Biochemical Basis of APOBEC3 Deoxycytidine Deaminase Activity on

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Biochemical basis of APOBEC3 deoxycytidine deaminase activity on diverse DNA substrates Madison B Adolph, Robin P Love, and Linda Chelico ACS Infect. Dis., Just Accepted Manuscript • DOI: 10.1021/acsinfecdis.7b00221 • Publication Date (Web): 18 Jan 2018 Downloaded from http://pubs.acs.org on January 20, 2018

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ACS Infectious Diseases

Biochemical basis of APOBEC3 deoxycytidine deaminase activity on diverse DNA substrates Madison B. Adolph, Robin P. Love, and Linda Chelico* Department of Microbiology and Immunology, College of Medicine, University of Saskatchewan, 107 Wiggins Road, Saskatoon, Saskatchewan, S7N 5E5, Canada Email: [email protected]

The Apolipoprotein B mRNA editing complex (APOBEC) family of enzymes are single-stranded polynucleotide cytidine deaminases. These enzymes catalyze the deamination of cytidine in RNA or single-stranded DNA, which forms uracil. From this eleven member enzyme family in humans, the deamination of single-stranded DNA by the seven APOBEC3 family members are considered here. The APOBEC3 family has many roles such as, restricting endogenous and exogenous retrovirus replication and retrotransposon insertion events and reducing DNA-induced inflammation. Similar to other APOBEC family members, the APOBEC3 enzymes are a double-edged sword that can catalyze deamination of cytosine in genomic DNA, which results in potential genomic instability due to the many mutagenic fates of uracil in DNA. Here we discuss how these enzymes find their single-stranded DNA substrate in different biological contexts such as during Human Immunodeficiency Virus (HIV) proviral DNA synthesis, retrotransposition of the LINE-1element, and the “off-target” genomic DNA substrate. The enzymes must be able to efficiently deaminate transiently available single-stranded DNA during reverse transcription, replication, or transcription. Specific biochemical characteristics promote deamination in each situation to increase enzyme efficiency through processivity, rapid enzyme cycling between substrates, or oligomerization state. The use of biochemical data to clarify biological functions and alignment with cellular data is discussed. Models to bridge knowledge from biochemical, structural, and single molecule experiments are presented.

Keywords: restriction factor, mutagenesis, DNA replication, HIV-1, processivity, enzyme mechanisms

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Overview of APOBEC family The Apolipoprotein B mRNA editing complex (APOBEC) family of enzymes are named after the function of their namesake member, APOBEC1 (A1) and are single-stranded polynucleotide cytidine deaminases1. The enzymes can modify cytidine in either mRNA or single-stranded (ss) DNA to form uracil2-4. The primary substrate of family members other than A1 is ssDNA (Figure 1A). In RNA, where uracil has a coding function, these enzymes act as RNA editing tools, but the formation of uracil in DNA is a promutagenic lesion. Although uracil in DNA can be detrimental, during normal function the APOBEC family members expressed in lymphoid cells use the formation of uracil as a catalyst to enable evolution of antibody genes (Activation Induced Cytidine deaminase, AID) or inactivation of viral, retroelement, or other cytoplasmic inflammation-inducing DNA (APOBEC3) (Figure 1A)2, 5-6. These multiple functions are possible because uracil has multiple fates in ssDNA. Most commonly, uracil is recognized as a lesion by the base excision repair (BER) enzyme uracil-DNA glycosylase (UNG) and excised. Subsequent processing by AP endonuclease (APE) and polymerase gap filling is required to complete the repair7. If UNG removes uracils during replication, DNA damage tolerance mechanisms such as template switching can allow for error-free lesion bypass8. If left unrepaired, the uracil will template addition of adenine at a site that should have incorporated a guanine, thus creating a C/G→T/A transition mutation (Figure 1B)7. In B cells, the uracils formed by AID in antibody genes act as nucleating factors for error prone DNA repair to evolve and mature antibody genes or create double-stranded DNA breaks to facilitate immunoglobulin class switching (Figure 1B)5. In T cells, macrophages, and germ cells, various APOBEC3 (A3) members (A3A, A3B, A3C, A3D, A3F, A3G, and A3H) can use this same cytidine deamination activity in ssDNA intermediates of retroviruses, retrotransposons, and other foreign DNA to cause mutagenesis to the point of destroying gene function or inducing DNA repair mediated DNA degradation to block replication of foreign elements in these cells (Figure 1C)1, 9. The APOBEC enzymes must remain tightly regulated to avoid mutagenesis of the cell’s own genomic DNA in the wrong context (Figure 1A-B). Failure of this regulation leads to APOBEC-induced somatic mutagenesis and the C/G→T/A mutation signature is evident in multiple cancers10-11. Despite the risk of harboring these promutagenic enzymes, the APOBEC family is quite ancient and has been present since the emergence of jawed vertebrates, with AID being the ancestral member12. Despite APOBEC2 (A2) being the second APOBEC deaminase in evolution, catalytic activity has not been demonstrated for this enzyme, although it is required for proper muscle development in mice12-15. As more complex organisms evolved, duplications in the APOBEC family of genes occurred to create A1, likely from AID, and in placental mammals, the A3 enzymes appeared12, 15 (Figure 1D). In agreement with A1 evolving from AID, there is evidence that A1 from the green anole lizard cannot deaminate RNA, suggesting that the ancestral function of A1 was as a deoxycytidine deaminase16. The A3 enzymes increased from one member in mice to seven members in primates12, 15. This massive expansion of the A3 locus correlates with the increased number of primate retrotransposons and endogenous retroviruses and is thought to have been required to suppress excessive damage to the genome by these elements15, 17. With the majority of retrotransposons and endogenous retroviruses in humans inactivated, it is thought that the risk of A3 enzymes may outweigh the benefits of their protection17. Evidence of this comes from population stratifications, where there are deletions in the human A3 locus on chromosome 22 that removes A3B or sequence variations in A3 genes that decrease deaminase activity, such as some forms of A3H, A3D, and A3C18-20. Despite AID posing a risk to genomic integrity, this enzyme is indispensable to proper immunity and must be maintained21.


