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Taylor, J.‐S., Brockie, I. R., and O’Day, C. L. (1987). A building block for the sequence‐ specific introduction of cis‐syn thymine dimers into oligonucleotides: Solid‐phase synthesis of TpT{c,s}pTpT. J. Am. Chem. Soc. 109, 6735–6742. Todo, T., Kim, S.‐T., Hitomi, K., Otoshi, E., Inui, T., Morioka, H., Kobayashi, H., Ohtsuka, E., Toh, H., and Ikenaga, M. (1997). Flavin adenine dinucletide as a chromophore of the Xenopus (6‐4) photolyase. Nucleic Acids Res. 25, 764–768. Todo, T., Ryo, H., Yamamoto, K., Toh, H., Inui, T., Ayaki, H., Nomura, T., and Ikenaga, M. (1996). Similarity among the Drosophila (6‐4) photolyase, a human photolyase homolog, and the DNA photolyase‐blue‐light photoreceptor family. Science 272, 109–112. Todo, T., Takemori, H., Ryo, H., Ihara, M., Matsunaga, T., Nikaido, O., Sato, K., and Nomura, T. (1993). A new photoreactivating enzyme that specifically repairs ultraviolet light‐induced (6‐4)photoproducts. Nature 361, 371–374. Vande Berg, B. J., and Sancar, G. B. (1998). Evidence for dinucleotide flipping by DNA photolyase. J. Biol. Chem. 273, 20276–20284. Witmer, M. R., Altmann, E., Young, H., Begley, T. P., and Sancar, A. (1989). Mechanistic studies on DNA photolyase. 1. Secondary deuterium isotope effects on the cleavage of 20 ‐deoxyuridine dinucleotide photodimers. J. Am. Chem. Soc. 111, 9264–9265. Wulff, D. L., and Rupert, C. S. (1962). Disappearance of thymine photodimer in ultraviolet irradiated DNA upon treatment with a photoreactivating enzyme from baker’s yeast. Biochem. Biophys. Res. Commun. 7, 237–240. Zhao, X., Liu, J., Hsu, D. S., Zhao, S., Taylor, J.‐S., and Sancar, A. (1997). Reaction mechanism of (6‐4) photolyase. J. Biol. Chem. 272, 32580–32590.

[10] Genetic and In Vitro Assays of DNA Deamination By HEATHER A. COKER, HUGH D. MORGAN , and SVEND K. PETERSEN‐MAHRT Abstract

The DNA deaminase family encompasses enzymes that have been highly conserved throughout vertebrate evolution and which display wide‐ranging positive effects upon innate and adaptive immune system and development. Activation‐induced cytidine deaminase was identified as a DNA mutator after its necessity in the successful development of high‐ affinity B cells via somatic hypermutation, class switch recombination, and gene conversion was determined. APOBEC3 exhibits the ability to deaminate retroviral first strand cDNA in a variety of viral infections, including HIV and hepatitis. Recent work has highlighted the potential importance of activation‐induced cytidine deaminase (AID) and APOBEC1 in epigenetic reprogramming, and also the role that AID and the APOBECs may

METHODS IN ENZYMOLOGY, VOL. 408 Copyright 2006, Elsevier Inc. All rights reserved.

0076-6879/06 $35.00 DOI: 10.1016/S0076-6879(06)08010-4

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have in the development of cancer. In addition to the known activities of these members of the protein family, there are still other deaminases, such as APOBEC2, whose targets and functions are as yet unknown. This chapter provides the details of two assays that have proved to be invaluable in elucidating the exact specificities of deaminases both in vitro and in Escherichia coli. The application of these assays to future studies of the deaminase family will provide an indispensible tool in determining the potentially diverse functions of the remainder of this family of enzymes. Introduction

