The retinitis pigmentosa-mutated RP2 protein exhibits exonuclease activity and translocates to the nucleus in response to DNA damage

E XP E RI ME N TA L CE L L RE S E A RCH 3 1 2 ( 2 00 6 ) 1 3 2 3 –13 3 4

a v a i l a b l e a t w w w. s c i e n c e d i r e c t . c o m

w w w. e l s e v i e r. c o m / l o c a t e / y e x c r

Research Article

The retinitis pigmentosa-mutated RP2 protein exhibits exonuclease activity and translocates to the nucleus in response to DNA damage Jung-Hoon Yoon a , Junzhuan Qiu b , Sheng Cai c , Yuan Chen c , Michael E. Cheetham d , Binghui Shen b , Gerd P. Pfeifer a,⁎ a

Division of Biology, Beckman Research Institute of the City of Hope, 1500 East Duarte Road, Duarte, CA 91010, USA Department of Radiation Biology, Beckman Research Institute of City of Hope, Duarte, CA 91010, USA c Division of Immunology, Beckman Research Institute of City of Hope, Duarte, CA 91010, USA d Division of Molecular and Cellular Neuroscience, Institute of Ophthalmology, UCL, London EC1V 9EL, UK b

ARTICLE INFORMATION

ABS T R AC T

Article Chronology:

Retinitis pigmentosa (RP) is a genetically heterogeneous disease characterized by

Received 15 November 2005

degeneration of the retina. Mutations in the RP2 gene are linked to the second most

Revised version received

frequent form of X-linked retinitis pigmentosa. RP2 is a plasma membrane-associated

21 December 2005

protein of unknown function. The N-terminal domain of RP2 shares amino acid sequence

Accepted 22 December 2005

similarity to the tubulin-specific chaperone protein co-factor C. The C-terminus consists of a

Available online 2 February 2006

domain with similarity to nucleoside diphosphate kinases (NDKs). Human NDK1, in addition to its role in providing nucleoside triphosphates, has recently been described as a 3′

Keywords:

to 5′ exonuclease. Here, we show that RP2 is a DNA-binding protein that exhibits

Retinitis pigmentosa

exonuclease activity, with a preference for single-stranded or nicked DNA substrates that

RP2

occur as intermediates of base excision repair pathways. Furthermore, we show that RP2

Nucleoside diphosphate kinase

undergoes re-localization into the nucleus upon treatment of cells with DNA damaging

DNA damage

agents inducing oxidative stress, most notably solar simulated light and UVA radiation. The

Exonuclease

data suggest that RP2 may have previously unrecognized roles as a DNA damage response factor and 3′ to 5′ exonuclease. © 2005 Elsevier Inc. All rights reserved.

Introduction Retinitis pigmentosa (RP) is the major form of heritable blindness and occurs in approximately 1 of 3700 individuals [1]. X-linked retinitis pigmentosa (XLRP) is the most severe form of RP. Children with XLRP initially present with night blindness. Disease progression is rapid, with patients functionally blind at the age of 40 to 45 years. The disorder is genetically heterogeneous with several loci on the X chromo-

some being implicated. The RP2 gene was originally identified by positional cloning, and its link to XLRP was confirmed by disease-associated mutations [2–4]. Mutations in RP2 are linked to the second most common cause of XLRP and occur in 10–20% of the XLRP patients. Because mutations in RP2 are strongly associated with the disease, alteration or loss of protein function is likely to account for the development of Xliked retinitis pigmentosa. However, the function of the RP2 protein is currently unknown.

⁎ Corresponding author. Fax: +1 626 358 7703. E-mail address: [email protected] (G.P. Pfeifer). 0014-4827/$ – see front matter © 2005 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2005.12.026

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RP2 has sites for N-terminal acyl modification by myristoylation and palmitoylation and is targeted to the cytoplasmic face of the plasma membrane in cultured cells and in cells of the human retina [5–8]. Sequence analysis suggests that RP2, which consists of 350 amino acid residues, contains two major domains. The N-terminal domain of RP2 shares amino acid sequence similarity with the tubulin-specific chaperone protein co-factor C. Like co-factor C itself, RP2 stimulates the GTPase activity of tubulin in the presence of co-factor D [9]. The C-terminal domain of RP2 (residues 235–350) has sequence similarity to nucleoside diphosphate kinases (NDKs). The majority of the reported RP2 mutations are expected to result in a truncated protein with loss of the Cterminal domain [2,8,10,11]. This may indicate that the Cterminal domain has an important function. Nucleotide diphosphate kinases are involved in the transfer of phosphate groups from ATP to nucleoside and deoxynucleoside diphosphates [12]. This gene family has eight members in the human genome. In addition to their kinase function, there is increasing evidence that some mammalian NDK proteins have direct DNA processing functions. In particular, NDK2 has been shown to be involved in regulation of transcription, by binding to and activating a nucleasehypersensitive element in the c-myc promoter [13]. Certain NDK proteins, such as NDK1, NDK2, and E. coli NDK can cleave DNA sequences with unusual structures in vitro [14–16]. A recent study has shown that NDK1 is directly involved as the DNA cleavage component in a complex that promotes cytotoxic T-lymphocyte-mediated apoptosis [17], and this protein has been characterized biochemically as a 3′ to 5′ exonuclease [18,19]. In order to help elucidate the function of RP2, we have carried out biochemical studies with purified recombinant human RP2 protein. We observe that RP2, similar to human NDK1, contains an unexpected exonuclease activity. We also report that RP2 is translocated from its cytoplasmic plasma membrane localization into the nuclear compartment upon treatment of cells with DNA damaging agents, most notably long wave ultraviolet radiation.

Materials and methods Reagents and oligonucleotides Oligonucleotides for PCR primers and substrates containing uracil (U) were purchased from IDT (Coralville, IA). E. coli uracil DNA glycosylase (UDG) and restriction enzymes were purchased from New England Biolabs (Beverly, MA). All chemicals and reagents were obtained from Sigma (St. Louis, MO).

