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Is Sda a protein, or is it a protein domain of DnaA?

Is Sda a protein, or is it a protein domain of DnaA?



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I initially thought that a domain was a specific part of a protein, with it given tertiary structure, to which a given molecule is able to bind. (I think I recall phrases such as "the haem binding domain of protein X… " being used in lectures?)

Having consulted Wikipedia on protein domains, I see the definition is a bit more subtle:

A protein domain is a conserved part of a given protein sequence and (tertiary) structure that can evolve, function, and exist independently of the rest of the protein chain. Each domain forms a compact three-dimensional structure and often can be independently stable and folded.

I can understand this, however the reason why I started questioning what a 'domain' actually refers to was because if it's use in reference to Sda in control of endosporulation of bacteria. In my lecture notes, it is stated that "KinA is bound and destabilised by Sda, a DnaA target,"

From this I was under the impression that Sda is a protein (although to the best of my knowledge DnaA only binds DNA, so I do not know why Sda would be target of DnaA. Anyhow,) on the other hand Wikipedia states "the protein domain Sda is short for suppressor of dnaA or otherwise known as sporulation inhibitor A.", which seems to me to suggest that Sda is a part of DnaA whose modifications allow DnaA suppression?

Also, on Wikipedia, it is later written "Sda protein domain is a checkpoint which prevents the formation of spores." How can a protein, if Sda is one, be a checkpoint?

On the other hand my lecture notes later talk about the regulation of Sda levels, so it again is referred to as a separate protein!

I would very much appreciate if someone could explain what 'domain' means in this context.


Short answer

Wikipedia has unnecessarily made an already confusing situation much worse. Sda is its own protein, it is regulated by DnaA, and it prevents sporulation.

Full answer

I don't think your misunderstanding is based on the meaning of 'domain' - the definition you give sounds quite reasonable - but rather the use of the term domain with respect to Sda and the naming of Sda itself, which I agree is quite confusing and made much worse by Wikipedia in this case.

Sda is a protein that inhibits KinA activity (Burkholder et al., 2001) and is a completely separate protein from DnaA. I don't see where you jumped from your notes to the idea that Sda is a domain of DnaA except for from Wikipedia: I don't know why Wikipedia thinks Sda is a protein domain, it is a protein. It is a very small protein, so perhaps it would be appropriate to think of it all as one domain, but there is not a single result for "Sda protein domain" on Google Scholar - no one else is using this terminology. A full Google search only returns a few (~150) results, most of which are direct copies/translations from Wikipedia, or Wikipedia is a direct copy of them. Wikipedia can be useful, but if it starts to cause you confusion, don't assume it is correct.

The phrasing "KinA is bound and destabilised by Sda, a DnaA target" means that Sda binds KinA and destabilizes it, and notes that Sda is a target of DnaA; since DnaA is a transcription factor, you can infer that this means that DnaA controls expression of Sda in some way.

In biology a checkpoint just refers to a place that a process can be halted/arrested, often in reference to the cell cycle. Sda is considered a checkpoint because when Sda is expressed, it prevents cells from sporulating by indirectly preventing activation of Spo0A (Burkholder et al., 2001).

Another source of confusion is that the name "suppressor of dnaA" on Wikipedia is WRONG, but further, molecular biologists sometimes have an impenetrable way of naming things, owing to the complexity of control of gene expression and the circuitous way that new proteins are identified and understood.

The correct name for sda is "suppressor of dnaA1". dnaA1 is NOT DnaA - dnaA1 is a MUTANT ALLELE of DnaA that results in cells that are completely unable to sporulate. But! sda is also NOT Sda: sda is a MUTANT ALLELE that produces non-functional Sda (the protein).