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In this article we will focus on discussing the benefits and risks of A3 enzymes. Although the antiviral functions of A3 enzymes were discovered 15 years ago, an understanding of their biochemical mechanisms and structural basis for activity has only become clear in recent years. Thus focusing on biochemical mechanisms, we will review our understanding of how A3 enzymes can both protect and harm the genome. A3 enzymes restrict the replication of Human Immunodeficiency Virus Type 1 (HIV-1) The discovery and greatest understanding of A3 enzymes is in the field of HIV-1 restriction. HIV-1 is an enveloped virus with the nucleic acid genome being contained within a capsid. The capsid also contains all the proteins required for synthesis and integration of the double-stranded (ds) DNA genome, reverse transcriptase, integrase, and nucleocapsid22. When the HIV-1 fuses with a target CD4+ T cell, the capsid is released into the cytoplasm (Figure 2A). The deoxynucleotide triphosphates (dNTP) from the host cell diffuse into the cytoplasm and reverse transcription begins23. Due to the impermeability of the capsid to cellular proteins at this early stage, if A3 enzymes are to access the HIV1 (-) DNA synthesized by reverse transcriptase, they must be present in the viral capsid. Thus, the model for HIV-1 restriction by A3 enzymes in CD4+ T cells comes with a requirement that they become encapsidated into newly formed virus particles in a producer cell24. The A3 enzymes preferentially bind cellular and viral RNAs with the same sequence affinity as HIV-1 nucleocapsid domain of Gag that packages the HIV-1 RNA25-26. Encapsidation is also facilitated by A3 oligomerization on RNA although the reason of why is not entirely understood27. It could simply be that oligomerization is a way of hedging bets for encapsidation, since there is no guarantee of an A3 in each virion. Usually, the encapsidation of A3 enzymes is blocked by the HIV-1 protein Vif (virus infectivity factor) that causes the polyubiquitination and degradation of A3 enzymes through the 26S proteasome28. Vif can specifically interact with A3 enzymes and acts as a substrate receptor by interacting with components of a Cullin RING ligase 5 E3 ubiquitin ligase complex, and results in a complex capable of binding and polyubiquitinating an A3 (Figure 2A)28-32. In HIV-1 infected people, the suppression of A3 enzymes by Vif is not complete since proviral genomes have been found to contain footprints of A3 deaminase activity33-34. However, the inhibition is clearly sufficient enough to enable ongoing viral replication. From cell culture studies with ∆Vif HIV-1, only A3D, A3F, A3G, and A3H are able to encapsidate and restrict HIV-1 replication35. Although all four of these A3s have variants in the human population, A3H is the most notable since it is the most variable and has seven major haplotypes (hap), but only hap II, hap V, and hap VII are able to restrict HIV-1 replication36-37. The encapsidated A3 enzymes are in a competitive situation where they must deaminate cytosines in an ssDNA substrate that is not available initially and only available for a finite time after synthesis (Figure 2B-C). To form the (-) DNA, the HIV-1 RNA genome is primed by a tRNAlys,3 that is unfolded and annealed to a primer binding site (PBS) by the HIV-1 nucleic acid chaperone, nucleocapsid38. After reverse transcription and degradation of the RNA by the RNaseH domain associated with the HIV-1 reverse transcriptase, nucleocapsid facilitates a strand transfer to another complementary repeat (R) region and full synthesis of the (-) DNA can begin. During synthesis of the (-) DNA, the RNaseH activity of reverse transcriptase will cleave the RNA at internal sites until the RNA cleavage leads to dissociation22. The nucleocapsid is abundant in the capsid and protects the HIV-1 genetic material by binding the RNA or ssDNA at approximately one nucleocapsid every seven nucleotides39. Nucleocapsid facilitates reverse transcriptase by melting secondary structure in the template40. Although not fully understood, the mechanism of nucleic acid chaperoning by nucleocapsid involves the protein rapidly binding and unbinding the RNA or DNA40. Thus, A3 enzymes can compete with nucleocapsid for access to the (-) DNA (Figure 2C). However, there are still physical obstacles the 3 ACS Paragon Plus Environment


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enzymes must overcome such as annealed RNA fragments, the reverse transcriptase, and constraints such as finding the deamination motif. Each A3 deaminates cytosine within a preferred sequence. A trinucleotide motif is the preferred substrate, although a dinucleotide motif can also be deaminated. For A3G this is 5'CCC (underlined C deaminated) and for other A3 enzymes it is 5'TC, with each enzyme having a specific preference, e.g., 5'TTC for A3F and 5'CTC for A3H41-43. The C→U deaminations catalyzed on the (-) DNA strand become G→A mutations when reverse transcriptase is forced to use uracil as a template during (+) DNA strand synthesis (Figure 1C)44-47. Throughout this process, the A3 enzymes also directly disrupt proviral DNA synthesis by oligomerizing on the template DNA resulting in a “roadblock” for the reverse transcriptase48-51. In addition, it has been specifically shown for A3G that it can interact with reverse transcriptase directly and inhibit polymerase activity48, 52. The double-stranded proviral genome is imported into the nucleus and either the uracils cause DNA repair mediated cleavage of the provirus or the provirus is integrated through the action of HIV-1 integrase and host protein interactions52 (Figure 1C, Figure 2A). Depending on the number and type of mutations induced by the A3 enzymes, the integrated provirus may be functionally inactivated. An early observation with A3G was that it was able to induce stop codons41. The ability to induce stop codons arises from the single codon used to code for tryptophan (Figure 3A). In the (-) DNA orientation the tryptophan antisense codon contains a deamination motif for A3G (Figure 3A). Deaminations of cytosine within the antisense codon can result in two possible stop codons (Figure 3A). Depending on the surrounding sequence context, A3F may also be able to induce a stop codon (Figure 3A). However, in proviral DNA sequences, the A3G-induced tryptophan mutation to a stop codon is recovered more often than the A3F-induced tryptophan mutation to a stop codon51, 53 (Figure 3B). A3F is more likely to induce a variety of missense (amino acid changing) mutations than A3G that primarily causes mutations at glycine codons (Figure 3B)51, 53. The A3G-induced mutations at glycines tend to be nonconservative and inactivating mutations whereas the A3F-induced missense mutations have less inactivation ability51, 53-55. Similar data has been found using in vitro substrates for A3A that also deaminates at 5'TTC56. Accordingly, there is evidence that 5'TC modifying A3 enzymes may contribute more than A3G to HIV-1 evolution in the form of immune escape or drug resistance, rather than inducing HIV-1 inactivation55, 57-62. Most commonly labs have studied A3-induced mutagenesis of a ∆Vif HIV-1 virus. In these situations, the A3 “hypermutation” arises where enzymes such as A3G can induce 5 or more mutations per kilobase44, 46-47. Although hypermutated and integrated proviruses have been isolated from HIV-1 infected patients, it is unknown the number of replication cycles that the provirus underwent before becoming hypermutated and nonfunctional63-64. In the presence of Vif, the A3 enzymes are depleted from cells to such an extent that viruses can replicate although a low level of mutations can persist. Measurements of the number of A3induced mutations in a single round of infection are lacking and need to be determined in order to identify whether A3 deaminations in a wild type virus infection induces hypermutation and viral inactivation, a low level of mutation that leads to virus evolution, or has a neutral effect. All of these possibilities have been characterized in different labs using different experimental systems34, 59, 65-68. The variability may be due to which A3 and the amount that was encapsidated (Figure 3). The number of A3-induced mutations can be determined by encapsidation levels, which are indirectly determined by Vif-mediated A3 degradation, but can also be determined by the inherent properties of the A3 enzymes to bind and oligomerize on cellular and viral RNA.