Construction of plasmids containing NDK proteins, human RP2 and human RP2 mutants A PCR fragment of the E. coli NDK gene was prepared with E. coli genomic DNA, Pfu turbo DNA polymerase (Stratagene, La Jolla, CA) and two primers, forward primer 5′-GGTCGGGATCCGATGGCTATTGAACGT (BamHI site underlined) and reverse primer 5′-GTGCTCGAGTTAACGGGTGCGCGG (XhoI site underlined). To construct plasmids containing human NDKs,

human NDK cDNA clones were obtained from the ATCC (Manassas, VA). Human NDK1 and NDK2 cDNAs were directly subcloned into plasmid pET-28a(+) (Novagen, Madison, WI). For human full-length RP2 and the RP2 deletion mutant containing only the NDK domain, PCR products containing appropriate restriction sites were prepared from an EST clone (ATCC) using Pfu turbo DNA polymerase and specific primers for in frame insertion. The PCR products were cloned into plasmid pET-28b(+). The ligated plasmids were transformed into bacterial competent cells BL21 (DE3) (Stratagene, La Jolla, CA). To construct the point mutants of human RP2, the primers containing mutated sequences were purchased from IDT (Coralville, IA) and the pET-28a(+) vectors containing point mutations of human RP2 were prepared using the Quikchange™ site-directed mutagenesis kit (Stratagene, La Jolla, CA). All plasmids were isolated and sequenced to confirm the cDNA sequence. These expression vectors generate an N-terminal 6x-histidine-tagged open reading frame.

Purification of recombinant human RP2 proteins Recombinant 6x-His-tagged RP2 proteins were purified from E. coli under native conditions using the QIAexpressionist kit (Qiagen, Valencia, CA). The bound proteins were released from the Ni-NTA-agarose column with elution buffer (50 mM sodium phosphate, pH 8.0, 0.1% Triton X-100, 500 mM NaCl, and each of 100 mM, 150 mM and 250 mM imidazole, respectively). Purified RP2 was dialyzed against storage buffer (20 mM HEPES, pH 7.9, 20 mM KCl, 5 mM beta-mercaptoethanol and 40% glycerol). E. coli NDK and human NDK1 and NDK2 were prepared using similar methods. DEAE sepharose (Amersham Pharmacia Biotech, Piscataway, NJ) chromatography was used for additional purification of the RP2 protein. The 250 mM imidazole fraction of human RP2 was dialyzed overnight at 4°C against loading buffer (20 mM Tris–HCl, pH 8.2, 0.5 mM EDTA, 5 mM betamercaptoethanol, and 5% glycerol). Bound proteins were eluted with a linear gradient of 0–500 mM NaCl in loading buffer. The eluted human RP2 protein fractions were dialyzed against storage buffer. Additional gel filtration chromatography of RP2 was carried out with ACA54 columns (Amersham Pharmacia Biotech). This final purified fraction was used in all enzymatic assays of RP2.

Phosphoenzyme and enzyme coupled kinase assay Purified E. coli NDK and human RP2 were tested for kinase activity using a gel-based assay. For phosphoenzyme formation, 500 ng of each protein was used in an autophosphorylation reaction with 1 μM gamma-32P-ATP for 10 min at 37°C in 10 μl reaction buffer (20 mM HEPES, pH 7.9, 20 mM KCl, 2 mM MgCl2, and 20% glycerol). The reaction was stopped by adding SDS gel loading buffer, and the samples were loaded onto a 15% SDS-polyacrylamide gel. After electrophoresis, the wet gel was autoradiographed and then stained with Coomassie blue. The kinase activity of NDKs and human RP2 was determined by an enzyme coupled assay as previously described [15]. Briefly, 1 ml reactions in buffer containing 50 mM Tris–

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HCl, pH 7.4, 50 mM KCI, 6 mM MgCl2, 0.1 mM phosphoenolpyruvate, 0.1 mg/ml NADH, 2 U of pyruvate kinase, 2.5 U of lactate dehydrogenase, and 1 mg/ml BSA (bovine serum albumin) in plastic disposable cuvettes were incubated at room temperature in the presence of ATP as the donor of phosphate and dTDP as the phosphate acceptor nucleotides. The reaction was initiated by the addition of 200 ng of purified NDKs or RP2. NADH oxidation, which produced a decrease in absorbance at 340 nm, was measured by UV spectrophotometry. Nucleoside diphosphate kinase activity of the proteins was determined by calculation of the decreasing value of absorbance per minute at the initial rates where a typical reaction rate is produced.

Exonuclease assays The exonuclease activity of purified human RP2 was tested with a standard 30-mer single-stranded oligonucleotide, 5′CTCGTCAGCATCATGATCATACAGTCAGTG-3′. To verify the substrate specificity, various structurally different oligonucleotide substrates were tested [20]. The oligonucleotides used to make these substrates were 5′-TTGAGGCAGAGTCC (O-1), 5′-GGACTCTGCCTCAA (O-2), 5′-GGACTCT-GCCTCAAG (O-3), 5′-GGACTCTGCCTCAAGACG (O-4), 5′-CACGTTGACTACCGTC (O-5), 5′-GGACTCTGCCTCAAGACGGTAGTCAA-CGTG (O-6), and 5′-GATGTCAAG-CAGTCCTAAGTTTGAGGCAGAGTCC (O-7). The 5′ end labeling of each top strand was carried out with T4 polynucleotide kinase and ATP. The end-labeled oligonucleotides were annealed to their complementary bottom strands in annealing buffer (10 mM Tris– HCl, pH 8.0, 1 mM EDTA, 30 mM NaCl, and 40 mM KCl). The standard assay was performed in a 30 μl volume with

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reaction buffer (20 mM HEPES, pH 7.9, 20 mM KCl, and 2 mM MgCl2), 5 pmol of DNA substrate, and human RP2 (100 ng protein) at 37°C for 1 h. The exonuclease activity of human RP2 mutants was tested with the 5′ end-labeled singlestranded 30-mer in the standard assay as described above. The reactions were terminated by phenol/chloroform extraction and ethanol precipitation. The DNA pellets were dissolved in formamide loading dye buffer. The cleavage of the oligomers was analyzed by 16% denaturing polyacrylamide gel electrophoresis. Double-stranded oligonucleotides containing a U/A base pair were also used. The 5′-labeled 30-mer top strand oligonucleotide was 5′-CTCGTCAGCATCAUGATCATACAGTCAGTG-3′. The top strand was annealed to the bottom strand. The end-labeled double-stranded oligonucleotide containing uracil was first incubated with E. coli UDG (10 U) for 20 min at 37°C. After directly adding 50 ng of human apurinic/apyrimidinic endonuclease 1 (APE1), the mixtures were incubated for an additional 20 min at 37°C before RP2 protein was added. The samples were incubated for an additional 30 min at 37°C. The cleaved DNA was analyzed as described above.

