Simultaneous Fluorescence Imaging of the Activities of DNases and 3

Sep 10, 2013 - Real-time fluorescence imaging of the activity of nucleases in living cells has been a difficult issue because of unintended degradatio...
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Simultaneous Fluorescence Imaging of the Activities of DNases and 3′ Exonucleases in Living Cells with Chimeric Oligonucleotide Probes Xin Su,† Chen Zhang,† Xiaocui Zhu, Simin Fang, Rui Weng, Xianjin Xiao, and Meiping Zhao* Beijing National Laboratory for Molecular Sciences, MOE Key Laboratory of Bioorganic Chemistry and Molecular Engineering, College of Chemistry and Molecular Engineering, Peking University, Beijing, 100871, China S Supporting Information *

ABSTRACT: Real-time fluorescence imaging of the activity of nucleases in living cells has been a difficult issue because of unintended degradation of the natural oligonucleotides by nontarget nucleases or interactions with other proteins. In this work, we demonstrate two types of highly selective, sensitive, and robust oligonucleotide probes for simultaneous imaging of the activities of two different nucleases in living cells. The probes consist of the desired substrate structure of the target nuclease and partially phosphorothioate modified backbone labeled with fluorophore and quencher for protection from undesired degradation by other nucleases and signal transduction. Upon reaction with the target nuclease, the initially fluorescence quenched probe was cleaved and the fluorophore was separated from the quencher, giving out strong fluorescence signals. Two nucleases, DNase I and Exonuclease III, were employed as model enzymes to demonstrate the concept. In vitro studies proved that the two probes could discriminate their respective target nucleases in serum with high resistance to other coexisting enzymes. The lower limits of detection for DNase I and Exonuclease III were observed to be 40 U/L and 2.0 U/L, respectively. By labeling the two probes with different fluorophores and quenchers, simultaneous visualization of the activities of DNases and 3′ exonucleases was achieved in both HeLa cells and the suspension cells of Arabidopsis thaliana. The developed approaches may greatly facilitate the studies on the intracellular functions of the two nucleases and other related biological processes. The probe design concept may also be further adapted to the detection of many other nucleases.

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important waste-management nuclease with responsibility for degrading extracellular DNA. The deficiency of DNase I may lead to autoimmune disease that affects multiple organ systems such as systemic lupus erythematosus.7 Enzymes with 3′ exonuclease activities, such as E. coli exonuclease III (Exo III) and Mre 11 protein, prefer to catalyze the stepwise removal of mononucleotides from the 3′-hydroxyl end of duplex DNAs.8 They play vital roles in maintaining genome stability. Some of them are also involved in processing and degradation of a variety of RNAs.8 Organisms that lack 3′ exonucleases are found to be more susceptible to cancer, especially under stress conditions.8a In this work, we attempt to construct efficient probes for fluorescence imaging of the above two nucleases in living cells. First, we designed a novel end-capped oligonucleotide fluorescent probe for the detection of DNase I activity (see Probe D3 in Scheme 1A). The probe has five phosphorothioate (PS) internucleoside linkages at both the 3′-end and 5′-end. The fluorophore (FAM) and quencher (BHQ1) are labeled at

luorescence imaging is an attractive technique for in situ and real-time monitoring of the enzymatic reactions in living cells.1 Imaging intracellular enzyme activity not only helps to elucidate the in vivo enzyme functions and related biological pathways but also favors disease diagnosis and drug screening that use enzymes as the biomarkers.1,2 Nucleases are enzymes that cleave the phosphodiester bonds between the nucleotide subunits of nucleic acids. Because of the important functions of nucleases for living organisms, considerable efforts have been devoted to the development of nucleic acid fluorescent probes for various nucleases.1−3 However, most of the available probes are subjected to degradations by various nontarget enzymes in biological backgrounds, which may bring about false positive signals. It remains a critical challenge to image the intracellular activities of a specific nuclease in living cells. Deoxyribonuclease I (DNase I) is an important endodeoxyribonuclease that hydrolyzes single or double-stranded DNA with no nucleotide sequence specificity.4 It is ubiquitously expressed in mammalian tissues and plays essential roles in DNA degradation in cell apoptosis.5 In vivo, the enzymatic activity of DNase I is inhibited by the binding of G-actin and the formed actin-DNase I complex is involved in the formation of cytoskeleton.6 As a secreted protein, it also acts as an © 2013 American Chemical Society

Received: August 17, 2013 Accepted: September 10, 2013 Published: September 10, 2013 9939

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fluorophores, we successfully achieved simultaneous fluorescence imaging of the two enzymes in living cells.