I'll quote from the title of the table in Burkholder et al:

sda Mutations Suppress the Sporulation Defect of dnaA1 Mutants

Or, stated again, in dnaA1 mutants, there is a defect in DnaA which prevents sporulation. The researchers searched for mutations that would reverse this effect. They found one, and named it sda, because it reverses (suppresses) the effect of the mutation they were studying. The dnaA1 mutant's effects on sporulation seem to be caused by an increased expression of Sda, which is why the sda mutant reverses the effect.

References


Burkholder, W. F., Kurtser, I., & Grossman, A. D. (2001). Replication initiation proteins regulate a developmental checkpoint in Bacillus subtilis. Cell, 104(2), 269-279.


CBS Domain Protein TA0289

When TA0289 was purified, it showed a compelling reddish-purple color, which may give a hint to its function. The color is caused by the binding of iron ions in the metal-binding site. This is similar to the small redox protein rubredoxin, which contains a similar iron-binding site and shows a similar beautiful color when purified. With testing, MCSG researchers found that TA0289 can transport electrons, suggesting that it may perform this function inside cells. The CBS domain links the two chains into a stable dimer, so its role may be primarily structural. Although the reason that TA0289 needs to be a dimer is still a mystery. the two iron binding sites are too far apart to transfer electrons to one another. You can take a look at this protein structure in the PDB entry 2qh1.

CBS Domain Protein TA0289 (PDB entry 2qh1)

TA0289 is a dimer of identical subunits. The two CBS domains, colored darker blue here, link the two chains together. The two zinc ribbon domains, colored turquoise, extend on opposite sides of the complex. The four cysteine amino acids in each zinc ribbon coordinate an iron atom, shown here with a reddish sphere.

References

  1. M. Proudfoot, S. A. Sanders, A. Singer, R. Zhang, G. Brown, A. Binkowski, L. Xu, J. A. Lukin, A. G. Murzin, A. Joachimiak, C. H. Arrowsmith, A. M. Edwards, A. V. Savchencko, A. F. Yakunin (2008) Biochemical and structural characterization of a novel family of cystathionine beta-synthase domain proteins fused to a Zn ribbon-like domain. J. Mol. Biol. 375, 301-315.

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PDB-101 helps teachers, students, and the general public explore the 3D world of proteins and nucleic acids. Learning about their diverse shapes and functions helps to understand all aspects of biomedicine and agriculture, from protein synthesis to health and disease to biological energy.

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Response to DNA damage: Dual role of extramitochondrial cytochrome C

Domain architecture and N-end structure of ANP32B. (A) Schematic representation of ANP32B domain organization. The N-end structured domain of ANP32B is colored in blue, whereas the histone chaperone LCAR is represented in yellow. The four LRRs are colored in red. ANP32B Nuclear Localization Signal (NLS) is represented in orange and the histone chaperone Nuclear Export Signal (NES) is marked on the image. ANP32B Thr244 residue is represented in grey. (B) Ribbon representation of ANP32B (PDB: 2ELL [12]) N-terminal domain following the color scheme described in A. ANP32B structure is rotated 90° around the horizontal axes in each view. (For interpretation of the references to color in this figure legend, the reader is referred to the Web version of this article.)

Living beings are continuously exposed to harmful agents, both exogenous (ultraviolet radiation, polluting gases, etc.) and endogenous (secondary products of cellular metabolism) that can affect DNA integrity. That's why cells are endowed with a series of molecular mechanisms whose purpose is to identify and signpost possible damage to the genetic material for speedy repair. These mechanisms are precisely regulated because they are key to cell survival. In extreme situations of massive and irreparable damage, cells enter a phase of controlled dismantling called "programmed cell death." Among the events that take place during this process is the massive delivery to the cytoplasm of a mitochondrial protein called cytochrome C. Under homeostatic conditions, this protein plays a role in energy metabolism within the mitochondria. However, in situations of irreparable damage, it directs the controlled and orchestrated destruction of the cell from the cytoplasm.

