Nucleic acid determinants for selective deamination of DNA over RNA by activation-induced deaminase

Abstract

Activation-induced deaminase (AID), a member of the larger AID/APOBEC family, is the key catalyst in initiating antibody somatic hypermutation and class-switch recombination. The DNA deamination model accounting for AID’s functional role posits that AID deaminates genomic deoxycytosine bases within the immunoglobulin locus, activating downstream repair pathways that result in antibody maturation. Although this model is well supported, the molecular basis for AID’s selectivity for DNA over RNA remains an open and pressing question, reflecting a broader need to elucidate how AID/APOBEC enzymes engage their substrates. To address these questions, we have synthesized a series of chimeric nucleic acid substrates and characterized their reactivity with AID. These chimeric substrates feature targeted variations at the 2′-position of nucleotide sugars, allowing us to interrogate the steric and conformational basis for nucleic acid selectivity. We demonstrate that modifications to the target nucleotide can significantly alter AID’s reactivity. Strikingly, within a substrate that is otherwise DNA, a single RNA-like 2′-hydroxyl substitution at the target cytosine is sufficient to compromise deamination. Alternatively, modifications that favor a DNA-like conformation (or sugar pucker) are compatible with deamination. AID’s closely related homolog APOBEC1 is similarly sensitive to RNA-like substitutions at the target cytosine. Inversely, with unreactive 2′-fluoro-RNA substrates, AID’s deaminase activity was rescued by introducing a trinucleotide DNA patch spanning the target cytosine and two nucleotides upstream. These data suggest a role for nucleotide sugar pucker in explaining the molecular basis for AID’s DNA selectivity and, more generally, suggest how other nucleic acid-modifying enzymes may distinguish DNA from RNA.

In the life of the cell, enzymes must sort through a complex milieu of biomacromolecules to identify their proper substrates and perform their intended function. For enzymes that target nucleic acids, distinguishing DNA from RNA presents a notably critical challenge. Nucleic acid selectivity is particularly pertinent for activation-induced deaminase (AID), the key enzyme that governs antibody maturation via somatic hypermutation (SHM) and class-switch recombination (CSR). AID accomplishes these tasks by purposefully mutating cytosine to uracil within immunoglobulin genes. Generation of deoxyuracil activates downstream base excision, mismatch repair, and recombination pathways, ultimately enhancing antibody affinity for antigen or altering antibody isotype (1–3).

When AID was initially discovered, the closest known homolog was apolipoprotein B mRNA editing enzyme, catalytic polypeptide 1 (APOBEC1), an RNA deaminase that introduces a premature stop codon in the mRNA of apolipoprotein B (4). From this observation, it was assumed that AID was also an RNA deaminase; however, several lines of evidence have subsequently challenged this initial assumption in favor of a DNA deamination model. Early studies showed that overexpressed AID is capable of mutating the Escherichia coli genome and that mutations could be enhanced by inhibiting or eliminating uracil DNA glycosylase (UDG), a base excision repair enzyme that specifically targets uracil within DNA (5, 6). In mice and humans, UDG and the DNA mismatch repair enzyme Msh2 are required for SHM and CSR, providing physiological evidence for the significance of AID-generated deoxyuracil in antibody maturation (7–10). The evolving model of DNA deamination has been subsequently bolstered by in vitro biochemical studies of purified AID. The enzyme was shown to carry out deamination of single-stranded DNA oligonucleotides without observable activity on single-stranded RNA, double-stranded DNA, or RNA–DNA hybrids (11–14). Most recently, RNA sequencing has failed to demonstrate any AID-dependent RNA editing in activated B-cells (15), and AID-dependent accumulation of deoxyuracil has been demonstrated within the immunoglobulin locus (16). This direct observation of uracil in genomic DNA provides the strongest evidence to date in favor of the DNA deamination model, and this model is now widely accepted.