Scheme 1. (A) Schematic Representation of the Probes for Measurement of the Activity of DNasesa and (B) Schematic Representation of Probe E and Its Control Probe Probe E-C for Selective Detection of the 3′ Exonucleases Activityb

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MATERIALS AND METHODS In Vitro Enzymatic Reactions. All the in vitro enzymatic reactions were carried out in 200 μL sealed PCR tubes. The fluorescence intensity of the reaction solution was monitored in real time on Rotor-Gene Q (Qiagen, Germany). The thermal program was 250 cycles at 37 °C with 5 s per cycle, and fluorescence was measured at the end of each cycle. The excitation and emission wavelengths are 470 and 510 nm for FAM and 530 and 550 nm for TAMRA, respectively. The fluorescence gain level is 10. The reaction buffers for different enzymes are listed in Table S1 in Supporting Information. The concentration of the oligonucleotide probes used for the enzymatic reactions was fixed at 200 nM unless otherwise stated. Delivery of Probes into HeLa Cells by Using SLO. Probes were delivered into HeLa cells by using a reversible permeabilization method with streptolysin O (SLO). Adherent cells grown in 96-well plates were incubated at 37 °C with a mixture of 1.6 U/mL SLO (about 0.09 U SLO per 104 cells) and 1.0 μM probe in 100 μL of DPBS for 5 min for cell permeabilization and probe delivery. After incubation, the medium was changed to regular growth medium and incubated at 37 °C for 30 min prior to fluorescence imaging. Delivery of Probes into Arabidopsis thaliana Suspension Cells by Using Electroporation. An ECM 830 electroporation system (Harvard Apparatus, USA) was employed for delivery of the probes into Arabidopsis thaliana suspension cells. The electroporation buffer contains 137 g/L sucrose, 2.4 g/L HEPES, 6 g/L KCl, and 0.6 CaCl2 g/L (pH 7.2). The suspension cells were distributed in electroporation buffer containing 0.5 μM probes. The electroporation process involves two variables. The field strength is 550 V/cm (220 V adding on 4 mm electroporation cuvette) and the pulse length is 15 ms. After this process, the cells were rinsed with fresh culture medium and then transferred to 96-well plates for fluorescence imaging. Fluorescence Imaging. An Olympus IX 71 inverted fluorescence microscope (Japan), equipped with an EvolveEMCCD camera (Photometrics, USA), was used for cell imaging. The fluorescence signals were detected by using WIBA filter with excitation 460−490 nm and emission 510−550 nm for FAM and WG filter with excitation 510−550 nm and emission 590 nm for TAMRA. The cells in the 96-well plate were observed by a 100× objective (HeLa cell) and a 40× objective (Arabidopsis thaliana cell) of the microscopy. Images were acquired by EMCCD (exposure time 10 ms and EM gain 100) and analyzed by ImageJ software.

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Probe D3 can selectively detect the DNases activity in complex biological samples. Probe D3-C is the control probe of Probe D3 to monitor the background signals. bThe red wavy line in the probes indicates that the internucleoside linkages have been phosphorothioate modified.

the fifth base from the 5′-end and fourth base from the 3′-end, respectively. The normal hairpinned sequence between the two labels provides the substrate of DNase I, while the phosphorothioated stem at the ends of the probe acts as the signal transducer and the blocker against various exonucleases. In the absence of DNase I, the hairpin structure is intact and the fluorescence of the fluorophore is quenched by the quencher through fluorescence resonance energy transfer (FRET). Upon addition of DNase I, the unmodified hairpin structure is quickly digested. The resultant short phosphorothioated duplex fragment bearing the fluorophore and the quencher immediately dissociates from each other and leads to strong fluorescence emission. The probe showed high selectivity for DNase I over other nontarget nucleases. Then we further constructed another fluorescent probe for 3′ exonucleases (see Probe E in Scheme 1B). In Probe E, all the internucleoside linkages from the 5′-end to the 3′-end have been PS modified, leaving one unmodified phosphodiester bond between the 3′-end nucleoside and the labeled fluorophore to provide the only cleavable site for 3′ exonucleases. The quencher is labeled at the fourth base from the 3′-end, so the intact probe displays a quenched fluorescence in its free state. In the presence of 3′ exonucleases, the fluorophore is quickly cleaved and releases into the free solution, resulting in a dramatic fluorescence enhancement. To demonstrate the suitability of the above two probes in complex environments, we first applied them for direct measurement of the activity of the two target enzymes in serum samples without any cleanup or preconcentration steps. Then the probes were further introduced into living cells or apoptotic cells for in situ and real-time imaging of the in vivo activities of DNases and 3′ exonucleases, respectively. Finally, by labeling the two probes with two different types of

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RESULTS AND DISCUSSION Construction of End-Capped Oligonucleotide Fluorescent Probes for Selective Detection of DNase I Activity. To develop a substrate probe that is exclusively processed by the target nuclease in complex biological backgrounds such as serum or intracellular environment, a crucial point is to protect the probe from degradation by other coexisting enzymes. Previous study has proved that phosphorothioate (PS) modification of internucleoside linkages at either end of the oligonucleotide sequence can provide protection against 3′- and 5′-exonucleases.3e,9 Taking advantage of this 9940

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Figure 1. (A) Selectivity of Probe D3 (200 nM) to DNase I (1.2 U/mL) over other possible coexisting nucleases (Exo III, 50 U/mL; Exo I, 10 U/ mL; T7 exo, 100 U/mL; λ exo, 2.5 U/mL; SSB, 500 nM; S1 Nuclease, 2000 U/mL). (B) Comparison of the fluorescence responses of Probe D3 (200 nM) and Probe D3-C (200 nM) to 2.0 U/mL DNase I. (C) Time courses of the reaction of Probe D3 (200 nM) with DNase I at different concentrations. (D) Linear working range of the assay: 40−500 U/L, R2 = 0.997.