NMR analysis 13

C/15N labeled full-length RP2 and the C-terminal domain of RP2 were produced by growing E. coli cells in modified M9 media supplemented with trace mineral and vitamin mix using 13C-glucose and 15NH4Cl as the carbon and nitrogen sources, respectively. The proteins were purified with Ni-NTA columns, and the samples for NMR spectroscopy contained 0.5

Fig. 1 – Similarity of human RP2 with nucleoside diphosphate kinases. (A) The sequence alignment of E. coli NDK, human NDK1, NDK2, and RP2 protein is shown. Alignment was performed with the Clustal W program. (B) Domain structures of NDKs and human RP2.

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to 1 mM protein, 20 mM sodium phosphate buffer (pH 7.0), and 0.02% sodium azide in 90% H2O/10% D2O. The DNA– protein complex was obtained by titration to the protein with 20 mM unlabeled single-stranded DNA of the sequence 5′-CTCGTCAGCATCATGATCATACAGTCAGTG-3′. All NMR spectra were acquired at 25°C on a Brucker Avance 600 spectrometer equipped with pulse shaping, pulsed field gradient, and a cryoprobe. The program NMRPIPE [21] was used for Fourier transformation, and the program NMRVIEW [22] was used for spectra analysis.

Cell lines and antibodies HeLa cervical carcinoma and ARPE-19 retinal pigment epithelial cells were purchased from the ATCC and grown in DMEM with 10% fetal bovine serum, 2 mM glutamine, 100 U/ml penicillin, and 100 mg/ml streptomycin. Rabbit antisera against HDAC2, beta-tubulin, and H-CAM, normal rabbit IgG, horseradish peroxidase-conjugated goat anti-rabbit IgG, and horseradish peroxidase-conjugated donkey anti-mouse IgG were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). An anti-peptide polyclonal rabbit serum and a sheep antiserum against RP2 were as described before [7].

Immunohistochemistry and Western blot analysis HeLa or ARPE-19 cells were grown on glass coverslips (40–50% confluence) and treated with various DNA damaging agents such as H2O2 (5 or 10 mM, 20 min at 37°C), cisplatin (10–20 μM, 20 min at 37°C), solar simulated UV radiation for 1, 5, or 10 min emitted from a 1000-W solar UV simulator (Oriel Inc., Stratford, CT), UVB light emitted from sunlamps (556 J/m2), and UVA light emitted from two 360-nm black lights (Sylvania 15W F15T8, 1.13 kJ/m2/min). The UV dose emitted from the UV

lamps was measured with a UVX radiometer (Ultraviolet Products, Upland, CA). For DNA damaging agent treatment, the cell culture medium was removed, and cells were treated in PBS. After treatment, the DMEM culture medium was added back, and cells were incubated for an additional 20 min. Cells were fixed in 3.7% (v/v) paraformaldehyde in PBS for 10 min at room temperature and were then washed with PBS. Cells were permeabilized with 0.2% Triton X-100 in PBS for 15 min at room temperature and incubated in blocking buffer (PBS containing 1% bovine serum albumin and 0.2% Tween 20) for 1 h at room temperature. Sheep anti-human RP2 antibody (1:500 dilution) was added in blocking buffer and incubated for 1 h at room temperature. Cells were washed 3 times with PBS containing 0.2% Tween 20 for 5 min each. Alexa Fluor 488 or Alexa Fluor 568-conjugated anti-sheep secondary antibody (1 μg/ml; Molecular Probes, Eugene, OR) was added to the cells and incubated in blocking buffer for 1 h at room temperature followed by three washes as described above. Nuclear counterstaining was performed with 0.2 μg/ml 4′,6′-diamidino-2phenylindole (DAPI) in PBS for 20 min. The cells were washed five times with PBS, mounted with Fluoromount G (Southern Biotechnology Associates; Birmingham, AL), and visualized with a fluorescence microscope (Olympus IX81). For Western blot analysis, HeLa or ARPE-19 cells (1–2 × 106 cells) were treated with various DNA damaging agents for 20 min. Using a nuclear extract preparation kit (Chemicon, Temecula, CA), cellular and nuclear extracts were prepared from DNA damaging agent treated cells. Equivalent amounts (approximately 10 μg) of prepared cytoplasmic and nuclear extracts were mixed with SDS loading buffer and heated at 95°C for 3 min. The proteins were separated on a 15% SDS-polyacrylamide gel and transferred to a PVDF membrane. The blots were probed with polyclonal antibodies against human RP2 (rabbit anti-RP2 peptide antibody), HDAC2, H-CAM,

Fig. 2 – Purified human RP2 protein lacks nucleoside diphosphate kinase activity. (A) The recombinant proteins, obtained after Ni-NTA-agarose and DEAE sepharose chromatography, were resolved on an SDS-polyacrylamide gel and stained with Coomassie Blue. (B) Phosphoenzyme assay. Catalytically active E. coli NDK contains a histidine phosphotransferase activity. This could be demonstrated by a gel-based assay. Recombinant RP2 lacks such activity. (C) Enzymatic assay for nucleoside diphosphate kinase activity. A decrease in absorbance at 340 nm is indicative of the presence of enzymatic activity. This could be demonstrated for E. coli NDK and human NDK1 and NDK2 but not for the human RP2 protein.

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and β-tubulin followed by appropriate secondary antibodies conjugated with horseradish peroxidase. The signals were detected using ECL-Plus (Amersham).

Results Purified human RP2 does not contain nucleoside diphosphate kinase activity The human RP2 protein contains a domain at its C-terminus that has similarity to nucleoside diphosphate kinases (Fig. 1).