Protein‐directed DNA deamination is an exciting new field in molecular biology: whereas we have been aware of the occurrence of spontaneous deamination of cytosine for some time, identification of proteins that deaminate cytosine to form uracil in DNA has only occurred in recent years (Harris et al., 2002; Petersen‐Mahrt et al., 2002). Evolutionarily the most ancestral, activation‐induced cytidine deaminase (AID) is required to form a functional immune system (Conticello et al., 2005; Muramatsu et al., 2000). Subsequently, other members of the family of DNA deaminases have been shown to be important for the innate immune system, in addition to influencing the spread of retrotransposable elements (Esnault et al., 2005; Sheehy et al., 2002). As deaminases can introduce mutations in DNA, it is also perhaps not surprising that they are now being implicated in the development of cancer. More recently, the expression of AID during early embryogenesis has implicated a role of DNA deaminases in the reprogramming of the epigenetic status of the organism (Morgan et al., 2004). Due to their emerging importance in the fields of immunology, virology, cancer, and epigenetics, it is vital that one has efficient tools with which to study deaminase activity. In the past, we have developed both an Escherichia coli and an in vitro‐based assay to determine the broad activity, as well as the fine specificity, of these enzymes. This chapter describes in detail how to carry out both such assays. Escherichia coli Assay of Deamination

Assays of DNA deaminase activity in E. coli predate in vitro assays requiring purified proteins. The E. coli assay exploits the ability of the deaminase to mutate DNA bases of the endogenous E. coli genome, scored using particular genes as selectable markers. Verifying the exact nature of the mutation introduced into that gene can provide further additional

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information on the deaminase. Suppressing DNA repair systems in E. coli can also lend support to the action of the deaminase, as may modification of the assay to incorporate exogenous factors. Mutation Creates a Selectable Marker There are several genes that, when mutated, generate positive selectable markers (please see Miller, 1972). Due to the possible intrinsic specificities or preferences of a deaminase, it may mutate only a subset of the sites available in a particular gene. Alternatively, a deaminase may not mutate any sites in a particular marker so more than one selectable marker may need to be investigated. It is also possible to modify the system to such an extent as to allow for selection and mutation on an exogenous plasmid (Beale et al., 2004; Pham et al., 2003; Ramiro et al., 2003; Sohail et al., 2003). This chapter focuses on the E. coli RNA polymerase gene (rpoB) as a marker for deaminase‐mediated mutations, using rifampicin (rif) selection. Deaminase cDNA in an Expression Vector The deaminase cDNA is transferred to an expression vector containing an inducible promoter. Many such vectors exist; the classic combination of a tac promoter and lacIq repressor is combined in the pTrc99A vector (ampR, ColE1 origin of replication). Its multiple cloning site allows the required deaminase cDNA to be inserted downstream of the promoter and other necessary bacterial transcription and translation elements. Some IPTG inducible systems exhibit low levels of expression in the absence of isopropyl‐ ‐D‐thiogalactoside (IPTG), even with lac operon repression. This may be minimized by the addition of 0.2% (w/v) glucose to solid or liquid media. Bacterial Strain Choice and Relevant Repair Pathways The product of cytosine deamination, uracil, is predominantly removed via the uracil DNA glycosylase (UDG/ung)‐dependent base excision repair pathway. The presence of this pathway acting on uracil may therefore have the consequence of affecting the number of C–T transitions observed in the assay. There is, however, the opportunity for assaying DNA deaminase activity in E. coli strains lacking ung (Harris et al., 2002; Petersen‐Mahrt et al., 2002). For example, activity may be assayed in KL16 E. coli compared to BW310 (lacking ung, but otherwise considered the same genotype as KL16). Evidence that DNA deaminase activity is more severe