Table 1. Sequences of the Oligonucleotide Probes Used in the Study sequences (from 5′ to 3′)a

probe Probe Probe Probe Probe Probe Probe Probe Probe Probe

D1 D2 D3 D3-C D4 D4-C D0 E E-C

FAM-C*C*C*C*TCCCCGCACCAATAGGGTGCGGGGA*G*G*G*G-BHQ1 C*A*A*C*T*CA(dT-FAM)CCACCTGGGA(dT-BHQ1)G*A*G*T*T*G C*A*A*C*(dT-FAM)*ACATCACTCGGATG(dT-BHQ1)*A*G*T*T*G C*A*A*C*(dT-FAM)*A*C*A*T*C*A*C*T*C*G*G*A*T*G*(dT-BHQ1)*A*G*T*T*G C*A*A*C*(dT-TAMRA)*ACATCACTCGGATG(dT-BHQ2)*A*G*T*T*G C*A*A*C*(dT-TAMRA)*A*C*A*T*C*A*C*T*C*G*G*A*T*G*(dT-BHQ2)*A*G*T*T*G Dabcyl-GCTTCCTGTAATTCAATAAGCTGGAAGC-FAM A*T*C*A*T*C*T*T*T*A*C*G*C*A*A*G*A*(dT-BHQ1)*G*A*T-FAM A*T*C*A*T*C*T*T*T*A*C*G*C*A*A*G*A*(dT-BHQ1)*G*A*T*-FAM

a The self-complementary parts are underlined. The phosphorothioated nucleotides (at 3′ side) are indicated with an asterisk after the nucleotides. FAM is fluorescein. Dabcyl is 4-((4-(dimethylamino)phenyl)azo)-benzoyl. BHQ1 is Black Hole Quencher 1 and BHQ2 is Black Hole Quencher 2. TAMRA is tetramethylrhodamine. The self-complementary parts are underlined and shown in italic. Probe D3-C, Probe D4-C, and Probe E-C are the negative control probes for Probe D3, Probe D4, and Probe E, respectively.

via a phosphodiester bond without PS modification, which might have been recognized as a cleavable site by Exo III. After BHQ1 was removed, the fluorescence of FAM was quenched by the stacked guanines at the 5′-end through photoinduced electron transfer mechanism,3a,e,11 so the fluorescence plateau was much lower than that of curve a in Figure S1A in the Supporting Information. To address this issue, we moved the fluorophore to the sixth base from 5′ end and the quencher to the fifth base from 3′ end (see Probe D2 in Figure 1A and Table 1). From curve b in Figure S1B in the Supporting Information, the false positive signal caused by Exo III was completely inhibited. However, curve b in Figure S1A in the Supporting Information shows that the increase rate of fluorescence intensity of Probe D2 is much slower than that of Probe D1. This is most probably because the two labels have hampered the enzyme to access the substrate backbone or some of the

property, we designed a type of end-capped oligonucleotide fluorescent probe for selective detection of DNase I. Probe D1 in Figure 1A shows our preliminary design, in which the fluorophore and the quencher were labeled at the 5′ end and 3′ end, respectively. The probe shows rapid and strong fluorescence increase after degradation by DNase I (see curve a in Figure S1A in the Supporting Information) and expected resistance to T7 exonuclease (T7 Exo), which catalyzes the removal of nucleotides from duplex DNA in the 5′ to 3′ direction.10 However, when the probe was incubated with Exo III, a small increase of fluorescence intensity was also observed (see curve a in Figure S1B in the Supporting Information). After careful examination of the sequence and the modifications of Probe D1 (see Table 1), we found that the fluorescence increase was caused by the removal of the quencher by Exo III. The quencher (BHQ1) was labeled at the 3′ end of the probe 9941

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Figure 2. (A) Selectivity of Probe E (200 nM) to Exo III (0.5 U/mL) over other possible coexisting nucleases (DNase I, 1.2 U/mL; T7 exo, 100 U/ mL; λ exo, 2.5 U/mL; Exo I, 10 U/mL; ALP, 1.0 U/mL; T4 pol exo++, 30 U/mL; KF exo++, 50 U/mL). (B) Comparison of the fluorescence responses of Probe E (200 nM) and Probe E-C (200 nM) to 0.5 U/mL Exo III. (C) Time courses of the reaction of Probe E (200 nM) with Exo III at different concentrations. (D) Linear working range of the assay: 2.0−400 U/L, R2 = 0.995.

