Smith 1999; Bibikova et al. 2002; Urnov et al. 2005; Miller et al. 2007; Szczepek et al. 2007; Mino et al. 2009; Mori et al. 2009; Carroll 2011a,b; Gabriel et al. 2011; Handel and Cathomen 2011; Pattanayak et al. 2011; Ramalingam et al. 2011, 2013; Handel et al. 2012; Bhakta et al. 2013; Pingoud et al. 2014), but PvuII (CAG↓CTG) has also been used for this purpose (Schierling et al. 2012).

Nonspecific (“off-target”) cleavage can be reduced by mutations in the dimerization surface (Miller et al. 2007; Szczepek et al. 2007), but according to Steve Halford the off-target problem might well be due to dimerization between a specific and a nonspecific ZFN (Halford et al. 2011).

The fusion construct with PvuII (CAG↓CTG) was slightly better than that with FokI (Schierling et al. 2012; Pingoud et al. 2014), but fusions with the transcription activator-like effector (TALE) proteins, where one module recognizes one base (Fig. 6A,B; Pingoud et al. 2014), were an improvement on engineered nuclease contructs. These proteins contain many (up to 35) nearly identical repeats of ∼34 aa. The 13th residue in each repeat recognizes the DNA base. The repeats form a superhelix around the DNA, following the track of the major groove for several turns. The individual repeats are left-handed two-helix bundles that, one after the other, juxtapose the 13th amino acid of each repeat to adjacent bases in one strand of the DNA (Deng et al. 2012; Mak et al. 2012, 2013).


FIGURE 6. Engineering of Type II REases as tools for gene targeting. (A) Engineered highly specific endonucleases that can be used for gene targeting by introducing a double-strand break into a complex genome and thereby stimulating homologous recombination (Yanik et al. 2013). With the exception of engineered homing endonucleases (“meganucleases”) in which the function of DNA binding and DNA cleavage is present in the same polypeptide chain (Galetto et al. 2009), the other engineered nucleases consist of separate DNA-binding (green) and DNA-cleavage (blue) modules. ZFNs and TALENs usually have the nonspecific cleavage domain of the restriction endonuclease FokI as DNA-cleavage module, but the restriction endonuclease PvuII can also be used for this purpose (Schierling et al. 2012; Yanik et al. 2013). PvuII has also been employed in triple-helix-forming oligonucleotide (TFO)-linked nucleases (Eisenschmidt et al. 2005) and in protein fusions (with catalytically inactive I-SceI) (Fonfara et al. 2012) as DNA-cleavage module. ZFNs, TALENs, and TFO-linked nucleases are programmable, as are the RNA-mediated nucleases (Jinek et al. 2012) modified after Pingoud and Wende (2011). (Reprinted from Yanik et al. 2013.) (B) TALE-PvuII fusion proteins. (a) Scheme of the architecture of TALE–PvuII fusion proteins. (Left) wtPvuII, a homodimer in which the DNA-binding module of a TALE protein is fused via a linker of defined length. (Right) scPvuII, a monomeric nuclease in which the DNA-binding module of a TALE protein is fused via a linker of defined length. (b) Model of a TALE–wtPvuII fusion protein. The fusion protein is a dimer of identical subunits, each composed of a PvuII subunit and a TALE protein. This model was constructed by aligning the structures of the individual proteins PDB 1pvi (Cheng et al. 1994) and PDB 3ugm (Mak et al. 2012) on a DNA composed of the PvuII recognition site and two TALE target sites upstream of and downstream from the PvuII recognition site, separated by 6 bp. The carboxyl termini of the PvuII subunits and the amino termini of the TALE protein are separated by ∼3 nm. This distance must be covered by a peptide linker of suitable length. The image was generated with PyMol. (Reprinted from Yanik et al. 2013.)


In the case of PvuII, the DBD of a TALE protein is fused via a linker of defined length to the homodimeric REase (Fig. 6B). Wild-type PvuII (wtPvuII) is shown on the left in Figure 6Ba, and a variant of PvuII as a TALE-linked monomer (scPvuII) on the right in Figure 6Ba. A model of a TALE-PvuII fusion protein was constructed using the structures PDB 1pvi (Cheng et al. 1994) and PDB 3ugm (Mak et al. 2012) on a DNA composed of the PvuII recognition site and two TALE target sites upstream of and downstream from the PvuII recognition site, separated by 6 bp (Fig. 6Bb). The fusion protein is a dimer of identical subunits, each composed of a PvuII subunit and a TALE protein.

TALE-based nucleases (engineered TALE nucleases [TALENs]), based on FokI and PvuII, proved much better tools for genome manipulations than did ZFNs (Miller et al. 2011; Perez-Pinera et al. 2012b; Joung and Sander 2013; Yanik et al. 2013), but they also have some off-target activity. Profiling of 30 different unique TALENs for the ability of potential off-target cleavage using in vitro selection and high-throughput sequencing resulted in 76 predicted off-target substrates in the human genome, 16 of which were accessible and modified by TALENs in human cells (Guilinger et al. 2014a). This analysis allowed the construction of a TALEN variant with ∼10× lower off-target activity in human cells (Guilinger et al. 2014a).

In 2014, FokI was fused to Cas9, which cleaves dsDNA at a sequence programmed by a short single-stranded guide RNA (Guilinger et al. 2014b). Unfortunately, genome editing by Cas9 can also result in off-target DNA recognition. Fusions of catalytically inactive Cas9 and FokI nuclease (fCas9) modified target DNA sites with >140-fold higher specificity than wild-type Cas9 and with an efficiency similar to that of paired Cas9 “nickases” that cleave only one DNA strand each. The specificity of fCas9 was at least fourfold higher than that of paired nickases and may be a good strategy for highly specific genome-wide editing (Guilinger et al. 2014b). Use of very long (up to 10 kb) homologous flanking arms for break repair also improves targeting (Baker et al. 2017).

Nickases (Nicking Enzymes)

Another approach to gene targeting has been the use of nickases. Precise incisions in genomic DNA are required for (faithful) homologous recombination, but dsDNA breaks would activate the error-prone, nonhomologous end-joining (NHEJ) pathway. This led to the idea that a nicking domain that would cut only one DNA strand might work better than a cleavage domain, and could be used for DNA repair studies and other DNA manipulations (e.g., terminal labeling, genome mapping, and DNA amplification) (Chan et al. 2011; Xiao et al. 2011). The large subunits of some heterodimeric REases (e.g., some Type IIT and IIS) can function as nicking enzymes when separated from their normal partner (Higgins et al. 2001; Heiter et al. 2005; Yunusova et al. 2006; Xu et al. 2007), whereas dimeric enzymes can be mutated to generate a single catalytic site (Stahl et al. 1996; Wende et al. 1996; Morgan et al. 2000; Simoncsits et al. 2001; Heiter et al. 2005). Examples are BbvCI (Heiter et al. 2005), BspD6I (GACTC [4/6]) (Kachalova et al. 2008), BsrDI (GCAATG [2/0]) (Xu et al. 2007), Mva1269I (Armalyte et al. 2005), and BtsCI (GGATG [2/0]) (Too et al. 2010). Such nickases have been used in fusions with zinc fingers, TALE proteins, and methyl CpG binding domains (for further details, see Boch et al. 2009; Moscou and Bogdanove 2009; Hockemeyer et al. 2011; Gabsalilow et al. 2013; Mussolino et al. 2014; Pingoud et al. 2014; Ramalingam et al. 2014; Thanisch et al. 2014; Dreyer et al. 2015; Rogers et al. 2015).

Control of Restriction of Type II Enzymes

Control by C Proteins

Expression of the MTase gene and methylation of the host DNA before synthesis of the REase is essential after entry of a Type II system into the cell. In 1992, the Blumenthal laboratory provided the first evidence for temporal control in a subset of R-M systems, the plasmid-based PvuII system of Proteus vulgaris (Tao et al. 1991; Tao and Blumenthal 1992), soon followed by that in the BamHI system (Ives et al. 1992; Sohail et al. 1995).

A small C gene upstream of, and partially overlapping with, the REase gene is coexpressed from pres1, located within the MTase gene, at low level with the REase after entry of the self-transmissible PvuII plasmid into a new host, whereas the MTase gene is expressed at normal levels from its own two promoters pmod1 and pmod2 located within the C gene (Fig. 7).


FIGURE 7. Intricate control of restriction in the operons of the Type II R-M systems of PvuII and Esp1396I by controlling C proteins (Loenen et al. 2014b). A small C gene upstream of, and partially overlapping with, R is coexpressed from pres1, located within the M gene, at low level with R after entry of the self-transmissible PvuII plasmid into a new host, whereas M is expressed at normal levels from its own two promoters pmod1 and pmod2 located within the C gene. A similar C protein operates in Esp1396I, but in this case the genes are convergently transcribed with transcription terminator structures in between, and M is expressed from a promoter under negative control of operator OR, when engaged by the C protein in a manner similar to that of the PvuII system. C proteins keep both R and M under control and have been tentatively identified in more than 300 R-M systems. See the text for further details. (Reprinted from Loenen et al. 2014b.)


The C protein binds to two palindromic DNA sequences (C boxes) upstream of the C and REase genes: OL, associated with activation, and OR, associated with repression. Low basal expression from the pvuIIC promoter leads to accumulation of the activator, which enhances transcription of the C and REase genes (Tao et al. 1991; Tao and Blumenthal 1992; Bart et al. 1999; Knowle et al. 2005; Williams et al. 2013). After this initial low-level expression of C·PvuII protein from the weak promoter pres1, positive feedback by high-affinity binding of a C protein dimer to the distal OL site later stimulates expression from the second promoter pres, resulting in a leaderless transcript and more C and R protein. The proximal site OR is a much weaker binding site, but C protein bound at OL enhances the affinity of OR for C protein, and at high levels of C protein, the protein–OR complex down-regulates expression of C and R. In this way, C protein is both an activator and negative regulator of its own transcription.

The regulation is similar to gene control in phage lambda: Differential binding affinities for the promoters in turn depend on differential DNA sequence and dual symmetry recognition. C proteins belong to the helix-turn-helix family of transcriptional regulators that include the cI and cro repressor proteins of lambdoid phages. In the wake of PvuII and BamHI, other R-M systems were discovered that were controlled by C proteins, including BglII (A↓GATCT) (Anton et al. 1997), Eco72I (CAC↓GTG) (Rimseliene et al. 1995), EcoRV (Zheleznaya et al. 2003), Esp1396I (CCANNNN↓NTGG) (Cesnaviciene et al. 2003; Bogdanova et al. 2009), SmaI (CCC↓GGG) (Heidmann et al. 1989), and AhdI (McGeehan et al. 2005). In the case of Esp1396I, the genes are convergently transcribed with transcription terminator structures in between, and the MTase gene is expressed from a promoter under negative control of operator OR, when engaged by C protein in a manner similar to that of the PvuII system (Fig. 7). C·Esp1396I controls OR, OL, and OM in a similar manner as described above. In this way, C proteins keep both R and M under control. This delay of REase expression depends on the rate of C-protein accumulation, and this may help explain the ability of C-regulated R-M systems to spread widely (Williams et al. 2013). By September 2013, REBASE listed 19 characterized C proteins, as well as 432 putatives (http://rebase.neb.com/rebase/rebase.html). The organization of the genes in the system and regulatory details differ from system to system, and some C proteins are fused to their REase genes (http://rebase.neb.com/rebase/rebase.html; Tao et al. 1991; Tao and Blumenthal 1992; Semenova et al. 2005; Bogdanova et al. 2009; Liang and Blumenthal 2013). Whether R-M systems as a whole evolved in concert with C proteins remains to be investigated.

The first structures of C proteins without DNA appeared in 2005: C·AhdI from Geoff Kneale's laboratory (McGeehan et al. 2005) and C·BclI from Ganesaratinam (Bali) K. Balendiran's laboratory in collaboration with NEB (Sawaya et al. 2005). These structures resembled those of helix-turn-helix DNA-binding proteins, as expected. The details of the interactions between C proteins and their C boxes in the DNA came later with the studies on the AhdI operon, and the structures of C·AhdI, C·Esp1396I, and C·BclI (Marks et al. 2003; McGeehan et al. 2004, 2005, 2006; Streeter et al. 2004; Sawaya et al. 2005; Callow et al. 2007; Papapanagiotou et al. 2007; Bogdanova et al. 2008; Ball et al. 2009). With the structure and further experiments, the mechanism behind the genetic switch could be elucidated (McGeehan et al. 2008, 2012; Ball et al. 2009, 2012). C·Esp1396I bound as a tetramer, with two dimers bound adjacently on the 35-bp operator sequence OL + OR (McGeehan et al. 2008). This cooperative binding of dimers to the DNA operator controls the switch from activation to repression of the C and R genes. The existence of C proteins explained the difficulty to introduce some R-M genes into E. coli (e.g., BamHI [Brooks et al. 1989]; see Loenen et al. 2014b for further details). The C genes belong to different incompatibility groups, which exclude unrelated R-M systems (called “apoptotic mutual exclusion”): For example, the pvuIIC and bamHIC genes define one exclusion group and prevent entry of ecoRVC due to premature activation of the EcoRV REase gene (Nakayama and Kobayashi 1998).

In 2016, the group of Iwona Mruk in Gdansk reported an unexpected regulatory variation on the above theme (Rezulak et al. 2016). The C·Csp231I gene regulates expression of the REase gene like other C-regulated R-M systems, but there is additional novel control. Separate tandem promoters drive most transcription of the Csp231I REase gene, a distinctive property not seen in other tested C-linked R-M systems. Further, the C protein only partially controls REase expression, yet plays a role in viability of the cells within the population by affecting stability and propagation. Deletion of the C gene led to high REase activity and resulted in loss of these cells in mixed cultures with wild-type R-M cells.

Transcriptional Control: The Case of EcoRI

The transcriptional control discussed above via C proteins has been found for many Type II enzymes, but not all Type II enzymes have such multiple (convergent) promoters and controlling C proteins. A prime example is EcoRI, whose enzymatic activity is controlled in a different way until methylation is complete. The Tokyo group of Ichizo Kobayashi investigated the intricate control of the EcoRI gene, ecoRIR (Liu and Kobayashi 2007; Liu et al. 2007; Mruk et al. 2011). This gene is upstream of the modification gene, ecoRIM. The M gene can be transcribed from two promoters within ecoRIR, allowing expression of the MTase gene with and without ecoRIR, as there is no transcription terminator between the two genes. In addition, the ecoRIR gene has two reverse promoters. These convergent promoters negatively affect each other, as in lambda (Ward and Murray 1979). Transcription from the reverse promoter is terminated by the forward promoters and generates a small antisense RNA. The presence of the antisense RNA gene in trans reduced lethality mediated by cleavage of undermethylated chromosomes after loss of the EcoRI plasmid (postsegregational killing) (Heitman et al. 1989; Mruk et al. 2011). This can be viewed as programmed cell death in prokaryotes. Kobayashi compares R-M systems with toxin/antitoxin (TA) systems composed of an intracellular toxin (the REase) and an antitoxin (the MTase) that neutralizes its effect. These systems would limit the genetic flux between lineages with different sequence-specific DNA methylation (“epigenetic identity”) but would require intricate control of restriction activity (reviewed in Mruk and Kobayashi 2014).