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in the absence of ung supports deaminase action on cytidine in DNA. This type of genetic determinism can also be useful when analyzing other potential DNA mutators, where the E. coli assay may be adapted for the analysis of mutations derived from noncytosine‐based lesions, using strains with deletions in alternative repair pathways (such as nfi‐1 to detect dA deamination (Guo et al., 1997)). Expression of Additional Proteins Other activities or factors may be required for deaminase activity that are not normally present in E. coli, and the ability to simultaneously express more than one protein in E. coli allows one to develop an assay that can incorporate these factors. The second factor is introduced most easily using a second expression vector, with both vectors then being independently selected and their expression verified. In order to assay deaminase activity on methylcytosine, it was necessary to introduce methylcytosine at the potential target sites (Morgan et al., 2004). An inducible expression cassette of the SssI DNA methyltransferase (which methylates CpG residues in a similar fashion to mammalian methylation), generously made available by New England Biolabs, was subcloned into part of pACYC177 containing the p15A ori (compatible with the ColE1 ori of pTrc99A) and kanamycin resistance (Morgan et al., 2004). Mammalian‐ like CpG methylation is not tolerated in strains of bacteria containing the Mcr/Mrr restriction system so the introduction of SssI requires using a strain lacking both restriction systems (e.g., ER1821 [New England Biolabs]). We established that SssI was functioning by methylation‐sensitive restriction enzyme analysis of plasmids in E. coli: the plasmid was harvested from the SssI containing E. coli and digested with a methylation‐ sensitive restriction enzyme. Transformation of Mcr
/Mrr
E. coli with this digested plasmid (with undigested plasmid as a control) then indicated the relative extent of plasmid methylation by SssI. Electrotransformation of Competent E. coli Many E. coli strains suited to protein expression, or possessing other characteristics required by the assay, may not be well suited to chemical transformation, but even two plasmids can be introduced into most strains using electrotransformation (ensure that the DNA is in a low salt solution). It is preferable to transform both plasmids at once to minimize the drift that makes the assay less reproducible following two separate rounds of transformation. At this stage, glucose should be present to prevent

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induction. Overnight incubation of these plates at less than 37 (e.g., 30–32 ) ensures continuous, equivalent growth. Bacterial Cultures Induced to Express Protein A general outline of the E. coli mutation screen is provided in Fig. 1. Colonies from the plates (as described earlier) are used to set up individual cultures for the induction of protein expression. The frequency of deamination affects the number of cells acquiring resistance, as does the number of cell divisions after the mutation arises. Before the cultures reach saturation, the number of cells in the starting culture should be at a low titer and consistently similar between the different samples. To avoid potential effects of the deaminase (or other factor) on the growth rate of the cells, counting the number of viable cells at the end of the culture (and in some instances during growth) is vital. However, if growth retardation is so severe that the viable count variability among the experimental groups is too extreme, then the use of an 8‐h starter culture (without induction, but also without glucose, which would inhibit subsequent induction) followed by an overnight induced culture may be required. Some laboratories have even been able to use a 90‐min induction protocol for the mutation assay (Martomo et al., 2005). Eight to 12 cultures per condition are generally a good experimental setup. For each culture, an entire colony is transferred to a 1.5‐ml microcentrifuge tube containing 1 ml of YT or LB media and vortexed. This is then diluted 104‐fold, and 10 l is used to inoculate 3 ml of media containing 1 mM IPTG (for induction) and the appropriate plasmid selection antibiotics. If a starter culture is used, then less dilution is warranted: a 1‐ml starter culture is inoculated for 8 h, followed by the addition of 2 ml IPTG‐containing media. Cultures are incubated at 37 for 16 h with vigorous shaking. For optimal activity, both time and temperature (e.g., 18 for deaminases from cold water fish [Conticello et al., 2005]) can be varied. Counting Viable Cells and Mutated Cells Mutation frequency is determined by establishing the number of rifR colonies vs the number of viable colonies per milliliter. To determine the number of viable cells in the culture, around 50–200 l of a 106 dilution is plated on LB plates (with or without plasmid antibiotic selection). It can be sufficient to count only six to eight plates per condition to obtain an accurate viable count. To determine the number of cells acquiring

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FIG. 1. Schematic of the E. coli mutation screen. An outline of the E. coli mutation assay using a mutable gene (rpoB) as a selection marker for deaminase activity (as described in detail in text).