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Fig. 1A indicates the degree of similarity between E. coli NDK, human NDK1 and NDK2, and the RP2 protein. Human NDK1 and NDK2 are closely related to the E. coli protein. There is 19% identity and 43% similarity between human NDK2 and the Cterminal NDK-like domain of RP2 (residues 250 to 350), with several conserved sequence blocks. The conserved catalytic histidine residue in the three NDKs (position 117 in E. coli NDK) is replaced by a phenylalanine in RP2, and part of the substratebinding segments for the NDK activity is deleted in the sequence of RP2. E. coli NDK, human NDK1 and NDK2, and human RP2 were expressed and purified as recombinant histidine-tagged

Fig. 3 – Exonuclease activity of purified human RP2 and co-elution of RP2 with exonuclease activity. (A) A single-stranded 30-mer oligonucleotide was labeled at the 5′ end. Ni-NTA-agarose column-purified full-length human RP2 was eluted from a DEAE sepharose column, and the fractions were assayed for exonuclease activity (top panel) and protein content by SDS-PAGE and Coomassie blue staining (bottom panel). No, no enzyme. (B) The Ni-NTA-agarose column-purified C-terminal fragment of RP2, containing the NDK-like domain only, was eluted from a DEAE sepharose column, and the fractions were assayed for exonuclease activity (top panel) and protein content by SDS-PAGE and Coomassie blue staining (bottom panel). No, no enzyme. (C) The peak DEAE sepharose column fractions of full-length RP2 were dialyzed and subjected to gel filtration column chromatography. Individual fractions were assayed for exonuclease activity (top panel) and protein content by SDS-PAGE and Coomassie blue staining (bottom panel).

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proteins from E. coli. We tested the recombinant proteins for potential nucleoside diphosphate kinase activity, both by gelbased detection of a phosphohistidine intermediate and by an enzymatic assay (Fig. 2). Only E. coli NDK and human NDK1 and NDK2 displayed measurable nucleoside diphosphate kinase activities. RP2 did not show autophosphorylation in the gelbased assay and was inactive in the standard dNDP substrate assay.

The 3′ to 5′ exonuclease activity of human RP2 Since NDK1 has been reported to have exonuclease activity [18,19], we searched for exonuclease activity in the recombinant human RP2 protein. We used a 30-mer single-stranded oligonucleotide substrate that was labeled at its 5′ end. Interestingly, human RP2 showed substantial 3′ to 5′ exonuclease activity (Fig. 3A). We found that human RP2 required 1 to 2 mM Mg2+ for optimal reactivity (data not shown). A relatively high amount of RP2 (at least 0.6 pmol of protein per reaction with 5 pmol of DNA) was required to detect significant cleavage with single-stranded DNA substrates. Since the proteins were purified as recombinant proteins from E. coli, we were concerned that the enzyme preparations were potentially contaminated with E. coli proteins having exonuclease activity. To exclude this possibility, we purified human RP2 using three consecutive procedures consisting of Ni-NTA-agarose, DEAE sepharose, and gel filtration columns. In all cases, the RP2 protein band, detected on SDS gels, exactly co-purified with the 3′ to 5′ exonuclease activity (Fig. 3). This argues against the co-purification of an RP2-associated contaminant. In order to test if the 3′ to 5′ exonuclease activity of RP2 is contained in the C-terminal NDK domain alone, we created a recombinant protein consisting of only amino acids 230–350. This protein fragment exhibited only very weak exonuclease activity (Fig. 3B). However, its activity still co-purified with the fractionated protein band, suggesting that this activity is intrinsic to this RP2 fragment. The full-length RP2 protein has a much more pronounced exonuclease activity (Fig. 3A), suggesting that residues in the N-terminal and central domain of RP2 are important for this activity or that correct folding of the protein is required for the activity. The interaction between RP2 and the single-stranded DNA substrate was characterized by NMR chemical shift perturbation. When a protein forms specific interactions with another molecule, such as DNA, the amino acid residues at the binding interface will inevitably experience changes of their environments. Such changes of environments would result in specific changes of chemical shifts of the nuclei of the amino acid residues at the binding interface. The amino acid residues that are not located at the binding interface would not experience such changes of environment and thus would not exhibit any changes of chemical shifts. Due to the large size of full-length RP2 protein (350 residues), most of the resonances were very broad and had low intensity in the 1H-15N TROSY spectrum (Fig. 4, black peaks). However, there were some sharp signals, which correspond to residues that are unstructured and highly dynamic. Sequence analysis indicated that the N-terminal tubulin-binding domain and the C-terminal

Fig. 4 – NMR study with purified human RP2 and a singlestranded oligonucleotide substrate. Superimposed 1H-15N TROSY spectra of full-length RP2, free and bound to DNA. Resonances in black correspond to the spectrum of free RP2 while those in red correspond to the DNA-bound protein when the protein to DNA molar ratio was 1:1. Only a representative region of the spectra is shown.

NDK-like domain are connected by a linker, which is likely flexible and unstructured and some of these residues will produce such sharp resonances. Upon titration of the DNA to the protein, 1H-15N HSQC spectra showed significant chemical shift changes for some of these sharp signals (Fig. 4, red peaks). Since both the protein and DNA are in the identical buffer at identical pH before titration, these changes should only arise from the presence of DNA at or close to the binding sites of the protein. Similar chemical shift perturbation experiments were also performed using the C-terminal NDK domain of RP2, however, no significant chemical shift changes were observed (data not shown). These data indicate that DNA binds specifically to the full-length RP2 but not to the fragment that only contains the NDK domain, and that key DNA-binding sites of the protein include amino acid residues that are flexible and unstructured. To further confirm the exonuclease activity of human RP2, we created and purified several human RP2 mutants and tested their exonuclease activity. Since acidic amino acids, in particular Asp (D) and Glu (E), having carboxylic acid residues commonly associated with divalent cation binding, are often essential for nuclease activity [20,23], we chose several acidic amino acid residues, E234, E283, E328, and D336, which are conserved in the NDK domain (Fig. 1A). We also included a mutant corresponding to the single reported RP-associated missense mutation in the NDK-like domain of RP2 (L253R) [24]. The human RP2 mutants along with the wild-type protein were employed to test the exonuclease activity. The purified mutants are shown in Fig. 5B. The mutants almost completely lost the 3′ to 5′