The authors of this paper suggest that extramitochondrial cytochrome C plays a dual role in response to cell damage. In the initial stages of damage, a limited amount of cytochrome C is able to reach the cell nucleus without accumulating in the cytoplasm and, therefore, without triggering programmed cell death. Once in the nucleus, cytochrome C interacts with the protein ANP32B, a histone chaperone that helps maintain DNA structure and inhibits the activity of the enzyme (phosphatase) PP2A, a major promoter of DNA damage repair. Thus, cytochrome C "hijacks" ANP32B as a means of activating PP2A and facilitating DNA repair. When damage to the genetic material cannot be repaired by the cellular mechanisms, cytochrome C "floods" both the cytoplasm and the nucleus, leading to the irremediable death of the cell. "What's the point of continuing to build a house that's going to be demolished?"


Characteristic structural features

Orc1-5 as well as Cdc6 have conserved AAA+ folds, including Walker A and Walker B ATP-binding domains, characteristic of ATP-dependent clamp-loading proteins, which allow ring-shaped protein complexes to encircle duplex DNA (see Figure 1). Sensor-1 and Sensor-2 motifs are also found within the AAA+ fold and are believed to detect whether ADP or ATP is bound and to contribute to ATPase activity [18]. These domains are located centrally, in the case of Orc1 and Orc2, and towards the amino termini in Cdc6, Orc3, Orc4, and Orc5. Near the carboxyl termini of these proteins a winged-helix domain is present that mediates DNA binding [14, 15, 17]. Somewhat surprisingly, structural studies of archaeal Orc1 suggest that the AAA+ domain also contributes to its association with origin sequences [14, 15]. Interestingly, Cdc6 has been shown to act like an additional ORC subunit, associating with the complex in the G1 phase of the cell cycle and inducing a conformational change that increases its sequence specificity for DNA binding [19, 20]. When Cdc6 is bound to ORC, a ring-like structure is predicted with structural similarities to the Mcm2-7 helicase complex that ORC-Cdc6 loads onto DNA in an ATP-dependent manner [19, 21].

As mentioned above, sequence similarity has been identified for Orc1 and Sir3, with a particularly high degree of conservation between their amino-terminal 214 amino acids (50% identical, 63% similar), which includes a BAH (bromo-adjacent homology) protein-protein interaction domain [6, 22]. Sir3 is required for transcriptional silencing of telomeres and mating-type loci, functions that are also ORC-dependent [3, 5, 23], as discussed below. Although formally a member of ORC, Orc6 contains none of the aforementioned structural features, and shows no evidence of a common evolutionary origin with Orc1-5. It is nevertheless considered an ORC protein as its association with the other five subunits is required to promote the initiation of DNA replication. Relative to other ORC subunits, Orc6 is poorly conserved between budding yeast and metazoan eukaryotes [11] (see Figure 2). Nevertheless, a number of important domains specific to Orc6 have been identified in S. cerevisiae, including an amino-terminal 'RXL' docking sequence (amino acids 177-183) which mediates an interaction with the S-phase cyclin Clb5 [24], and a carboxy-terminal region (the last 62 amino acids) which associates with the other ORC subunits. Both ends of Orc6 (amino-terminal 185 amino acids, carboxy-terminal 165 amino acids) interact with Cdt1, another replication factor required to load Mcm2-7 onto DNA [25]. In both human and Drosophila cells, Orc6 plays a role in cytokinesis, and studies with the latter organism have identified a carboxy-terminal domain that interacts with the septin Pnut, a component of the septin ring that forms in cell division, as well as an amino-terminal domain that is important for DNA binding [26–29]. Interestingly, structural modeling of Drosophila Orc6 revealed that the amino terminus, but not the carboxyl terminus, is homologous to the human transcription factor TFIIB, raising the possibility that proteins involved in replication and transcription may have coevolved [27].