Control by the Cognate MTase

M·Ecl18kI and M·SsoII are two MTases that act as transcription factors and activate expression of their respective REase genes via binding to the regulatory site in the promoter region of these genes (Karyagina et al. 1997; Shilov et al. 1998; Fedotova et al. 2009). The amino-terminal region of M·Ecl18kI performs the regulatory function, but is also important for methylation activity. Loss of methylation activity per se does not prevent the MTase from performing its regulatory function and even increases its affinity to the regulatory site. However, the presence of the methylation domain is necessary for M·Ecl18kI to perform its regulatory function (Burenina et al. 2013).


Type I Families and Diversity

As discussed in Chapter 7, a single Type I common ancestor is likely, given the high sequence similarity of confirmed (biochemically analyzed) and putative enzymes and irrespective of the host within subclasses up to 80%–99%, between subclasses ∼20%–35%. This section is based on two reviews published in 2014 (Loenen et al. 2014a,b), and the reader is referred to these for more details and references. By 2013, ∼50% (1140/2145) of sequenced bacterial and archaeal genomes in REBASE carried one or more hsdR, hsdM, and hsdS genes and 40% appeared to have none, whereas the remainder had some but not all three genes or disrupted or scrambled genes (http://rebase.neb.com/rebase/rebase.html). On average, cells had two systems, although, for example, Desulfococcus oleovorans has eight systems. Type I enzymes could undergo specificity changes via TRD exchanges by homologous recombination, unequal crossing-over, or transposition (Chapter 6, Fig. 6). Domain shuffling is not limited to Type IA enzymes but can be observed between members of the same or different families (Loenen et al. 2014a). Within Type I families, HsdS subunits have the same organization, but between families they have different amino and carboxyl termini (circular permutations; see Loenen et al. 2014a for details). Circular permutation of HsdS of EcoAI indicated structural, but not necessarily functional equivalence, as different permutations resulted in an active R-M system, active in methylation only, or inactive, indicating that the HsdS termini interact with the HsdM and HsdR subunits (Janscak and Bickle 1998). Some bacteria have only one or two hsdR and hsdM genes but many hsdS genes (up to 22 in Mycoplasma sp.!) allowing multiple specificity changes providing protection against invaders (Sitaraman and Dybvig 1997; Dybvig et al. 1998; Loenen et al. 2014a). Shuffling of those 22 hsdS genes could easily result in more than 500 new specificities, “a defensive repertoire reminiscent of the immunoglobulins of higher organisms” (Loenen et al. 2014a). The advent of SMRT sequencing, which allows the localization of methylated bases, has led to a breakthrough in the determination of Type I recognition sites (Eid et al. 2009; Flusberg et al. 2010; Korlach et al. 2010; Clark et al. 2012; Korlach and Turner 2012), which may generate renewed interest in these “sophisticated molecular machines” (Murray 2000). SMRT sequencing not only led to an exponential increase in the number of known Type I recognition specificities (rising from approximately 40 biochemically characterized specificities in 2011 to more than 1100 by 2017) but also the discovery of Type I enzymes that produce m6A on one strand and m4C on the other (Morgan et al. 2016).

Single-Molecule Studies of EcoKI and EcoR124I

AFM and single-molecule studies, together with improved biochemical and biophysical methods, revealed new details about translocation by EcoKI via the motor domains that belong to superfamily 2 (SF2) (Neaves et al. 2009). Mutational analysis of the DEAD-box, RecA-domain-like, motifs of EcoR124I showed long-range effects of various mutations—for example, nuclease mutants could lower translocation and ATP usage rate, there could be a decrease in the off rate, and/or there could be slower restart and turnover (Sisakova et al. 2008a,b). Dimerization appeared to occur preferentially on two-site DNA, whereas DNA looping could occur in the absence of ATP hydrolysis. Would this be a common way to bring distant DNA regions together? Would this mean that SF2-dependent enzyme complexes in higher organisms also use such looping (which are involved in DNA repair, replication, recombination, chromosome remodeling, and RNA metabolism; for discussion, see, e.g., Tuteja and Tuteja 2004; Singleton et al. 2007; Fairman-Williams et al. 2010; Ramanathan and Agarwal 2011; Umate et al. 2011)?

Single-molecule studies using magnetic tweezers were designed to analyze single translocating molecules of EcoR124I in real time (Seidel et al. 2004, 2005, 2008; Stanley et al. 2006; Seidel and Dekker 2007). These experiments provided details on the rate of DNA translocation, as well as the processivity and ATP dependence of the HsdR motors. New facts emerged that may be of consequence for the studies on the eukaryotic SF2-dependent complexes mentioned above: (1) The two motors could work independently and the enzyme tracked along the helical pitch of the DNA on torsionally constrained molecules; (2) translocation could stop and restart by disassembly and reassembly, and (3) the HsdR subunits released the DNA roughly every 500 bp during this process (whereas the MTase remained attached to the recognition site); in other words, about four times over a distance of 2 kb. Concomitantly with this stop and restart process, the enzymes consumed vast amounts of ATP. A translocation block by collision with another HsdR or the presence of supercoiled DNA resulted in cleavage. The enzyme remained at the site but could be displaced by other proteins (e.g., E. coli RecBCD) (Bianco and Hurley 2005).

Type I Enzyme Atomic Structure

In the absence of crystals, the DNA recognition complex of EcoKI, the trimeric M·EcoKI (M2S1), had been modeled on 3D structures of other MTases (Chapter 7, Fig. 10). This model suggested a common origin of Type I and Type II MTases. Would this ancestral MTase combine with one or two HsdR molecules allowing translocation to the site of cleavage? Did translocation involve contacts with nonspecific DNA adjacent to the recognition site in a cleft in HsdR, which would close and reopen using ATP, Mg2+, and probably SAM, to fuel and control the conformational changes? Other questions remained to be answered: Would HsdR touch one strand or both strands of the dsDNA via backbone contacts, and what about step size, or the amount of DNA transported per physical step? And why no cleavage during the initial translocation? Was the translocation rate too high, or the catalytic PD region in the wrong conformation to contact the DNA? One thing seemed certain: Two REases were needed for dsDNA breaks—one for each strand. The answer to some of these questions came when finally the first structures of Type I enzymes appeared.

The Structure of the M·EcoKI (M2S1) Complex

The first crystal structures of HsdS subunits appeared in 2005 and 2010 (Calisto et al. 2005; Kim et al. 2005). The two TRDs are in inverted orientations, which makes the S subunit functionally symmetric (Fig. 8A, bottom right; see Loenen et al. 2014a for discussion). Each TRD consists of a globular DBD and an α-helical dimerization domain. The long α-helices (D1 and D2) encoded by the two conserved regions of the hsdS gene associate to form an antiparallel coiled-coil dimerization helix between the two variable HsdS specificity domains (S1 and S2) (Calisto et al. 2005; Kim et al. 2005). Amino acid side chains down their lengths interlock “like tines of a zipper” and form a hydrophobic core that holds the two helices together and separates the globular specificity domains by a fixed distance. S1 and S2 each recognize one-half of the recognition sequence. Each TRD also associates with one HsdM subunit to form an M2S trimer. Neither HsdS nor HsdM subunits bind to DNA alone, but the EcoKI trimer methylates both strands of the recognition sequence—the “top” strand of the 5′ half-sequence (Am6AC) and the “bottom” strand of the 3′ half-sequence (CGm6AC) of the bipartite AACNNNNNNGTCG—thus protecting the host DNA during replication (Fig. 8A).



FIGURE 8. (A) Model of the M·EcoKI MTase (PDB ID: 2Y7H). The S subunit is composed of two TRDs in inverted orientations. Each TRD comprises a globular DBD and an α-helical dimerization domain. The N-TRD (green) and C-TRD (orange) are specific for the two halves of the recognition sequence (AACNNNNNNGCAG). Zipper-like association of the helices separates the globular domains by a fixed distance and reverses the orientation of the C-TRD. Each TRD also associates with one M subunit (identical, but shown here in different shades of blue for clarity) to form an M2S trimer, which methylates both strands and protects the resident DNA during DNA replication. (B) Structure of the Type IA HsdS protein S-ORF132P (PDB ID: 1YF2). Structure of the Type IA HsdS protein S-MjaXI (PDB ID: 1YF2). The upper diagram shows the domain organization of the protein; arrows represent DBDs, and curly lines represent dimerization α-helices. The amino acid sequence of the protein is shown below, with the domains in corresponding colors. Below this are three views of the structure, from three perpendicular directions: “sideways,” “end-on,” and “above.” The panels on the left depict the protein; those on the right depict the protein with modeled DNA positioned approximately as it is bound. The DNA was taken from PDB ID: 2Y7H and transferred by structural alignment of the S subunits. (A,B, Reprinted from Loenen et al. 2014a.)


The structure and sequence of the S subunit from Methanocaldococcus jannaschii, S·MiaXI, is shown in the top part of Figure 8B. The recognition sequence of this protein is unknown. It is closely related to the Type IA family of EcoKI. Below this are three views of the structure from three perpendicular directions. The panels on the left show the protein on its own, and those on the right a model of the protein bound to DNA.

The structure of the trimeric M·EcoKI (M2S1) was resolved in 2009, thanks to the product of the 0.3 gene product of phage T7, T7 Ocr (Chapter 6), which proved to be a DNA mimic (see section Antagonists of Type I Action: Antirestriction, starting on page 230). Ocr was used to stabilize the otherwise labile MTase complex (Kennaway et al. 2009). Many different single M·EcoKI-Ocr complexes were imaged in the EM, allowing a reconstruction of the 3D complex to a resolution of 18 Å. A model of M·EcoKI is shown in Figure 8A, which depicts the location of the specificity domains of the S subunit in relation to the two M subunits (see Kennaway et al. 2009 for further details).

The Structure of the EcoKI and EcoR124I (R2M2S1) Complexes

The first data on the crystal structure of EcoR124I HsdR were published by Lapkouski et al. (Lapkouski et al. 2007, 2009). This suggested how the pentamer might be assembled and how the motors might translocate dsDNA (Lapkouski et al. 2009). The PD motif was found opposite the translocation domain, which would allow coupling of translocation to restriction. A model was proposed (Lapkouski et al. 2009) based on this HsdR structure, a DNA path across the subunit, and an early, incomplete model of the MTase core (Obarska et al. 2006). This model has much in common with the later model by Kennaway et al. (Kennaway et al. 2012) but differs in the orientation of the HsdR with respect to the MTase core and the path taken by the DNA (see below). Soon afterward, crystal data appeared on the amino-terminal fragment of a putative Type I enzyme from Vibrio sp. This contained three globular domains with an endonuclease core and the ATPase site close to the probable DNA-binding site for translocation. The authors suggested the involvement of a linker helix in the transition from DNA motor protein to nuclease (Uyen et al. 2008, 2009).

In 2012, years of efforts by David Dryden and coworkers finally paid off and the structure of the pentameric EcoKI and EcoR124I (R2M2S1) was elucidated by computer-assisted EM single-particle reconstructions (Kennaway et al. 2012). Single-particle analysis of negative stain EM images showed large differences between DNA-bound (Fig. 9A) and unbound EcoR124I enzymes (Fig. 9B), with their longest dimensions being ∼18 nm versus ∼22–26 nm, respectively. (The smaller particles were the R1M2S1 form and were analyzed separately as described later.) EcoR124I with DNA was in a closed conformation, whereas the enzyme alone was in an open form without DNA. Apparent twofold symmetry was visible in many image averages. Using these data, a 3D reconstruction was generated (Fig. 9A) of EcoR124I bound to a 30-bp dsDNA fragment with the unmethylated recognition site and the enzyme on its own (Fig. 9B; see Kennaway et al. 2012 for details).


FIGURE 9. Gallery of Type I RM structures and conformations determined by EM and single-particle analysis. (A) EcoR124I + DNA (closed state) negative stain EM. (B) EcoR124I without DNA (open state) negative stain EM. (C) EcoKI + DNA negative stain EM. For each 3 × 3 panel, the top rows are image averages, the middle rows are their corresponding reprojections, and the bottom rows are 3D surface views of the 3D reconstruction (bars, 200 Å); on the right is a larger 3D surface perspective view. See Kennaway et al. 2012 for further details. (Reprinted from Kennaway et al. 2012, with permission from Cold Spring Harbor Laboratory Press.)


EcoR124I without DNA was highly extended and more flexible, which allowed a low-resolution (∼3.5-nm) 3D reconstruction of the enzyme (Fig. 9B). Most particles (∼80%) appeared to have their twofold axis roughly normal to the plane of the carbon film, but ∼5% were seen to be folded up into the closed state, indicating a dynamic equilibrium between states in the absence of cognate DNA. Some very thin connections between the domains were “likely pivot points for flexing to allow the enzyme to close up” (Kennaway et al. 2012). These data showed that the subunits strongly moved in a manner to allow entry of the DNA substrate (Fig. 9A,B). This change from an extended structure to a more compact form in the presence of DNA had been seen previously for the MTase (Kennaway et al. 2009; see Kennaway et al. 2012 for discussion).

Negatively stained particles of EcoKI with DNA bound (Fig. 9C) appeared smaller than EcoR124I with DNA (∼16 nm long) and appeared to be more rounded and variable than EcoR124I. The 3D reconstruction of EcoKI with a 75-bp fragment of dsDNA indicated a compact structure with many features similar to EcoR124I with DNA, including recognizable density for the five subunits in a matching arrangement, suggesting a common architecture for Type I enzymes. However, the EcoKI particles were compact and appeared to be identical with and without DNA, and not elongated, as seen for EcoR124I without DNA. The dynamic equilibrium between open and closed forms apparently favored the closed form for EcoKI under the conditions used for EM.

Scattering experiments were used to construct a model showing the location of HsdR and the MTase (Kennaway et al. 2012). In the elongated structure of EcoR124I, the two HsdR subunits were located toward the extreme ends on either side of the MTase core. Fortunately, a fraction of the enzyme existed as a tetramer (R1M2S1). In negative stain EM, the particles of EcoR124I with (Fig. 10A) or without (Fig. 10B) DNA were not 100% homogeneous, and further analysis showed a large missing region at the extremity of the smaller particles, which had to be the location of one of the HsdR subunits. The existence of tetrameric complexes with only one HsdR was consistent with previous biochemical data on EcoR124I (Janscak et al. 1998).


FIGURE 10. 2D difference images from EM data show the position of the HsdR in the EcoR124I complex. (A) Difference imaging between image averages of large (left) and small (right) particles in the EcoR124I + DNA negative stain EM data set reveals a large “negative density” region (red contour at −2.5σ), consistent with a missing HsdR in the small particles. (B) Difference imaging of HsdR in the open state of EcoR124I (without DNA). Although the relative flexibility of the open complex gives rise to a less well-defined difference map, a region of negative density consistent with a missing HsdR is visible nevertheless (red contour). The deduced atomic structure of each EM particle is shown below the EM image. (Adapted from Kennaway et al. 2012, with permission from Cold Spring Harbor Laboratory Press.)