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Fig. 5 – Exonuclease activity of purified human RP2 mutants. (A) Exonuclease activity of purified human wild type and mutant RP2 proteins was tested with a single-stranded 30-mer oligonucleotide and equal amounts (100 ng) of purified proteins. (B) Ni-NTA column-purified human wild type and mutant RP2 proteins separated on an SDS-PAGE gel. exonuclease activity (Fig. 5A). The mutants exhibited less than 5% of wild-type activity according to densitometric scanning of the cleavage products. L253R had partially

(∼50%) reduced activity. These data confirm that the 3′ to 5′ exonuclease activity of human RP2 is not due to contamination by bacterial nucleases but is an intrinsic

Fig. 6 – Exonuclease activity of human RP2 protein with various DNA substrates. Oligonucleotide substrates with singlestranded, double-stranded, or partially single-stranded configurations were incubated with no protein (none) or with human RP2 (100 ng). The cleavage products were separated on polyacrylamide gels. (A) Composition of the substrates. The asterisks indicate the positions of the label. (B) Cleavage assays. Exonuclease activity of human RP2 was most pronounced with single-stranded substrates or with substrates containing single-stranded 3′ overhangs.

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activity of this protein, and that the disease-associated mutation has reduced exonuclease activity.

Substrate specificity of the exonuclease activity

Fig. 7 – Enzymatic activity of human RP2 protein in presence of base excision repair proteins. Cleavage of an abasic site-containing oligonucleotide by combined reaction with UDG, human APE1, and RP2 protein. The abasic site was created by UDG that excised uracil from a double-stranded oligonucleotide containing a U/A mispair. The abasic site was then cleaved by human APE1 followed by reaction with RP2 protein. Human RP2 showed specific exonuclease activity emanating from the cleaved abasic site.

In order to further substantiate the exonuclease activity present in human RP2, we used a series of oligonucleotide substrates containing double-stranded, single-stranded, or partially single-stranded DNA. Human RP2 was tested in these assays and exhibited significant exonuclease activity with several substrates (Fig. 6). The 3′ to 5′ exonuclease activity was most pronounced with single-stranded DNA substrates or with substrates containing 3′ single-stranded overhangs (e.g., substrates S1, S8, S10). We noticed that a substrate with a 3′-terminal mismatch (S4) was cleaved only slightly better than a substrate without mismatch (S3) (Fig. 6B), suggesting that this exonuclease probably does not have a major role in supporting proofreading activity. We hypothesized that the 3′ to 5′ exonuclease activity of human RP2 might be involved in DNA processing events in concert with other repair pathways, specifically in base excision repair. To test this hypothesis, we investigated human RP2 activity in the presence of a DNA glycosylase and hAPE1, which cleaves abasic sites. First, we found that human RP2 by itself has very weak or no activity towards cleaving abasic sites (data not shown). We used a doublestranded oligonucleotide containing a U/A base pair as a substrate (Fig. 7). E. coli UDG was incubated with the oligonucleotide substrate containing U/A to produce an abasic site. Human APE1 was then added to the mixture to cleave the abasic site followed by the addition of human RP2. RP2 effectively cleaved the double-stranded oligonucleotide substrate to produce a 3′ to 5′ exonuclease cleavage pattern initiated from the AP cleavage site produced by hAPE1 (Fig. 7). These findings suggest that for optimal exonuclease activity, human RP2 might require the coordination with other proteins such as a DNA glycosylase and APE1.

Fig. 8 – Western blot analysis of RP2 in HeLa and ARPE-19 cells. (A) Cytoplasmic and nuclear extracts from DNA damaging agent-treated HeLa cells were separated by SDS-PAGE. Endogenous RP2 in both cytoplasmic (C) and nuclear (N) fractions was detected with anti-human RP2 antibody. HDAC2 was used as a control for the nuclear protein fraction and as a loading standard. H-CAM was used as a membrane/cytoplasmic specific and loading control. Cells were lysed 20 min after DNA damage induction. The “increased ratio” refers to the relative increase in the percentage of nuclear localized protein in damaging agent-treated cells relative to untreated cells. These values were corrected for loading differences using the nuclear and cytoplasmic loading controls. (B) Localization of human RP2 in untreated and DNA damaging agent-treated ARPE-19 retinal epithelial cells. HDAC2 and β-tubulin were used as nuclear and cytoplasmic specific loading controls, respectively.

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Nuclear translocation of human RP2 in response to DNA damage If human RP2 has DNA exonuclease activity, one would expect that this protein should be localized in the nucleus. What is puzzling in this regard is that a predominant plasma membrane localization has been reported for human RP2 [5–8]. However, it is possible that RP2 is transported to the nucleus upon a stimulus and then carries out repair and/or other DNA processing reactions only under specific circumstances. Interestingly, human NDK1/NM23-H1 is found in an ER-associated complex with hAPE1 and other proteins and is translocated to the nucleus upon a pro-apoptotic stimulus triggered by cytotoxic T lymphocytes [17] where it carries out a DNA cleavage reaction. DNA damaging agents also trigger nuclear translocation of human NDK1 [19].

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On the basis of these findings and the data described above, we hypothesized that, when cells are treated with DNA damaging agents, human RP2 may be translocated from the plasma membrane to the nucleus. To investigate RP2 cellular localization, we carried out Western blot and immunohistochemistry studies. HeLa cervical carcinoma and ARPE-19 retinal pigment epithelial cells were treated with various DNA damaging agents including cisplatin, H2O2, and different sources of UV radiation. Cytoplasmic and nuclear extracts were prepared from DNA damaging agent treated cells. Nuclear translocation of RP2 was analyzed by immunoblot of cytosolic and nuclear fractions with anti-RP2 antibody and control antibodies (HDAC2 for the nuclear fraction and betatubulin or H-CAM for the cytoplasmic or membrane fractions). Fig. 8 shows that most of the RP2 protein was predominantly, but not exclusively, in the cytoplasmic/membrane fractions before treatment in both cell types. Intriguingly, nuclear

Fig. 9 – DNA damage-induced nuclear translocation of RP2 in HeLa cells. Untreated and DNA damaging agents-treated HeLa cells were fixed and stained with anti-human RP2 antibody 20 min after DNA damage induction. The signals were visualized by Alexa Fluor 568-conjugated secondary antibody (middle). Control nuclear staining was performed with DAPI (left). Merged images are also shown (right).