Biochemistry of A3-mediated HIV-1 restriction


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At a cursory assessment of the HIV-1 and A3 interface, it would seem that a key factor in increasing mutations is increasing the number of A3 molecules encapsidated. However, the A3 enzymes cannot guarantee their encapsidation into each virus because the encapsidation is dependent on the nucleocapsid domain of HIV-1 Gag binding the same RNA to which the A3 enzyme is bound and directing its virion encapsidation25-26. In the absence of Vif there is an estimated 7 ± 4 molecules of A3G encapsidated69. In the presence of Vif this is decreased to 0.3 to 0.8 molecules of A3G encapsidated70. These estimated numbers must be considered in addition to the finite time that the HIV-1 (-)DNA remains single-stranded and emphasizes that in order to inactivate HIV-1, an A3 needs to efficiently deaminate ssDNA. For DNA modification enzymes, efficiency generally means that the enzyme is processive and can deaminate multiple cytosines in a single enzyme-substrate encounter. For enzymes that do not use an energy source to move on DNA, they can search the DNA for a specific target motif using facilitated diffusion71-72. Facilitated diffusion describes how enzymes can use the electrostatic interaction with DNA to facilitate their Brownian motion driven diffusion71-72. The diffusion can be described by different terms depending on whether the enzyme diffuses along the phosphate backbone (sliding), diffuses along the DNA without always making direct contact with the phosphate backbone (jumping or hopping), or moving through a doubly-bound state (intersegmental transfer) (Figure 4)1, 71-73. The sliding and hopping movements are efficient for searching < 20 nt of DNA and for A3 enzymes that slide, this movement is essential for finding the deamination motif1. The jumping and intersegmental transfer facilitates movements on DNA > 20 nt, but at each place the enzyme lands, there is no local search unless the enzyme can also undergo sliding53, 73-74. Thus, if an enzyme only has sliding or only has a three-dimensional search mechanism such as hopping, jumping, or intersegmental transfer, the search will be inefficient1, 51, 53, 74-75. To put a quantitative measurement on this observation, if A3G that can slide and jump and A3H hap II that can slide, jump, and undergo intersegmental transfer are compared to A3F and A3C that can only jump or undergo intersegmental transfer, respectively, the number of mutations per kb of HIV-1 proviral DNA is 3 to 6 (A3F) or 100 to 200 (A3C) fold less51, 53, 74-75. By mutagenesis of processivity domains to make A3F and A3C able to slide, it was found that the enzymes were still not as efficient as A3G or A3H hap II at deaminating proviral DNA53, 74-76. For A3F this appeared to be because the enzyme bound ssDNA with such high affinity that the actual movement and search of (-) DNA was limited, despite the acquired ability to slide on oligonucleotides in vitro53. For A3C, the enzyme was unable to encapsidate as efficiently as A3G and despite mutagenesis enabling A3C to slide processively to deaminate cytosines, the processivity was still 2- to 4- fold less than A3G76. Thus, A3G and A3H hap II appear to have the correct balance of search components to find their target sequence compared to other A3 enzymes and are readily encapsidated. Notably, A3F is also encapisdated efficiently into virions, but the lesser amount of mutations in comparison to A3G and A3H emphasizes the importance of processivity53. Recently it was shown that A3F and A3G heterooligomerize in cells and this results in more efficient mutagenesis of HIV-1 when A3F and A3G were coencapsidated51. More studies should assess the extent to which A3s can mix and match their capabilities so that we can fully understand their action during a natural HIV-1 infection. Processivity has been questioned as being relevant in a virion since the imposed confines of the capsid would increase the apparent concentration of the components. For instance, if an enzyme were to dissociate from the ssDNA often, it could quickly rebind the ssDNA without escaping into the bulk solution. To test this, one could use A3A, the only nonprocessive A3, but it is not normally encapsidated into HIV-1 virions77. However, using an in vitro system that recapitulated proviral DNA synthesis steps of HIV-1, it was found that A3A was able to induce mutations at a level equal to A3G in regions of the cDNA that were single-stranded the longest, but A3A-induced mutations were lower than A3G in 5 ACS Paragon Plus Environment


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regions near the second strand DNA primer, a region single-stranded for the shortest time56 (Figure 2BC). These data demonstrated that processivity was required to speed up the search for cytosines in regions that are not single-stranded for extended periods of time, but that an ability to cycle on and off DNA substrates and a high specific activity can, for the most part, approximate the efficiency imparted by processivity56, 78-79. This is in contrast to A3C that has a similar processivity to A3A, but has a 50- to 300- fold lower specific activity, and is less mutagenic in the same in vitro assay51, 56, 80. The interesting aspect of A3 processivity is that this feature is important despite the many obstacles that the enzymes encounter on the RNA or (-) DNA. In vitro experiments have shown that jumping is important not only for restarting the search process at a distal site, but also for enabling the enzymes to directly transverse over a physical obstacle such as an RNA/DNA hybrid without dissociating from the DNA and remaining processive53 (Figure 4). Further, A3G (and presumably A3F that has a higher affinity for ssDNA) can stay bound to ssDNA in excess of 10 min81-84. This enables A3G to outcompete nucleocapsid that has a fast on and off rate for ssDNA (Figure 2C). There is even evidence that A3G can bind with such high affinity that it acts as a roadblock to the reverse transcriptase and inhibits completion of proviral DNA synthesis in a timely manner48-49. A history of protecting genome integrity Although some A3 enzymes have evolved the biochemical characteristics necessary to restrict HIV-1 replication their role in protecting genome integrity predates immunodeficiency viruses. More of the A3 family members are able to restrict retrotransposons than HIV-1 and retrotransposon pressure on primate genomes is thought to have caused the expansion of the A3 locus (Figure 1D)15. Retrotransposons replicate through an RNA intermediate and are divided into two groups, long terminal repeat (LTR) retrotransposons (also known as endogenous retroviruses) and non-LTR retrotransposons9. LTR retrotransposons (endogenous retroviruses) constitute approximately 10% of the human genome9. Owing to the accumulation of mutations, endogenous retroviruses, such as HERV-K, have become largely inactive in humans. Reconstruction of endogenous retrovirus genomes has led to the observation that they accumulated many C/G→T/A mutations and were inactivated by A3 enzyme activity85. When active, endogenous retroviruses were single-strand (+) RNA viruses that infected germ cells for vertical transmission, but otherwise replicated like HIV-1 and underwent reverse transcription in the cytoplasm9. Non-LTR retrotransposons constitute approximately 30% of the human genome9. Three types of non-LTR retrotransposons are still active in humans, the long interspersed element-1 (LINE-1), the short interspersed element (SINE) Alu, and the composite retrotransposon SINE-VNTR-Alu (SVR)9, 86. Since Alu and SVR depend on LINE-1 activity for their retrotransposition, the LINE-1 is the model used to study A3-mediated restriction of retrotransposon activity. All A3s except A3G can restrict LINE-1 retrotransposition to various degrees9, 87. The LINE-1 RNA is reverse transcribed in the nucleus, and a cellular RNaseH degrades the LINE-1 RNA and exposes the (-) single-stranded DNA, enabling A3 deamination of cytosines9, 88. Although multiple labs demonstrated that A3 enzymes could restrict LINE-1 retrotransposition, it was not known until recently whether this was due to a deamination dependent or deamination independent mechanism9, 88. Since A3 enzymes bind RNA with high affinity, it was thought that the A3 enzymes would act as a roadblock to reverse transcription or bind the LINE-1 RNA in the cytoplasm and cause it to accumulate in cytoplasmic RNA processing bodies (P-bodies), preventing nuclear import89-90. This was supported by DNA sequencing of integrated LINE-1 genomes that rarely found evidence of cytosine deamination and that many A3 enzymes bind RNA in cells and purify from cells in a Ribonucleoprotein (RNP) mass9, 91. However, if A3 deaminations occurred in the 6 ACS Paragon Plus Environment

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nucleus, the uracils formed on the exposed single-stranded (-) DNA would be recognized by the BER enzyme UNG and excised. If the uracils were numerous enough, this would result in DNA breaks and degradation of the LINE-1 (-) DNA due to the action of APE that acts downstream of UNG7 (Figure 1C). DNA repair mediated degradation may have more time to occur since LINE-1 relies on cellular polymerases and DNA repair enzymes to copy the (-) DNA to form a dsDNA LINE-1 cassette that is integrated into the genome88. By using an UNG inhibitor, it was demonstrated that A3A mediated inhibition of LINE-1 involved deamination of cytosines in LINE-1 single-stranded (-) DNA, and that the primary fate of the LINE-1 was degradation, not integration of a mutated retrotransposons88, 92. However, it does not seem to be a common mechanism for other A3s since A3H uses a deamination independent mechanism to inhibit LINE-192. Thus, although there are some parallels with HIV-1 restriction in that reverse transcription is needed to generate the ssDNA substrate, the restriction of LINE-1 by A3 enzymes still needs to be addressed to determine if A3s other than A3A can deaminate LINE-1 (-) DNA and identify the mechanism responsible for the deamination-independent restriction of LINE-1 retrotransposition. Interestingly, the fate of deaminated LINE-1 (-) DNA is different than for HIV-1 or endogenous retroviruses. Perhaps due to the longer time that the LINE-1 (-) DNA may remain single-stranded owing to a dependence on cellular rather than self-encoded mechanism of (+) DNA synthesis, the A3-catalyzed uracils result in LINE-1 degradation more often than for HIV-1 or endogenous retroviruses. Off-target somatic mutagenesis induced by A3 enzymes Despite these benefits of A3 enzymes for suppression of retroelements, HIV-1 restriction, and even lesser characterized roles in restricting replication of Hepatitis B virus93-94, Cytomegalovirus95, and Human Papilloma virus96, there is evidence that there is a cost to this defense system in the form of offtarget A3-catalyzed deaminations that occurs in our genomes during our lifetime. This happens when the expression of A3 enzymes occurs in the wrong cell, at the wrong time, and deamination occurs in the wrong gene, such as a tumor suppressor97. Usually, redundant DNA repair mechanisms can remove uracils and negate most of these promutagenic lesions7. However, with the development of Next Generation Sequencing technology to obtain greater sequencing depth, it is clear that many cancer genomes have a bias of C/G→T/A mutations and most cancer cells or tumors show overexpression of A3B or A3H hap I mRNA, suggesting that some uracils persist and develop into mutations10-11, 98-102. Additionally, the abasic site resulting from BER of A3-catalyzed uracils may induce replication stress or an error-prone polymerase can mediate base insertion opposite the abasic site, resulting in C→G or C→A mutations103-104 (Figure 1B). Several lines of evidence demonstrate that A3 enzymes provide tumor cells with a “just right” rate of mutagenesis97. For example, one report found that women diagnosed with Estrogen Receptor positive breast cancer that also have high A3B mRNA expression are associated with poor survival105. The resulting increase in mutation rate has also been shown to lead to tamoxifen therapy resistance106. A3B is the most well studied in cancer mutagenesis and was identified to contribute to not only breast cancer progression but also head/neck, lung, bladder, cervical, and ovarian cancer11, 102, 107-108. Since A3 enzymes each deaminate cytosine in a specific nucleotide sequence context, A3B activity was identified by retroactive sequence analysis in combination with in vitro analysis of the preferred nucleotide sequence context11, 102. The A3B “footprint” is mutations at 5'ATCA (underlined base becomes mutated)102, 109. Additionally, studies have found the A3A mutational footprints of 5'TTCA or 5'CTCA in tumors, but no corresponding A3A expression which suggested that A3A is upregulated early, but later inactivated, perhaps due to being the most active deaminase that could cause cell death through its activity over time, rather than a selective advantage109-111. In addition 7 ACS Paragon Plus Environment