<p>This section provides any useful information about the protein, mostly biological knowledge.<p><a href='/help/function_section' target='_top'>More. </a></p> Function i

Activates ribosomal RNA transcription, as well other genes. Plays a direct role in upstream activation of rRNA promoters. Binds to a recombinational enhancer sequence that is required to stimulate hin-mediated DNA inversion. Prevents initiation of DNA replication from oriC. Binds to hundreds of transcriptionally active and inactive AT-rich sites, approximately half its binding sites are in non-coding DNA, which only accounts for about 10% of the genome (PubMed:16963779).

<p>Manually curated information for which there is published experimental evidence.</p> <p><a href="/manual/evidences#ECO:0000269">More. </a></p> Manual assertion based on experiment in i


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Protein interactions are fundamentally characterized as stable or transient, and both types of interactions can be either strong or weak. Stable interactions are those associated with proteins that are purified as multi-subunit complexes, and the subunits of these complexes can be identical or different. Hemoglobin and core RNA polymerase are examples of multi-subunit interactions that form stable complexes.

Transient interactions are expected to control the majority of cellular processes. As the name implies, transient interactions are temporary in nature and typically require a set of conditions that promote the interaction, such as phosphorylation, conformational changes or localization to discrete areas of the cell. Transient interactions can be strong or weak, and fast or slow. While in contact with their binding partners, transiently interacting proteins are involved in a wide range of cellular processes, including protein modification, transport, folding, signaling, apoptosis and cell cycling. The following example provides an illustration of protein interactions that regulate apoptotic and anti-apoptotic processes.


Heavy BAD protein–protein interaction. Panel A: Coomassie-stained SDS-PAGE gel of recombinant light and heavy BAD-GST-HA-6xHIS purified from HeLa IVT lysates (L), using glutathione resin (E1) and cobalt resin (E2) tandem affinity. The flow-through (FT) from each column is indicated. Panel B: Schematic of BAD phosphorylation and protein interactions during cell survival and cell death (i.e., apoptosis). Panel C: BAD protein sequence coverage showing identified Akt consensus phosphorylation sites (red box). Panel D: MS spectra of stable isotope-labeled BAD peptide HSSYPAGTEDDEGmGEEPSPFr.

Proteins bind to each other through a combination of hydrophobic bonding, van der Waals forces, and salt bridges at specific binding domains on each protein. These domains can be small binding clefts or large surfaces and can be just a few peptides long or span hundreds of amino acids. The strength of the binding is influenced by the size of the binding domain. One example of a common surface domain that facilitates stable protein–protein interactions is the leucine zipper, which consists of α-helices on each protein that bind to each other in a parallel fashion through the hydrophobic bonding of regularly-spaced leucine residues on each α-helix that project between the adjacent helical peptide chains. Because of the tight molecular packing, leucine zippers provide stable binding for multi-protein complexes, although all leucine zippers do not bind identically due to non-leucine amino acids in the α-helix that can reduce the molecular packing and therefore the strength of the interaction.

Two Src homology (SH) domains, SH2 and SH3, are examples of common transient binding domains that bind short peptide sequences and are commonly found in signaling proteins. The SH2 domain recognizes peptide sequences with phosphorylated tyrosine residues, which are often indicative of protein activation. SH2 domains play a key role in growth factor receptor signaling, during which ligand-mediated receptor phosphorylation at tyrosine residues recruits downstream effectors that recognize these residues via their SH2 domains. The SH3 domain usually recognizes proline-rich peptide sequences and is commonly used by kinases, phospholipases and GTPases to identify target proteins. Although both SH2 and SH3 domains generally bind to these motifs, specificity for distinct protein interactions is dictated by neighboring amino acid residues in the respective motif.