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localization of RP2 was significantly increased when cells were treated with DNA damaging agents. Depending on cell type and the DNA damaging agent used, the nuclear translocation level was different. Simulated solar UV light and UVA radiation were most effective, both in ARPE-19 and HeLa cells. Hydrogen peroxide (H2O2) also induced nuclear translocation, but cisplatin was relatively ineffective (Fig. 8). The nuclear translocation of RP2 was also detected by immunohistochemistry with anti-RP2 antibody and fluorescence microscopy. In untreated cells, most of the human RP2 was localized near the plasma membrane (Figs. 9 and 10, no treatment). However, when cells were treated with DNA damaging agents, the anti-RP2 antibody stained both cytoplasmic and nuclear regions (Figs. 9, 10). In some cells treated with the damaging agents, a larger focal or dot signal in the nucleus was detected. RP2 was translocated to the nucleus within 20 min after DNA damage treatment. Hydrogen peroxide, UVA, and simulated sunlight were more effective in inducing nuclear translocation than were UVB or cisplatin. These results indicate that in response to DNA damage, human RP2 might participate in DNA processing reactions in the nucleus. The treatment conditions used did not produce any evidence for apoptotic DNA damage, either by nuclear fragmentation detected by DAPI staining or by DNA laddering detected by agarose gel analysis (not shown).

Fig. 10 – Localization of RP2 in ARPE-19 cells. Untreated and DNA damaging agents-treated ARPE-19 cells were fixed and stained with anti-human RP2 antibody 20 min after DNA damage induction. The signals were visualized by Alexa Fluor 488-conjugated secondary antibody (right). Control nuclear staining was performed with DAPI (left).

Discussion In mammalian cells, NDK1 (NM23-H1) and NDK2 (NM23-H2) have been reported to possess nuclease activities [13– 16,18,19]. Moreover, human NDK1 has been implicated in cytotoxic T-lymphocyte-initiated apoptotic DNA fragmentation [17]. However, the detailed reaction mechanism and the biological role of the nuclease activity of NDK proteins is still unclear. In this report, we have demonstrated that the human RP2 protein, which contains a domain similar to E. coli NDK and several mammalian NDK proteins, contains intrinsic 3′ to 5′ exonuclease activity. Human RP2 tightly co-purified with the 3′ to 5′ exonuclease activity throughout several column chromatographic procedures. A series of RP2 mutants showed no exonuclease activity. The exonuclease activity of RP2 was particularly pronounced when the reaction was part of a base excision repair step involving a DNA glycosylase and APE1 and acting on abasic sites. The data, therefore, suggest that RP2 may be involved in DNA repair processes. Our in vitro data point to a potential role of RP2 in base excision repair processing of abasic sites, although an involvement of RP2 in DNA repair in vivo remains to be proven, a task that may be difficult to achieve, technically, given the redundancy of 3′ to 5′ exonuclease activities in mammalian cells. There are other 3′ to 5′ exonucleases that appear to participate in base excision repair, at least in vitro, such as TREX [25], WRN [26], NDK1 [19], and APE1 contains a 3′ to 5′ exonuclease activity itself [27–30], although this was not obvious under our conditions (Fig. 7). RP2 efficiently cleaved single-stranded substrates or substrates having 3′ single-stranded overhangs. This raises the possibility that the RP2 protein may potentially function in double-strand break repair processes where it may be involved in trimming 3′ single-stranded overhangs before ligation can occur. This possibility requires further investigation. Although we have not detected apoptotic cells under the conditions we used (where RP2 is translocated into the nucleus), we cannot currently exclude the possibility that RP2, under certain circumstances, may function as an apoptotic nuclease in response to DNA damage. Its role in DNA repair or apoptosis may depend on the level of DNA damage sustained by the cell. It has been shown that other proteins involved in DNA repair, such as APE1 [31], also move to the nucleus upon DNA damage. Further studies are required to substantiate a more specific role of mammalian RP2 protein in DNA repair pathways. The data are consistent with the idea that oxidative stress, induced by H2O2, UVA, or simulated sunlight, may be mechanistically responsible for the translocation process, since these were the agents most effective in causing nuclear localization of RP2. Treatment with agents that produce DNA damage predominantly repaired by nucleotide excision repair, such as UVB and cisplatin, were relatively inefficient in inducing nuclear translocation (Figs. 8–10). Oxidative DNA damage is repaired by base excision repair, for example through the action of OGG1 DNA glycosylase [32–37]. UVA and blue light are known to induce oxidative DNA damage [38–40] and, through photosensitizers such as rhodopsin, alltrans retinal, cytochrome C, hemoglobin, or melanin, may induce damage to the DNA of cells in the retina [41–45]. Our

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initial report on new functional characteristics of RP2 suggests the intriguing possibility that the degeneration of the retina in patients with RP2 mutations is a consequence of inefficient repair of such DNA damage. [12]

Acknowledgments [13]

This work was supported by grants from the National Institute of Environmental Health Sciences (ES06070) to G.P. P and by a grant from the National Cancer Institute (CA85344) to B.S.