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mutations that confer resistance, between 1 and 1000 l of induced culture is plated onto LB low salt plates containing rifampicin (100 g/ml), ensuring that plates containing rifampicin are protected from light during storage. One can use the same spreader to first spread a diluted viable sample and then the corresponding undiluted original culture without sterilization. For all spreading, the plated volume should be between 50 and 300 l, potentially requiring dilution or concentration (pulse spin) of the induced culture. Optimal counting accuracy, after an overnight incubation at 37 , is achieved with 100–200 colonies per plate. For rifampicin, a further 8‐ to 16‐h room temperature incubation helps visualize smaller colonies. Colonies should always be counted in an identical way, with respect to time of incubation and size of colony. The mutation frequency under each condition can be determined from the median colony number of rifR cells per viable cell; this minimizes the effect of an early mutation in a culture resulting in ‘‘jack‐pot’’ plate counts. Because the possibility exists that the protein of interest is toxic, one should ensure that the viable count on LB, and LB containing the plasmid selection antibiotic, is equivalent. Scoring on the antibiotic ensures that mutations are detected per cell harboring the plasmid. Determining the Sequence Context of Mutations To investigate the qualitative nature of mutations introduced into the selectable gene, colonies are picked from plates containing the selecting agent (e.g., rifampicin). To avoid the ‘‘jackpot’’ effect, only two randomly chosen colonies from each plate are analyzed. The two colonies are respotted onto a master plate also containing rifampicin to ensure that the colony is truly rifR and also to reduce the number of plates to be stored. After overnight growth the chosen colonies can be analyzed by conventional sequencing after polymerase chain reaction (PCR) amplification of the implicated gene, e.g., rpoB (Beale et al., 2004; Conticello et al., 2005; Harris et al., 2002; Petersen‐Mahrt et al., 2002). For amplification of rpoB, the oligonucleotides rpopcrf (50 ‐TTG GCG AAA TGG CGG AAA ACC‐ 30 ) and rpopcrr (50 ‐CAC CGA CGG ATA CCA CCT GCT G‐30 ) are used: a 25‐l PCR is performed on a 1/3000 dilution of the colony. The product is purified and resuspended in 125 l of TE; 5 l is then used for sequencing with rposeqf (50 ‐CGT TGG CCT GGT ACG TGT AGA GCG‐30 ) and rposeqr (50 ‐GGC AAG TTA CCA GGT CTT CTA CG‐30 ). If only a single site within the target gene is of interest, one can use a simple PCR reaction for the analysis (Morgan et al., 2004). A primer is chosen so that the 30 nucleotide matches only the mutation of interest. In practice, it is necessary to establish stringent PCR conditions using E. coli

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genomic DNA known to contain the mutation. A control PCR should also be carried out to ensure that DNA was present in the original template solution. Both PCRs can be carried out in one reaction tube as a duplex PCR. A crude template solution can be made by transferring a portion of the colony to be analyzed directly into 50 l of Milli‐Q H2O, vortexing, and then using 3 l of this as template DNA, along with a 95 denaturing and lysing step for 3 min during the initial PCR cycle. Mutations arising in the selectable marker can in themselves be informative. The spectrum of transitions and transversions should change in the presence of a cytidine deaminase. This increase should be even more extreme with the disruption of DNA repair pathways (Petersen‐Mahrt et al., 2002). Sites chosen for mutation, compared to those available for mutation, may also give an indication of the sequence specificity of the deaminase. In Vitro Assay of Deamination

The in vitro assay requires the use of deaminases that can be produced and purified to a fairly high, pure yield (microgram quantities at least). Even though a number of different methods for producing DNA deaminases have been introduced in the literature (particularly that using baculovirus) (Beale et al., 2004; Bransteitter et al., 2003; Chaudhuri et al., 2003; Muramatsu et al., 1999; Petersen‐Mahrt and Neuberger, 2003; Ramiro et al., 2003; Yu et al., 2004), this chapter focuses on the production of tagged DNA deaminases in E. coli, with this having the added advantage of enabling concomitant analysis of a mutator phenotype in the assay described earlier. The following protocols have been used to produce and analyze functional human AID, APOBEC1, APOBEC3G, and also chicken AID, although this chapter concentrates on the production and activity of human AID. Preparation of Deaminases for Use in the In Vitro Assay We habitually use a C‐terminal his‐tagged human AID plasmid under the control of a T7 promotor with kanamycin resistance. We have also successfully modified this procedure to enable protein to be prepared from up to 20 liters of E. coli culture, with protein generated and purified with a N‐terminal MBP tag. In the past we also have used untagged protein, purified via ion‐exchange chromatography (Petersen‐Mahrt and Neuberger, 2003). This allowed us to separate single‐stranded DNase activity from the DNA deamination enzymes. To investigate DNA deamination from a crude extract on a single‐stranded substrate, we recommend using a random‐labeled, heat‐denatured DNA. As shown in