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or alternatively, A3H hap I is able to induce mutations in breast and lung cancer cells and there is a high association of the A3H hap I allele in an A3B-/- genetic background101. The A3H hap I footprint is 5'CTCA and overlaps with A3A75, 101. Thus more tumor based studies are needed to differentiate the activity of A3A and A3H hap I. Importantly, these enzymes that induce somatic mutagenesis must not only be able to localize to the nucleus but must be able to deaminate transiently available ssDNA created during the processes of transcription or replication. For a related family member AID, that normally deaminates the immunoglobulin genes to enable antibody maturation and class switching5, the targeting of AID to the correct genomic region and maximum access to both strands of the genomic DNA requires many interacting partners112. Although AID can access other regions of the genome, it is thought that interactions with transcription machinery occurs to facilitate its specific role in deaminating cytosine in immunoglobulin genes113-114. These protein-protein interactions may not only assist in targeting immunoglobulin genes during B-cell activation, but may be required for AID to catalyze deaminations since its catalytic rate is slow115. The slow catalytic rate of AID may be a protective mechanism against AID-catalyzed “off-target” activity115-116. In contrast, in vitro studies indicate that the A3 enzymes relevant to cancer mutagenesis are able to deaminate cytosines during transcription or replication without the requirement for interacting partners80. While no interacting partners have been identified, there is the potential that a partner may exist that facilitates this mechanism. Interestingly, in an understudied area of A3 biology, it has been proposed that A3s in the cytoplasm of cells can deaminate incoming foreign DNA or stress associated cytoplasmic dsDNA released from mitochondria6, 117-118. This cytoplasmic dsDNA is transcribed into dsRNA by RNA polymerase III to activate RIG-I inflammatory pathways6. A3 deamination presumably occurs on the ssDNA generated during transcription and serves to induce degradation through the action of cytoplasmic UNG or prevent reannealing of the DNA strands to thwart infection or inflammation6. This transcription-associated deamination also appears to be in the absence of any targeting mechanisms. In contrast to AID and possible anti-inflammatory roles of A3 enzymes, studies in yeast and analysis of TCGA sequences have identified that A3 enzymes primarily deaminate ssDNA on the lagging strand during DNA synthesis, deaminate less on the leading strand, and do not show significant amounts of deamination during transcription119-123. These studies are based on the analysis of strand biases that would place deaminations either on the lagging strand or nontranscribed strand of DNA during transcription. During DNA replication, recombination, and transcription, the ssDNA generated is protected by ssDNA binding proteins Replication Protein A (RPA) or RAD51 to avoid degradation by cellular nucleases and to promote DNA processing124-125. A3 enzymes must be able to breach this protective barrier to access the ssDNA. Under normal replication conditions, the leading strand polymerase and helicase move together with the helicase in front of the polymerase, but if the polymerase is blocked by a lesion in the DNA or imbalances in dNTP pools, then the helicase can become uncoupled and will keep unwinding DNA without concomitant synthesis, which generates long stretches of ssDNA126-127. This ssDNA is then bound by the ssDNA binding protein RPA to protect from nucleases or other chemical damage and A3s must compete with RPA to access to the ssDNA substrate126-127. Thus, in a normal cell, an A3 must access ssDNA between the polymerase and helicase, which are usually traveling along the DNA rapidly127-128 (Figure 5). Although there are ssDNA gaps on the lagging strand, which is probably why this strand is favored by A3 enzymes, the access to the ssDNA requires the A3 to displace RPA (Figure 5). However, a hallmark of cell transformation is replication stress that would cause the uncoupling of the polymerase and facilitate A3 deamination127, 129-130. This ssDNA would be stably bound by RPA awaiting replication restart or displaced by RAD51 during homologous recombination. Thus, it is notable that A3B is not problematic until a cellular alteration occurs that 8 ACS Paragon Plus Environment

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increases A3B expression and increases replication stress104, 121, 130. In contrast, the ssDNA generated on the nontranscribed strand during transcription is accessible at multiple times during the cell cycle of a normal or transformed cell, especially during RNA polymerase stalling and preinitiation113-114. Although A3 enzymes would still need to access transiently available nontranscribed ssDNA that interacts with the RNA polymerase, the ssDNA is solvent accessible131. Despite this, the A3 enzymes do not use ssDNA generated during transcription as a major substrate, in contrast to AID. This may be due to the lack of protein-protein interactions that recruit A3 enzymes to sites of transcription or a lack of temporal resolution in cellular studies. Biochemistry of A3-induced somatic mutagenesis For DNA repair enzymes, much like the A3 enzymes that restrict HIV-1 replication, processivity is important for efficiently scanning the genome for lesions. However, in a study examining A3 deamination activity using in vitro transcription and replication model systems the opposite was found where A3A, a non-processive enzyme, was equally effective in deamination during replication as A3B and A3H that were processive enzymes80. The important characteristic was that the enzymes rapidly cycled between ssDNA substrates80. However, processivity was not detrimental to enzyme activity. A3B and A3H cycled between substrates and maintained processivity whereas A3A simply cycled on and off substrates rapidly80. Notably, the ability to cycle between ssDNA substrates was not only important to access naked ssDNA being replicated, but also ssDNA bound by RPA80. Despite the high affinity and long-lived association of RPA with ssDNA, the RPA can be displaced by a mechanism known as facilitated dissociation132-133. This is where excess RPA or RAD51 was found to be able to exchange with bound RPA on the ssDNA. In vitro analysis of A3 enzymes competing with RPA saturated ssDNA for DNA binding showed that rapid cycling was required for RPA facilitated dissociation80. Namely, A3A, A3B, and A3H hap I had comparable specific activities to each other and A3G, however, the specific activity of A3G was decreased 10-fold in the presence of RPA, in contrast to A3A, A3B, and A3H that had at most a 2-fold decrease in specific activity80. This observation is interesting since A3G is one of the most efficient deaminases for HIV-1 restriction, but the decreased frequency of A3G cycling between ssDNA substrates was detrimental in the presence of RPA80-84. The processive A3 enzymes remained processive in the presence of RPA, but this processivity decreased approximately 2-fold, suggesting that RPA acted as a physical block that had to be transversed, much like RNA/DNA hybrids during HIV-1 replication53, 80. These data demonstrate that during replication stress where larger amounts of ssDNA accumulate, that RPA is not a protective barrier against A3A, A3B, and A3H hap I. In general, in vitro experiments using model replication and transcription substrates for A3 enzymes have agreed with yeast-based and tumor cell studies. For example A3B-induced mutations were found to correlate with the lagging strand of replication, not the ssDNA on the nontranscribed strand generated during mRNA transcription80, 119. Biochemical analysis using an in vitro T7 RNA polymerase system showed that A3B, that formed tetramers, was unable to access cytosines during active transcription due to a size limitation imposed by oligomerization, although R-loops were accessible80. Notably, a study found that A3B could deaminate during transcription of tRNA genes, but these transcription bubbles are larger than those of mRNA, in agreement with transcription having a size-dependent selectivity80, 134. For A3A, there was an association of mutations in a yeast system with the nontranscribed strand, in agreement with in vitro transcription data, but this was minor and the majority of mutations correlated with lagging strand of replication80, 119. A3H is able to deaminate during transcription in vitro, but there are no corresponding cellular studies. However, A3H hap I was determined through clonal sequence analysis to induce mutations in lung cancers that arise early and accumulate slowly101. This is in contrast 9 ACS Paragon Plus Environment