The result of two or more proteins that interact with a specific functional objective can be demonstrated in several different ways. The measurable effects of protein interactions have been outlined as follows:

  • Alter the kinetic properties of enzymes, which may be the result of subtle changes in substrate binding or allosteric effects
  • Allow for substrate channeling by moving a substrate between domains or subunits, resulting ultimately in an intended end product
  • Create a new binding site, typically for small effector molecules
  • Inactivate or destroy a protein
  • Change the specificity of a protein for its substrate through the interaction with different binding partners, e.g., demonstrate a new function that neither protein can exhibit alone
  • Serve a regulatory role in either an upstream or a downstream event

Usually a combination of techniques is necessary to validate, characterize and confirm protein interactions. Previously unknown proteins may be discovered by their association with one or more proteins that are known. Protein interaction analysis may also uncover unique, unforeseen functional roles for well-known proteins. The discovery or verification of an interaction is the first step on the road to understanding where, how and under what conditions these proteins interact in vivo and the functional implications of these interactions.

While the various methods and approaches to studying protein–protein interactions are too numerous to describe here, the table below and the remainder of this section focuses on common methods to analyze protein–protein interactions and the types of interactions that can be studies using each method. In summary, stable protein–protein interactions are easiest to isolate by physical methods like co-immunoprecipitation and pull-down assays because the protein complex does not disassemble over time. Weak or transient interactions can be identified using these methods by first covalently crosslinking the proteins to freeze the interaction during the co-IP or pull-down. Alternatively, crosslinking, along with label transfer and far–western blot analysis, can be performed independent of other methods to identify protein–protein interactions.

Common methods to analyze the various types of protein interactions

MethodProtein–protein interactions
Co-immunoprecipitation (co-IP)Stable or strong
Pull-down assayStable or strong
Crosslinking protein interaction analysisTransient or weak
Label transfer protein interaction analysisTransient or weak
Far–western blot analysisModerately stable

Co-immunoprecipitation (co-IP) is a popular technique for protein interaction discovery. Co-IP is conducted in essentially the same manner as an immunoprecipitation (IP) of a single protein, except that the target protein precipitated by the antibody, also called the "bait", is used to co-precipitate a binding partner/protein complex, or "prey", from a lysate. Essentially, the interacting protein is bound to the target antigen, which is bound by the antibody that is immobilized to the support. Immunoprecipitated proteins and their binding partners are commonly detected by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) and western blot analysis. The assumption that is usually made when associated proteins are co-precipitated is that these proteins are related to the function of the target antigen at the cellular level. This is only an assumption, however, that is subject to further verification.

Co-immunoprecipitation of cyclin B and Cdk1. The Thermo Scientific Pierce Protein A/G Magnetic Beads bind to Cdk1 antibody complexed with Cdk1. Cyclin B is bound to the Cdk1, and is captured along with its binding partner.


Characteristic structural features

The structures of SET-domain proteins that are currently known include: the crystal structures of two members of the SUV39 family, Neurospora crassa Decrease in DNA methylation 5 (DIM-5) [15, 16] and S. pombe CLR4 [17] four structures of human SET7/9 in various configurations [18–21] a nuclear magnetic resonance (NMR) structure of a viral protein that contains only the SET domain [22] and a structure of the non-histone protein methyltransferase Rubisco LSMT, an unclassified member of the superfamily [23, 24]. These structures revealed that the SET domain forms a novel β fold not seen in any other previously characterized AdoMet-dependent methyltransferases (reviewed in [25]). The fold has a series of curved β strands forming several small sheets, packed together with pre-SET (or N-SET) and post-SET (or C-SET) domains or regions (Figure 3). The pre-SET domain of SUV39-family proteins (see Table 2) contains nine invariant cysteine residues that are grouped into two segments of five and four cysteines separated by various numbers of amino acids (CXCX5CX4CXC-XN-CX3CXCX3C, where N is 46 in DIM-5 and 28 in CLR4). The nine cysteines of the pre-SET domain of DIM-5 coordinate three zinc ions to form an equilateral triangular cluster, Zn3Cys9 (Figure 3a). The SET domain, which may have evolved through the duplication of a three-stranded unit [26], is folded in all the solved structures into several small β sheets surrounding a knot-like structure by threading of the carboxyl terminus through an opening of a short loop formed by a preceding stretch of the sequence (Figure 3). This remarkable 'pseudoknot' fold brings together the two most-conserved sequence motifs of the SET domain (RFINHXCXPN and ELXFDY see Figure 1) to form an active site in a location immediately next to the pocket where the methyl donor binds and to the peptide-binding cleft.