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the past, using random‐labeled DNA, the assay is not dependent on the absence of DNase activity (Petersen‐Mahrt and Neuberger, 2003), but due to the number of complex interactions and variation in substrate, the presence of DNase activity does not lend the assay to the study of the enzyme in such a controlled fashion as the in vitro oligonucleotide assay. Growth. Transform E. coli of the C41(DE3) strain (Miroux and Walker, 1996) with the protein expression plasmid and plate onto an appropriate selective LB plate (all subsequent bacterial growth takes place using nonselective media). Once the transformed bacteria have grown overnight at 37 , take one colony and resuspend thoroughly in 1 ml 2 TY media. Use this to further inoculate 30 ml 2 TY, which is then grown, shaking at 37 until reaching an OD600 of 0.4. At this stage, 10 ml of the culture is used to inoculate each of three, 2‐liter flasks, each containing 1 liter of 2 TY. These are grown, shaking at 37 , until reaching an OD600 of between 1.0 and 1.5. The flasks should then be immersed in iced H2O for 15 min prior to induction with a final concentration of 0.4 mM IPTG. In addition, at this stage add ZnCl2 to a final concentration of 10 M. Importantly, holes are made in the foil lids of the flasks to ensure that sufficient air exchange is able to take place during protein production. The cultures should then be incubated, shaking at 250 rpm at 16 for 16 h. Extraction. The 3 liters of bacteria is harvested by centrifugation at 3500g, 10 min at 4 , the supernatant is removed carefully but thoroughly, and the pellets are resuspended each in approximately 100 ml of ice‐cold H2O before combining. Bacteria are again centrifuged at 3500g, for 10 min at 4 , before the supernatant is immediately removed and the pellet resuspended in 100 ml of extraction buffer (20 mM MES [pH 6.0], 100 mM NaCl, 50 mM KCl, 5 mM ‐mercaptoethanol, 1.6 mM CHAPS, 300 mM L‐arginine HCl, filtered and adjusted to pH 6) along with six complete EDTA‐free protease inhibitor cocktail tablets (Roche) and 150 mg RNase A (Sigma), which has been heated previously to 80 for 10 min in extraction buffer. Keeping the supernatant on ice, sonicate until the cells are completely disrupted (this needs to be optimized depending on the machine used, but we have found that up to 25 min in total, with 15‐s pulses at an amplitude of 50, is sufficient). The suspension is then centrifuged at 100,000g, 30 min at 4 , and the supernatant is filtered using a prechilled 0.22‐m Stericup (Millipore). In an initial study we determined that greatest protein solubility was obtained using this low pH 6 extraction buffer. Furthermore, adding 300 mM L‐arginine HCl, as detailed earlier, appeared to increase the stability of the protein. The addition of RNase A both at this stage and