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to A3B induced mutations that were absent early in tumor formation, but then suddenly arose and were maintained at a high level101. Although speculative, since A3B somatic mutagenesis appears to require prior genomic instability, unlike for A3H hap I that could act early in lung cancer mutations, we propose that A3H hap I may be able to induce this constant low level of mutagenesis by deaminating ssDNA generated during transcription80. Based on biochemical and cellular data, A3B would be unable to deaminate ssDNA generated during mRNA transcription, unless stable R-loops were to form80. In the yeast study and other cancer genome studies, transcription may not have been observed as a dominant form of ssDNA substrate due to the studies being an end-point analysis where only accumulated but not temporal mutations were analyzed119-123. Determining during initial A3-induced mutagenesis how and when A3 enzymes can deaminate genomic DNA will identify if they cause cell transformation, perhaps through a slow continuous mutagenesis such as that observed for A3H hap I, or contribute to a mutator phenotype in already transformed cells, such as that observed for A3B101. Basis for divergent activity Both HIV-1 replication and genomic DNA replication and transcription processes are dynamic, but the A3 enzymes use different biochemical characteristics to access these substrates1, 80. These differences may arise due to the nonprocessive HIV-1 reverse transcriptase creating an environment where an A3 enzyme can more freely scan and maintain long residence times on ssDNA135. Even the HIV-1 nucleic acid chaperone, nucleocapsid, cycles rapidly and can be easily displaced40. Not only is eukaryotic replication faster than HIV-1 proviral DNA synthesis, but the competition with RPA is more difficult since RPA is not as readily displaced and requires that an excess of RPA be present132. Thus, an A3 enzyme being able to cycle on and off ssDNA or between regions of the ssDNA to maintain an interaction as the ssDNA changes as well as being able to promote facilitated dissociation of RPA, rather than remain in contact and slide along the ssDNA, is more advantageous. These biochemical characterizations identify that some A3 enzymes are inherently better at deaminating cytosines in specific types of ssDNA. Thus, in both the HIV-1 and cancer mutagenesis situations, the availability of ssDNA is necessary, but alone not sufficient for A3-induced mutagenesis in cells because there is also a dependence on the inherent biochemical properties of the enzymes, such as processivity, cycling, and specific activity1, 80, 115. All of these features are influenced by the surface charge of the enzyme that mediates the electrostatic interactions with ssDNA1, 115. The lack of a requirement for processivity in regards to DNA binding protein interactions is unexpected considering the number of processive DNA repair and replication proteins136-140. However, these other proteins that are normally functional in the genome usually bind and scan dsDNA, which is less dynamic than ssDNA in the genome.

Structural and Single-molecule assessments of A3 interactions with DNA The biochemical analysis of A3 enzymes is still an expanding field. After the initial discovery of A3G in 2002 and the biochemical analysis of A3G published in 2006, there was a gap of 6 years before the biochemical analysis of A3A24, 56, 141-142. Purification and analyses of other A3 enzymes followed and currently only A3D remains to be fully characterized biochemically51, 53, 75, 80. The difficulty comes from the fact that most of the enzymes do not fold or express well in E. coli and some do not even express well in 293T or Sf9/baculovirus systems. Due to a lack of consistent approaches between labs, the field currently has biochemical and single molecule studies using primarily 293T or Sf9/baculovirus derived 10 ACS Paragon Plus Environment

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A3 enzymes and structural studies using A3 enzymes produced from E. coli. Several labs have overcome the limitations of the E. coli system by truncating and mutating the A3 enzymes to express better115. While there is no doubt that the structural information is useful, it leaves the field in a place where it is difficult to converge the observations made from these two areas because there is a paucity of full length unmutated structures that can be directly compared to biochemical studies using the native enzymes143-144. For example, structural studies have shown that A3B, but not A3A, has a closed catalytic pocket115, 145146 . The closed catalytic pocket has been proposed to reduce off-target mutations by APOBEC family enzymes115-116. However, these conclusions for A3B were made from crystal structures using the catalytic C-terminal domain (CTD) and not the full-length enzyme115, 145-146. The full length A3B is 10to 40- fold more active than the A3B CTD, depending on the location of the truncation and experimental system80, 147. Thus, it is not known whether increased A3B activity in the full-length enzyme is due to different catalytic pocket accessibility, if increased processivity in full length A3B enables a more efficient search for potential deamination motifs, if the N-terminal domain (NTD) can control substrate access of the CTD, or a combination of all of these factors. Further, structural studies have demonstrated that A3A and A3B CTD bend short ssDNA substrates bound to the active site145, 148-149. A3A and the A3B CTD are not processive and it has not been determined if a processive A3 enzyme also bends the ssDNA in the active site56, 80. This should be considered since it is difficult to imagine an enzyme sliding or jumping along ssDNA while bending each nucleotide it encounters. However, based on singlemolecule data from a study with A3G we know that A3G scans ssDNA very rapidly and bidirectionally, but when A3G encounters a deamination motif, the enzyme movement slows down and “hovers” over the deamination motif 84. If this observation can be extrapolated to include structural observations, a model can be envisioned where bending of ssDNA, which has been observed in singlemolecule, bulk biochemical, and structural studies, serves to slow down the diffusion to enable deaminations to take place82, 84 145, 148-149. This slow diffusion and deamination could exist in between moments where the enzyme is processively searching ssDNA by fast diffusion mechanisms. Single molecule experiments have also been able to answer questions regarding oligomerization in the absence of a native A3 oligomer crystal structure. For example, AFM, single-molecule optical tweezers studies, and biochemical studies with A3G have demonstrated that A3G oligomerization is dynamic49, 81-83, 150. A3G is primarily a dimer or monomer in solution, depending on the salt and protein concentrations82, 151. Single molecule studies have shown that monomers bind ssDNA and then they rapidly oligomerize83. However, only monomers dissociate from ssDNA and oligomers remain bound in excess of 15 min83. The oligomers that remain bound are not as mobile on ssDNA as monomers82-83. This is seemingly in contrast with a large body of biochemical and virological data suggesting that A3G is processive, is a dimer or larger oligomer, and that dimerization is required for the jumping component of A3G processivity53, 74, 151. Although there are different buffer conditions and timescales between single molecule and deamination experiments, the data can be resolved if jumping by A3G is imagined as a breaking up of the oligomer and re-nucleation of the monomers at another site on the same DNA (Figure 4E). Without this ability to oligomerize again at the new site, the enzyme would simply fall off the DNA and this would be observed in bulk studies as an inability to jump74, 83. This working model could not be obtained from bulk biochemical studies alone that only view an average of all these events. For A3C, biochemical experiments were able to determine that dimer dissociation was also required for intersegmental transfer, but the dimer properties of A3C are much less dynamic than A3G51 (Figure 4FG). Thus, in the future if more single molecule studies are conducted with other A3 enzymes we could have a more integrated idea of their structure and function at a molecular level.