[14]

REFERENCES [15] [1] J.A. Boughman, P.M. Conneally, W.E. Nance, Population genetic studies of retinitis pigmentosa, Am. J. Hum. Genet. 32 (1980) 223–235. [2] U. Schwahn, S. Lenzner, J. Dong, S. Feil, B. Hinzmann, G. van Duijnhoven, R. Kirschner, M. Hemberger, A.A. Bergen, T. Rosenberg, A.J. Pinckers, R. Fundele, A. Rosenthal, F.P. Cremers, H.H. Ropers, W. Berger, Positional cloning of the gene for X-linked retinitis pigmentosa 2, Nat. Genet. 19 (1998) 327–332. [3] A.J. Mears, L. Gieser, D. Yan, C. Chen, S. Fahrner, S. Hiriyanna, R. Fujita, S.G. Jacobson, P.A. Sieving, A. Swaroop, Protein-truncation mutations in the RP2 gene in a North American cohort of families with X-linked retinitis pigmentosa, Am. J. Hum. Genet. 64 (1999) 897–900. [4] A.J. Hardcastle, D.L. Thiselton, L. Van Maldergem, B.K. Saha, M. Jay, C. Plant, R. Taylor, A.C. Bird, S. Bhattacharya, Mutations in the RP2 gene cause disease in 10% of families with familial X-linked retinitis pigmentosa assessed in this study, Am. J. Hum. Genet. 64 (1999) 1210–1215. [5] J.P. Chapple, A.J. Hardcastle, C. Grayson, L.A. Spackman, K.R. Willison, M.E. Cheetham, Mutations in the N-terminus of the X-linked retinitis pigmentosa protein RP2 interfere with the normal targeting of the protein to the plasma membrane, Hum. Mol. Genet. 9 (2000) 1919–1926. [6] J.P. Chapple, C. Grayson, A.J. Hardcastle, T.A. Bailey, K. Matter, P. Adamson, C.H. Graham, K.R. Willison, M.E. Cheetham, Organization on the plasma membrane of the retinitis pigmentosa protein RP2: investigation of association with detergent-resistant membranes and polarized sorting, Biochem. J. 372 (2003) 427–433. [7] C. Grayson, F. Bartolini, J.P. Chapple, K.R. Willison, A. Bhamidipati, S.A. Lewis, P.J. Luthert, A.J. Hardcastle, N.J. Cowan, M.E. Cheetham, Localization in the human retina of the X-linked retinitis pigmentosa protein RP2, its homologue cofactor C and the RP2 interacting protein Arl3, Hum. Mol. Genet. 11 (2002) 3065–3074. [8] U. Schwahn, N. Paland, S. Techritz, S. Lenzner, W. Berger, Mutations in the X-linked RP2 gene cause intracellular misrouting and loss of the protein, Hum. Mol. Genet. 10 (2001) 1177–1183. [9] F. Bartolini, A. Bhamidipati, S. Thomas, U. Schwahn, S.A. Lewis, N.J. Cowan, Functional overlap between retinitis pigmentosa 2 protein and the tubulin-specific chaperone cofactor C, J. Biol. Chem. 277 (2002) 14629–14634. [10] D. Sharon, M.A. Sandberg, V.W. Rabe, M. Stillberger, T.P. Dryja, E.L. Berson, RP2 and RPGR mutations and clinical correlations in patients with X-linked retinitis pigmentosa, Am. J. Hum. Genet. 73 (2003) 1131–1146. [11] M.G. Miano, F. Testa, F. Filippini, M. Trujillo, I. Conte, C. Lanzara, J.M. Millan, C. De Bernardo, B. Grammatico, M.

[16]

[17]

[18]

[19]

[20]

[21]

[22]

[23]

[24]

[25]

[26]

[27]

[28]

[29]

1333

Mangino, I. Torrente, R. Carrozzo, F. Simonelli, E. Rinaldi, V. Ventruto, M. D'Urso, C. Ayuso, A. Ciccodicola, Identification of novel RP2 mutations in a subset of X-linked retinitis pigmentosa families and prediction of new domains, Hum. Mutat. 18 (2001) 109–119. M.L. Lacombe, L. Milon, A. Munier, J.G. Mehus, D.O. Lambeth, The human Nm23/nucleoside diphosphate kinases, J. Bioenerg. Biomembr. 32 (2000) 247–258. E.H. Postel, S.J. Berberich, S.J. Flint, C.A. Ferrone, Human c-myc transcription factor PuF identified as nm23-H2 nucleoside diphosphate kinase, a candidate suppressor of tumor metastasis, Science 261 (1993) 478–480. E.H. Postel, B.M. Abramczyk, M.N. Levit, S. Kyin, Catalysis of DNA cleavage and nucleoside triphosphate synthesis by NM23-H2/NDP kinase share an active site that implies a DNA repair function, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 14194–14199. D. Ma, Z. Xing, B. Liu, N.G. Pedigo, S.G. Zimmer, Z. Bai, E.H. Postel, D.M. Kaetzel, NM23-H1 and NM23-H2 repress transcriptional activities of nuclease-hypersensitive elements in the platelet-derived growth factor-A promoter, J. Biol. Chem. 277 (2002) 1560–1567. M.N. Levit, B.M. Abramczyk, J.B. Stock, E.H. Postel, Interactions between Escherichia coli nucleoside-diphosphate kinase and DNA, J. Biol. Chem. 277 (2002) 5163–5167. Z. Fan, P.J. Beresford, D.Y. Oh, D. Zhang, J. Lieberman, Tumor suppressor NM23-H1 is a granzyme A-activated DNase during CTL-mediated apoptosis, and the nucleosome assembly protein SET is its inhibitor, Cell 112 (2003) 659–672. D. Ma, J.R. McCorkle, D.M. Kaetzel, The metastasis suppressor NM23-H1 possesses 3′-5′ exonuclease activity, J. Biol. Chem. 279 (2004) 18073–18084. J.-H. Yoon, P. Singh, D.-H. Lee, J. Qiu, S. Cai, T.R. O'Connor, Y. Chen, B. Shen, G.P. Pfeifer, Characterization of the 3′ to 5′ exonuclease activity found in human nucleoside diphosphate kinase 1 (NDK1) and several of its homologues, Biochemistry 44 (2005) 15774–15786. J. Qiu, J.H. Yoon, B. Shen, Search for apoptotic nucleases in yeast: role of Tat-D nuclease in apoptotic DNA degradation, J. Biol. Chem. 280 (2005) 15370–15379. F. Delaglio, S. Grzesiek, G.W. Vuister, G. Zhu, J. Pfeifer, A. Bax, NMRPipe: a multidimensional spectral processing system based on UNIX pipes, J. Biomol. NMR 6 (1995) 277–293. B.A. Johnson, Using NMRView to visualize and analyze the NMR spectra of macromolecules, Methods Mol. Biol. 278 (2004) 313–352. B. Shen, J.P. Nolan, L.A. Sklar, M.S. Park, Essential amino acids for substrate binding and catalysis of human flap endonuclease 1, J. Biol. Chem. 271 (1996) 9173–9176. Y. Wada, M. Nakazawa, T. Abe, M. Tamai, A new Leu253Arg mutation in the RP2 gene in a Japanese family with X-linked retinitis pigmentosa, Invest. Ophthalmol. Visual Sci. 41 (2000) 290–293. D.J. Mazur, F.W. Perrino, Excision of 3′ termini by the Trex1 and TREX2 3′-N5′ exonucleases. Characterization of the recombinant proteins, J. Biol. Chem. 276 (2001) 17022–17029. B. Ahn, J.A. Harrigan, F.E. Indig, D.M. Wilson III, V.A. Bohr, Regulation of WRN helicase activity in human base excision repair, J. Biol. Chem. 279 (2004) 53465–53474. K.M. Chou, Y.C. Cheng, An exonucleolytic activity of human apurinic/apyrimidinic endonuclease on 3′ mispaired DNA, Nature 415 (2002) 655–659. C. Cistulli, O.I. Lavrik, R. Prasad, E. Hou, S.H. Wilson, AP endonuclease and poly(ADP-ribose) polymerase-1 interact with the same base excision repair intermediate, DNA Repair (Amst.) 3 (2004) 581–591. D.M. Wilson III, Properties of and substrate determinants for the exonuclease activity of human apurinic endonuclease Ape1, J. Mol. Biol. 330 (2003) 1027–1037.