Although the DNA is not visible in these experiments, the authors were able to use T7 Ocr, which binds very tightly to the DNA-binding site of Type I enzymes (Atanasiu et al. 2002; Walkinshaw et al. 2002). EcoR124I-Ocr complexes adopted a closed conformation and further analysis revealed the position of a banana-shaped object running through the center of the enzyme but tilted at an angle relative to the long axis of the 3D map. This banana-like shape matched well with the structure of the Ocr protein (Walkinshaw et al. 2002). This orientation of Ocr in the EM map plus the structural models of the MTase core of EcoKI (Kennaway et al. 2009) and of EcoR124I (Kennaway et al. 2012) allowed only one possible orientation of the MTase with the dimerization helix of the S subunit exposed to the solvent. This was in agreement with previous observations that this region could accommodate small (Gubler and Bickle 1991) and large (Kannan et al. 1989) amino acid insertions, and even a fusion with green fluorescent protein (Chen et al. 2010), without loss of function. Moreover, limited proteolysis indicated preferential cleavage within the dimerization helix (Webb et al. 1995), and hence surface exposure of this region.

In these studies the carboxyl terminus of the R subunit of EcoR124I (aa residues 893–1038) was not visible, but could be modeled using known crystal structures (Kennaway et al. 2012). Using these crystal structures, the EM data, and scattering analyses, atomic models of complete R subunits for EcoKI and EcoR124I were constructed, backed up by the plethora of published biochemical data on these enzymes.

The authors proposed a model that fit the data and gave the location and directionality of the DNA motor domains by aligning these with those of the dsDNA-bound SWI2/SNF2 chromatin remodeling translocase from S. solfataricus (Lapkouski et al. 2009). The direction of DNA translocation of this translocase was known and imposed a similar directionality on each HsdR, and because these had to pull DNA in toward the MTase core of the Type I enzyme, the orientation of each HsdR relative to the MTase core became defined. Based on DNA footprinting experiments (Mernagh et al. 1998; Powell et al. 1998) and the known minimum length of 45 bp of DNA required for ATP hydrolysis (Roberts et al. 2011), the assumption was made that the DNA path between the DNA bound to the HsdR and the DNA bound to the core MTase could not be longer than ∼40 bp. This meant that the motor domains of the HsdR had to have their DNA-binding sites close to the DNA-binding site of the MTase core. Placement of the HsdR on either side of the MTase and interacting directly with DNA was further supported by the length of the structure of another DNA mimic protein, ArdA (Nekrasov et al. 2007; McMahon et al. 2009), which occupies the entire DNA-binding site on Type I enzymes. This then allowed the complete structures for the closed forms of EcoR124I and EcoKI to be constructed as shown in Figure 11.



FIGURE 11. Atomic models of EcoR124I + DNA, EcoR124I, and EcoKI + DNA docked into the EM map densities. (A) Two views of the EcoR124I + DNA model showing the MTase core closed around DNA (green; DNA bound to each HsdR is not shown for clarity). Adenine bases are flipped out into the active sites of each of the two HsdM (light and dark blue), induced by an ∼45° bend in the DNA. (Yellow) HsdS, (red) HsdR, with the β-sheets of the RecA-like motor domains (orange). (Gray) Residues missing from the crystal structures (the 44 and 152 carboxy-terminal residues of HsdM and HsdR, respectively) were modeled de novo. The carboxy-terminal regions of HsdM extend down to bind at the coiled coil of HsdS, and the HsdR carboxy-terminal domains fill some empty density next to the amino terminus of HsdM. (B) A model for EcoKI bound to DNA (colors as in A). HsdS and HsdM from the MTase structure (PDB ID: 2y2C) were docked in as a single rigid body; HsdR modeled on those from EcoR124I (PDB ID: 2w00) (see Supplemental Material of Kennaway et al. 2012) and placed in a position analogous to the EcoR124I model. (C) The model of EcoR124I in the open conformation (i.e., without DNA; colors as in A). Although the EM map is at a lower resolution, a full atomic model could be built, aided by the EcoR124I + DNA model, SANS data, and 2D difference imaging. HsdM and HsdR swing out as a unit away from HsdS. The predicted hinge regions in the carboxyl termini of the HsdM (gray) and their connections to HsdS are not well resolved. (Reprinted from Kennaway et al. 2012, with permission from Cold Spring Harbor Laboratory Press.)


Placement of the R subunit of EcoR124I forced two large kinks in the DNA to allow the DNA to thread through the MTase core (Fig. 11A). This kinked path shortens the through-space end-to-end distance of a duplex bound to the enzyme by ∼10 nm, in line with AFM measurements of complexes of EcoR124I on DNA that showed that binding of the enzyme shortened the length of a long linear DNA molecule by ∼11 nm (van Noort et al. 2004). AFM measurements of EcoKI bound to DNA also showed a pronounced kink (Walkinshaw et al. 2002; Neaves et al. 2009), and circular dichroism analysis of EcoR124I also indicated a large structural distortion to the DNA when bound (Taylor et al. 1994).

A fit of subunits into the EcoKI EM density map (Fig. 11B) corresponded closely to that of EcoR124I in the closed state. The thin protrusions at either side of the EM envelope for EcoKI could fit the long coiled-coil amino-terminal extensions of unknown function predicted in the R subunit of EcoKI but absent in EcoR124I (Fig. 11B). Significant sequence differences existed between the two enzymes, and this might account for other structural differences, although the overall architecture remained unchanged.

An optimal fit of subunits into the lower-resolution open EcoR124I map (Fig. 11C) was obtained by moving and rotating each HsdM–HsdR pair as a single rigid body away from HsdS. A relatively simple ∼90° rotation and an ∼80° twist around a pivot point near the carboxyl terminus of HsdM were sufficient to move between open and closed states. It had previously been shown that HsdR and HsdM can form a complex (Dryden et al. 1997), supporting movement of the two subunits as a rigid body. The carboxy-terminal residues of EcoKI HsdM were disordered in the crystal (PDB ID: 2ar0) (Kennaway et al. 2009) and were sensitive to proteolysis (Cooper and Dryden 1994) and could play the role of the flexible linker proposed. Proteolytic removal of this region inhibited assembly of the pentamer (Powell et al. 2003).

Taken together, the data showed how Type I enzymes assemble, bind, and distort DNA before the initiation of ATP-driven DNA translocation. Although these EM and small-angle scattering structures were of low resolution, the proposed atomic models are in agreement with the extensive data from biochemical, biophysical, and genetic studies (Murray 2000, 2002; Loenen 2003; Tock and Dryden 2005). These provided several further constraints on the subunit orientations and gave confidence in the atomic models shown in Figure 11. These make it clear that there is an equilibrium between open and closed forms of the Type I enzymes, with the equilibrium constant depending on the particular enzyme and the presence or absence of DNA (and presumably the cofactors SAM and ATP). EcoKI prefers to be closed whether DNA is present or not and must therefore transiently open up to allow DNA access to the MTase core. EcoR124I appears to prefer an open form in the absence of DNA but is closed with DNA bound.

It would appear possible for the Type I enzymes to reach the closed “initiation” complex with the S-shaped DNA path via different routes. The model shown in Figure 12A is based on the EcoR124I structure but David Dryden (pers. comm.) states that “it would also apply to other Type I enzymes if we could actually ever see the open conformation—which we did not for EcoKI but this does not mean that it does not exist transiently.” The open form can bind DNA nonspecifically using HsdR (left side of Fig. 12A) and diffuse along the DNA until the MTase core recognizes a target sequence or dissociates. The trigger for closing and formation of the initiation complex would be most likely the recognition of the target sequence by the MTase core. Alternatively, the closed form of the enzyme must open up transiently to allow DNA to enter the MTase core, followed by closing of the core around the DNA (right side of Fig. 12A) and diffusion of the enzyme on the DNA until it either recognizes its DNA target sequence or reopens and dissociates. Starting the process of target sequence location and recognition via this pathway means that the motor domains of the HsdR will have to rely on the inherent flexibility of DNA for them to grasp it and force it into the S-like shape shown in the initiation complex.


FIGURE 12. Schematic of large-scale conformational change and initiation of DNA looping and translocation by EcoR124I. (A) Type I enzymes exist in a dynamic equilibrium between open and closed states (movement is shown by orange arrows, and pivot points in carboxy-terminal regions of HsdM are indicated by pink dots). DNA (green) binding to form encounter complexes can occur nonspecifically to the HsdR (red) or via the target sequence to the MTase core (HsdM is in light and dark blue, and HsdS is in yellow). Complete closure of the enzyme and bending of the DNA around the HsdR produces the initiation complex for DNA translocation. (B) The predicted complete path of the DNA (green dots) through the atomic model of EcoR124I with segments of bound DNA. This is the proposed initiation complex (from Fig. 10A). During active translocation, the DNA would then form expanding loops from each side (light-green dots for DNA, and the direction of translocation is shown by black arrows). The inset shows the initiation complex turned 90° to the main panel. (Reprinted from Kennaway et al. 2012, with permission from Cold Spring Harbor Laboratory Press.)


The introduction of sharp bends in the DNA would require considerable energy to be expended by the enzyme. This may come from the transition between open and closed forms of the RM enzyme, but it may also require the hydrolysis of ATP by the HsdR. The models suggest that once the enzyme has closed around DNA and the motor of an HsdR subunit has a good grip on a segment of DNA, further hydrolysis of ATP (required for translocation and cleavage, although not DNA binding) would push the segment bound to the motor toward the central MTase core, as indicated by large arrows in Figure 12A. Because the MTase core is also tightly bound to the DNA target sequence, DNA at the bend between the segments bound to the motor and to the MTase core would twist and perhaps even buckle, forming the small loop shown in Figure 12B. Formation of this highly strained loop is certain to be energetically unfavorable, in agreement with translocation measurements for Type I enzymes in which it appears that much ATP is used in abortive attempts to initiate translocation (Seidel et al. 2008). Once the loop has formed, further DNA translocation would occur as the motors pump DNA toward the MTase core. Single-molecule experiments make it clear that the motors can work independently (Seidel et al. 2004, 2008), perhaps explaining why early EM studies showed both single- and double-looped structures (Yuan et al. 1980; Endlich and Linn 1985). In light of the large changes occurring upon DNA binding, it is possible that the actively translocating enzyme undergoes further changes in structure (e.g., in the presence of ATP). One may speculate that this great flexibility would allow the enzyme to accommodate the stresses built up during the extensive DNA translocation periods observed for these molecular machines. It is noteworthy, in this respect, that a process of deassembly of the enzymes occurs after DNA cleavage, and some of the subunits—although not all and depending on the particular Type I enzyme—can be reused (Roberts et al. 2011; Simons and Szczelkun 2011).

As mentioned above, the earlier model of Lapkouski et al. (Lapkouski et al. 2009) differs from this new model in two respects: namely, the orientation of the HsdR with respect to the MTase core, and the path taken by the DNA. Previously (Lapkouski et al. 2009), the interface of HsdR with the MTase core was not defined when compared with the new models. More importantly, the DNA was proposed to bend across the motor domains of HsdR, so that it came near to the endonuclease domain in the same HsdR and could be cleaved. If this model was correct (Lapkouski et al. 2009), the partially assembled R1M2S1 form of EcoR124I should have been able to cleave DNA, which was, however, not the case (Janscak et al. 1998). The current model suggests that the endonuclease domain of one HsdR is in proximity to DNA translocated by the other HsdR (Fig. 12B). This would explain the absence of DNA cleavage by partially assembled R1M2S1 forms of EcoR124I, despite the fact that such an assembly translocates DNA effectively (Janscak et al. 1998; Seidel et al. 2004, 2008). Thus, the current models are a significant improvement on the previously published models (Davies et al. 1999; Lapkouski et al. 2009).

Last, the structural models presented can be compared with the structures of complex Type II REases enzymes in groups IIB and IIG (Roberts et al. 2003), which cleave at defined distances either on both sides (IIB) or only one side (IIG) of the target sequence. As mentioned above, these classes are Type I-like combined R-M systems, with an amino-terminal endonuclease PD domain directly fused to a γ-class MTase domain in a single protein, but without motor domains (Dryden 1999; Nakonieczna et al. 2009; Shen et al. 2011). Whereas Type IIB enzymes have a single HsdS-like subunit with two TRDs, the Type IIG enzymes BpuSI and MmeI have only one TRD. The Type IIB enzyme is effectively a dimeric Type IIG. Thus, a Type IIB enzyme is like a motorless Type I system, and a Type IIG system is like one-half of a motorless Type I RM enzyme. Figure 13 compares the relative locations of one endonuclease domain, one HsdM, and the HsdS from the closed form of EcoR124I with the structures of MmeI and BpuSI (Nakonieczna et al. 2009; Shen et al. 2011). It can be seen how fusion of the endonuclease domain from HsdR to the start of HsdM in EcoR124I would move it to the same location as observed in the Type IIG REases and lead to cleavage downstream from the target sequence. Thus, the proposed role of gene fusions in the evolution of different groups of Type II R-M systems (Mokrishcheva et al. 2011) can be extended to include the evolution of the Type I systems.


FIGURE 13. Structural evolution of Type IIG enzymes from a Type I enzyme undergoing fusion of the carboxyl terminus of an endonuclease domain from HsdR via deletion of the motor domains, to the amino terminus of HsdM. (Left) Part of EcoR124I, with one endonuclease domain from HsdR (red), one HsdM (green is the amino-terminal domain, and blue is the MTase catalytic domain), and HsdS (yellow with two TRDs). DNA bound to the MTase core is shown, but DNA bound to HsdR is omitted for clarity. The dashed line shows how the end of the endonuclease domain could join with the amino terminus of HsdM to form a structure similar to the type IIG structures shown on the right. The catalytic motifs in the endonuclease domain and HsdM are shown in spacefill. (Middle) Model of MmeI with bound DNA with the same colors for the equivalent domains (Nakonieczna et al. 2009; coordinates from ftp://genesilico.pl/iamb/models/RM.MmeI). (Right) Crystal structure of BpuSI (PDB ID: 3s1s) with the same coloring of domains as in the other structures and with an inserted extra domain shown in gray (Shen et al. 2011). DNA is absent in this structure, and one can see that the endonuclease domain would be blocking the DNA-binding site on the TRD. Shen et al. (2011) proposed that the endonuclease domain would twist away to allow DNA sequence recognition. (Reprinted from Kennaway et al. 2012, with permission from Cold Spring Harbor Laboratory Press.)


Additional Roles for Type I Enzymes

In 1977, Werner Arber wondered whether REases might have additional functions in the cell (Arber 1977). The finding that a Type I enzyme could cleave a replication fork at its branch may indicate that the answer is yes (Ishikawa et al. 2009). This probably happens when the enzyme travels along the DNA and encounters a replication fork, halts, and cuts (Ishikawa et al. 2009). Possibly in line with this, the groups of Hirotada Mori in Nara and Chieko Wada in Kyoto identified protein–protein interactions in E. coli (Arifuzzaman et al. 2006) between the EcoKI subunits and some other proteins, in a comprehensive pull-down assay using a His-tagged library of ORFs. Some of these interactions may be just a matter of sticky proteins, but others could be of in vivo relevance—for example, ATP-dependent helicase HrpA, the replicative helicase DnaB, DNA polymerase III DnaE, or CTP synthase PyrG, which may point to potential fine-tuning of R-M activity with DNA replication and primary metabolism. Such cross talk between (endo)nucleases and primary metabolism may well be universal, although likely to be much more complex in eukaryotic systems. As such, the E. coli system remains a useful model system to study such complex processes.