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later in the assay itself appears to increase the activity of the protein (Bransteitter et al., 2003). Purification. Transfer the filtered supernatant into two 50‐ml Falcon tubes. Add 1 ml of Ni‐NTA beads (Qiagen), prewashed in extraction buffer, to each tube. Incubate the beads with protein at 4 , rotating for 90 min. Centrifuge briefly to pellet beads (1000g, 1 min at 4 ) before loading the beads onto a Poly‐Prep chromatography column (Bio‐Rad) fitted with a 21‐gauge needle and prewashed with extraction buffer at 4 . Wash the beaded column at 4 with 10 ml of 30 mM imidazole and then elute the protein at 4 with 10 ml of 150 mM imidazole. Concentrate the eluate using a centrifugal concentrator (Vivaspin) with a Mr 10,000 cutoff (or appropriate for your protein of interest). Follow the manufacturer’s instructions to concentrate the eluate to 150 l or a suitable concentration. We have found that deaminases lose activity when stored at ‐80 and, over time, even when stored in liquid nitrogen. We would recommend that the concentrated protein be snap‐frozen in small aliquots and stored in liquid nitrogen for use within 10 months. Assay of Deaminase Activity Oligonucleotides. Using an enzyme assay that is reliant on the presence of free cytosine or cytidine is not only inaccurate, but also does not reflect the true activity of the enzyme. This is predominantly due to the possible contamination of the protein preparation by E. coli cytidine deaminases (Beale et al., 2004; Petersen‐Mahrt and Neuberger, 2003). The basis of the in vitro assay is the deamination of cytosine within labeled oligonucleotides and then the subsequent readout of deamination of those oligonucleotides by cleavage at the sites of deamination. There are no real restrictions on the design of the oligonucleotides; the design simply reflects the particular questions that you wish to answer of the deaminase. The oligonucleotide may contain single or multiple potential target sites. Multiple target sites allow for analysis of the fine specificity of deaminase activity; we found this to be of particular use not only when comparing the preference of AID and APOBEC1 for cytosine compared to methylcytosine but also in determining the exact sequence motif preferences of the deaminases (Morgan et al., 2004). We have found 40‐mers to be the most practical length of oligonucleotide if assaying two sites simultaneously, and it is important that the oligonucleotides are HPLC purified to ensure that any prematurely truncated oligonucleotides are removed. Resuspension of the oligonucleotides in TE as usual is perfectly sufficient. An oligonucleotide containing uracil is a useful control for the efficiency/UDG dependence of the reactions in each experiment.

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The most convenient in vitro assay that we have established depends on 50 biotinylation of the oligonucleotides to enable subsequent purification by streptavidin‐coated magnetic beads. The 30 end of the oligonucleotide is labeled for visualization; in the past, we have used 32P‐labeled oligonucleotides, although we now use 30 ‐fluorescein‐labeled oligonucleotides most frequently. We have also used other fluorescent dyes, such as JOE and TAMRA (PerkinElmer Life Sciences Inc.). While oligonucleotides can be purchased with these labels already attached, end labeling provides a simple means of labeling aliquots of an oligonucleotide with different dyes for the potential of a dual or multiple color readout (although it can be difficult to minimize sufficiently the bleed through of fluorescent emission between the laser channels). Oligonucleotide Deamination. The exact quantity of a deaminase needed for optimal activity in the in vitro assay needs to be determined for each protein preparation by conducting a pilot experiment titrating a range of concentrations. We have found, however, that protein made by our method is generally used at between 0.05 and 0.5 l. If a protein preparation is particularly good, it may therefore be necessary to dilute the protein further using extraction buffer, e.g., 1/10, and snap freeze again. Refreezing once, in this manner, does not appear to affect the deaminase activity. It is important to use a control protein in each experiment to ensure that spontaneous deamination of the oligonucleotides does not occur during the procedure. Two microliters of reaction buffer (40 mM Tris pH 8, 40 mM KCl, 50 mM NaCl, 5 mM EDTA, 1 mM dithiothreitol, 10% glycerol), 2.5 mM EDTA, 5 g RNase A (Sigma) (heated previously to 80 for 10 min in 0.1 M Tris, pH 8), and 2.5 pmol 50 ‐biotinylated, 30 ‐FITC‐labeled oligonucleotide should be combined to give a final 10‐l reaction once the deaminase is added. The mix is then heated to 90 for 3 min before quenching on ice. (The reaction buffer keeps for up to 6 months if refrigerated, but the dithiothreitol should be replenished monthly.) The deaminase is thawed on ice and then the appropriate amount is added to the ice‐cold reaction mix for each sample in 0.5‐ml tubes. Each reaction should be incubated at 37 for 15 min for optimal activity, although we have previously tested other temperatures and incubation times, and it is worth noting that deaminases can still demonstrate (albeit reduced) activity at 18 (Conticello et al., 2005). Each reaction is terminated by adding 100 l of H2O and then heating at 90 for 4 min before placing on ice. Purification of Oligonucleotides. Use of biotin‐labeled oligonucleotides enables the use of streptavidin‐coated magnetic beads for purification of the oligonucleotides. We have modified the purification step to use