Despite a compelling body of evidence in favor of the DNA deamination model, several key questions remain unanswered regarding AID’s mechanism of action. AID’s promiscuous binding interactions with RNA (12, 13) demonstrate the lack of any molecular explanation for AID’s nucleic acid selectivity. In fact, purified AID requires RNase treatment before DNA deamination activity can be observed, suggesting that RNA can competitively bind in the enzyme’s nucleic acid binding site (12). A similar pattern of DNA deamination despite RNA binding is seen with APOBEC3 homologs of AID that contribute to restriction of retroviruses such as HIV. In this case, RNA binding appears critical for the appropriate packaging of these restriction factors within their retroviral targets (17–19). Thus, there remains the persistent and unanswered question of how AID can freely interact with both RNA and DNA yet retain catalytic specificity for DNA.

The lack of insight regarding AID’s nucleic acid selectivity is emblematic of a larger gap in understanding how AID and other APOBECs engage their nucleic acid substrates for deamination (20). Although X-ray crystallography and NMR solution studies have revealed the structures of at least five different APOBEC homologs—including the potentially genotoxic APOBEC3A and the catalytic domain of the anti-HIV factor APOBEC3G—no structure of any AID/APOBEC family member bound to nucleic acid has yet been reported (21). Biochemical studies have provided some indirect evidence that explains how AID/APOBECs interact with nucleic acid substrates. Specifically, regarding nucleobase interactions, multiple reports have identified an 11-amino-acid “hotspot recognition” loop within AID that targets deaminase activity to WRC trinucleotide repeats (W = A/T at −2 position, R = A/G at −1 position) (22–24). Beyond recognition of the hotspot nucleobases, interactions with the nucleic acid backbone have also been shown to influence DNA deamination with the HIV restriction factor APOBEC3G (25).

Given the established and emerging roles of AID and other APOBECs in adaptive and innate immunity, as well as DNA damage and oncogenesis (26–30), there is a pressing need to elucidate the mechanisms by which AID and its homologs specifically recognize and deaminate cytosine. In this study, we directly address the mechanisms of substrate recognition by interrogating the molecular basis of AID’s selectivity for DNA over RNA. We have found that AID’s deaminase activity is governed by the 2′-substituents of the nucleotide sugar at both the target cytosine and the two upstream nucleotides, localizing the key nucleic acid determinants of selectivity. Our results provide a mechanistic rationale for AID’s DNA selectivity, offer clarity for the mode of hotspot targeting and engagement, and provide insights into how other nucleic acid-modifying enzymes may specifically discern DNA from RNA.

Results

2′-Substituents of the Target Cytosine Regulate AID Deamination.

RNA is distinguished from DNA by the presence of the 2′-(R)-hydroxyl group in the sugar of the nucleotide-building blocks. Given that this hydroxyl group introduces a steric constraint and alters the conformational preferences of the nucleotide, we reasoned that this 2′-substituent of the target cytosine itself could be an important determinant of deamination selectivity. To test this hypothesis, we synthesized chimeric substrates that contain 2′-modifications in a single target cytosine embedded within an otherwise DNA backbone: DNA with a single 2′-deoxycytidine (D-dC), 2′-ribocytidine (D-rC), 2′-fluororibocytidine (D-frC), 2′-arabinocytidine (D-aC), or 2′-fluoroarabinocytidine (D-faC) (Fig. 1A).

Fig. 1.

The 2′-substitution of target cytosine is sufficient to disrupt AID deaminase activity. (A) Chimeric substrate design. Sequence and cytosine conformers (xC) incorporated at target position. (B) Restriction-endonuclease-based deamination assay. End-labeled substrates were incubated with AID, duplexed to a complementary strand, and incubated with the restriction endonuclease FspBI. FspBI cleaves cytosine-containing substrates but not uracil products. Assay validation with all substrates is shown in Fig. S1B. (C) Restriction-endonuclease–based assay of chimeric substrates in the presence of AID with a 1-h enzyme incubation. (D) Chimeric substrates were incubated with AID for various time points over the course of 1 h. Total product formation is graphed as a function time (n = 3 replicates; SD shown).