negative control probe of Probe D3 to rule out other likely sources of background signals. The difference between the signals obtained by Probe D3 and Probe D3-C reflects the DNase I activity in complex biological samples. To evaluate the effects of the PS modifications in Probe D3 on the affinity of DNase I for the substrate, we synthesized a reference probe without any PS-modifications (see Probe D0 in Table S1 in the Supporting Information). We performed kinetic analysis of the hydrolytic reactions of Probe D3 and Probe D0 by DNase I under the same conditions (see Figure S3 in the Supporting Information). The kinetic constants of the two probes were observed to be Km = 136 nM and Vmax = 1.70 nM/s for Probe D3 and Km = 158 nM and Vmax = 1.96 nM/s for Probe D0, respectively, indicating that the end-capped PSmodification and the two labels have no significant influences on the hydrolytic kinetics of DNase I. Figure 1C shows the time courses of the hydrolytic reactions of 200 nM Probe D3 at different concentrations of DNase I. The initial reaction rates obtained from the linear section of the time curves show a good linear relationship with the DNase I concentration in the range from 40 to 500 U/L (Figure 1D). The detection limit was found to be 40 U/L. To investigate the applicability of Probe D3 in detecting DNase I activity in complex environments, we incubated 200 nM Probe D3 and Probe D3-C with the same amount of 800fold diluted fresh mouse serum samples, respectively. Probe D3 gave out strong fluorescence in the diluted serum sample, while no fluorescence signals were observed for Probe D3-C, indicating that there were no significant interfering substances in the serum samples. The DNase I activity in the unspiked and spiked serum samples measured by Probe D3 were shown in Figure S4 in the Supporting Information. The original

degradation products by DNase I is still in a hybridized state, failing to separate the two labels from each other. So we further designed Probe D3, in which the fluorophore and quencher were both labeled at the base on the last phosphorothioated nucleotide from the two ends. From curve c in Figure S1A in the Supporting Information, the fluorescence increase of Probe D3 is as fast as that of Probe D1, while curve c in Figure S1B in the Supporting Information shows that the false positive signal from Exo III has been efficiently eliminated. Then we examined the possible nonspecific interactions of Probe D3 with other nucleases, including Exonuclease I (Exo I),12 Lambda Exonuclease (λ exo),13 T4 gene 32 protein (a single-strand DNA-binding protein, SSB),14 and S1 Nuclease.15 No significant fluorescent signals were observed when Probe D3 was incubated with these proteins, indicating that they will not affect the detection of DNase I in real samples (see Figure 1A). We also tested the fluorescence response of Probe D3 to its complementary strand. As shown in Figure S2 in the Supporting Information, the fluorescence remained unchanged in the presence of the complementary strands. These results may be explained by the strong intramolecular hydrogen bonds in the long stem of Probe D3, especially the PS modified part, which cannot be opened by the complementary strand or the SSB or the cleavage of the loop by S1 Nuclease. As the intracellular environment is very complex, we further synthesized a uniformly PS modified control probe (referred to as Probe D3-C) with the same sequence as Probe D3 to monitor the background signals originating from other nontarget interactions inside the cells (see Scheme 1A and Table 1). Figure 1B shows that Probe D3-C has no response to DNase I, indicating that the uniform PS-modification completely blocked the degradation by DNase I. So Probe D3-C can be used as a 9942

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found to be 0.63 ± 0.02 U/mL with an average recovery of 110 ± 10% (n = 4). From above results, we confirm that Probe E has strong capability in direct measurement of 3′ exonucleases in a complex background. This new probe offers a more sensitive and robust tool for direct fluorescence imaging of 3′ exonucleases in biological samples in comparison with our previous reported method.3e Fluorescence Imaging of DNases Activity in Living Cells and Apoptotic Cells. DNase I is a type of Ca2+/Mg2+dependent endonuclease acting at neutral pH conditions. In recent years, a number of other cation dependent and noncation-dependent endonucleases have been identified, which are responsible for the DNA cleavage under different circumstances.16 In cells, Probe D3 may also be digested by other endonucleases which belong to the same family as DNase I. Thus, the increase of fluorescence intensity of Probe D3 corresponds to the total activity of DNases in the tested samples. As oligonucleotide probes cannot pass cell membranes by themselves, to demonstrate the feasibility of fluorescence imaging of the intracellular activity of these DNases under a fluorescence microscope by using Probe D3, we adopted a reversible permeabilization approach to deliver the probes into the cells by using Streptolysin O (SLO).17 SLO is a membranedamaging protein which can generate pores on the cell membrane within only a few minutes. Thus, by incubation of Probe D3 and SLO with the cells, the probes can readily diffuse into the cytoplasm of living cells through the tiny pores. Figure 3A displays the intracelluar fluorescence images of DNases activity at 30 min after the probes were introduced. Heterogeneously distributed fluorescence signals could be