In an interesting report, Marie Weiserova and colleagues used immunoblotting to show that HsdR is phosphorylated on threonine in vivo only when coproduced with the MTase subunits (HsdM and HsdS) (Cajthamlova et al. 2007). HsdR lacks this phosphorylation when introduced in the cell by itself. Is this as yet unexplained phosphorylation of EcoKI HsdR another way of restriction control or genome maintenance (e.g., involved in recruiting other enzymes to the DNA) (Cajthamlova et al. 2007)? It certainly warrants further investigation.

On a different track, the Type I ecoprrI system contains an additional gene, originally identified by the group of Tom Bickle (Tyndall et al. 1994; Kaufmann 2000). The protein encoded by prrC proved to be a member of the group of latent anticodon nucleases (ACNs), which are proteins that may be linked to stress responses, and have been called RNA-based innate immunity systems that distinguish self from non-self (Jain et al. 2011).

Most studies on Type I systems concerned enzymes from strains that could be cultured in the laboratory. However, with the advent of whole-genome sequencing, many Type I systems have been identified in benign as well as pathogenic bacteria, in which they may limit genetic exchange between species and contribute to genome and strain stability. Because of the presence in their genomes of sometimes large numbers and different types of REases, these enzymes have proven useful for strain typing of, for example, methicillin-resistant Staphylococcus aureus (MRSA) bacteria (Lindsay 2010). REases such as SauI can limit horizontal transfer through conjugation, phage transduction, or transposition of antibiotic-resistance genes or virulence factors, but they cannot completely prevent it (Waldron and Lindsay 2006; Veiga and Pinho 2009). Knowledge of the target sites of different SauI members in a wide range of S. aureus lineages will aid studies on these pathogens (Cooper et al. 2017).

In 2011, the Kobayashi group showed that within a gene, stretches of amino acids can move from one position to another (Furuta et al. 2011). The authors suggest that such lateral domain movements within genes may be a novel common route to generate new specificities. Finally, much attention has been paid to pathogens that use phase variation, as discussed in Part E.

Antagonists of Type I Action: Antirestriction

This section is a shorter version of the text by Loenen et al. (Loenen et al. 2014a). Note that this type of antirestriction is rather different from the ClpXP-mediated proteolysis mentioned before (Chapter 7; see also, e.g., Simons et al. 2014), or the lambda Ral protein (or the analogous prophage protein Lar), which alleviates restriction by changing EcoKI from a maintenance into a de novo MTase (Chapter 6; see also, e.g., Loenen 2003). Antirestriction (anti-R) and anti-restriction-modification (anti-RM) systems in phage, plasmids, and transposons enhance their survival in a new host. Much of the early seminal work on anti-R by the T-uneven phages T7 and T3 was carried out by the groups of Bill Studier (Studier 1975, 2013; Studier and Movva 1976; Dunn and Studier 1981; Mark and Studier 1981; Bandyopadhyay et al. 1985; Moffatt and Studier 1988) and Detlev Kruger (Kruger et al. 1977a,b,c, 1983). These phages inject a small part of their DNA carrying the 0.3 gene (which encodes Ocr; see next page). The Ocr protein is produced and inactivates EcoKI and EcoBI, before the remainder of their DNA enters the cell. The best-known “artful dodger” of host restriction is probably phage T4, which encodes multiple functions to escape host defenses, some of them useful to genetic engineers (e.g., polynucleotide kinase) (Kruger and Bickle 1983; Bickle and Kruger 1993; Miller et al. 2003; Rifat et al. 2008; Petrov et al. 2010). Other work on anti-RM was carried out by the groups of Tom Bickle and about 20 papers (mainly in Russian) by Belogurov and Zavil'gel'skii spanning nearly 30 years (reviewed in Kruger and Bickle 1983; Bickle and Kruger 1993; Zavil'gel'skii 2000; Thomas et al. 2003; Tock and Dryden 2005; Dryden 2006), whereas in the United Kingdom the Wilkins laboratory solved the riddle of control of anti-R of the self-transmissible IncI plasmid, thus identifying novel transcriptional regulation from a single-stranded promoter (see the next section; Althorpe et al. 1999; Bates et al. 1999; Wilkins 2002; Nasim et al. 2004). During evolution, new mechanisms and countermechanisms between anti-RM and RM appeared continuously (Kruger and Bickle 1983; Bickle and Kruger 1993; Zavil'gel'skii 2000; Wilkins 2002; Thomas et al. 2003; Putnam and Tainer 2005; Tock and Dryden 2005; Dryden 2006; Zavil'gel'skii and Rastorguev 2009). T7 Ocr proved to be a DNA mimic with a large negatively charged patch on its surface (Atanasiu et al. 2001; Walkinshaw et al. 2002); see the next section. In the wake of Ocr, the structures of several other anti-RM proteins have been elucidated: ArdA from Tn916 from Enterococcus faecalis (Serfiotis-Mitsa et al. 2008), ArdB from E. coli CFT073 (Oke et al. 2010), and KlcA, an ArdB homolog from plasmid pBP136 from Bordetella pertussis (Serfiotis-Mitsa et al. 2010); see the subsection The Structure of ArdA.

Whereas Ocr seems to be confined to phage (particularly T7 and its relatives), ArdA and ArdB proteins are usually encoded by conjugative plasmids and transposons (Gefter et al. 1966; Hausmann 1967; Chilley and Wilkins 1995; Chen et al. 2014). In T7, Ocr is synthesized for the first 2 min of infection, before entry of the remainder of the phage DNA (Gefter et al. 1966; Hausmann 1967, 1988; Hausmann and Messerschmid 1988a,b; Moffatt and Studier 1988; Garcia and Molineux 1996; Molineux 2001). This completely inhibits the resident Type I enzymes, and the rest of the T7 genome can safely enter the cell. Interestingly, Ocr could bind in vitro to E. coli RNA polymerase (Ratner 1974), leading to the possibility that Ocr has another role in the cell, like other “moonlighting” proteins (Mani et al. 2015; Jeffery 2016) as, for example, reported for SF2 proteins involved in the skin disease Xeroderma pigmentosum (Le May et al. 2010; Kuper et al. 2014; Compe and Egly 2016). These SF2s are involved in nucleotide excision repair, but also in transcription, leading to active demethylation of CpG islands (Le May et al. 2010).

In the case of ArdA of plasmid ColIb-P9, ssDNA (which is resistant to restriction) enters the cell and forms an unusual promoter via a dsDNA hairpin, which allows transcription of the ardA gene (Chilley and Wilkins 1995; Althorpe et al. 1999; Bates et al. 1999; Nasim et al. 2004). Production of ArdA or ArdB rapidly inhibits the resident Type I enzymes. This novel transcription method may well be more common and deserves further research. Importantly, genome sequencing projects indicate that ard genes are widespread and often accompanied by antibiotic-resistance genes. The impact of combined transfer of these genes on the rate of spread of resistance in bacterial populations will be obvious.

The Structure of Ocr

Ocr is a striking example of DNA mimicry by a protein (reviewed in Loenen et al. 2014a). It is a dimer of two monomers of 116 aa and shaped like a banana with a length of ∼7.5 nm and 2–2.5 nm thick (Dunn et al. 1981; Atanasiu et al. 2001; Blackstock et al. 2001; Walkinshaw et al. 2002; Zavil'gel'skii et al. 2009). In this way, it mimics the shape and surface charge of a section of B-form DNA (Fig. 14). Each monomer contains several α-helices, a long loop, and unstructured flexible amino and carboxyl termini, respectively. The thinness of the structure means that it has a minimal hydrophobic core with many aromatic amino acids, which may resemble the aromatic core of another DNA mimic, Qnr (Hegde et al. 2005). Despite the small core, Ocr is very stable to heat and chemical denaturation (Atanasiu et al. 2001). On the surface of each monomer are 34 negatively charged amino acids and only 6 positive amino acids (∼12 of each would be expected for a typical globular protein of this size), although not all of these are required for activity (Stephanou et al. 2009a,b; Kanwar et al. 2016). The negative surface charges are spaced at roughly the same separation as the phosphate groups on 24 bp of B-form DNA containing a bend in its center, which explains its affinity for Type I enzymes (Atanasiu et al. 2002; Su et al. 2005).


FIGURE 14. Superimposition of two 12-bp B-DNA molecules on the T7 Ocr dimer. The phosphate groups of 12 bases in each DNA dodecamer overlap with 12 carboxyl groups on each Ocr monomer, thus mimicking the shape and surface charge of B-form DNA. Ocr is shown as a blue ribbon with amino (N) and carboxyl (C) termini indicated and the dimer interface shown as a red line. Phosphate groups are colored yellow (phosphorus) and purple (oxygen). The carboxyl groups are colored red (oxygen) and black (carbon). The sugar backbones of the DNA chains are colored in two shades of green with the base pairs omitted for clarity. Vectors for the DNA helical axes are drawn as black lines. (Reprinted from Atanasiu et al. 2002.)


The Structure of ArdA

ArdA is also a mimic of B-form DNA, like T7 Ocr (Fig. 15; McMahon et al. 2009). In the crystal structure, ArdA is a dimer but it can exist both as a dimer and as a higher multimer in solution (Serfiotis-Mitsa et al. 2008). ArdA from Tn916 from E. faecalis is 166 aa long with a very small dimer interface (like Ocr). The dimer is an elongated bent cylinder of ∼15 nm × 2 nm. The thinness of the structure again means that it has a minimal hydrophobic core, but unlike Ocr, ArdA is not very stable to denaturation (Serfiotis-Mitsa et al. 2008). The fold of ArdA is completely different from that of Ocr: Each ArdA monomer has three small, loosely packed domains, suggesting a flexible structure. The domain folds have been found in other protein structures with a mix of α-helices, β-strands, and loops. The surface of each monomer is covered with numerous carboxyl groups such that the dimer mimics ∼42 bp of bent B-form DNA. The flexibility of the structure may indicate that the ArdA protein can mold itself to the contorted S-shaped DNA-binding groove on the Type I enzyme, with different domains interacting with the different R, M, and S subunits (Kennaway et al. 2009, 2012).


FIGURE 15. Protein inhibitors of Type I R–M enzymes. (Top to bottom) DNA model (hydrogen atoms omitted) from PDB: 2Y7H displayed on the same scale as the proteins for structural comparisons; T7 Ocr (PDB: 1S7Z and 2Y7C), ArdA (PDB: 2W82) from Tn916 of E. faecalis (Davies et al. 1999), and ArdB (PDB: 2WJ9) from a pathogenicity island of E. coli CFT073, respectively. All three proteins are homodimeric. Their subunits are identical, but are displayed here in different colors. (Reprinted from Loenen et al. 2014a.)


The Structure of ArdB

The structures of two members of the ArdB family have been solved by both crystallography (ArdB from a pathogenicity island in E. coli CFT073) and NMR spectroscopy (KlcA from B. pertussis plasmid pBP136 [Oke et al. 2010; Serfiotis-Mitsa et al. 2010]). The ArdB and KlcA amino acid sequences are close homologs with >30% sequence identity. Klc genes form part of the kor operon involved in a regulatory network of these promiscuous plasmids (Larsen and Figurski 1994). The two ArdB structures are clearly very different from those of Ocr and ArdA and are globular proteins with a novel fold. They are neither elongated nor possess significant charged patches, so they are unlikely to cause anti-R via DNA mimicry (see Loenen et al. 2014a for discussion).

Effect of Protein Inhibitors Ocr and Ard on Restriction and Modification

The effectiveness of Ocr, ArdA, and ArdB in inhibiting Type I R-M systems has been tested in vivo with the classical efficiency of plating test (e.o.p. test; see, e.g., Chapter 1) comparing the titer of phage on an r+ host carrying an anti-RM gene on a plasmid versus the strain lacking the plasmid (Walkinshaw et al. 2002; Serfiotis-Mitsa et al. 2008, 2010; Zavil'gel'skii and Rastorguev 2009; Zavil'gel'skii et al. 2009, 2011). Active anti-R enhances the number of recovered phages, which are then tested for modification by comparing the e.o.p. on the r+ versus r strain: anti-M activity leads to a lower e.o.p. on the former. A novel calibrated in vivo titration assay was designed for EcoKI by the group of Zavil'gel'skii (Zavil'gel'skii et al. 2009), which allows antirestriction proteins to be distinguished based on the quantitative differences seen at different expression levels of Type I enzymes ( Zavil'gel'skii and Rastorguev 2009; Zavil'gel'skii et al. 2009, 2011).

The plating assays show that Ocr blocks all Type I R-M systems (Walkinshaw et al. 2002). This is a direct consequence of the extremely strong binding of Ocr to the DNA-binding groove in the MTase core of the enzymes (Atanasiu et al. 2002). ArdA (Serfiotis-Mitsa et al. 2008) and ArdB/KlcA (Serfiotis-Mitsa et al. 2010) block restriction in all Type I families. ArdA discriminates between restriction and modification (Nekrasov et al. 2007). ArdA of plasmid R16 preferentially targets the restriction function of EcoKI (Thomas et al. 2003). This minimal anti-M effect is due to the binding of ArdA to the MTase core being of similar or weaker strength than DNA binding to the core. This weak binding is sufficient to prevent restriction but not modification. ArdA and ArdB differ in their propensity to block modification (Walkinshaw et al. 2002; Zavil'gel'skii and Rastorguev 2009; Zavil'gel'skii et al. 2009, 2011; Serfiotis-Mitsa et al. 2010). ArdB/KlcA wild variants all have strong restriction inhibition but weak effect on modification for four Type I classes (Serfiotis-Mitsa et al. 2010). ArdB also shows little or no anti-M effect in vivo and, in line with this, no interaction has been observed in vitro between ArdB and the MTase core. Furthermore, although ArdB causes anti-R in vivo, no effect could be demonstrated in vitro on restriction. Therefore, the mechanism of anti-R used by ArdB is indirect and requires further investigation. David Dryden states (Loenen et al. 2014a): “Our understanding of anti-RM is still in its infancy. Aside from the three systems described above, few others have been studied beyond their initial discovery. Given their synergistic role with restriction and modification in regulating horizontal gene transfer and the resistome (Wright 2010; Stern and Sorek 2011), this deficiency in our knowledge needs to be addressed.”