Traditional biochemical assays for deamination rely upon UDG for detection of reaction products. Because UDG does not recognize several nucleotide conformers used in our chimeric substrates, we developed an alternative assay to screen for deamination and found that DNA restriction endonucleases were well suited to the task (Fig. 1B). Specifically, the restriction endonuclease FspBI was surprisingly tolerant to the presence of a chimeric non-DNA base within its target and could accurately discriminate between a C:G match or a U:G mismatch within its recognition sequence (Fig. S1 A and B). Of note, the isoschizomer BfaI also tolerated these same chimeric substrates (Fig. S1 C and D), suggesting that other DNA endonucleases may be able to recognize chimeric non-DNA substrates.

As expected, a 1-h incubation of AID with the D-dC substrate yielded robust deamination (Fig. 1C). D-rC differs from D-dC only by the presence of a 2′-hydroxyl at the target cytosine; the presence of this single molecular substitution was sufficient to compromise deamination of the otherwise DNA substrate. Incubation of AID with D-frC yielded similarly negligible deamination, demonstrating that both 2′-hydroxyl and 2′-fluoro substitutions with (R)-stereochemistry impair deaminase activity. By contrast, AID more efficiently deaminated cytosine substrates when the 2′-hydroxyl and 2′-fluoro substituents were present in epimeric, or inverted, configurations in D-aC and D-faC. The striking sensitivity of AID to the addition of a single 2′-(R)-hydroxyl at the target cytosine was also apparent when we examined alternative hotspot or coldspot sequences, a DNA sequence derived from the mouse Sα switch region consensus sequence, or a more physiologically relevant dsDNA bubble substrate (Fig. S2) (31, 32). Notably, we used a truncated form of AID lacking its terminal exon (residues 181–198) in our experiments. This version of AID has previously been shown to retain identical sequence targeting and has increased enzymatic activity compared with the full-length enzyme (22), thereby increasing the dynamic range of the enzyme’s reactivity and permitting the study of less favored substrates. As expected, when we also assayed the chimeric substrates with full-length AID, the enzyme demonstrated similar preferences although overall enzymatic activity was diminished (Fig. S3).

To obtain a more rigorous comparison between substrates, we repeated our deaminase assays at multiple early time points for kinetic analysis and under extended-incubation conditions. The trend observed with the 1-h incubation was reflected in different initial rates of turnover at early time points (Fig. 1D). For a quantitative comparison of substrate reactivity, we further determined product formation as a function of enzyme concentration with extended 12-h incubations. We focused on a range where product formation was linearly dependent upon enzyme concentration and compared the overall substrate reactivity by normalization to those values obtained with D-dC (Table 1 and Fig. S4A). Based on the lower limits of detection for D-rC deamination (Fig. S4C), the D-dC substrate was deaminated over 500-fold more efficiently than the D-rC substrate, highlighting AID’s striking degree of sensitivity to the 2′-(R)-hydroxyl at the target cytosine. Comparing epimers of the same substituent (D-rC vs. D-aC for 2′-hydroxyl; D-frC vs. D-faC for 2′-fluoro), the arabinosyl epimers D-aC and D-faC were preferred by more than an order of magnitude.

Table 1.

AID deamination and binding of chimeric DNA substrates

Substrate	Deaminase activity, nM product per nM enzyme	Relative deaminase activity	Kd, nM
D-dC	1.130 ± 0.027	1.000	66 ± 3
D-rC	<0.003	<0.002	56 ± 4
D-frC	0.018 ± 0.003	0.016	72 ± 6
D-aC	0.103 ± 0.003	0.091	69 ± 10
D-faC	0.688 ± 0.017	0.609	65 ± 7

For each value, SD is shown (n = 3 replicates). Graph for derivation of deaminase activity is shown in Fig. S4A. Values for D-rC are based on assay detection limits (Fig. S4C). Graph of substrate binding is shown in Fig. S5B.