concentration of DNase I in mouse serum was found to be 86 ± 3 U/mL with an average recovery of 115 ± 13% (n = 4). These results proved that Probe D3 is well suited for direct measurement of serum samples without the need for any sample cleanup steps. The probe also displayed great potential for the detection of DNase I activity in other complex biological samples. Construction of PS-Modified Oligonucleotide Fluorescent Probes for Selective Detection of 3′ Exonucleases. 3′ Exonucleases prefer to catalyze the stepwise removal of mononucleotides from the 3′-hydroxyl end of duplex DNAs. However, the natural phosphodiester bonds between two nucleotide is also subjected to degradation by endonucleases such as DNase I. Inspired by aforementioned results of Probe D1, we designed a novel hairpined probe for 3′ exonucleases by labeling the fluorophore (FAM) at the 3′ end via a phosphodiester bond (see Probe E in Scheme 1B). In Probe E, all the internucleoside phosphodiester bonds have been phosphorothioated for protection against degradation by other nucleases, thus the phosphodiester bond between the fluorophore and the 3′-terminal nucleoside provids the only cleavable site for 3′ exonucleases. The quencher was labeled at the fourth base from 3′ end, which efficiently quenched the fluorescence of the fluorophore in its initial state. In the presence of 3′ exonucleases, the phosphodiester bond between the fluorophore and the 3′-terminal nucleoside is cleaved and the fluorophore is released to the free solution, leading to strong fluorescence emission. As the nucleotides near the 5′ end were all phosphorothioate modified, the 5′ exonucleases won′t affect the signal of the probe. Besides, since the quencher is labeled at a very close position to the 3′ end, little fluorescence change will occur even if the probe is opened by SSB or its complementary sequence. Accordingly, Probe E was designed to be slightly shorter than Probe D3. Moreover, because of the presence of the fluorophore at the 3′ end, alkaline phosphatase (ALP) didn’t induce any signal increase of the probe. The influences of two exonuclease-proficient polymerases were also tested. The signal increases of Probe E in the presence of 30 U/mL T4 pol exo++ and 50 U/mL KF exo++ were both lower than 10% of that generated by 0.5 U/ mL Exo III. Figure 2A summarizes the fluorescence responses of Probe E to various proteins with reference to the target Exo III, which demonstrates the high selectivity of Probe E to 3′ exonucleases against other nontarget proteins. A control probe for Probe E (referred to as Probe E-C) was also synthesized which has the same sequences as Probe E but with all the phosphodiester bonds PS-modified, including those internucleoside linkages and the one between the fluorophore and the 3′-terminal nucleoside. Figure 2B proves that Probe E-C has no response to Exo III. Figure 2C shows the time courses of exonucleolytic reactions of 200 nM Probe E at different concentrations of Exo III. The initial reaction rates of the time curves show good linear relationship with the Exo III concentration in the range from 2.0 to 400 U/L (Figure 2D). The detection limit was found to be 2.0 U/L. To test if Probe E can be used to directly detect the activity of 3′ exonucleases in biological samples, we incubated 10-fold diluted mouse serum with 200 nM of Probe E and Probe E-C, respectively. Probe E produced notable fluorescence signals, while little fluorescence response was observed for Probe E-C, indicating that the complex matrixes in serum have no significant influence on the performance of the probe. The original concentration of 3′ exonucleases in mouse serum was

Figure 3. (A) In vivo fluorescence images of DNases activity in HeLa cells detected with Probe D3 (1.0 μM). The probes were delivered into the cytoplasm of HeLa cells based on a reversible permeabilization approach by using SLO. The images were captured at 30 min after probe delivery. (B) Control experiments with Probe D3-C under the same conditions as in part A. (C) Fluorescence images of DNases activity in apoptotic HeLa cells induced by etoposide. The cells were treated with 68 μM etoposide at 37 °C for 20 h, then incubated with 1.0 μM Probe D3 for 30 min. (D) Control experiment with Probe D3-C under the same conditions as in part C. (A−D) left, bright field; middle, fluorescence image; and right, merged picture. 9943

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Figure 4. (A) In vivo fluorescence images of 3′ exonucleases activity in HeLa cells detected with Probe E (1.0 μM). The probes were delivered into the cytoplasm of HeLa cells by using SLO. The images were captured at 30 min after probe delivery: left, bright field; middle, fluorescence image; and right, merged picture. (B) Control experiments with Probe E-C under the same conditions as part A. (C) HeLa cells stained with DAPI and Probe-E to show the location of the 3′ exonucleases: (a) bright field, (b) fluorescence image of 3′ exonucleases, (c) fluorescence image of DAPI to show the cell nucleus, (d) merged picture of parts a−c. (D) Fluorescence images of 3′ exonucleases activity in HeLa cells treated with114 μM mirin for different time periods.

observed within the cells. The signals in the cytoplasm are relatively weak, most likely because DNase I is inactivated by combination with actin as cytoskeletal structures. The signals around the cellular membranes are slightly higher, most likely indicating the secretion of DNases from the cells to the culture medium. To confirm that the cells are alive after the SLO treatment, we stained the cells with Hoechst 33342 (a bluefluorescence dye for staining of live cells) and propidium iodide (a red-fluorescence dye for staining of dead cells), respectively. From the images shown in Figure S5 in the Supporting Information, the cells are intact with normal cellular morphology after the SLO treatment. We also assayed the viability of HeLa cells after SLO treatment by using Cell Counting Kit-8 (CCK-8). The results shown in Figure S6 in the Supporting Information demonstrate that there is no significant difference between the SLO-treated and normal HeLa cells. Control experiments were performed by using the control probe Probe D3-C (see Figure 3B), which shows negligible fluorescence signals, suggesting that the fluorescence signals of Probe D3 were generated by DNases within the cells. It has been reported that DNase I and other related DNases have critical roles in the cell apoptotic process. We measured the activity of DNases in HeLa cells treated with etoposide, an antitumor drug that inhibits topoisomerase II and induces cell apoptosis.5a HeLa cells were treated with 68 μM etoposide for 20 h followed by incubation with Probe D3 in fresh medium for 30 min without the addition of SLO. The fluorescence images of DNases activity in etoposide-treated HeLa cells were depicted in Figure 3C, which clearly indicated the uptake of the probes by the apoptotic cells. Figure 3C (left) shows the morphology of the cells in apoptosis. It can be seen that the fluorescence signals in the nucleus of the apoptotic cells with destructed morphology are much higher than those in the less affected cells, indicating high DNases activity in late apoptotic stage for gene degradation. To confirm the uptake of Probe D3