Representative examples of SET-domain-containing structures. (a) Neurospora crassa DIM-5 (Protein DataBank (PDB) code 1PEG.pdb) (b) human SET7/9 (1O9S.pdb). The pre-SET, SET, and post-SET domains in DIM-5 and the N-SET, SET, and C-SET domains in SET7/9 are indicated. The pseudoknot formed by two conserved SET motifs and the bound histone H3 peptide are also illustrated. The reaction byproduct AdoHcy is in stick representation and the zinc ions are shown as balls. N, amino terminus C, carboxyl terminus.

The post-SET region of DIM-5 contains three conserved cysteine residues, arranged CXCX4C, that are essential for its histone lysine methyltransferase activity [15]. The structure of DIM-5 in a ternary complex with an H3 K9 peptide and AdoHcy [16] reveals that, as expected from their arrangement, these three post-SET-domain cysteines coordinate a zinc ion tetrahedrally together with cysteine 244 of the SET-domain signature motif RFINHXCXPN in the pseudoknot near the active site (Figure 3a). Consequently, a narrow channel is formed to accommodate the side chain of the target lysine. Three ternary structures - SET7/9 in complex with a peptide containing histone H3 K4 [21], DIM-5 in complex with a histone H3 K9 peptide [16], and Rubisco LSMT in complex with a free lysine [24] - reveal that the target lysine is inserted into a narrow channel so that the target nitrogen would be in close proximity to the methyl donor AdoMet at the opposite end of the channel.

Close examination of the region carboxy-terminal to the SET domain in many proteins, including members of the SUV39, SET1, and SET2 families, suggests that the post-SET-domain metal center observed in DIM-5 is universal among all those members of the superfamily that have the cysteine-rich post-SET domain. For almost all SET-domain proteins, there appears to be an absolute correlation between the presence of the post-SET domain and a cysteine corresponding to Cys244 of DIM-5 near the active site. Comparison of DIM-5 with SET7/9 [19, 21] and the Rubisco LSMT [23, 24], two SET-domain proteins that do not have Cys-rich pre-SET and post-SET domains, reveals a remarkable example of convergent evolution. In particular, as in DIM-5, these two enzymes rely on residues carboxy-terminal to the SET domain for the formation of lysine channel, but they do so by packing of an α helix, rather than a metal center, onto the active site.


1. Introduction

Consistent protein nomenclature is indispensable for communication, literature searching and entry retrieval. A good protein name is one which is unique, unambiguous, can be attributed to orthologs from other species and follows official gene nomenclature where applicable. The process of associating a name with a protein sequence has various components: sequence function identification/prediction, choosing a name and applying formatting. This document provides guidelines on naming choices and universal formatting. This does not include best practices on methods to be used for sequence function identification/prediction.


The Helix-Turn-Helix Motif Is One of the Simplest and Most Common DNA-binding Motifs

The first DNA-binding protein motif to be recognized was the helix-turn-helix. Originally identified in bacterial proteins, this motif has since been found in hundreds of DNA-binding proteins from both eucaryotes and procaryotes. It is constructed from two α helices connected by a short extended chain of amino acids, which constitutes the “turn” (Figure 7-13). The two helices are held at a fixed angle, primarily through interactions between the two helices. The more C-terminal helix is called the recognition helix because it fits into the major groove of DNA its amino acid side chains, which differ from protein to protein, play an important part in recognizing the specific DNA sequence to which the protein binds.