With a clear pattern of differential reactivity against chimeric DNA substrates, we wanted to determine whether these results reflected preferential catalysis or binding. Using fluorescence polarization, we determined that AID bound with a similar affinity across the series of substrates, suggesting that altered binding was not a sufficient explanation for selectivity (Table 1 and Fig. S5 A and B). Notably, because AID was purified as a recombinant fusion protein with maltose-binding protein (MBP), we examined binding with MBP alone and confirmed that substrate binding was specifically due to AID (Fig. S5C).

2′-Substituents of the Target Cytosine Regulate APOBEC1 Deamination.

The notion that AID could deaminate RNA originated from its homology to APOBEC1 (4). Despite this homology, AID acts on the immunoglobulin locus DNA, whereas APOBEC1 is physiologically known to target the apolipoprotein B mRNA. Given the similarities and contrasts between these two deaminases, we sought to determine APOBEC1’s sensitivity to 2′-substitution of the target cytosine. As with AID, we created a series of chimeric DNA substrates containing 2′-deoxycytidine, 2′-ribocytidine, and 2′-fluororibocytidine embedded within a sequence context preferred by APOBEC1 (A1-D-xC) (Table S1) (33). These substrates were incubated with APOBEC1 and evaluated using the FspBI restriction endonuclease-based assay for deamination.

We found that APOBEC1 readily deaminated the A1-D-dC substrate, in line with previous reports that have demonstrated the enzyme’s robust DNA deaminase activity (5, 13, 34). Notably, despite its physiological targeting of RNA, APOBEC1 deaminated A1-D-dC more efficiently than the A1-D-rC and A1-D-frC substrates, as reflected in the differential reactivity at early time points (Fig. 2 A and B). For a quantitative comparison of reactivity, as done for AID, we also determined product formation for each substrate as a function of APOBEC1 concentration with extended 12-h incubations and normalized it to the value obtained with A1-D-dC (Table 2 and Fig. S4B). APOBEC1 deaminated the A1-D-dC substrate 110-fold more efficiently than the A1-D-rC substrate and 20-fold more efficiently than the A1-D-frC substrate. As with AID, APOBEC1 bound all substrates with similar affinity, indicating that the differences in reactivity were not a function of altered binding (Table 2 and Fig. S5D).

Fig. 2.

The 2′-substitution of target cytosine disrupts APOBEC1 deaminase activity. (A) Restriction-endonuclease–based assay of chimeric substrates in the presence of APOBEC1 with a 1-h incubation. (B) Chimeric substrates were incubated with APOBEC1 for various time points over the course of 1 h. Total product formation is graphed as a function of time (n = 3 replicates; SD shown).

Table 2.

APOBEC1 deamination and binding of chimeric DNA substrates

Substrate	Deaminase activity, nM product per nM enzyme	Relative deaminase activity	Kd, nM
A1-D-dC	0.336 ± 0.019	1.000	57 ± 9
A1-D-rC	0.003 ± 0.001	0.009	44 ± 10
A1-D-frC	0.016 ± 0.001	0.048	45 ± 7

For each value, SD is shown (n = 3 replicates). Graph for derivation of deaminase activity is shown in Fig. S4B. Graph of substrate binding is shown in Fig. S5D.

Minimal DNA Requirements for AID Deamination.

Analysis of the chimeric DNA substrates demonstrated that RNA-like 2′-substitutions to the target cytosine are sufficient to compromise AID’s deaminase activity underlying AID’s specificity for DNA. To gain further insight into the molecular determinants of AID’s deamination activity, we wanted to determine whether the inverse were also true: Would removal of the 2'-substituent from the target cytosine of an RNA substrate rescue deamination?