by the apoptotic cells, we performed a control study by incubation of Probe D3 with HeLa cells without etoposide treatment in the absence of SLO. From the results shown in Figure S7 in the Supporting Information, Probe D3 could not enter the normal cells with intact morphology. Then we further conducted a control study in which Probe D3-C was incubated with the apoptotic cells without SLO treatment. It can be seen from Figure 3D that fluorescence signals of the control probes are negligible. These results demonstrate the great potential of Probe D3 in visualization of DNases release and DNA degradation in cell apoptosis. Fluorescence Imaging of 3′ Exonucleases in HeLa Cells. The 3′ exonuclease-specific probe (Probe E) was delivered into living HeLa cells with the assistance of SLO. Figure 4A shows bright fluorescence signals of Probe E within HeLa cells, while the control experiments with Probe E-C performed in the same manner showed little fluorescence signal increase (Figure 4B). To further investigate whether the 3′ exonucleases distribute in the cytoplasm or nucleus, we stained the nucleus with DAPI, which preferentially stained dsDNA and produced about 20-fold fluorescence enhancement. Then we delivered Probe E into the cytoplasm of the cells by incubation with SLO. Figure 4C shows that the fluorescence signals of Probe-E was mainly in cytoplasm. Mirin (Z-5-(4-hydroxybenzylidene)-2-imino-1,3-thiazolidin4-one) has been reported to be able to inhibit the 3′ exonuclease activity of Mre11.18 We treated HeLa cells with mirin to investigate its in vivo inhibition effect on the activity of Mre11. HeLa cells were cultured in the medium containing 114 μM mirin. After different exposure time, the medium was replaced with fresh culture medium containing SLO and Probe E. From the images shown in Figure 4D, Probe E emits much weaker fluorescence signals in mirin-treated cells in comparison with those in the untreated cells. With the increase of the mirin treatment time, the fluorescence signals in cells further 9944

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decreased, indicating that the 3′ exonuclease activity of Mre 11 had been significantly inhibited by mirin. For comparison, we also observed the fluorescence images of 3′ exonucleases activity in HeLa cells without mirin treatment at different time periods. From the results shown in Figure S8 in the Supporting Information, in the absence of mirin, the fluorescence signals within the cells actually turned brighter with the increase of incubation time. This confirms that the fluorescence decrease shown in Figure 4D has been caused by the specific inhibition effect of mirin. These results demonstrate that Probe E can be used to screen inhibitors of 3′ exonucleases in situ and in vivo. Simultaneous Fluorescence Imaging of Two Nucleases in Living Cells. From above results, the two probes (Probe D3 and Probe E) showed exceptional selectivity to their respective target nucleases. Next, we attempt to detect the two types of nucleases simultaneously by labeling the two probes with different fluorophores that have separate excitation and emission wavelengths. Probe D4 was synthesized by replacing FAM and BHQ1 in Probe D3 with TAMRA and BHQ2, respectively. A corresponding negative control probe (Probe D4-C, see Table 1) was also prepared to monitor the background signals. Probe D4 and Probe E were then incubated with a mixture solution of DNase I and Exonuclease III to examine possible cross interference between the two reaction systems. Figure S9 in the Supporting Information shows the responses of the two probes and their respective control probes to the nuclease activities in the mixture solution. The results demonstrated that the two probes responded to their target nucleases by generation fluorescence signals at different emission wavelengths with little cross interference. Then Probe D4 and Probe E targeting their respective nucleases were simultaneously delivered into HeLa cells by incubation with SLO. Figure 5A shows the images of the activity of DNases and 3′ exonucleases within the HeLa cells. For comparison, the probes were also separately delivered into the same batch of cells for verification of the fluorescence signals. Figure 5D shows the negative control experimental results obtained with Probe D4-C and Probe E-C. From Figure 5A−D, 3′ exonucleases mainly exist in the cytoplasm, while DNases distributed both in cytoplasm and in nucleus. Interestingly, a previous study reveals that both exonuclease III and DNase I stimulate the ability of Endonuclease G (Endo G), an apoptotic nuclease that is capable of inducing DNA cleavage in cells, to generate dsDNA cleavage products at physiological ionic strength in vitro. Our method offers a useful tool for in vivo investigation of such processes. Besides, the probes can also be employed to investigate the major factors that control the nucleotide hydrolysis by DNase I during the cell cycle. In our previous work, we have observed secretion of large amount of 3′ exonucleases by suspension cells of Arabidopsis thaliana. In this work, we further explored the intracellular activity of 3′ exonucleases in these plant cells. To deliver Probe E into the suspension cells of Arabidopsis thaliana, we adopted electroporation, a rapid and reversible approach to permeabilize cell membrane. In a very short time (15 ms), the probes were introduced into the suspension cells. From Figure 5E, bright fluorescence signals of Probe E can be observed within the cells. We also tried to observe DNase-like activities in the cells with Probe D4. However, no remarkable signal changes of Probe D4 were observed in Arabidopsis thaliana suspension cells, indicating low activity of free DNases in these plant cells. The viability of Arabidopsis thaliana cells after electroporation