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streptavidin‐coated 96‐well plates (Roche) and also streptavidin‐coated magnetic beads in a 96‐well format, but from the point of view of reproducibility we have found that using individual tubes is best. Eight microliters of resuspended streptavidin‐magnetic bead slurry (Dynal) per reaction is washed twice in TEN buffer (10 mM EDTA, 50 mM Tris, pH 7.5, 1 M NaCl) before resuspending in 800 l TEN per reaction and adding to each protein sample and then transferring to a 1.5‐ ml microcentrifuge tube. The samples are mixed by rotating at room temperature for 15 min. Use of a Dynal microcentrifuge magnet enables the washing of the bead‐bound oligonucleotides: an incubation of 3 min on the magnet is sufficient. The supernatant is then carefully removed by pipette before removing the tube from the magnet and resuspending the beads in 750 l, prewarmed, 70 TEN. The bead‐bound oligonucleotides are then washed by rotating for 5 min at room temperature, before repeating the wash procedure in exactly the same manner with TEN, and then finally with 100 l room temperature TE, pH 8. The tubes are subject to a pulse spin in a benchtop microcentrifuge before placing on the magnet and removing the supernatant for the final time. Removal of Uracil from Oligonucleotides. Cleavage of the oligonucleotides at the site of deamination of cytosine to uracil provides the readout of AID activity. The bead‐bound oligonucleotides are resuspended in 10 l of UDG reaction mix (1 UDG buffer, excess UDG [1 l, 1 U, New England Biolabs]) at 37 for 1 h, mixing the samples gently after 30 min. Contaminant apyriminidic endonuclease within commercial sources of UDG means that a separate cleavage step of the UDG‐generated abasic site is not necessary. Twenty microliters of fuscin formamide [0.1% fuscin dye (Sigma), 10 mM Tris, pH 8, in formamide] should then be added to terminate each reaction. Use of TDG to Resolve a T:G or U:G Mismatch. The flexibility of the in vitro assay means that when we investigated deaminase activity on methylcytosine (Morgan et al., 2004), we adopted a different approach to generate a readout of deamination. The deamination of methylcytosine generates a thymine residue and so this event would not be recognized by UDG. However, the resulting T:G base pair is acted on by TDG. Indeed, as TDG recognizes both T:G and U:G mismatches, even an oligonucleotide designed to contain both cytosine and methylcytosine target sites may be processed using TDG and a reverse complementary oligonucleotide. After purification, the bead‐bound oligonucleotides are resuspended in TDG reaction mix, containing 1 buffer (Trevigen) and an excess (5 pmol) of the reverse complement oligonucleotide. Each sample is then heated to 90 for 3 min and the oligonucleotides are allowed to anneal by cooling over 30 min in a hot block to room temperature. An excess of TDG (1 l,

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FIG. 2. In vitro DNA deamination of FITC‐labeled oligonucleotides. Five FITC‐labeled oligonucleotides each containing different, multiple deamination target motifs were incubated with AID, annealed with their reverse complement oligonucleotide, and then cleaved at U:G or T:G mismatches with TDG (described in detail in text). Samples were then subject to 20% PAGE‐urea and visualized with a fluorescent scanner, enabling the relative efficiency of enzyme activity at each motif to be determined. This allows the fine specificity of deaminase activity to be analyzed; it is evident, for example, from lane 2, that the substrate AGCme (motif 1) is preferred to ATC (motif 2). C* indicates an unmethylated cytosine.