To address this question, we synthesized chimeric oligonucleotides consisting of 2′-fluororibonucleotides (2′-F-RNA) with a varied number of DNA nucleotides embedded at the target cytosine and neighboring positions (Fig. 3A). 2′-F-RNA was selected for its stability relative to RNA. As DNA endonucleases were intolerant of these predominantly non-DNA substrates, we designed a primer-extension assay for deamination using reverse transcriptase (RT) (Fig. 3B). RT tolerated both 2′-F-RNA and our chimeric templates, properly incorporating the chain-terminator ddATP opposite the 0 position of the uracil product controls and the −2 position of cytosine substrates during primer extension (Fig. S6 A and B).

Fig. 3.

Rescue of deamination of chimeric 2′-F-RNA oligonucleotides requires a DNA patch from positions −2 to 0. (A) Chimeric substrate sequences and design. Control substrates consist entirely of DNA (D-control) or 2′-F-RNA (F-control). (B) Reverse-transcriptase–based deamination assay. A Cy5-labeled primer extended with reverse transcriptase using a nucleotide pool containing ddATP results in truncation of extension with deaminated substrates. Deaminated products yield a primer extension to 10 bp, whereas unreacted substrates yield extension to 12 bp. Assay validation with all substrates is shown in Fig. S6B. (C) Chimeric substrates incubated with AID for 1 h and evaluated via the reverse transcriptase-based assay. Asterisk denotes band associated with deaminated product.

When incubated with AID for 1 h, the entirely DNA control (D-control) was readily deaminated whereas the entirely 2′-F-RNA substrate (F-control) was not, as expected given AID’s selectivity against RNA (Fig. 3C). DNA at the target cytosine alone [F-d(0)] rescued deamination negligibly compared with the all-DNA control, indicating that removing additional 2′-substituents from neighboring nucleotides may be necessary to rescue deamination. Expanding beyond the target cytosine, removing the 2′-F from the −1 position [F-d(−1:0)] also yielded negligible, although slightly increased, deamination. On the other hand, expansion of the chimeric DNA patch from the −2 to the 0 position [F-d(−2:0)] fully rescued deamination to the level seen with the all-DNA control, as did further expansion of the DNA patch to the −3 to +1 positions [F-d(−3:+1)]. These findings were confirmed using an independent UDG-based assay for deamination (Fig. S6 C and D).

To evaluate the contribution of binding to deamination, we determined AID’s binding affinity for 2′-F-RNA chimeras and DNA/2′-F-RNA controls by fluorescence polarization measurements. AID showed a similar affinity for all of the substrates, with a slight preference for the predominantly 2′-F-RNA substrates over the all-DNA control (Table 3 and Fig. S5E).

Table 3.

AID binding of 2′-F-RNA chimeric substrates

Substrate	Kd, nM
D-control	32 ± 4
F-control	14 ± 1
F-d(0)	11 ± 1
F-d(−1:0)	16 ± 1
F-d(−2:0)	18 ± 1
F-d(−3:+1)	19 ± 1

SD is shown (n = 3 replicates). Graph of substrate binding is shown in Fig. S5E.

Discussion

In this report, we have demonstrated that AID’s deaminase activity is strongly influenced by the 2′-substituents of its target nucleotides. Introducing a single 2′-(R)-OH at the target cytosine nucleotide of an otherwise DNA substrate was sufficient to disrupt deaminase activity by at least 500-fold. Inversely, removing the 2′-fluoro substituents from the target cytosine and two upstream nucleotides rescued deamination of a 2′-F-RNA substrate. AID’s intrinsic preference for DNA is consistent with its physiologic DNA deaminase activity, and, although we did not address AID's in vivo preference, these data indicate that 2′-substitution of the nucleotide sugar is an important molecular determinant of selectivity for AID’s deaminase activity.