Figure 5. (A) Simultaneous fluorescence imaging of the activities of DNases and 3′ exonucleases in HeLa cells with Probe D4 (1.0 μM) and Probe E (1.0 μM). The probes were delivered into the cells by using SLO. The images were captured 30 min after probe delivery. From left to right: bright field; fluorescence image of the activities of 3′ exonucleases; fluorescence image of the activities of DNases; merged picture to visualize the fluorescence images of the activities of the two nucleases within HeLa cells. (B) Control experiments with Probe D4 and Probe E-C under the same conditions as part A to confirm the results for DNases. (C) Control experiments with Probe D4-C and Probe E under the same conditions as part A to confirm the results for 3′ exonucleases. (D) Negative control experiments with Probe D4-C and Probe E-C under the same conditions as part A to show the background signals. (E) Simultaneous fluorescence imaging of the activities of DNases and 3′ exonucleases in suspension cells of Arabidopsis thaliana with Probe D4 (1.0 μM) and Probe E (1.0 μM). The probes were delivered into the cells by using electroporation. The images were captured at 30 min after probe delivery.

treatment was also examined by using the CCK-8. From the data shown in Figure S10 in the Supporting Information, the cell viability before and after the electroporation treatment is generally the same. Above results proved that Probe D4 and Probe E are capable of simultaneously visualizing the intracellular activities of DNases and 3′ exonucleases in different types of living cells, which may benefit in vivo mechanistic studies of the nucleases and facilitate the screening of their respective inhibitors in situ and in real time.

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CONCLUSIONS In summary, we have developed two novel oligonucleotide fluorescent probes which are capable of discriminating DNases and 3′ exonucleases from other nucleases with high selectivity and sensitivity. The probes consist of both natural and PS modified nucleotides, which may preserve the substrate structure of target nucleases while protecting other parts of the probe from being nonspecifically digested. Probe D3/Probe D4 and Probe E have been proven to be highly efficient tools for the research on DNases and 3′ exonucleases activities, respectively. The increases in fluorescence intensity of theses probes directly reveal the respective enzyme activities. Using the developed probes, we successfully detected the activities of DNases and 3′ exonuclease in serum samples without the need 9945

dx.doi.org/10.1021/ac402615c | Anal. Chem. 2013, 85, 9939−9946

Analytical Chemistry

Article

Dodge, A. H.; Sinicropi, D.; Shak, S.; Lazarus, R. A. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 8225−8229. (5) (a) Liu, Q. Y.; Ribecco, M.; Pandey, S.; Walker, P. R.; Sikorska, M. Ann. N.Y. Acad. Sci. 1999, 887, 60−76. (b) Samejima, K.; Earnshaw, W. C. Nat. Rev. Mol. Cell. Biol. 2005, 6, 677−688. (6) Chhabra, D.; Bao, S.; dos Remedios, C. G. Cell. Res. 2002, 12, 207−214. (7) Hakkim, A.; Furnrohr, B. G.; Amann, K.; Laube, B.; Abed, U. A.; Brinkmann, V.; Herrmann, M.; Voll, R. E.; Zychlinsky, A. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 9813−9818. (8) (a) Shevelev, I. V.; Hubscher, U. Nat. Rev. Mol. Cell. Biol. 2002, 3, 364−376. (b) Chekanova, J. A.; Shaw, R. J.; Wills, M. A.; Belostotsky, D. A. J. Biol. Chem. 2000, 275, 33158−33166. (c) Paull, T. T.; Gellert, M. Mol. Cell 1998, 1, 969−979. (9) (a) Schreiber, G.; Koch, E. M.; Neubert, W. J. Nucleic Acids Res. 1985, 13, 7663−7672. (b) Uhlmann, E.; Ryte, A.; Peyman, A. Antisense Nucleic Acid Drug Dev. 1997, 7, 345−350. (10) Kerr, C.; Sadowski, P. D. J. Biol. Chem. 1972, 247, 305−310. (11) (a) Knemeyer, J. P.; Marme, N.; Sauer, M. Anal. Chem. 2000, 72, 3717−3724. (b) Stohr, K.; Hafner, B.; Nolte, O.; Wolfrum, J.; Sauer, M.; Herten, D. P. Anal. Chem. 2005, 77, 7195−7203. (12) Breyer, W. A.; Matthews, B. W. Nat. Struct. Biol. 2000, 7, 1125− 1128. (13) (a) Perkins, T. T.; Dalal, R. V.; Mitsis, P. G.; Block, S. M. Science 2003, 301, 1914−1918. (b) Subramanian, K.; Rutvisuttinunt, W.; Scott, W.; Myers, R. S. Nucleic Acids Res. 2003, 31, 1585−1596. (14) Williams, K. R.; Konigsberg, W. J. Biol. Chem. 1978, 253, 2463− 2470. (15) Desai, N. A.; Shankar, V. FEMS Microbiol. Rev. 2003, 26, 457− 491. (16) (a) Mukae, N.; Enari, M.; Sakahira, H.; Fukuda, Y.; Inazawa, J.; Toh, H.; Nagata, S. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 9123−9128. (b) Baker, K. P.; Baron, W. F.; Henzel, W. J.; Spencer, S. A. Gene 1998, 215, 281−289. (17) (a) Duncan, J. L.; Buckingham, L. Infect. Immun. 1977, 18, 688− 693. (b) Bhakdi, S.; Tranum-Jensen, J.; Sziegoleit, A. Infect. Immun. 1985, 47, 52−60. (c) Meehl, M. A.; Caparon, M. G. Mol. Microbiol. 2004, 52, 1665−76. (d) Nitin, N.; Bao, G. Bioconjugate Chem. 2008, 19, 2205−2211. (18) Dupre, A.; Boyer-Chatenet, L.; Sattler, R. M.; Modi, A. P.; Lee, J. H.; Nicolette, M. L.; Kopelovich, L.; Jasin, M.; Baer, R.; Paull, T. T.; Gautier, J. Nat. Chem. Biol. 2008, 4, 119−125.