Figure 7-13

The DNA-binding helix-turn-helix motif. The motif is shown in (A), where each white circle denotes the central carbon of an amino acid. The C-terminal α helix (red) is called the recognition helix because it participates in sequence-specific recognition (more. )

Outside the helix-turn-helix region, the structure of the various proteins that contain this motif can vary enormously (Figure 7-14). Thus each protein “presents” its helix-turn-helix motif to the DNA in a unique way, a feature thought to enhance the versatility of the helix-turn-helix motif by increasing the number of DNA sequences that the motif can be used to recognize. Moreover, in most of these proteins, parts of the polypeptide chain outside the helix-turn-helix domain also make important contacts with the DNA, helping to fine-tune the interaction.

Figure 7-14

Some helix-turn-helix DNA-binding proteins. All of the proteins bind DNA as dimers in which the two copies of the recognition helix (red cylinder) are separated by exactly one turn of the DNA helix (3.4 nm). The other helix of the helix-turn-helix motif (more. )

The group of helix-turn-helix proteins shown in Figure 7-14 demonstrates a feature that is common to many sequence-specific DNA-binding proteins. They bind as symmetric dimers to DNA sequences that are composed of two very similar “half-sites,” which are also arranged symmetrically (Figure 7-15). This arrangement allows each protein monomer to make a nearly identical set of contacts and enormously increases the binding affinity: as a first approximation, doubling the number of contacts doubles the free energy of the interaction and thereby squares the affinity constant.

Figure 7-15

A specific DNA sequence recognized by the bacteriophage lambda Cro protein. The nucleotides labeled in green in this sequence are arranged symmetrically, allowing each half of the DNA site to be recognized in the same way by each protein monomer, also (more. )


Mammalian DNA Methyltransferase Structural Themes

The SRA Domain of UHRF1 Flips 5-Methylcytosine Out of the DNA Helix

An accessory protein UHRF1 (ubiquitin-like, containing PHD and RING finger domains 1) ( Figure 4(a) ) targets Dnmt1 to hemimethylated replication forks (and presumably repair sites). The crystal structure of the SET and RING associated (SRA) domain of UHRF1 in complex with DNA containing a hemimethylated CpG site was determined. They reveal that the SRA domain flips the 5-methylcytosine (5mC) completely out of the DNA helix ( Figure 4(b) ) and is positioned in a binding pocket. The structure also suggests an explanation for the preference for hemimethylated sites. In the major groove side, a backbone carbonyl oxygen is close to the C5 ring carbon of the unmethylated cytosine, forming a C = O … H-C hydrogen bond. The addition of a methyl group to C5 of the unmethylated cytosine would cause a steric clash between the methyl group and SRA.

Figure 4 . UHRF1. (a) Schematic representation of UHRF1 – a multi-domain protein. (b) Structure of SRA-DNA complex. The 5mC flips out and is bound in a cage-like pocket involving hydrogen bonding and van der Waals interactions (inserted).

DNA methyltransferases or DNA repair enzymes use base flipping to gain access to a DNA base to perform chemistry on it, but the SRA domain probably uses base flipping to increase its protein–DNA interface and to prevent the SRA domain from linear diffusion away from the site on the DNA. This may be particularly important for the SRA domain, as its recognition sequence is only two base pairs. We suggest that the SRA–DNA interaction (through recognition and flipping of the 5mC) serves as an anchor to keep UHRF1 at hemimethylated CpG site where it recruits Dnmt1 for maintenance methylation, and perhaps other proteins such as DNA repair enzymes for mismatch repair. The 5mC base flipping by the SRA domain might also provide a more general mechanism to distinguish the methylated parental strand from the unmethylated daughter strand, an ability particularly important for mismatch repair if an error occurs during DNA replication. Supporting this hypothesis, the expression of UHRF1 is deregulated in cancer cells, and mouse UHRF1-null cells are more sensitive to DNA-damaging agents and DNA replication arrest.


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