Intriguingly, when assayed against a similar panel of chimeric DNA substrates, APOBEC1 also demonstrated a high degree of sensitivity to the 2′-substituent of the target cytosine. Similar to AID, APOBEC1 showed a strong preference for deamination of A1-D-dC. However, in contrast to AID, APOBEC1 also showed detectable, although diminished, activity against A1-D-rC. On one hand, these data align well with recent studies on ancestral homologs of APOBEC1 that suggest that the enzyme was initially a DNA deaminase and evolved physiological RNA deaminase activity only recently (35). On the other hand, the discordance between APOBEC1’s intrinsic preference for DNA and its known physiological RNA deaminase activity also demonstrates that an enzyme’s intrinsic preferences alone do not dictate its physiological activity. Thus, our study also leaves open the possibility that AID, too, could overcome its intrinsic preferences within the cell or that APOBEC1 may possess an as-yet-undescribed cellular function against DNA. We speculate that the targeting protein partner ACF could play a role in allowing APOBEC1 to overcome its inherent preference for DNA, although this remains an important area for future study.

What is the molecular basis of the deaminases’ preference for DNA over RNA? Biophysically, there are two closely related principal mechanisms by which the 2′-OH substitution may enable enzymes to distinguish between the two nucleic acids. The first method of discrimination is steric exclusion, wherein steric interactions with the 2′-OH of RNA exclude the target nucleotide. A “steric gate” mechanism has been observed in DNA polymerases (36) and base excision repair enzymes, including uracil DNA glycosylase (37). Although the 2′-OH provides a basis for steric exclusion, it also results in conformational differences between the nucleotide sugar of RNA and DNA, known as sugar pucker. DNA prefers a C(2′)-endo (south) conformation, whereas RNA is more restricted to an alternative C(3′)-endo (north) conformation (Fig. 4A) (38). For enzymes that modify nucleic acids, alternative sugar puckers can impact the reactivity or positioning of the substrate in the active site by altering the angular projection of the base from the sugar-phosphate backbone, with RNA ligases serving as one example (39).

Fig. 4.

Potential role of nucleotide sugar pucker as a determinant of selectivity for DNA. (A) Conformational twist of nucleic acid sugar exists as an equilibrium between two dominant conformations: C(2′)-endo or C(3′)-endo. DNA prefers C(2′)-endo, whereas RNA prefers C(3′)-endo. (B) Speculative, mechanistic model for nucleic acid targeting and selectivity. Upon AID binding to its hotspot WRC target (W = A/T, R = A/G), a preferred C(2′)-endo conformation from −2 to 0 facilitates productive active-site interactions, which can be antagonized by a single C(3′)-endo-promoting substituent at the target nucleotide.

AID’s relative reactivity with the chimeric DNA substrates suggests that sugar pucker of the target nucleotide is critical for catalysis. Specifically, the cytosine conformers that prefer the C(2′)-endo (south) conformation—dC, aC, and faC (40)—were readily deaminated. Among these three substrates, D-dC was most favored over D-faC and D-aC, correlating reactivity with smaller 2′-(S)-substituents. This observation suggests that steric exclusion of the 2′-substituent may provide a secondary factor for discrimination. Further highlighting the importance of these nucleic acid determinants, the conformers that prefer the C(3′)-endo (north) conformation—rC and frC (41)—were disfavored for deamination, even when embedded in a nucleic acid substrate that is otherwise entirely DNA. Prior studies have shown that chimeric nucleotides embedded in DNA independently retain their sugar pucker and perturb local helix formation (42, 43), indicating that the cytosine conformers used in this study are likely to retain their expected north–south conformations in our chimeric DNA substrates. Taken together, reactivity with our chimeric substrates supports the notion that an isolated, disfavored sugar pucker at the target cytosine is sufficient to wdisrupt AID’s deaminase activity and underlies selectivity for DNA (Fig. 4B). More broadly, this model implicates sugar pucker as a potential discrimination mechanism by which APOBECs and other cellular enzymes may distinguish DNA from RNA at the level of individual nucleotides.