Type I Single Protein

Type ISP (Type I single protein) are similar to Type IIL REases (like, e.g., MmeI), but have an additional helicase-ATPase domain, which is essential for restriction (Smith et al. 2009a,b,c). The prototype ISP REases are two plasmid-encoded R-M systems in Lactococcus lactis, LlaGI and LlaBIII. LlaBIII is of commercial importance as protection against phage infections in milk fermentations. An α-helical coupler domain connects the Mrr-like PD domain–cum-SF2 helicase region to the m6A γ-MTase-TRD domain. With the exception of the TRD region, LlaBIII is >95% homologous to LlaGI (Sisakova et al. 2013). Structural and single-molecule studies indicated a conformational change after binding to the recognition site; the enzyme leaves the site and translocates DNA without looping, at ∼300 bp/sec at 25°C, consuming one to two ATP per base pair (Chand et al. 2015; Kulkarni et al. 2016). As the PD and helicase/ATPase domains are upstream of the direction of translocation, these results indicated that the enzyme could not simply dimerize via its nuclease domain like Type I enzymes (Chand et al. 2015; Kulkarni et al. 2016). Together with data from single cleavage sequence analysis, this led to the proposal that DNA cleavage occurred as a result of multiple nicks by colliding enzymes, roughly halfway between sites, with the nuclease domains distal (Chand et al. 2015). Recent experiments show that translocation activates the nuclease domains via distant interactions of the helicase or MTase-TRD, without requiring direct nuclease dimerization (van Aelst et al. 2015).

Sequence analyses of 552 Type ISP enzymes showed structurally well-conserved elements involved in target recognition of LlaGI and LlaBIII, although the primary sequences of the TRDs were not that well conserved (cf. Type II enzymes) (Kulkarni et al. 2016). This led to a partial consensus code for target recognition by this class of enzymes with specificity changes due to residues that contacted the bases as well as novel contacts (Kulkarni et al. 2016).



As detailed in the previous chapters, research on the Type III REases was basically limited to the enzymes from phage P1 and plasmid P15 by the groups of Bickle, Rao, Kruger and Reuter, and later Szczelkun (McClelland 2004), whereas the structure of EcoP15I would be finally published by Aneel Aggarwal and coworkers (see the next section). The EcoP15I R-M complex acted as MTase in the absence of ATP, and as MTase or REase in the presence of ATP, depending on the methylation state of the recognition site. Important questions remained: Why was the MTase a dimer of two Mod subunits, as methylation occurred on only one strand of the recognition sequence? Did this play a role in the stability of the complex? There appeared to be quite convincing evidence for a translocation and cutting mechanism similar to that of the Type I enzymes, but why the differences with respect to the interaction with ATP and SAM, and the location of the cut site? Did cleavage (always) occur after DNA tracking with ATP as an energy source, and collision by two complexes? Did this involve one Res subunit or two Res subunits, and did this occur in one or both directions? Would collision result in a conformational change and cleavage 25–27 bp downstream from one of the two sites?

This part of the chapter is based on the historical perspective by Rao et al. (Rao et al. 2014), recent papers that addressed the role of ATP hydrolysis in long-distance communication between sites before cleavage could occur, and the different models based on 1D diffusion and/or 3D-DNA looping. Although it had been known that Type III enzymes needed two sites in a specific head-to-head orientation for cleavage, later evidence indicates that the sites can also be in a tail-to-tail configuration (van Aelst et al. 2010). Moreover, new data provided evidence for the trimeric nature of EcoP15I, EcoP1I, and PstII: one Res (and not two) and two Mod subunits (Butterer et al. 2014). The long-awaited structure appeared of the first Type III REase, which is also the first structure of a dimeric MTase, EcoP15I,which shed unexpected new light on the interactions of this Mod2 dimer with the Res subunit (Gupta et al. 2015); see the next page. Finally, whole-genome sequencing data indicated Type III R-M systems in many sequenced genomes, in which a role for these enzymes in “phase variation” is being unraveled with respect to pathogenicity and virulence of clinically relevant organisms, such as H. influenzae and biofilm formation. After its initial discovery in the 1970s by Andrzej Piekarowicz and Stuart Glover in H. influenzae (Glover and Piekarowicz 1972; Piekarowicz and Glover 1972; Piekarowicz 1974; Piekarowicz and Kalinowska 1974; Piekarowicz et al. 1974, 1975, 1976, 1981, 1986; Jablonska et al. 1975; Piekarowicz and Baj 1975; Kauc and Piekarowicz 1978; Piekarowicz and Brzezinski 1980; Brzezinski and Piekarowicz 1982; Piekarowicz 1982, 1984), phase variation has become an important phenomenon and has also been studied in, for example, Neisseria sp. (Piekarowicz et al. 1988; Kwiatek and Piekarowicz 2007; Adamczyk-Popławska et al. 2009; Kwiatek et al. 2010); see the video from Andrzej's talk at the aforementioned CSHL meeting in 2013 (Piekarowicz 2013), and in Part E.

The Structure of EcoP15I

Different types of experiments addressing the composition of the EcoP15I R-M complex and the mechanism of DNA translocation and looping resulted in different models and much controversy (Peakman and Szczelkun 2004; Raghavendra and Rao 2004; Reich et al. 2004; Crampton et al. 2007a,b; Moncke-Buchner et al. 2009; Ramanathan et al. 2009; van Aelst et al. 2010; Dryden et al. 2011; Szczelkun 2011; Wyszomirski et al. 2012; Schwarz et al. 2013; Rao et al. 2014). Was this related to differences in the composition of the complex (ResMod2 or Res2Mod2 [Rao et al. 2014]) or perhaps even multimeric complexes present in the preparation? There was evidence for translocation and DNA looping by EcoP15I, but was looping essential and/or could ATP drive 1D diffusion of the enzyme on the DNA? In the absence of crystals, Aneel Aggarwal and coworkers used small-angle X-ray scattering (SAXS) and analytical ultracentrifugation to analyze the structure of the EcoP15I R-M complex and the dimeric Mod2MTase (Gupta et al. 2012). Whereas the MTase was relatively compact, the R-M complex was an elongated crescent shape of ∼218 Å. Their data were in line with a model in which the MTase dimer was lodged between Res subunits. Did this mean that the Res subunits would come together and form a sliding clamp around the DNA in order to cut the DNA? Three years later (2015), the first crystal structures of the EcoP15I complex with DNA (and AMP) were published. These results came as a surprise and led to novel insights into the way in which the helicase and modification domains of EcoP15I interacted with DNA and each other (Gupta et al. 2015).

DNA Recognition by EcoP15I

Until this DNA-EcoP15I cocrystal structure appeared, by necessity the interpretation of data on dimeric MTase-DNA structures was limited to those obtained with monomeric MTases (all without DNA) (Gupta et al. 2015). The structure was a big surprise: One EcoP15I Mod subunit, ModA, turns out to be involved in specific recognition of the bases in the target site, whereas the other subunit, ModB, has the target adenine (CAGCAG) in its catalytic cleft, which is rotated 180° out of the DNA helix (Fig. 16).


FIGURE 16. Overall structure of EcoP15I/DNA/AMP complex (PDB: 4ZCFI). (A) The domain arrangements of Mod and Res subunits. (B) An overall view of the Mod and Res subunits (Mod2Res1) bound to DNA and AMP. The two Mod protomers, ModA and ModB, are shown in cyan and blue, respectively, whereas the Res subunit is shown in magenta. The DNA is shown in gray, with the exception of the extrahelical adenine base (yellow). The AMP molecule is shown in yellow. ModA recognizes DNA through base-specific interaction from its TRD (TRDA) and interacts with Res through its MTase domain and CTD. The TRD of ModB (TRDB) does not enter the DNA major groove. CTD of ModA (CTDA) interacts with the Res subunit, whereas the CTD of ModB (CTDB) is exposed to solvent. ModA and ModB dimerize via their NTDs (NTDA/B) and central MTase domains (MTaseA/B). AMP binds in a cleft between the RecA1 and RecA2 motor domains of the Res subunit. The endonuclease domain that ensues the helical spacer is disordered and labeled in a dashed box. The proximity of TRDA of ModA and Pin domain of Res (interdomain distance B14 Å) is highlighted by a double-headed arrow. The intervening loops in the structure that are not modeled because of weak density are represented by colored dashes. (Reprinted from Gupta et al. 2015.)


In contrast to γ-class MTases (Type I HsdM or Type II M·TaqI) or α-class MTases (Type II M·FokI, EcoDam), in which the TRD is adjacent to the active site cleft, EcoP15I Mod belongs to the β-class MTases, where the TRD lies far off this cleft (Gupta et al. 2015). The authors suggested that, by extension, a similar division of labor (ModA subunit for recognition, ModB subunit for modification) may be used by other, mainly dimeric, β-class MTases (e.g., M·RsrI) (Thomas and Gumport 2006) and even also apply to other DNA or RNA m6A-MTases in other organisms, including mammalian cells (Gupta et al. 2015). Such RNA methylation is very common in both nucleus and cytoplasm and, for example, is implicated in RNA metabolism (transcription/splicing) and stem cell development (possibly involving the β-class MTases METTL3/METTL14) (Gupta et al. 2015). Is this division of labor universal? Is it active in all kingdoms, exemplified, for example, by the plant de novo MTase, DRM2 (domains rearranged MTase 2, which methylates only one DNA strand, like EcoP15I) or SPOUT RNA MTases (in which the RNA binds in a cleft between two monomers, and the target base is in the catalytic pocket of one monomer) (Gupta et al. 2015)?

Restriction by EcoP15I

During the early 1990s, helicases had been considered ATP-dependent DNA and RNA unwinding enzymes, but this view was subsequently challenged by data on Type I and other enzymes indicating translocation without strand separation. It became clear that the specificity of helicases or translocases for different substrates was dictated by additional regions in between the RecA motor domains and/or the amino- or carboxy-terminal flanking regions (Singleton et al. 2007). For example, true helicases contain a wedge-like domain between the RecA domains to disrupt the base pair for unwinding (Singleton et al. 2007). During unwinding or translocation, the motors consume ATP with every step, but why did some enzymes consume very little ATP while traveling long distances on the DNA? The answer came from single-molecule fluorescent microscopy studies. These indicated that ATP hydrolysis of EcoP15I bound to its target site did not result in DNA translocation: The energy generated induced a conformational change that resulted in long-range diffusion of the enzyme on the DNA (Schwarz et al. 2013). Such ATP-triggered change of state, which allows sliding, was named “molecular switching” of the enzyme, which could happen on DNA as well as RNA (Szczelkun et al. 2010; Schwarz et al. 2013; Szczelkun 2013), but could probably also cause other events such as protein–protein interactions, for example, for the clamp loader (Kelch 2016). Subsequent kinetic studies support this notion of two distinct ATPase phases, a rapid consumption of ∼10 ATP inducing a conformational change and a slower phase related to the rate of dissociation of the enzyme from the recognition site (Toth et al. 2015).

The novel EcoP15I structure revealed three new substructures, two within the RecA1 segment and one between RecA1 and RecA2: a loop, a β-hairpin-like “Q-arm,” and a novel “Pin” domain (Fig. 16; Gupta et al. 2015). These are scattered through the RecA-like domain. Two are in the amino-terminal half and the Pin domain is an insertion that divides RecA1 half (motifs I-III) from RecA2 (motifs IV–VI) (see Supplementary Fig. 5 of Gupta et al. 2015 and Fig. 1a in Mackeldanz et al. 2013). The motor domains bound dsDNA and facilitated DNA sliding via this specialized Pin domain. The Pin domain adopted a tertiary structure that extended toward the ModA TRD subunit and interacted with the translocating strand of the DNA duplex (Gupta et al. 2015). The DNA was severely distorted from B form at two sites along its axis (i.e., where the adenine was ejected from the recognition site and near the ModA-Res interface). The first distortion was due to intrusion of ModA into the DNA major groove, whereas at the ModA-Res junction the DNA was bent ∼24° toward the minor groove, in the direction of the ModA TRD and Res Pin domain. The result was a reduction in the distance between these domains to <14 Å that “may facilitate an interaction between the two domains when EcoP15I assumes a diffusive or sliding state on DNA” (Gupta et al. 2015). Therefore, the EcoP15I motor domain interacted predominantly with the translocating strand. The semiclosed configuration of the EcoP15I motor domain differed from the more open structure of another translocase, S. solfataricus SF2 translocase, which might indicate that EcoP15I was in an intermediate state, following ATP hydrolysis but before AMP dissociation (see Gupta et al. 2015 for further discussion).

With this structure containing AMP in hand, could one predict what might happen in the presence of ATP? How might the enzyme slide? According to Aneel Aggarwal and coworkers (Gupta et al. 2015), the ModA TRD might move from the DNA major groove to the Pin domain and in this way adopt a “nonspecific” conformation that would result in DNA sliding. Such a movement of ∼40° would be possible because of a flexible linker between the TRD and MTase domain, which would prevent the Pin domain reaching the DNA. This structural model is in agreement with single-molecule studies, which suggest that the ResMod2 complex moves along the DNA (like Type ISP, but unlike Type I, in which the M2S complex remains bound to the recognition site). This trimeric complex would slide along the DNA until it collides with another bound complex, which would make it cleavage competent (Schwarz et al. 2013).



Like the Type IIM REases, Type IV modification-dependent REases (MDEs) recognize a variety of DNA modifications at cytosine or adenine bases (http://rebase.neb.com/rebase/rebase.html; Carlson et al. 1994; Roberts et al. 2003). This section is based on two recent reviews (Loenen and Raleigh 2014; Weigele and Raleigh 2016), and the reader is referred to these for more details and references. Over the years many papers were published reporting phages protecting their DNA against a wide range of Type I, II, and III REases by base modification such as methylation. This did not protect them against Type IV enzymes that preferentially or exclusively attack modified Cs and As. In the laboratory strain E. coli K12, McrA and McrBC recognize hm5C DNA and m5C, but the pathogenic E. coli CT596 also carries the gmrS and gmrD (glucose-modified hm5C restriction) genes allowing restriction of ghm5C DNA in, for example, wild-type T4 DNA (Bair and Black 2007). Interestingly, this activity in turn could be inhibited by T4 IPI*, a small protein encoded by this champion dodger of bacterial defense systems (Bair et al. 2007; Rifat et al. 2008).

Another Type IV REase, Mrr (modified DNA rejection and restriction), was identified in E. coli K12, because it caused cloning trouble by recognizing both m5C and m6A (Heitman and Model 1987; Waite-Rees et al. 1991). In addition to m5C, m6A, and m4C, many other modifications exist in both prokaryotes and eukaryotes with probable roles in defense and stress situations, in part via regulation of replication and transcription (Freitag and Selker 2005; Lobner-Olesen et al. 2005; Zhou et al. 2005; Borst and Sabatini 2008; Kaminska and Bujnicki 2008; Low and Casadesus 2008; Iyer et al. 2009; Kriaucionis and Heintz 2009; Ou et al. 2009; Tahiliani et al. 2009; Xu et al. 2009; Prohaska et al. 2010; Wang et al. 2011; Loenen and Raleigh 2014). The Type IV REases are hard to identify as they lack a cognate MTase, and therefore one cannot use Pacific Biosciences (PacBio) SMRT sequencing to find the recognition sites. Their discovery relies on a genetic system and suitable phages or plasmids as challengers. Cell extracts to digest DNA in vitro and analyze on gel are of little use because of severe degradation of the DNA (Eid et al. 2009; Korlach and Turner 2012; Roberts et al. 2015).