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Conclusions Despite different functions and substrates, A3 enzymes share many similar features with other DNA binding and modification enzymes. For example, many DNA repair enzymes scan DNA rapidly by facilitated diffusion and slow down to hover over their target motif or lesion, much like what has been observed for A3G84, 136, 139-140. Where a DNA repair enzyme is charged with being an efficient enzyme that searches dsDNA for lesions that need to be repaired, A3 enzymes must search for deamination motifs before the ssDNA is covered up by the complementary strand or other DNA binding proteins. Considering the data indicating that some A3 enzymes can actively compete with RPA and are not just opportunistic in how they access ssDNA in the genome, perhaps more parallels can be drawn between A3 enzymes and proteins involved in genomic DNA binding and modification. Merging the insights from A3 biochemical studies and the existing understanding of DNA replication proteins may offer key insights into mechanisms of A3-induced somatic mutagenesis. What is most unique about A3 enzymes, but remains understudied is their relationship with cellular RNA. It has been observed that they bind RNA in the cytoplasm of cells, purify from cells as part of an RNP complex, and when purified in vitro in the presence of RNaseA, still never fully dissociate from RNA fragments144, 152-153. Recent A3H crystal structures have shown that a duplex RNA bridges two A3H molecules together to form a dimer in the absence of any protein-protein interactions144, 153. If this is unique to A3H remains to be determined. All the A3s that bind RNA in cells require RNaseA treatment in vitro to be fully active27, 141. If future research can determine if RNA binding is merely a mechanism to suppress off-target deamination activity, has a biological function, or is required for A3 structural stability, we can obtain a holistic understanding of A3 biology and function.


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139. Min, J. H.; Pavletich, N. P., Recognition of DNA damage by the Rad4 nucleotide excision repair protein. Nature 2007, 449 (7162), 570-575. DOI: 10.1038/nature06155. 140. Sugasawa, K.; Shimizu, Y.; Iwai, S.; Hanaoka, F., A molecular mechanism for DNA damage recognition by the xeroderma pigmentosum group C protein complex. DNA Repair (Amst) 2002, 1 (1), 95-107. 141. Chelico, L.; Pham, P.; Calabrese, P.; Goodman, M. F., APOBEC3G DNA deaminase acts processively 3' --> 5' on single-stranded DNA. Nat Struct Mol Biol 2006, 13 (5), 392-399. DOI: 10.1038/nsmb1086. 142. Iwatani, Y.; Takeuchi, H.; Strebel, K.; Levin, J. G., Biochemical activities of highly purified, catalytically active human APOBEC3G: correlation with antiviral effect. J Virol 2006, 80 (12), 59926002. DOI: 10.1128/JVI.02680-05. 143. Kitamura, S.; Ode, H.; Nakashima, M.; Imahashi, M.; Naganawa, Y.; Kurosawa, T.; Yokomaku, Y.; Yamane, T.; Watanabe, N.; Suzuki, A.; Sugiura, W.; Iwatani, Y., The APOBEC3C crystal structure and the interface for HIV-1 Vif binding. Nat Struct Mol Biol 2012, 19 (10), 1005-1010. DOI: 10.1038/nsmb.2378. 144. Bohn, J. A.; Thummar, K.; York, A.; Raymond, A.; Brown, W. C.; Bieniasz, P. D.; Hatziioannou, T.; Smith, J. L., APOBEC3H structure reveals an unusual mechanism of interaction with duplex RNA. Nat Commun 2017, 8 (1), 1021. DOI: 10.1038/s41467-017-01309-6. 145. Shi, K.; Carpenter, M. A.; Banerjee, S.; Shaban, N. M.; Kurahashi, K.; Salamango, D. J.; McCann, J. L.; Starrett, G. J.; Duffy, J. V.; Demir, O.; Amaro, R. E.; Harki, D. A.; Harris, R. S.; Aihara, H., Structural basis for targeted DNA cytosine deamination and mutagenesis by APOBEC3A and APOBEC3B. Nat Struct Mol Biol 2017, 24 (2), 131-139. DOI: 10.1038/nsmb.3344. 146. Shi, K.; Carpenter, M. A.; Kurahashi, K.; Harris, R. S.; Aihara, H., Crystal Structure of the DNA Deaminase APOBEC3B Catalytic Domain. J Biol Chem 2015, 290 (47), 28120-28130. DOI: 10.1074/jbc.M115.679951. 147. Siriwardena, S. U.; Guruge, T. A.; Bhagwat, A. S., Characterization of the Catalytic Domain of Human APOBEC3B and the Critical Structural Role for a Conserved Methionine. J Mol Biol 2015, 427 (19), 3042-3055. DOI: 10.1016/j.jmb.2015.08.006. 148. Harjes, S.; Jameson, G. B.; Filichev, V. V.; Edwards, P. J. B.; Harjes, E., NMR-based method of small changes reveals how DNA mutator APOBEC3A interacts with its single-stranded DNA substrate. Nucleic Acids Res 2017, 45 (9), 5602-5613. DOI: 10.1093/nar/gkx196. 149. Kouno, T.; Silvas, T. V.; Hilbert, B. J.; Shandilya, S. M. D.; Bohn, M. F.; Kelch, B. A.; Royer, W. E.; Somasundaran, M.; Kurt Yilmaz, N.; Matsuo, H.; Schiffer, C. A., Crystal structure of APOBEC3A bound to single-stranded DNA reveals structural basis for cytidine deamination and specificity. Nat Commun 2017, 8, 15024. DOI: 10.1038/ncomms15024. 150. Shlyakhtenko, L. S.; Lushnikov, A. Y.; Miyagi, A.; Li, M.; Harris, R. S.; Lyubchenko, Y. L., Atomic force microscopy studies of APOBEC3G oligomerization and dynamics. J Struct Biol 2013, 184 (2), 217-225. DOI: 10.1016/j.jsb.2013.09.008. 151. Salter, J. D.; Krucinska, J.; Raina, J.; Smith, H. C.; Wedekind, J. E., A hydrodynamic analysis of APOBEC3G reveals a monomer-dimer-tetramer self-association that has implications for anti-HIV function. Biochemistry 2009, 48 (45), 10685-10687. DOI: 10.1021/bi901642c. 152. Smith, H. C., RNA binding to APOBEC deaminases; Not simply a substrate for C to U editing. RNA Biol 2016, 1-13. DOI: 10.1080/15476286.2016.1259783. 153. Shaban, N. M.; Shi, K.; Lauer, K. V.; Carpenter, M. A.; Richards, C. M.; Salamango, D.; Wang, J.; Lopresti, M. W.; Banerjee, S.; Levin-Klein, R.; Brown, W. L.; Aihara, H.; Harris, R. S., The Antiviral



and Cancer Genomic DNA Deaminase APOBEC3H Is Regulated by an RNA-Mediated Dimerization Mechanism. Molecular cell 2018, 69 (1), 75-86 e79. DOI: 10.1016/j.molcel.2017.12.010.