Moving beyond the target cytosine, our study shows that the 2′-substituents play a critical role at neighboring nucleotide positions as well. Removal of the 2′-fluoro substituent from the −1 and −2 positions was necessary to rescue full deamination of 2′-F-RNA chimeric substrates. Our finding regarding the significance of the −2 to 0 positions aligns well with the identification of the hotspot recognition loop within AID that specifically targets deaminase activity to the cytosine of WRC trinucleotide hotspots (22–24), as well as nucleoside analog interference studies with the homolog APOBEC3G (25). Taken together, the most significant determinants of nucleic acid recognition—sugar, backbone, and nucleobase recognition—appear to be confined to this critical trinucleotide patch spanning the −2 to 0 positions.

The results from this study demonstrate the significance of the 2′-substituent of nucleic acid substrates as a molecular determinant of AID’s selectivity for DNA and imply a recognition mechanism for targeted deamination involving multiple nucleotides. Consistent with current biochemical, structural, and single-molecule–based studies (3, 22–25, 31, 44, 45), we speculate the following model for deamination. After nucleic acid binding, the nucleobases for hotspot targets are recognized by the enzyme. Subsequently, permissive sugar puckers in a DNA-like conformation, dictated by the 2′-substituents at the −2, −1, and 0 positions, stabilize the enzyme-substrate complex and facilitate cytosine entering the active site, ultimately resulting in substrate deamination (Fig. 4B). Beyond explaining AID’s selectivity for DNA, this mechanistic framework for targeting may inform further studies focused on understanding the potential dynamic structural changes in both the enzyme and the substrate that drive AID/APOBEC-catalyzed targeted deamination.

Materials and Methods

Protein Expression and Purification.

Human AID (amino acids 1–181 or full length) and mouse APOBEC1 were cloned downstream of an N-terminal MBP in a pET41 expression plasmid. BL21-DE3 E. coli (Novagen) were transformed with expression constructs and grown to OD₆₀₀ 0.6, and protein expression was induced by addition of 1 mM IPTG (Sigma). Following induction, cells were transferred to 16 °C and incubated for 18 h before cultures were pelleted.

Proteins were purified essentially as described previously (46). Bacterial pellets were resuspended in 150 mM NaCl, 50 mM Tris⋅HCl, pH 7.5, and 10% (vol/vol) glycerol with EDTA-free protease inhibitors (Roche), and cells were lysed in a microfluidizer processor. Following removal of the insoluble fraction of the cellular lysates by centrifugation, the soluble fraction was added to amylose resin (New England Biolabs) and incubated at 4 °C for 1 h. Resin was washed in a high-salt buffer (750 mM NaCl, 50 mM Tris⋅HCl, pH 7.5, 10% glycerol) and then a low-salt buffer (150 mM NaCl, 50 mM Tris⋅HCl, pH 7.5, 10% glycerol), and proteins were eluted with maltose-containing elution buffer (10 mM maltose, 150 mM NaCl, 50 mM Tris⋅HCl, pH 7.5, 10% glycerol). Following elution, enzyme was dialyzed overnight at 4 °C in a storage buffer containing 75 mM NaCl, 50 mM Tris⋅HCl, pH 7.5, and 10% glycerol.

Oligonucleotide Substrates.

All oligonucleotides were synthesized using standard phosphoramidite chemistry either at the University of Calgary DNA Synthesis Core Facility (Calgary, Alberta, Canada) or using an ABI 394 Synthesizer (Applied Biosystems). Phosphoramidite building blocks and reagents were obtained from Glen Research or Metkinen Chemistry and used according to the manufacturer’s recommendations. Following synthesis, oligonucleotides were deprotected and purified using Glen-Pak DMT-ON columns, according to the manufacturer’s recommendations. Oligonucleotides were PAGE-purified, and masses were confirmed (Table S1). Oligonucleotides used in Fig. 3 were synthesized on 6-fluorescein (FAM) CPG columns and therefore contain a 3′-FAM label. Oligonucleotides used in Figs. 1 and 2 were synthesized without 3′-FAM; however, following synthesis, oligonucleotides were enzymatically 3′-end–labeled with ddUTP-12-FAM (Enzo Life Sciences) by incubation with Terminal Transferase (New England Biolabs) and purified using QIAquick Nucleotide Removal Kit (Qiagen).

