Elsevier

DNA Repair

Volume 27, March 2015, Pages 9-18
DNA Repair

A versatile new tool to quantify abasic sites in DNA and inhibit base excision repair

https://doi.org/10.1016/j.dnarep.2014.12.006Get rights and content

Highlights

Abstract

A number of endogenous and exogenous agents, and cellular processes create abasic (AP) sites in DNA. If unrepaired, AP sites cause mutations, strand breaks and cell death. Aldehyde-reactive agent methoxyamine reacts with AP sites and blocks their repair. Another alkoxyamine, ARP, tags AP sites with a biotin and is used to quantify these sites. We have combined both these abilities into one alkoxyamine, AA3, which reacts with AP sites with a better pH profile and reactivity than ARP. Additionally, AA3 contains an alkyne functionality for bioorthogonal click chemistry that can be used to link a wide variety of biochemical tags to AP sites. We used click chemistry to tag AP sites with biotin and a fluorescent molecule without the use of proteins or enzymes. AA3 has a better reactivity profile than ARP and gives much higher product yields at physiological pH than ARP. It is simpler to use than ARP and its use results in lower background and greater sensitivity for AP site detection. We also show that AA3 inhibits the first enzyme in the repair of abasic sites, APE-1, to about the same extent as methoxyamine. Furthermore, AA3 enhances the ability of an alkylating agent, methylmethane sulfonate, to kill human cells and is more effective in such combination chemotherapy than methoxyamine.

Introduction

Abasic sites are created in cellular DNA through water-mediated depurination or depyrimidination [1]. About 10,000 abasic sites are generated in this way in a human cell every day and these apurinic/apyrimidinic (AP) sites are considered to be the most commonly generated lesions in DNA [2]. AP sites are also created through the action of agents that react with DNA. For example, alkylation of 7-nitrogen of guanine destabilizes the glycosidic linkage and increases the rate of depurination [3]. Furthermore, many damaged bases are repaired via the base excision-repair (BER) pathway, which starts with the excision of the damaged base by a glycosylase creating an AP site [4]. Although the glycosylase action is normally coupled with other enzymes that process the AP sites, an imbalance in the repair enzymes may cause the AP sites to persist.

Replicative DNA polymerases cannot copy AP sites and the progress of the replication fork is blocked at AP sites causing single- and double-strand breaks. Alternately, AP sites may be copied by error-prone translesion-synthesis polymerases that cause base substitution mutations, but allow replication to continue [5]. The strand breaks resulting from unrepaired AP sites may be repaired error-free using homologous recombination, or may be repaired by a non-homologous end-joining process that creates small addition/deletion mutations. If unrepaired, the strand breaks lead to gross chromosome alterations such as translocations and cause cell death [6]. Thus creation of AP sites in the genome and their processing by cellular machinery has profound implications to genome integrity.

Many different techniques have been used to label, identify and quantify AP sites [7], [8], [9], [10], [11], [12], [13], [14]. It is difficult to use some of the techniques with a large number of samples because they either use equipment such as HPLC or use radioisotopes that are incompatible with a clinical setting. Consequently, the most commonly used method for the detection and quantification of AP sites is based on the reaction of an alkoxyamine called aldehyde-reactive probe (ARP) which reacts with the open form of deoxyribose sugar in AP sites forming an oxime and tagging the site with a biotin (Fig. 1A) [8], [10]. An advantage of the use of ARP in labeling AP sites is that multiple samples can be processed in parallel and the reaction products can be spotted on a membrane to create an ELISA-like assay. The biotin is subsequently bound with streptavidin that is linked with horseradish peroxidase [10] and incubated with chemiluminescent substrate, or directly bound to fluorescently tagged streptavidin [15] to obtain an optical readout. ARP has been used to determine AP sites in different mammalian tissues [16], to monitor changes in AP sites during aging [17] and AP sites generated as a result of treatment of cells with carcinogens [18]. It has also been adapted to quantify genomic uracils by excising uracils by uracil-DNA glycosylase to create AP sites followed by ARP treatment [19], [20], [21]. It has been used to determine uracil levels in normal and repair-deficient Escherichia coli cells [20], [21], [22], in normal mammalian tissue [15], [19] and in cancer cells [15].

However, ARP-based assays for AP sites suffer from several drawbacks. ARP contains biotin which is also present in cells, and hence fluorescent labeling of AP sites in living or fixed tissues using ARP results in considerable background (unpublished results). ARP is bulky (MW 331.4) and its reaction with AP sites is likely to be significantly hindered. The presence of biotin within ARP also necessitates use of a protein like streptavidin making the labeling scheme cumbersome and somewhat expensive. Finally, a detailed study of ARP reactivity with AP sites has shown that the reaction creates side products in addition to ARP linked to full-length DNA [23].

Another use of alkoxyamines is in combination chemotherapy. Alkylating agents such as temozolomide (TMZ) kill cancer cells by methylating DNA bases and backbone. However, cells can excise products of TMZ treatment such as 7-methylguanine and 3-methyladenine using DNA glycosylases, repair the resulting AP sites and suppress the effects of TMZ. The alkoxyamine that has been used in combination chemotherapy is methoxyamine (MX; Fig. 1A). MX potentiates the cytotoxic effects of TMZ by reacting with the AP sites created by DNA glycosylases and inhibiting the cleavage of AP sites by the AP endonuclease, APE-1 [24]. Multiple clinical trials using MX as one part of combination chemotherapy are underway or have been completed (Clinical trials.gov identifiers NCT00892385, NCT01658319 and NCT00692159).

To create a more versatile chemical for labeling AP sites, we synthesized a compound, AA3, that couples alkoxyamine chemistry with chemistry of 1,3-dipolar cycloaddition (click chemistry; [25]). The latter chemistry is a well-established bioorthogonal reaction that creates stable triazoles and has been used to label sugars, proteins, DNA and other biomolecules both in vitro and in situ [26], [27], [28]. We show here that AA3 can be used for labeling AP sites without the use of proteins or enzymes and for AP site quantification. Furthermore, we show that AA3 inhibits APE-1 about as well as MX, and is better than MX in a combination chemotherapy regimen.

Section snippets

Synthesis of O-2-propynylhydroxylamine hydrochloride (AA3)

AA3 was synthesized according to Scheme 1.

Propargyl bromide (11.3 mmol, Sigma–Aldrich) was added drop wise into a mixture of tert-butyl-N-hydroxycarbonate (3.7 mmol, Sigma–Aldrich) and sodium carbonate (7.4 mmol) in N,N-dimethylformamide. The reaction mixture was stirred overnight at 70 °C, washed with water and extracted with ethyl acetate three times. The combined organic solution was washed with saturated aqueous sodium chloride (50 mL), dried over sodium sulfate and concentrated. The crude

Design and synthesis of AA3

Methoxyamine (MX) and aldehyde-reactive probe (ARP) are well-known chemicals that react with AP sites (Fig. 1A). While MX does not allow tagging of AP sites, ARP is bulky, contains biotin as the only tag and requires proteins and enzymes for its use [8], [10]. To create a more versatile agent for labeling AP sites with good reactivity, we synthesized a small alkoxyamine with alkyne functionality (see Section 2). This chemical, AA3 (Fig. 1A) should react with AP sites in the same manner as MX,

Funding

National Institutes of Health, Wayne State University.

Conflict of interest

We plan to file a patent application based on some of the results in this manuscript.

Acknowledgements

The work reported here was supported by NIH grant GM 57200 (to A.S.B.), a Wayne State University graduate fellowship (to S.S.) and funds from Wayne State University (to both A.S.B. and Y-H Ahn).

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