of any cleanup or preconcentration steps. The lower limits of detection were 40 U/L for DNase I and 2.0 U/L for Exo III, respectively. More importantly, the probes enabled direct visualization of the activities of DNases and 3′ exonucleases in both HeLa cells and the suspension cells of Arabidopsis thaliana cells. The distribution of DNases in apoptotic cells and the in vivo inhibition of 3′ exonucleases by mirin were also investigated. Our method offers effective ways for monitoring the intracellular enzymatic reactions, which can provide realtime information of the physiological role of the target nucleases. Furthermore, the probe design concept may be further extended to the detection of many other nucleases, allowing highly selective detection of various nucleases in complex biological samples.

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ASSOCIATED CONTENT

* Supporting Information S

Supplementary methods, Supplementary Table S1, and Supplementary Figures S1−S10. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: 86-10-62758153. Author Contributions †

X.S. and C.Z. contributed equally to this work.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by National Natural Science Foundation of China (Grants 21375004, 91132717, and 21175007) and The Research Fund for the Doctoral Program of Higher Education of China (Grant 20110001110083). Funding for the open access charge was provided by the National Natural Science Foundation of China.

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REFERENCES

(1) Dai, N.; Kool, E. T. Chem. Soc. Rev. 2011, 40, 5756−5770. (2) Wang, K.; Tang, Z.; Yang, C. J.; Kim, Y.; Fang, X.; Li, W.; Wu, Y.; Medley, C. D.; Cao, Z.; Li, J.; Colon, P.; Lin, H.; Tan, W. Angew. Chem., Int. Ed. 2009, 48, 856−870. (3) (a) Song, C.; Zhao, M. Anal. Chem. 2009, 81, 1383−1388. (b) Song, C.; Zhang, C.; Zhao, M. Biosens. Bioelectron. 2009, 25, 301− 305. (c) Song, C.; Zhang, C.; Zhao, M. Biosens. Bioelectron. 2011, 26, 2699−2702. (d) Zhang, C.; Su, X.; Liang, Y.; Zhu, X.; Song, C.; Zhao, M. Biosens. Bioelectron. 2011, 28, 13−16. (e) Su, X.; Zhu, X.; Zhang, C.; Xiao, X.; Zhao, M. Anal. Chem. 2012, 84, 5059−5065. (f) Su, X.; Xiao, X.; Zhang, C.; Zhao, M. Appl. Spectrosc. 2012, 66, 1249−1262. (g) Leung, C. H.; Chan, D. S.; Man, B. Y.; Wang, C. J.; Lam, W.; Cheng, Y. C.; Fong, W. F.; Hsiao, W. L.; Ma, D. L. Anal. Chem. 2011, 83, 463−466. (h) He, H. Z.; Chan, D. S.; Leung, C. H.; Ma, D. L. Chem. Commun. 2012, 48, 9462−9464. (i) Ma, D. L.; He, H. Z.; Leung, K. H.; Zhong, H. J.; Chan, D. S.; Leung, C. H. Chem. Soc. Rev. 2013, 42, 3427−3440. (j) He, H. Z.; Chan, D. S.; Leung, C. H.; Ma, D. L. Nucleic. Acids. Res. 2013, 41, 4345−4359. (k) Leung, C.-H.; Zhong, H.-J.; He, H.-Z.; Lu, L.; Chan, D. S.-H.; Ma, D.-L. Chem. Sci. 2013, 4, 3781−3795. (l) Leung, K. H.; He, H. Z.; Ma, V. P.; Zhong, H. J.; Chan, D. S.; Zhou, J.; Mergny, J. L.; Leung, C. H.; Ma, D. L. Chem. Commun. 2013, 49, 5630−5632. (4) (a) Suck, D.; Oefner, C. Nature 1986, 321, 620−625. (b) Kabsch, W.; Mannherz, H. G.; Suck, D.; Pai, E. F.; Holmes, K. C. Nature 1990, 347, 37−44. (c) Ulmer, J. S.; Herzka, A.; Toy, K. J.; Baker, D. L.; 9946

dx.doi.org/10.1021/ac402615c | Anal. Chem. 2013, 85, 9939−9946