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Page 25 of 33 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure Legends Figure 1. Overview of human APOBEC cytidine deaminase functions. (A) During normal function, the APOBEC family members are each expressed in unique cell types. RNA editing by APOBEC enzymes can occur in the small intestine (A1) or in monocytes (A3A). In B-cells, AID deaminations initiate antibody diversification pathways for somatic hypermutation and class-switch recombination. In CD4+ T cells, A3 deamination of retroviral ssDNA intermediates restricts their replication. In CD4+ T cells and monocytes, A3s can restrict foreign DNA in order to reduce the DNA-induced inflammatory response. In germ cells and somatic cells, APOBEC enzymes may restrict retrotransposons through RNA binding or deamination of reverse transcripts. In various tissues, unregulated APOBEC expression can lead to unregulated deaminations in genomic DNA that can initiate genomic instability or cell transformation, which ultimately may lead to cancer. The APOBECs involved in each process are depicted below the figure. (B) Schematic of the fate of the uracil after cytosine deamination in genomic DNA. Without repair, replication of the U results in a T/A base pair instead of a C/G base pair. Alternatively, uracils created by cytosine deamination are removed by uracil DNA glycoslyase (UNG) to create an abasic site (X). This abasic site can be processed by AP endonuclease (APE) that the cleaves the DNA backbone at the abasic site to enable subsequent repair by Base Excision Repair (BER) polymerases, the Mismatch Repair (MMR) pathway, or depending on the number of uracils in the immediate area this can lead to DNA breaks. Alternatively, this lesion can be bypassed through error free mechanisms or gap-filling DNA synthesis can occur by high fidelity polymerases in an error free manner. Depending on cellular conditions, translesion synthesis polymerases may also synthesize DNA within the gap, resulting in error prone repair. (C) A3 enzymes deaminate single-stranded (-) DNA intermediates of retroelements, HIV-1, or endogenous retroviruses. The uracil formed by cytosine deamination in the (-) DNA strand serves as a template for the reverse transcriptase during (+) DNA synthesis, which leads to concomitant G→A mutation in the (+) DNA due to the absence of an unmodified template. In the host nucleus, the uracil can be excised by UNG/APE and depending on the number of uracils this can lead to degradation of the template or repair of uracil and integration of the retroelement or proviral DNA. (D) The APOBEC family in humans is composed of eleven members having either a single or double Zn2+-coordinating domain per polypeptide chain. Of the double domain enzymes, the N-terminal domain (NTD) is not catalytically active, but often serves as a processivity domain. The C-terminal domain (CTD) confers the catalytic activity.

Figure 2. Schematic overview of HIV-1 life cycle and reverse transcription. (A) Major steps involved in the HIV-1 replication cycle and influence of A3 enzymes and Vif. HIV-1 envelope glycoprotein binds to the cell surface receptor to trigger viral fusion. Uncoating of the viral capsid, coupled with reverse transcription, leads to the formation of a double-stranded DNA and a preintegration complex (PIC). The steps of reverse transcription (magnifying glass) are detailed in (B). The PIC-associated viral integrase (vertical box) mediates integration into the host genome. Proviral DNA (red) transcription is mediated by the host RNA polymerase II and HIV-1 Tat, producing viral mRNA transcripts (red) for viral protein production and viral genomic RNA. Protease-mediated maturation occurs after budding. Cellular A3s (purple) are unable to interact with viral RNA undergoing reverse transcription due to the presence of the capsid (hatched line) and must be encapsidated in the virus 25 ACS Paragon Plus Environment


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producer cell to deaminate proviral DNA. The action of A3 enzymes is thwarted by HIV-1 Vif that induces their degradation through the 26S proteasome. Vif (red square) interacts with CBFβ, Elongin C and Cullin 5, which enables recruitment of E3 ligase components (shown as a circle for simplicity) and degradation of A3 enzymes (purple). (B) The genomic RNA (red line) is primed for replication using a host tRNAlys3 primer. Synthesis of the minus strand (-) cDNA (blue line) generates an RNA/DNA hybrid that is a substrate for RNase H. RNase H degrades the genomic RNA strand (hatched red line), leaving the nascent (-) cDNA single-stranded. First strand transfer occurs when the (-) DNA hybridizes to the repeated (R) sequence at the 3' end of the viral RNA. The (-) DNA synthesis resumes and the reverse transcriptase associated RNaseH domain degrades the RNA strand (hatched line). Two purine-rich sequences called the polypurine tracts (PPT) are relatively resistant to RNaseH cleavage and serve as the primer for (+) DNA synthesis. HIV-1 has a central and 3' end PPT (cPPT and PPT). The (+) DNA synthesis enables removal of the tRNA primer. In a process called the second strand transfer, both the (+) and (-) DNA are extended until the entire DNA becomes double-stranded, eventually creating a dsDNA that has the same sequences (U3-R-U5) at both ends (called the long terminal repeats, LTRs). The nucleocapsid protein facilitates the strand transfer. Figure is adapted from Coffin et al.22 (C) Nucleocapsid (NC) (green) coats the DNA during (-) DNA elongation and (+) DNA synthesis, however the A3 (purple) is able to displace the transiently bound NC in order to access the ssDNA (dotted line) created through RNaseH degradation of the template strand.

Figure 3. Consequence of deaminations at 5′CC and 5′TC motifs. (A) The path to a stop codon through deamination of cytosine in the antisense codon for tryptophan in (-) DNA (5′CCA). (B) Deaminations by A3G at 5′CC motifs in the (-) DNA lead to codon changes in the (+) DNA primarily at glycine. Glycine can be changed to serine, glutamic acid, arginine, or lysine by A3G-induced mutations. The second highest number of mutations occur at tryptophan, which leads to a stop codon. The A3F deaminations at 5′TC motifs in the (-) DNA result in much more variable amino acid changes in the (+) DNA. Sequences of the protease gene of ∆Vif HIV-1 exposed to A3G or A3F were analyzed from Ara et al.51, 53.

Figure 4. DNA scanning by facilitated diffusion. (A) Sketch of the DNA showing the negative charge region, which is important for the electrostatic interaction of APOBEC3 enzymes with DNA and enables facilitated diffusion. (B) Sketch depicting the one-dimensional scanning by sliding of the enzyme (shown as a dimer). Sliding is important for a localized search of the substrate. Dotted line represents the diffusion motion of the enzyme. (C) Sketch representing the three-dimensional scanning by jumping. Jumping enables larger translocations, but lacks a local search processes. During jumping, the enzyme microscopically dissociates from the DNA without diffusing into the bulk solution. (D) Sketch demonstrating three-dimensional scanning by intersegmental transfer, which allows for larger translocations that are mediated by an enzyme having a doubly-bound state. An enzyme with two binding domains binds two regions of DNA simultaneously before dissociating from one region before moving to another. (E) Dynamics of A3G jumping consistent with single molecule experiments. A3G monomers bind to ssDNA and can slide rapidly. Over time the monomers oligomerize and the mobility of A3G on ssDNA decreases. The oligomers can spontaneously dissociate enabling each monomer 26 ACS Paragon Plus Environment

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subunit to jump to a distal place on the same ssDNA. The local search process that ensues is again slowed by oligomer formation. (F-G) Dynamics of intersegmental transfer of A3C as revealed by studies on A3C dimerization. (F) An intersegmental transfer can occur when the doubly bound state mediated by a dimer is broken through A3C dimer dissociation. (G) Stable dimers of A3C are unable to undergo intersegmental transfer.

Figure 5. Simplified view of a replication fork. At the replication fork, single stranded DNA in both the leading and lagging strand is protected by bound RPA. There is more RPA bound ssDNA in the lagging strand than the leading strand due to discontinuous synthesis of the lagging strand. In order for A3 enzymes to access the ssDNA formed in the lagging strand, they must be able to compete with RPA.



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For Table of Contents Use Only

In CD4+ T cells, APOBEC3 deoxycytidine deamination of retroviral ssDNA intermediates restricts their replication. However, outside of CD4+ T cells, unregulated APOBEC3 expression can lead to unregulated deoxycytidine deaminations in genomic DNA that can initiate genomic instability and cell transformation, which ultimately may lead to cancer. This “double-edged sword” metaphor is applicable to all APOBEC family members and this perspectives article highlights the beneficial and detrimental actions of the seven APOBEC3 family members.