2 U, Trevigen) is then added to each tube (final reaction volume 10 l), and the samples are incubated at 47 for 1 h. Twenty microliters of fuscin formamide is added to stop each reaction. Visualization and Quantification of Deamination Assay. In order to visualize the results of the assay, each sample is heated at 95 for 3 min and then quenched on ice before being subject to PAGE–urea electrophoresis on a prerun 15–20% gel at 200 V (for 1–2 h depending on the length of the oligonucleotide and the number of cleavage sites). The fluorescent signal is then detected using fluorescent scanners such as the FLA‐5000 (Fuji) (see Fig. 2). Quantification of the scanned image enables detailed comparisons of the deaminase activity of the different samples, allowing percentage conversion, estimates of protein turnover (when knowledge of microgram of protein per reaction is determined by Coomassie/Western blot studies), and further kinetic studies to be carried out. Percentage conversion may be calculated by the following equation: % conversion ¼ pixal volume of cleaved product (minus background)/(pixal volume of cleaved product (minus background) þ pixal volume of uncleaved substrate (minus background))  100. Conclusion

This chapter detailed the two main assays used to determine much of our biochemical knowledge of DNA deaminase activity. The E. coli assay has been of particular use in enabling analysis of the activity of deaminases

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when their purification has not been possible and also in providing an indication of possible in vivo deaminase activity. In contrast, the in vitro oligonucleotide assay provides the opportunity to compare the activity resulting from different posttranslational modifications made to the deaminases and, although this chapter only detailed the growth of deaminases in E. coli, also the activity of deaminases grown in different cell types. Furthermore, using a single oligonucleotide with a defined sequence has been instrumental in enabling us to identify the exact substrate and specificities of different DNA deaminases, its flexibility resulting from the huge range of different oligonucleotides that can be designed to answer specific questions. In addition, the ability to detect different lesions within the oligonucleotides by varying between the use of UDG or TDG expands the potential of this assay even further, providing us with ample tools with which to explore this family of enzymes, whose importance for the survival and evolution of higher organisms is becoming more and more apparent. Acknowledgments We acknowledge the contributions of Ian Watts to the development of the oligonucleotide assay and Reuben Harris for introducing the powers of E. coli genetics. We apologize to any contributors to the field that we have inadvertently omitted in this review due to space limitations.

References Beale, R. C., Petersen‐Mahrt, S. K., Watt, I. N., Harris, R. S., Rada, C., and Neuberger, M. S. (2004). Comparison of the differential context‐dependence of DNA deamination by APOBEC enzymes: Correlation with mutation spectra in vivo. J. Mol. Biol. 337, 585–596. Bransteitter, R., Pham, P., Scharff, M. D., and Goodman, M. F. (2003). Activation‐induced cytidine deaminase deaminates deoxycytidine on single‐stranded DNA but requires the action of RNase. Proc. Natl. Acad. Sci. USA 100, 4102–4107. Chaudhuri, J., Tian, M., Khuong, C., Chua, K., Pinaud, E., and Alt, F. W. (2003). Transcription‐targeted DNA deamination by the AID antibody diversification enzyme. Nature 422, 726–730. Conticello, S. G., Thomas, C. J., Petersen‐Mahrt, S. K., and Neuberger, M. S. (2005). Evolution of the AID/APOBEC family of polynucleotide (deoxy)cytidine deaminases. Mol. Biol. Evol. 22, 367–377. Esnault, C., Heidmann, O., Delebecque, F., Dewannieux, M., Ribet, D., Hance, A. J., Heidmann, T., and Schwartz, O. (2005). APOBEC3G cytidine deaminase inhibits retrotransposition of endogenous retroviruses. Nature 433, 430–433. Guo, G., Ding, Y., and Weiss, B. (1997). nfi, the gene for endonuclease V in Escherichia coli K‐12. J. Bacteriol 179, 310–316. Harris, R. S., Petersen‐Mahrt, S. K., and Neuberger, M. S. (2002). RNA editing enzyme APOBEC1 and some of its homologs can act as DNA mutators. Mol. Cell 10, 1247–1253.

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