Deaminase Incubations.

Deaminase assays were performed essentially as previously described (22). A 250-nM oligonucleotide was incubated with 500 nM AID or 1.5 µM APOBEC1, unless otherwise indicated. These highest concentrations of enzyme evaluated were chosen based on the stability of the purified enzymes. Reactions were incubated in the presence of 1× buffer DA (20 mM Tris⋅HCl, pH 8.0, 1 mM DTT, 1 mM EDTA) at 30 °C for times ranging from 1 min to 12 h as indicated, and enzymes were inactivated by incubation at 95 °C for 20 min.

Restriction Endonuclease-Based Assay for Deamination.

Following incubation with deaminase enzymes, oligonucleotides were annealed to an appropriate complementary strand at a concentration of 50 nM substrate to 100 nM complement. Oligonucleotides were annealed by incubation at 75 °C for 5 min and slow cooling to 37 °C, followed by 5 min of incubation at 37 °C. Duplexed DNA (20 nM) was incubated with 0.2 units/µL FspBI restriction endonuclease (Fermentas) at 37 °C for 3 h. Digestion reactions were quenched and denatured by addition of formamide (final concentration: 50% vol/vol) and incubation at 95 °C for 20 min. Samples were run on a denaturing 20% acrylamide/Tris-Borate-EDTA/urea PAGE at 50 °C and imaged on a Typhoon 9400 scanning gel reader (Amersham Biosciences). Substrate and product band intensities were quantified using QuantityOne (BioRad), and background intensities were subtracted. Total fraction of deamination was measured as the intensity of the product band divided by the sum of the intensities of both the product and the substrate bands. Relative deaminase activity was determined by the slope of a linear regression (GraphPad Prism), fit to the graph of product formation as a function of enzyme concentration. Linear regression for D-rC substrate did not significantly differ from zero; therefore, the lower limit of detection for D-rC deamination (Fig. S4C) is provided as a surrogate for deaminase activity against this substrate.

Reverse Transcriptase-Based Assay for Deamination.

Following incubation with AID, a 5′-Cy5-labeled primer (Table S1) was annealed to oligonucleotide substrate (166 nM substrate: 83 nM primer) by incubation at 75 °C for 5 min and slow cooling to 20 °C. A 50-nM primer/template duplex was extended by MuLV Reverse Transcriptase (New England Biolabs) in the presence of 1 mM ddATP and 100 µM dCTP/dGTP/dTTP at 25 °C for 4 h in polymerase buffer (10 mM Tris⋅HCl, pH 7.9, 10 mM MgCl2, 50 mM NaCl, 1 mM DTT). Primer extension reactions were quenched by addition of formamide (final concentration: 50% vol/vol) containing 1 µM TGC complement (Table S1) to denature primer-template duplexes and prevent reannealing. Samples were run on a denaturing PAGE gel, imaged using the Cy5 label for detection, and quantified as described above.

Fluorescence Polarization Analysis of Protein–Oligonucleotide Binding.

A 5-nM oligonucleotide substrate was incubated with increasing concentrations of protein under the same buffer conditions used in deaminase assays at room temperature (n = 3 replicates). Fluorescence polarization was measured using a Panvera Beacon 2000 Fluorescence Polarization system. Polarization values were graphed as a function of enzyme concentration in GraphPad Prism. A one-site binding, nonlinear regression was fit to the data, which yielded dissociation constants (K_d) and maximum polarization values for each substrate. For each substrate, values were normalized to maximum and minimum polarization values to yield total fraction of substrate bound.