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Figure 1. Overview of human APOBEC cytidine deaminase functions. (A) During normal function, the APOBEC family members are each expressed in unique cell types. RNA editing by APOBEC enzymes can occur in the small intestine (A1) or in monocytes (A3A). In B-cells, AID deaminations initiate antibody diversification pathways for somatic hypermutation and class-switch recombination. In CD4+ T cells, A3 deamination of retroviral ssDNA intermediates restricts their replication. In CD4+ T cells and monocytes, A3s can restrict foreign DNA in order to reduce the DNA-induced inflammatory response. In germ cells and somatic cells, APOBEC enzymes may restrict retrotransposons through RNA binding or deamination of reverse transcripts. In various tissues, unregulated APOBEC expression can lead to unregulated deaminations in genomic DNA that can initiate genomic instability or cell transformation, which ultimately may lead to cancer. The APOBECs involved in each process are depicted below the figure. (B) Schematic of the fate of the uracil after cytosine deamination in genomic DNA. Without repair, replication of the U results in a T/A base pair instead of a C/G base pair. Alternatively, uracils created by cytosine deamination are removed by uracil DNA glycoslyase (UNG) to create an abasic site (X). This abasic site can be processed by AP endonuclease (APE) that the cleaves the DNA backbone at the abasic site to enable subsequent repair by

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Base Excision Repair (BER) polymerases, the Mismatch Repair (MMR) pathway, or depending on the number of uracils in the immediate area this can lead to DNA breaks. Alternatively, this lesion can be bypassed through error free mechanisms or gap-filling DNA synthesis can occur by high fidelity polymerases in an error free manner. Depending on cellular conditions, translesion synthesis polymerases may also synthesize DNA within the gap, resulting in error prone repair. (C) A3 enzymes deaminate single-stranded (-) DNA intermediates of retroelements, HIV-1, or endogenous retroviruses. The uracil formed by cytosine deamination in the (-) DNA strand serves as a template for the reverse transcriptase during (+) DNA synthesis, which leads to concomitant G→A mutation in the (+) DNA due to the absence of an unmodified template. In the host nucleus, the uracil can be excised by UNG/APE and depending on the number of uracils this can lead to degradation of the template or repair of uracil and integration of the retroelement or proviral DNA. (D) The APOBEC family in humans is composed of eleven members having either a single or double Zn2+-coordinating domain per polypeptide chain. Of the double domain enzymes, the N-terminal domain (NTD) is not catalytically active, but often serves as a processivity domain. The C-terminal domain (CTD) confers the catalytic activity. 190x226mm (300 x 300 DPI)


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Figure 2. Schematic overview of HIV-1 life cycle and reverse transcription. (A) Major steps involved in the HIV-1 replication cycle and influence of A3 enzymes and Vif. HIV-1 envelope glycoprotein binds to the cell surface receptor to trigger viral fusion. Uncoating of the viral capsid, coupled with reverse transcription, leads to the formation of a double-stranded DNA and a pre-integration complex (PIC). The steps of reverse transcription (magnifying glass) are detailed in (B). The PIC-associated viral integrase (vertical box) mediates integration into the host genome. Proviral DNA (red) transcription is mediated by the host RNA polymerase II and HIV-1 Tat, producing viral mRNA transcripts (red) for viral protein production and viral genomic RNA. Protease-mediated maturation occurs after budding. Cellular A3s (purple) are unable to interact with viral RNA undergoing reverse transcription due to the presence of the capsid (hatched line) and must be encapsidated in the virus producer cell to deaminate proviral DNA. The action of A3 enzymes is thwarted by HIV-1 Vif that induces their degradation through the 26S proteasome. Vif (red square) interacts with CBFβ, Elongin C and Cullin 5, which enables recruitment of E3 ligase components (shown as a circle for simplicity) and degradation of A3 enzymes (purple). (B) The genomic RNA (red line) is primed for replication



using a host tRNAlys3 primer. Synthesis of the minus strand (-) cDNA (blue line) generates an RNA/DNA hybrid that is a substrate for RNase H. RNase H degrades the genomic RNA strand (hatched red line), leaving the nascent (-) cDNA single-stranded. First strand transfer occurs when the (-) DNA hybridizes to the repeated (R) sequence at the 3' end of the viral RNA. The (-) DNA synthesis resumes and the reverse transcriptase associated RNaseH domain degrades the RNA strand (hatched line). Two purine-rich sequences called the polypurine tracts (PPT) are relatively resistant to RNaseH cleavage and serves as the primer for (+) DNA synthesis. HIV-1 has a central and 3' end PPT (cPPT and PPT). The (+) DNA synthesis enables removal of the tRNA primer. In a process called the second strand transfer, both the (+) and (-) DNA are extended until the entire DNA becomes double-stranded, eventually creating a dsDNA that has the same sequences (U3-R-U5) at both ends (called the long terminal repeats, LTRs). The nucleocapsid protein facilitates the strand transfer. Figure is adapted from Coffin et al.22 (C) Nucleocapsid (NC) (green) coats the DNA during (-) DNA elongation and (+) DNA synthesis, however the A3 (purple) is able to displace the transiently bound NC in order to access the ssDNA (dotted line) created through RNaseH degradation of the template strand. 147x219mm (300 x 300 DPI)


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Figure 3. Consequence of deaminations at 5′CC and 5′TC motifs. (A) The path to a stop codon through deamination of cytosine in the antisense codon for tryptophan in (-) DNA (5′CCA). (B) Deaminations by A3G at 5′CC motifs in the (-) DNA lead to codon changes in the (+) DNA primarily at glycine. Glycine can be changed to serine, glutamic acid, arginine, or lysine by A3G-induced mutations. The second highest number of mutations occur at tryptophan, which leads to a stop codon. The A3F deaminations at 5′TC motifs in the (-) DNA result in much more variable amino acid changes in the (+) DNA. Sequences of the protease gene of ∆Vif HIV-1 exposed to A3G or A3F were analyzed from Ara et al.48-49. 120x126mm (300 x 300 DPI)



Figure 4. DNA scanning by facilitated diffusion. (A) Sketch of the DNA showing the negative charge region, which is important for the electrostatic interaction of APOBEC3 enzymes with DNA and enables facilitated diffusion. (B) Sketch depicting the one-dimensional scanning by sliding of the enzyme (shown as a dimer). Sliding is important for a localized search of the substrate. Dotted line represents the diffusion motion of the enzyme. (C) Sketch representing the three-dimensional scanning by jumping. Jumping enables larger translocations, but lacks a local search processes. During jumping, the enzyme microscopically dissociates from the DNA without diffusing into the bulk solution. (D) Sketch demonstrating three-dimensional scanning by intersegmental transfer, which allows for larger translocations that are mediated by an enzyme having a doubly-bound state. An enzyme with two binding domains binds two regions of DNA simultaneously before dissociating from one region before moving to another. (E) Dynamics of A3G jumping as revealed by single molecule experiments. A3G monomers bind to ssDNA and can slide rapidly. Over time the monomers oligomerize and the mobility of A3G on ssDNA decreases. The oligomers can spontaneously dissociate enabling each monomer subunit to jump to a distal place on the same ssDNA. The local search process that ensues is again slowed by oligomer formation. (F-G) Dynamics of intersegmental transfer of A3C as revealed by studies on A3C dimerization. (F) An intersegmental transfer can occur when the doubly bound state mediated by a dimer is broken through A3C dimer dissociation. (G) Stable dimers of A3C are unable to


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undergo intersegmental transfer. 151x168mm (300 x 300 DPI)



Figure 5. Simplified view of a replication fork. At the replication fork, single stranded DNA in both the leading and lagging strand is protected by bound RPA. There is more RPA bound ssDNA in the lagging strand than the leading strand due to discontinuous synthesis of the lagging strand. In order for A3 enzymes to access the ssDNA formed in the lagging strand, they must be able to compete with RPA. 83x29mm (300 x 300 DPI)


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Biochemical Basis of APOBEC3 Deoxycytidine Deaminase Activity on

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