It is not known whether all Type IV enzymes flip the methylated base out of the helix, but that may be common. Figure 17 shows the structure of McrBC compared to other base-flipping proteins.


FIGURE 17. McrB-N in comparison to other base-flipping proteins. (A) SRA domains SUVH5 (3Q0C) and UHRF1 (2ZKD) use loops extending from a crescent formed from two beta sheets to flip C or m5C from undeformed B-form DNA into a pocket (top row), whereas McrB-N (3SSC; bottom row) uses loops from one beta sheet to distort the DNA and flip the base. It resembles the human alkyladenine glycosylase (1BNK; bottom row) in bending the DNA toward the major groove, while flipping the base via the minor groove (Sukackaite et al. 2012). (B) The SRA-like hemimethylated m5C recognition domains. A ribbon model of the amino-terminal domain of the MspJI structure (4F0Q and 4F0P; top) compared with the SRA domain of URHF1 (PDB: 3FDE; bottom). The crescent shape formed by interacting beta sheets and helices αB and αC are the conserved features of the SRA domain highlighted here. Loops on the concave side of UHRF1 participate in flipping the base, and similar loops presumably do so for MspJI. Two of these vary in length among family members and may play roles in sequence context specificity (Horton et al. 2012). (Reprinted from Loenen and Raleigh 2014; A, originally adapted from Sukackaite et al. 2012; B, originally adapted from Horton et al. 2012.)


Fusions of DNA Binding and Cleavage Domains

All characterized Type IV proteins are fusions of various cleavage and DNA recognition domains. By 2014, six groups of enzymes had been identified that recognize modified DNA with low sequence selectivity, with the prototypes McrA, McrBC, Mrr, SauUSI, MspJI, PvuRts1I, and GmrSD (Loenen and Raleigh 2014). Five more groups have various fusions of the DBD and the cleavage domain, whereas they may be ATP- or GTP-dependent for recognition or cleavage (Weigele and Raleigh 2016). As PvuRts1I is qualitatively similar to MspJI family enzymes, and in turn to DpnI (i.e., typical IIM REases), it is debatable whether some enzymes should be classified as Type IIM or Type IV: The key feature discriminating IIM and IV enzymes is cleavage position, although this position is known (and fixed) for IIM enzymes (DpnI, PvuRts1I family, MspJI family), the cleavage position for the Type IV enzymes is either variable (McrBC) or unknown (McrA, Mrr).


McrA (recognition sequence YCGR) of E. coli K12 has been extensively analyzed by bioinformatics and mutagenesis. McrA has an amino-terminal DBD, and a carboxy-terminal HNH domain required for restriction in vivo (Bujnicki et al. 2000; Anton and Raleigh 2004). It recognizes m5C and hm5C (but not ghm5C), with a preference for C or T at the 5′ position, (Y>R)m5CGR; in vitro it binds m5CpG DNA, but does not restrict (Mulligan and Dunn 2008; Mulligan et al. 2010; Loenen and Raleigh 2014; Loenen et al. 2014b). Interestingly, a mutation in the DBD enabled in vivo discrimination between m5C (still recognized) and hm5C (not recognized) (Anton and Raleigh 2004). A protein with a similar HNH nuclease domain, but different DBD, in Streptomyces coelicolor A3, ScoA3McrA, cuts DNA modified by the E. coli Dcm protein (Cm5CWGG) or phosphorothioate (PT)-modified sites (or both) at a variable distance, and cleavage depends on Mn2+ or Co2+ (Gonzalez-Ceron et al. 2009; Xu et al. 2009; Liu et al. 2010).


McrBC of E. coli K12 has been well characterized (Raleigh 1992; Sutherland et al. 1992; Gast et al. 1997; Pieper et al. 1997, 2002; Stewart and Raleigh 1998; Stewart et al. 2000; Panne et al. 2001; Pieper and Pingoud 2002; Sukackaite et al. 2012; for details, see Loenen and Raleigh 2014; Weigele and Raleigh 2016). The mcrB gene also encodes an additional protein, McrB(S), which starts at an internal translation initiation codon (thus lacking the first 161 aa) and appears to have a regulatory function (Beary et al. 1997). Figure 18 shows a model for the assembly of the McrBC complex (Loenen and Raleigh 2014). McrBC is a GTP-dependent heteroheptamer in which McrC (with the PD nuclease motif) binds a complex of McrB with GTP and DNA. McrBC makes a double-strand cut near one Rm5C site but requires the cooperation of two sites or a translocation block. The sites may be on different daughters across a fork. These are separated by 30-3000 bp, may be on either strand, and minor cleavage clusters are found ∼40, ∼50, and ∼60 nt from the m5C (Pieper et al. 2002).


FIGURE 18. McrBC assembly model (Loenen and Raleigh 2014). Two proteins are expressed from mcrB in vivo. Both the complete protein (McrB-L) and a small one missing the amino terminus (McrB-S; top row) bind GTP, forming high-order multimers detected by gel filtration (second row). When visualized by scanning transmission EM, these appear as heptameric rings with a central channel. Rings of McrB-L in top views show projections that may correspond to the amino-terminal DBD (red segment). Both forms can then associate with McrC, judged again by gel filtration. McrB-L: GTP can bind to its specific substrate (RmC) in the absence of McrC (third row); in its presence, the substrate is cleaved (fourth row). GTP hydrolysis is required for cleavage (arrow): A supershifted binding complex forms in the presence of GTP-γ-S, but no cleavage occurs. Translocation accompanies GTP hydrolysis; dsDNA cleavage requires collaboration between two complexes, or a translocation block. The path of the DNA in the figure is arbitrary, as is the conformation of McrC. (Modified from Bourniquel and Bickle 2002, with permission from Elsevier Masson SAS.)



Mrr of E. coli K12 recognizes m6A and m5C (with uncertain specificity), which prevented cloning of some R-M systems (e.g., PstI [CTGm6AG], HhaII [Gm6ANTC], and StyLTI [CAGm6AG]) and other genes (Heitman and Model 1987; Kelleher and Raleigh 1991; Waite-Rees et al. 1991; Loenen and Raleigh 2014; Weigele and Raleigh 2016). Mrr contains a predicted variant of the PD motif in the carboxy-terminal domain, with a presumed amino-terminal winged-helix DBD, like the MspJI family (Loenen and Raleigh 2014; Weigele and Raleigh 2016).


The (d)ATP-dependent SauUSI (SCNGS) of S. aureus recognizes Sm5CNGS (S = C or G) (Loenen and Raleigh 2014). It has a PLD nuclease domain (like BfiI and other REases), a carboxy-terminal DBD, a helicase/translocase domain in between, and a cleavage domain (Xu et al. 2011) with a reaction mechanism resembling that of topoisomerases and transposases (Interthal et al. 2001; Sasnauskas et al. 2003, 2007), but not quite the same: PLD nucleases employ a covalent protein–DNA intermediate, but, unlike topoisomerases and some (e.g., serine) transposases, the covalent linkage is made by a histidine rather than serine or tyrosine.


PvuRts1I from P. vulgaris Rts1 restricted glucosylated T-even phages in vivo (Janosi et al. 1994; Loenen and Raleigh 2014). It recognizes (g)mC(N11-13/N9-10)G—that is, it cuts 11–13 nt downstream from a modified C, which is 20–23 nt before a G (hence, not very specific). It has approximately 20 active homologs, including AbaSI form Acinetobacter baumanii, which recognize m5C slightly and hm5C and ghm5C better, with differing preferences and weak and variable selectivity for the sequence surrounding the modified base (Szwagierczak et al. 2011; Borgaro and Zhu 2013). These enzymes require two modified sites for dsDNA cleavage ∼22 nt apart, with incisions ∼11–13 nt 3′ to the modified base on the one strand and 9–10 nt 3′ on the other. Bioinformatics, mutational evidence, and crystal structures indicate a carboxy-terminal SRA-like (SET and ring finger–associated) DBD, and an amino-terminal cleavage domain that is a divergent member of the PD family (Kazrani et al. 2014; Shao et al. 2014; Weigele and Raleigh 2016). The structure of AbaSI confirmed the results with PvuRts1I, with an amino-terminal nuclease domain resembling that of VSR, whereas the carboxy-terminal domain resembles SRA family members (Horton et al. 2014a; Weigele and Raleigh 2016).


The aforementioned GmrSD enzyme, encoded by the gmrS and gmrD genes from a pathogenic E. coli strain, also exists as a single fusion protein with similar characteristics (Bair et al. 2007; He et al. 2015; Machnicka et al. 2015). In vitro, the enzyme prefers ghm5C DNA to unmodified m5C. GmrD is the nuclease, whereas the GmrS domain has a ParB/Srx fold, present in conjugative plasmids (He et al. 2015; Machnicka et al. 2015; see Weigele and Raleigh 2016 for details). The enzyme has a strong preference for UTP over GTP and CTP (Loenen and Raleigh 2014; Weigele and Raleigh 2016).


MspJI from Mycobacterium sp. JLS, was the first member of a family of Type IV enzymes, which cleave with a four-base 5′ extension 12 nt from the m5C, and 16–17 nt on the opposite strand (Bujnicki and Rychlewski 2001; Zheng et al. 2010; Cohen-Karni et al. 2011; Horton et al. 2012, 2014b,c). Members recognize m5C and hm5C, but not ghm5C. MspJI has a SRA-like amino-terminal domain, which flips m5C out of the helix, like other SRA-like proteins. Swapping experiments with short protein loops contacting nearby bases, which varied among three members, significantly reduced selectivity for sequences flanking the modified base (Sasnauskas et al. 2015; Weigele and Raleigh 2016).

The endonuclease domain cuts a different DNA strand than the DNA bound by that polypeptide. This is reminiscent of M·EcoP15I in which the DBD of one subunit binds the recognition sequence, whereas the catalytic domains of, in this case, two other subunits, effect dsDNA cleavage. MspJI (and the isoschizomer AspBHI [Horton et al. 2014b,c]) can be thought of as Type IIS enzymes (albeit enzymes that bind only when the recognition site is modified) and their tetrameric structure and domain cooperation could be typical of many “normal” Type IIS enzymes, as well as the Type IIB, IIC, and IIG enzymes. The convention is to think of these acting as transient homodimers, but they might well act as tetramers (and even as multimers of tetramers, e.g., BcgI) instead.



Bacterial pathogens not only infect the host, but also try to maintain themselves (“colonize”) by hiding from the immune system and/or confusing it. One way of doing this is by hypermutation at simple sequence repeats or homopolymeric tracks located within the reading frame or in the promoter of a subset of genes (Moxon et al. 2006). This is due to polymerase slippage (called slipped-strand mispairing [SSM]), which could be an important evolutionary strategy (Levinson and Gutman 1987a,b). Genetic variation in the population of pathogenic bacteria via SSM would result in two or (many) more different phenotypes that allow the strain to evade the host immune system (Robertson and Meyer 1992). SSM changes the number of repeats or bases, which switches promoters “on” or “off” (e.g., by changing the distance between the -35 and -10 regions), causes frameshifts in coding regions, and/or dictates alternative usage of multiple translation initiation codons in different reading frames, thus altering or abolishing DNA recognition (see, e.g., van Ham et al. 1993; van Belkum et al. 1998; De Bolle et al. 2000; van der Woude and Baumler 2004; Srikhanta et al. 2005, 2010; Moxon et al. 2006; van der Woude 2006; Dixon et al. 2007).

The groups of Richard Moxon, Andrzej Piekarowicz, and Michael Jennings have studied repeat variation that created this so-called “phase variation” in three pathogens, H. influenzae, H. pylori, and Neisseria sp., revealing altered expression of up to 80 genes, including genes important for iron uptake, DNA repair, electron transport, amino acid transport, and growth (reviewed in Srikhanta et al. 2010) but also for MTases. In the latter cases, the MTases can function as an on–off switch for multiple genes that allow the pathogen to combat host immunity (De Bolle et al. 2000; Seib et al. 2002; Srikhanta et al. 2005, 2010; Casadesus and Low 2006; Moxon et al. 2006; Wion and Casadesus 2006; Marinus and Casadesus 2009). Various other clinical isolates also contain MTases with repeats of variable length, which are either lone MTases or associated with Type II systems (Kong et al. 2000; Xu et al. 2000a,b; Lin et al. 2001; Vitkute et al. 2001; Skoglund et al. 2007). These genetic systems have been called “phasevarions” or “phase-variable regulons,” and appear to be a common strategy to randomly switch between distinct cell types and create phenotypic heterogeneity in the bacterial pathogenic population (Weiser et al. 1990; van Ham et al. 1993; Hallet 2001; Srikhanta et al. 2010). Such methylation-driven alternative gene expression can be very complex because of the presence of multiple phase-variable genes (see Srikhanta et al. 2010 for further details).

Taken together, different types of variation may alter gene expression: (1) reversible changes (i.e., alternative states of the same genes result in expression or not, or expression of different genes [e.g., flagella or tail fibers], via inversion of promoters or amino termini, or slipped mispairing); (2) diversity selected changes, in which a rare genotype is favored because of changing circumstances (involving outside forces); and (3) replacement or addition variation, in which a gene is replaced by another gene or is added extra.

Variable Type II systems

Around the turn of the century, two dozen potential R-M systems were identified in two completely sequenced H. pylori strains, 26695 and J99, amounting for >4% of the total genome (i.e., much more than in other sequenced bacterial genomes) (Kong et al. 2000; Lin et al. 2001). Although nearly 90% of the R-M genes were present in both strains, <30% of the Type II R-M systems were functional in both strains with different sets active in each strain. An interesting observation was that all strain-specific R-M genes were active, whereas most shared genes were inactive. Did this indicate that these active strain-specific genes had been acquired recently via horizontal transfer from other bacteria? And did these pathogenic strains constantly acquire new R-M systems and inactivate and delete the old ones (Lin et al. 2001)? Were these multiple R-M systems a “primitive bacterial immune system, by alternatively turning on/off a subset of numerous R-M systems” (Kong et al. 2000)? Support for this idea came from other H. pylori strains, in which the R-M systems also proved to be highly diverse (Xu et al. 2000a). In addition, several REases had novel specificities: Hpy178III (TCNNGA), Hpy99I (GGWCG), and Hpy188I (TCNGA) (Xu et al. 2000a). The latter system was absent in the 26695 and J99 strains, whereas the GC content implied that the Hpy188I system had been recently introduced into the H. pylori genome (Xu et al. 2000b).

In 2016, phase variation of a Type IIG enzyme was reported in Campylobacter jejuni NCTC11168 (Anjum et al. 2016). This IIG protein methylates adenine in CCCGA and CCTGA sequences. Using both inhibition of restriction and PacBio-derived methylome analyses of mutants and phase variants, the cj0031c allele in this strain was demonstrated to alter site-specific methylation patterns and gene expression, which “may indirectly change adaptive traits” (Anjum et al. 2016).