1334

E XP E RI ME N TA L CE LL RE S E A RCH 3 1 2 ( 2 00 6 ) 1 3 2 3 –13 3 4

[30] D. Wong, M.S. DeMott, B. Demple, Modulation of the 3′-N5′exonuclease activity of human apurinic endonuclease (Ape1) by its 5′-incised Abasic DNA product, J. Biol. Chem. 278 (2003) 36242–36249. [31] C.V. Ramana, I. Boldogh, T. Izumi, S. Mitra, Activation of apurinic/apyrimidinic endonuclease in human cells by reactive oxygen species and its correlation with their adaptive response to genotoxicity of free radicals, Proc. Natl. Acad. Sci. U. S. A. 95 (1998) 5061–5066. [32] T.K. Hazra, J.W. Hill, T. Izumi, S. Mitra, Multiple DNA glycosylases for repair of 8-oxoguanine and their potential in vivo functions, Prog. Nucleic Acid Res. Mol. Biol. 68 (2001) 193–205. [33] P.A. van der Kemp, D. Thomas, R. Barbey, R. de Oliveira, S. Boiteux, Cloning and expression in Escherichia coli of the OGG1 gene of Saccharomyces cerevisiae, which codes for a DNA glycosylase that excises 7,8-dihydro-8-oxoguanine and 2,6-diamino-4-hydroxy-5-N-methylformamidopyrimidine, Proc. Natl. Acad. Sci. U. S. A. 93 (1996) 5197–5202. [34] K. Arai, K. Morishita, K. Shinmura, T. Kohno, S.R. Kim, T. Nohmi, M. Taniwaki, S. Ohwada, J. Yokota, Cloning of a human homolog of the yeast OGG1 gene that is involved in the repair of oxidative DNA damage, Oncogene 14 (1997) 2857–2861. [35] R. Lu, H.M. Nash, G.L. Verdine, A mammalian DNA repair enzyme that excises oxidatively damaged guanines maps to a locus frequently lost in lung cancer, Curr. Biol. 7 (1997) 397–407. [36] J.P. Radicella, C. Dherin, C. Desmaze, M.S. Fox, S. Boiteux, Cloning and characterization of hOGG1, a human homolog of the OGG1 gene of Saccharomyces cerevisiae, Proc. Natl. Acad. Sci. U. S. A. 94 (1997) 8010–8015.

[37] T.A. Rosenquist, D.O. Zharkov, A.P. Grollman, Cloning and characterization of a mammalian 8-oxoguanine DNA glycosylase, Proc. Natl. Acad. Sci. U. S. A. 94 (1997) 7429–7434. [38] M. Pflaum, C. Kielbassa, M. Garmyn, B. Epe, Oxidative DNA damage induced by visible light in mammalian cells: extent, inhibition by antioxidants and genotoxic effects, Mutat. Res. 408 (1998) 137–146. [39] A. Besaratinia, T.W. Synold, H.H. Chen, C. Chang, B. Xi, A.D. Riggs, G.P. Pfeifer, DNA lesions induced by UV A1 and B radiation in human cells: comparative analyses in the overall genome and in the p53 tumor suppressor gene, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 10058–10063. [40] B.F. Godley, F.A. Shamsi, F.Q. Liang, S.G. Jarrett, S. Davies, M. Boulton, Blue light induces mitochondrial DNA damage and free radical production in epithelial cells, J. Biol. Chem. 280 (2005) 21061–21066. [41] M. Rozanowska, J. Jarvis-Evans, W. Korytowski, M.E. Boulton, J.M. Burke, T. Sarna, Blue light-induced reactivity of retinal age pigment. In vitro generation of oxygen-reactive species, J. Biol. Chem. 270 (1995) 18825–18830. [42] M. Boulton, M. Rozanowska, B. Rozanowski, Retinal photodamage, J. Photochem. Photobiol., B 64 (2001) 144–161. [43] J. Shen, X. Yang, A. Dong, R.M. Petters, Y.W. Peng, F. Wong, P.A. Campochiaro, Oxidative damage is a potential cause of cone cell death in retinitis pigmentosa, J. Cell. Physiol. 203 (2005) 457–464. [44] M. Rozanowska, T. Sarna, Light-induced damage to the retina: role of rhodopsin chromophore revisited, Photochem. Photobiol. 81 (2005) 1305–1330. [45] R.J. Carmody, A.J. McGowan, T.G. Cotter, Reactive oxygen species as mediators of photoreceptor apoptosis in vitro, Exp. Cell Res. 248 (1999) 520–530.