Phase-Variable Type III systems

Phase-variable Type III mod genes have been identified in H. influenzae, H. pylori, and Neisseria sp., which contain tandem repeats that may be homopolymeric, or repeat tracks of 2, 3, 4, or 5 nt (Srikhanta et al. 2010). The mod-like gene of H. influenzae Rd has 40 AGTC repeats within its ORF. This mod gene was found in 21 out of 23 other H. influenzae strains, and in 13 of those the locus contained repeats of variable length (De Bolle et al. 2000). These repeats comprised a hypervariable region in the central region of the mod gene of 22 nontypeable H. influenzae strains, whereas the res gene was conserved (Bayliss et al. 2006). Moreover, similar mod genes with similar hypervariable regions were identified in pathogenic Neisseria sp., suggesting horizontal transfer of these genes between different species. This high phase variability of these MTases would not only protect against phage infections but might “also have implications for other fitness attributes of these bacterial species” (Bayliss et al. 2006). An example of phase variation that allows typing of these so-called untypeable H. influenzae (“NTHi”) strains is shown in Figure 19, redesigned from Figure 3 in Fox et al. 2007. Variation of the number of repeats generates ON/OFF switches. Switching within a clonal population results in subpopulations with and without RM activity. In Haemophilus and other taxa, this results in variable expression of distant genes as well, presumably regulated by the presence of modification in regulatory regions (e.g., Tan et al. 2016; see, for a review, Sanchez-Romero et al. 2015). The variable regulation at distant sites does not require activity of the restriction function (Fox et al. 2007). Also observed in this figure is the presence of variable segments (TRDs) that give distinct recognition specificities. These variations are generated by horizontal transfer between strains, species, and even genera (Bayliss et al. 2006). Further studies indicated that in H. influenzae Rd, phase variation of modA was calculated to occur at a frequency of 4 × 10−6 mutations/division/repeat unit for off-to-on switches and 7 × 10−6 mutations/division/repeat unit for on-to-off switches (De Bolle et al. 2000). The restriction phenotype of a Type III system in N. gonorrhoeae, NgoAXP, switched randomly because of a change in the number of pentanucleotides (CCAAC/G) present at the 5′ end of the coding region of the ngoAXPmod gene (Adamczyk-Popławska et al. 2009). The mod gene in another N. gonorrhoeae strain, FA1090, was linked to biofilm formation, adhesion to human cells and epithelial cell invasion, and hence to pathogenicity and systemic infection (Kwiatek et al. 2015). Various serotypes of Pasteurella haemolytica have CACAG repeats within the 5′ end of a Type III R-M system, repeats which could change in length upon serial subculture and most likely also occurred as a result of DNA SSM (Ryan and Lo 1999).


FIGURE 19. Phase variation. A representation of phase-variable methyltransferase genes present in different strains of nontypeable H. influenzae (NTHi). The prototypical strain of each NTHi strain in which each allele/arrangement is present is shown on the left side. Each modA gene is represented as a white arrow, with the DNA recognition domain (DRD) represented by a colored box. Downstream from each modA is the cognate restriction endonuclease gene, res, with the locations of the ATP-binding motif (TGxGKT), the ATP-hydrolysis motif (DEAH), and the endonuclease motif, PD · · · (D/E)XK, indicated above. Phase-variable modA genes are represented by the five most common modA alleles found in NTHi isolates from patients with middle ear infection (otitis media [OM]). These alleles—modA2, 4, 5, 9, and 10—all contain a simple-sequence repeat (SSR) tract; in this case the sequence AGCC repeats n times. The SSR tract is represented by a gray box in these alleles, with the number of AGCC repeats and the expression status of this number of repeats shown underneath each allele. For example, modA2 in NTHi strain 723 contains 16 AGCC repeats, which leads to expression (ON) of the gene. NTHi strain R539 contains a deletion of the entire mod-res region. NTHi strain 162 contains the modA7 allele that does not contain SSRs and therefore is not phase variable. ModA alleles that are not phase-variable contain the 12-nt sequence 5′-TCAGATAGTCAG-3′ in place of a SSR. In all cases, the genes flanking the mod-res locus are conserved: upstream is the gene rnhB (represented by the blue arrow to the left of each modA gene); downstream is the gene pcaC (represented by the yellow arrow to the right of each res gene). (Courtesy of John Atack.)


Phase-Variable Type I systems

Phase variation also occurs in Type I systems. HindI of H. influenzae has a (GACGA)4 repeat, and changes in the number of pentanucleotides (which is influenced by Dam methylation) within the coding sequence of hsdM were linked to protection against phage (Zaleski et al. 2005). In Mycoplasma pulmonis, two hsd loci each contain two hsdS genes with complex, site-specific DNA inversion systems (Dybvig et al. 1998). This generates a complete family of related hsdS genes with extensive sequence variations that recognize different DNA sequences, suggestive of additional roles in genome maintenance (Dybvig et al. 1998).

In the Type IC NgoAV system from N. gonorrhoeae, the length of tandem repeats of four amino acids were involved in the generation of a truncated or full-length HsdS protein, and only the long protein could complement other Type IC systems (Adamczyk-Popławska et al. 2011). Similar tetra-amino acid repeats (either TAEL, LEAT, SEAL, or TSEL) were identified in other Type IC systems in distantly related bacteria (Adamczyk-Popławska et al. 2003). Was this a common special characteristic of Type IC systems (Adamczyk-Popławska et al. 2003), and is there a tale to tell? Do these data shed light on the origin of the switch between the EcoR124I and EcoR124II systems mentioned in Chapter 6 (Fig. 6)? These proteins recognize GAA (N6) RTGC and GAA (N7) RTGC, respectively, and the presence of either six or seven bases between the two specific DNA regions was shown to relate to the presence of either two or three TAEL repeats in the respective HsdS subunits (Price et al. 1989). Price et al. (1989) suggested that the switch between the two specificities could be due to unequal crossing-over in these repeats (Price et al. 1989). Is this a coincidence or are these repeats perhaps the end result of prolonged SSM? Were these TAEL repeats once much longer, and were they slowly eliminated during continued growth under laboratory conditions, because they were no longer needed?

Such an assumption could fit in with LacZ fusion experiments by Bolle et al. (2000) with respect to the above-mentioned mod-like gene of H. influenzae Rd (with 40 AGTC repeats within its ORF). These authors fused a lacZ reporter to a chromosomal copy of mod downstream from the repeats, which resulted in high-level phase variation. Changing the number of repeats changed mutation rates. Phase variation occurred at a high frequency in strains with the wild-type number of repeats. Rates increased linearly with tract length over the range 17–38 repeat units. The majority of tract alterations were insertions or deletions of one repeat unit with a 2:1 bias toward contractions of the tract (De Bolle et al. 2000). This could be interpreted to mean that the shorter the track, the higher the chance that the track would become shorter faster. As under laboratory conditions the pressure is absent to protect against either phage or host immunity, the bacteria with shorter tracts would have a favorable advantage over bacteria with longer tracts. That advantage would eventually be lost once the number of repeats became very small, and the length of the spacer between the two recognition domains could no longer be decreased without loss of the DNA recognition function. The end result would be two active enzymes, EcoR124I and EcoR124II, with only two or three TAEL repeats left.

Monika Adamczyk-Popławska et al. (Adamczyk-Popławska et al. 2011) analyzed a poly(G) tract in the hsdS(NgoAV1) gene in N. gonorrhoeae. Deletion of 1 nt in this tract with seven guanines led to a frameshift at the 3′ end of the hsdS(NgoAV1) gene and fusion to a second downstream hsdS gene, hsdS(NgoAV2) (Adamczyk-Popławska et al. 2011). This resulted in a longer HsdS protein with two TRDs, rather than the original truncated HsdS protein with a single TRD derived from hsdS(NgoAV1) (Adamczyk-Popławska et al. 2011). Such a contraction of the poly(G) tract that caused this frameshift might well occur in vivo, as the authors found a minor subpopulation of cells that appeared to have only six guanines. Thus, it could be argued that the strain could switch this Type I system “on” (two TRDs = protection against phage and/or other foreign DNA) or “off” (one TRD and the possibility of DNA exchange via horizontal transfer).


It has proven very difficult to generate mutant or hybrid restriction enzymes with long DNA specificity sites that would serve as good tools for gene therapy. The new RNA-based method of genome editing using the CRISPR–Cas9 system may present a better alternative, but also has its drawbacks. To answer the question of how foolproof the CRISPR system is, Bull and Malik (2017) state in their discussion of a recent paper by Champer et al. (2017) that “the easy targeting of CRISPR, the very property that has led to its current popularity, may also be its downfall as a practical means to control populations or suppress disease transmission.” This would fit in with the research described in this book, which proves that organisms go to incredible lengths to keep genome alterations under tight control to avoid chromosomal instability and allow only a low level of heterogeneity.

The year of the 5th NEB meeting in Bristol in 2004 (Fig. 20) was also the year of the publication of the first and only book totally dedicated to restriction enzymes (edited by Alfred Pingoud [Pingoud 2004]). It reflects the growing realization of the importance of restriction-modification systems in “cells, not eppendorfs” (King and Murray 1994), and their role outside the laboratory in communities with benign and pathogenic bacteria and archaea.


FIGURE 20. The NEB meetings between 1988 and 2015. (Top left) The first four NEB meetings on restriction and modification enzymes. (Top right) The 5th NEB meeting on Restriction/Modification in Bristol (2004) was organized by Mark Sczcelkun, Bernard Connolly, and David Dryden. (Bottom left) The 6th NEB meeting on DNA Restriction and Modification in Bremen (2010) was organized by Albert Jeltsch, Alfred Pingoud, and Wolfgang Wende. (Bottom right) The 7th NEB meeting on DNA Restriction and Modification in Gdansk (2015) was organized by Iwona Mruk, Geoffrey Wilson, and Richard Morgan. (Courtesy of Rich Roberts.)


The large current number of restriction enzymes and the 50-odd structures indicate overlap between types and subtypes in different ways. The enzymes may share a common ancestor with three separate domains for DNA recognition, restriction, and methylation, whereas Type I and III enzymes also contain the ATP-dependent SF2-related molecular motor domains. The cooperation of restriction enzymes between sites with or without looping and with or without collision/stalling may be more the rule than the exception. Different catalytic nuclease domains, mainly with the PD · · · (D/E)XK motif, but also HNH, GIY-YIG, or PLD domains, seem to have been “mixed and matched” during the course of evolution. The structures indicate careful control of positioning of the nuclease domain and large conformational changes before cleavage can occur, plus a variety of other control systems to avoid rampant nuclease activity, which would result in genome instability. Such other control may inhibit synthesis of the restriction enzymes in the cell at the level of transcription at the operon promoter or via C proteins, at the level of translation and/or posttranslation, and finally via DNA mimics employed by various plasmids and phages that inhibit Type I enzymes. The structure of the EcoP15I Type III enzyme indicates a division of labor of the two modification subunits—one for DNA recognition and the other for methylation, which may be a more general mechanism and may also apply to EcoKI (see below). EcoP15I hydrolyses approximately 30 ATP molecules in two steps (a fast consumption of approximately 10 ATP molecules followed by a slower consumption of approximately 20 ATP, which switches the enzyme into another rather distinct structural state that can diffuse on DNA over long distances (Schwarz et al. 2013). This resembles the situation with EcoKI, in which ATP also acts as allosteric effector (Burckhardt et al. 1981). The big difference with EcoP15I is that the EcoKI methyltransferase complex remains bound to the recognition site, whereas the HsdR subunits translocate DNA and hydrolyze ATP, and the whole EcoP15I complex slides away from the site (thus consuming less ATP). Both the ATP-triggered thermal diffusion and ATP-dependent initiation of translocation are important observations for studies into the much more complex eukaryotic methyltransferases and helicases.

One of the most important applications of restriction enzymes with concomitant impact on science and society has been the development of the DNA fingerprinting technique by Alec Jeffreys. Alec started his “Zoo blots” in the 1970s next door to the author of this book in Leicester, where he continued his work on restriction fragment length polymorphisms (RFLPs), which research could fill a book by itself (Fig. 21; Jeffreys 2006).


FIGURE 21. The first DNA fingerprint developed by Alec Jeffreys in the Genetics Department in Leicester, September 10, 1984. (Courtesy of Alec Jeffreys.)


A final note about the intriguing story of EcoKI, hemimethylation, and cofactor SAM, a subject close to the author's heart (Loenen and Murray 1986; Loenen 2003, 2006, 2010, 2017, 2018): As already suggested in 1981 (Burckhardt et al. 1981), EcoKI has two types of binding sites for SAM—one for DNA recognition (the effector site) and the other for methylation (the methyl donor site). How does this relate to the ability of EcoKI to switch from a maintenance methyltransferase (with a strong preference for methylation of hemimethylated DNA) to a de novo methyltransferase? This ability is a property of Type IA enzymes (and not Type IC enzymes such as EcoR124I) that has been observed in the presence of the lambda Ral protein (Zabeau et al. 1980; Loenen and Murray 1986). How can a small protein like Ral cause such an important switch? Are there similar proteins to be discovered in eukaryotic systems? And what about the de novo methylation EcoKI mutants (e.g., the HsdM L113Q mutant made in Noreen Murray's laboratory) (Kelleher et al. 1991)? In contrast to wild-type EcoKI, which has two indistinguishable high-affinity SAM-binding sites, mutants like L113Q have only one high-affinity site, whereas the second site has a low affinity for SAM (Winter 1997). Is it the asymmetry of the complex with DNA that controls the switch between methylation and restriction? How the ability to bind only one SAM molecule properly turns L113Q into a de novo enzyme, without apparently affecting the methylation reaction as such, requires further investigation, as the ability to distinguish between unmethylated and hemimethylated DNA is of fundamental importance to cellular activities.


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http://library.cshl.edu/Meetings/restriction-enzymes/v-GeoffWilson.php Wilson G. 2013. The cloning efforts at NEB.

http://library.cshl.edu/Meetings/restriction-enzymes/v-Janulaitis.php Janulaitis A. 2013. Science and politics: three phases of commercialization at Fermentas.

http://library.cshl.edu/Meetings/restriction-enzymes/v-Lubys.php Lubys A. 2013. The cloning efforts at Fermentas.

http://library.cshl.edu/Meetings/restriction-enzymes/v-Roberts.php Roberts R. 2013. Many more REases at CSHL, the start of REBASE and more recent work.

http://library.cshl.edu/Meetings/restriction-enzymes/v-Studier.php Studier FW. 2013. Phage T7 is neither modified nor restricted by EcoKI or EcoBI, which led to the discovery of the T7 0.3 gene encoding Ocr, as well as to the collision model for Type I enzymes.

http://rebase.neb.com/cgi-bin/cryyearbar 2017. REBASE crystals. Restriction enzyme structures per year.

http://rebase.neb.com/rebase/rebase.html REBASE. The restriction enzyme database.

http://scop.mrc-lmb.cam.ac.uk SCOP. Structural classification of proteins.

https://www.knaw.nl/en/awards/laureates/dr-h-p-heinekenprijs-voor-biochemie-en-biofysica/alec-j-jeffreys-1950-groot-brittannia Koninklijke Nederlandse Academie van Wetenschappen site (Royal Dutch Academy of “Arts and Sciences”), which is hosting an Alec J. Jeffreys biography.


https://www.ncbi.nlm.nih.gov/pubmed/20011117 Gitschier J. 2009. DNA fingerprints: an interview with professor Sir Alec Jeffreys.


APPENDIX 1: Table of selected REases studied in recent years

Type II name and restriction site Sub-typea Detailsb Reference(s)c



Hybrid IIP with tetrameric M&S MTase, missing link Type I and II?

Marks et al. 2003; Pingoud et al. 2014



PLD REase cleaves poorly at high concentrations.

Heiter et al. 2015

CCTCAGC (−5/−2)


Two catalytic sites from different subunits, each cleaving own strand; useful as nicking enzyme.

Bellamy et al. 2005; Heiter et al. 2005

(10/12) CGA(N6)TGC (12/10)


Cleaves four bonds in concerted action.

Kong and Smith 1998; Marshall et al. 2007, 2011; Marshall and Halford 2010; Halford 2013; Smith et al. 2013a,b; Pingoud et al. 2014



Monomer localizes target site by 1D and 3D diffusion, nicks one DNA strand, turns 180°, and cleaves the second strand; the switch in orientation proceeds without dissociation into bulk solution (Sasnauskas et al. 2011); design of nicking endonucleases (Kostiuk et al. 2011).

Janulaitis et al. 1983; Sokolowska et al. 2007a; Kostiuk et al. 2011; Sasnauskas et al. 2011; Kostiuk et al. 2015

ACTGGG (5/4)


PLD. The first non-PD-(D/E)XK restriction enzyme identified (Sapranauskas et al. 2000). Well-studied; carboxy-terminal DBD resembles that of B3-like plant transcription factors; cleaves one strand at a time via covalent intermediate; catalyzes both DNA hydrolysis and transesterification reactions (Sasnauskas et al. 2007).

Sapranauskas et al. 2000; Lagunavicius et al. 2003; Sasnauskas et al. 2003, 2007, 2008, 2010; Zaremba et al. 2004; Gražulis et al. 2005; Marshall and Halford 2010; Golovenko et al. 2014; Pingoud et al. 2014



Gm5CNGC; relatives widespread requiring different numbers of m5C.

Xu et al. 2016

GGGAC (10/14)


RM·BpuSI has FokI-like carboxy-terminal domain; accompanied by two MTases.

Niv et al. 2007; Shen et al. 2011; Sarrade-Loucheur et al. 2013; Pingoud et al. 2014



Tetramer. Dimeric mutant still has same fidelity as tetramer. Member of CCGG family.

Gražulis et al. 2002; Zaremba et al. 2005, 2006, 2012; Manakova et al. 2012

GACTC (4/6)


N·BspD6I is a site-specific nickase in a heterodimeric complex with a catalytic subunit.

Kachalova et al. 2008

ACCTGC (4/8)


Detection relative orientation of two target sites.

Kingston et al. 2003



Monomeric REase cuts dsDNA recognition site in two independent binding events.

Rasko et al. 2010



Thermophilic REase with overlapping specificity to BamHI and BglII.

Townson et al. 2004, 2005

(9/12) ACNNNNNCTCC (10/7)


Domain swapping and circular permutation of TRD subdomains (or deletion) → active protein with altered specificity/poor protein yields or inactivity.

Xu et al. 2015



Tetramer. Importance of spatial rather than sequence conservation of aa at active center. Member of CCGG family.

Bozic et al. 1996; Skirgaila and Šikšnys 1998; Skirgaila et al. 1998; Šikšnys et al. 1999



Heterotetramer of R + H with extensive ATP hydrolysis. R protein is similar to BfiI.

Zaremba et al. 2014, 2015



Methylation-dependent; cuts as monomer, one strand at a time; amino-terminal PD domain and carboxy-terminal winged helix (wH) allosteric activator domain; both domains bind methylated DNA with sequence specificity.

Lacks and Greenberg 1975; Siwek et al. 2012; Mierzejewska et al. 2014; Pingoud et al. 2014



The first REase which flips out nucleotides from its target site to accommodate interrupted CCGG sequences in the conserved active site. Transient tetramerization triggers cleavage. Member of CCGG family.

Denjmukhametov et al. 1998; Bochtler et al. 2006; Tamulaitis et al. 2008; Fedotova et al. 2009; Protsenko et al. 2009; Zaremba et al. 2010; Burenina et al. 2013; Rutkauskas et al. 2014



GIY-YIG structure.

Mak et al. 2010; Pertzev et al. 1997; Ibryashkina et al. 2007; Orlowski and Bujnicki 2008; Mokrishcheva et al. 2011

CTGAAG (16/14)


Accompanied by one MTase (cuts 1½ turn away). Sequence specificity was altered by the methylation activity–based selection technique.

Janulaitis et al. 1992a,b; Rimseliene et al. 2003; Pingoud et al. 2014



The first structure of a restriction enzyme; one of the most extensively studied “classical” restriction enzyme. Studies on inhibition by Cu2+ ions; relaxed specificity and structure of mutants that cleave EcoRI star sites; role of flanking sequences; regulation ecoRIRM operon.

McClarin et al. 1986; Kim et al. 1990; Kurpiewski et al. 2004; Sapienza et al. 2005, 2007; Liu and Kobayashi 2007; Liu et al. 2007; VanderVeen et al. 2008; Ji et al. 2014; Sapienza et al. 2014;



For the first time it was shown that REase can be activated to cleave refractory DNA recognition sites (Kruger et al. 1988).The only demonstrated restriction enzyme which interacts with three recognition sites to effectively cleave one DNA site (Tamulaitis et al. 2006b). The first case of autoinhibition, a mechanism described for many transcription factors and signal transducing proteins (Zhou et al. 2004). Member of CCGG family.

Kruger et al. 1988; Zhou et al. 2002, 2003, 2004; Reuter et al. 2004; Kruger and Reuter 2005; Tamulaitis et al. 2006a,b, 2008; Shlyakhtenko et al. 2007; Gilmore et al. 2009; Golovenko et al. 2009; Szczepek et al. 2009



One of the most extensively studied “classical” restriction enzymes. Single molecule studies; tracking of single quantum dot labeled EcoRV sliding along DNA manipulated by double optical tweezers indicated that during sliding, EcoRV stays in close contact with the DNA.

Winkler et al. 1993; Kostrewa and Winkler 1995; Bonnet et al. 2008; Biebricher et al. 2009

GGATG (9/13)


Early, best-known IIS; two MTases fused in single protein; crystals indicate catalytic domain hidden behind DNA-binding domain; DNA-cleavage domain used for engineering purposes.

Sugisaki and Kanazawa 1981; Miller et al. 1985; Nwankwo and Wilson 1987; Mandecki and Bolling 1988; Kaczorowski et al. 1989; Kita et al. 1989a,b; Landry et al. 1989; Looney et al. 1989; Sugisaki et al. 1989; Goszczynski and McGhee 1991; Szybalski et al. 1991; Li et al. 1992, 1993; Li and Chandrasegaran 1993; Skowron et al. 1993, 1996; Waugh and Sauer 1993; Kim et al. 1994, 1996a,b, 1997, 1998; Waugh and Sauer 1994; Yonezawa and Sugiura 1994; Hirsch et al. 1997; Wah et al. 1997, 1998; Bitinaite et al. 1998; Leismann et al. 1998; Chandrasegaran and Smith 1999; Friedrich et al. 2000; Vanamee et al. 2001; Bibikova et al. 2002; Urnov et al. 2005; Catto et al. 2006, 2008; Gemmen et al. 2006; Bellamy et al. 2007; Miller et al. 2007; Szczepek et al. 2007; Vanamee et al. 2007; Mino et al. 2009, 2014; Mori et al. 2009; Sanders et al. 2009; Zhang et al. 2009; Guo et al. 2010; Imanishi et al. 2010; Klug 2010a,b; Carroll 2011a,b; Gabriel et al. 2011; Halford et al. 2011; Handel and Cathomen 2011; Li et al. 2011; Pattanayak et al. 2011; Ramalingam et al. 2011; Handel et al. 2012; Laurens et al. 2012; Pernstich and Halford 2012; Rusling et al. 2012; Bhakta et al. 2013; Ramalingam et al. 2013; Guilinger et al. 2014a; Pingoud et al. 2014



Monomeric REase with structural (but no sequence) similarity to MspI; back-to-back dimer with two active sites and two DNA duplexes bound on the outer surfaces of the dimer facing away from each other. The function of the base flipping is unclear and it seems to be part of great DNA distortions.

Yang et al. 2005; Horton et al. 2006

GGTGA (8/7)



Cymerman et al. 2006



GIY-YIG structure with highly specific recognition structure, in contrast with HEases or DNA repair enzymes.

Xu et al. 2000b; Kaminska et al. 2008; Orlowski and Bujnicki 2008; Sokolowska et al. 2011



The first member of a HNH family REase-mediated death benefits population; computer model, but no crystal structure available.

Chandrashekaran et al. 2004; Saravanan et al. 2004; 2007a,b; Gupta et al. 2010; Vasu et al. 2012, 2013; Vasu and Nagaraja 2013

CCTC (7/6)


Member of Type IIS with the HNH-type active site within carboxy-terminal domain, DNA recognition amino-terminal domain.

Kriukiene et al. 2005; Kriukiene 2006



Needs to interact with two copies of the recognition sequence for efficient cleavage of one. Real-time observation of DNA looping dynamics of NaeI and NarI compared.

Huai et al. 2000; van den Broek et al. 2006



Real-time observation of DNA looping dynamics of NaeI and NarI compared.

van den Broek et al. 2006

GAATGC (1/−1)


Monomeric REase with two EcoRI-like and FokI-like catalytic domains; design of nicking endonucleases (Armalyte et al. 2005).

Armalyte et al. 2005; Pingoud et al. 2014

TCCRAC (20/18)


No separate MTase (cuts two turns away, crystal); changes in the S domain alter recognition site for R and M (like Type I enzymes); altered specificities could be predicted.

Boyd et al. 1986; Tucholski et al. 1995, 1998; Nakonieczna et al. 2007, 2009; Morgan et al. 2008, 2009; Morgan and Luyten 2009; Callahan et al. 2011, 2016; Pingoud et al. 2014



Well-known 8-bp cutter; structure reveals unique metal binding fold (also in other putative endonucleases) with an iron atom within Cys4 motif

Lambert et al. 2008



“half-pipe”; cuts GTA/C; no true REase, as it flips all four purines out of the helix; DNA adenine glycosylase excises adenines.

Miyazono et al. 2007, 2014; Ishikawa et al. 2005; Watanabe et al. 2006; Pingoud et al. 2014; Kojima and Kobayashi 2015



HNH REase; 8–base pair rare-cutting homodimer, each subunit with two Zn2+-bound motifs surrounding a beta-beta-alpha-metal catalytic site.

Shen et al. 2010



Use nucleotide flipping as a part of its DNA recognition mechanism. Member of CCGG family.

Pingoud et al. 2003; Szczepanowski et al. 2008; Tamulaitis et al. 2008



The first characterized homotetrameric enzyme: Structures reveal two different binding states of SfiI: one with both DNA-binding sites fully occupied and the other with fully and partially occupied sites.

Wentzell et al. 1995; Viadiu et al. 2003; Embleton et al. 2004; Vanamee et al. 2005; Bellamy et al. 2007, 2008, 2009; Laurens et al. 2009



Filaments by cryoEM; Member of CCGG family; preferentially cleaves concertedly at two sites; assembles into homotetramers, then other molecules join to generate helical structures with one DNA-bound homodimer after another.

Laue et al. 1990; Tautz et al. 1990; Capoluongo et al. 2000; Bitinaite and Schildkraut 2002; Daniels et al. 2003; Hingorani-Varma and Bitinaite 2003; Šikšnys et al. 2004; Wood et al. 2005; Dunten et al. 2008, 2009; Park et al. 2010; Little et al. 2011; Lyumkis et al. 2013; Ma et al. 2013b; Horton 2015



Member of CCGG family; tested for applications such as genome surgery.

Kubareva et al. 1992, 2000; Pingoud et al. 2005, 2014; Sudina et al. 2005; Bochtler et al. 2006; Pingoud and Silva 2007; Fedotova et al. 2009; Schierling et al. 2010; Hien le et al. 2011; Abrosimova et al. 2013



8–base pair cutter of AT-rich site.

Dedkov and Degtyarev 1998; Shen et al. 2015



Cuts A:A and T:T mismatch in CAG and CTG repeats (useful for typing in Huntington's disease).

Ma et al. 2013a

ACGGA (11/9)


Monomeric bifunctional R-M with independent REase and MTase activities that can be uncoupled; resembles Type I, but lacks translocation domain (“half” Type I).

Skowron et al. 2003; Zylicz-Stachula et al. 2009, 2014

(8/13) CACNNNNNNTCC (12/7)


DNA recognition, cleavage, and methylation in one polypeptide.

Smith et al. 2014

Other Types d




Structure recognition domain McrBC; flips out the modified cytosine.

Sukackaite et al. 2012



Amino-terminal, atypical PD-(D/E)XK REase domain and carboxy-terminal SRA domain with potential pocket for a flipped hm5C or ghm5C. Epigenome studies (Szwagierczak et al. 2011; Wang et al. 2011). This one and other family members could also be classified as Type IIM (like DpnI). Also a Type IIS. Unlike Type IV (McrBC), PvuRts1I family enzymes cut at a fixed distance from the recognition site.

Janosi et al. 1994; Szwagierczak et al. 2011; Wang et al. 2011b; Kazrani et al. 2014; Shao et al. 2014; Zagorskaite and Sasnauskas 2014



Structure based on EM analysis.

Kennaway et al. 2012


Structure HsdR.

Csefalvay et al. 2015




Gupta et al. 2015

MspJI (and other family members, e.g., LpnPI, AspBHI, …)

IIM , IIS (methylation-dependent)

This is a large family of methylation-dependent (Type IIM and IIS) restriction enzymes. Structures of MspJI (apo- [Horton et al. 2012] and DNA bound, with a flipped base in the SRA domain (Horton et al. 2014c); structures of apo-LpnPI (Sasnauskas et al. 2015b), apo-AspBHI (Horton et al. 2014b) are also available. Base flipping in solution (Zagorskaite and Sasnauskas 2014). Applications in genome methylation studies (Zheng et al. 2010; Cohen-Karni et al. 2011).

Zheng et al. 2010; Cohen-Karni et al. 2011; Horton et al. 2012, 2014b,c; Zagorskaite and Sasnauskas 2014; Sasnauskas et al. 2015b

Update September 2017; courtesy of Vilnius group.

a Subtypes as listed on REBASE, with some adaptations by Pingoud et al. (2014) to indicate the overlap between different subtypes.

b The REases that were the first of their (sub)type (adapted from Horton 2015) are indicated in bold: For example, BcnI was the first Type II enzyme to be identified as a monomer (Sokolowska et al. 2007).

c References include older papers to the respective REases, where suitable: For example, SgrAI was the first REase to be shown to form filaments by cryoEM a few years ago (Lyumkis et al. 2013), but the enzyme was already first reported in 1990.

d See Parts B, C, and D within the chapter (on Type I, III, and IV, respectively) for details and other references.

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