Stereospecific PARP Trapping by BMN 673 and Comparison with Olaparib and Rucaparib

Poly(ADP-ribose) polymerase 1 (PARP1) and PARP2 detect DNA damage with great sensitivity (1–5). PARP1 is an abundant nuclear protein that binds damaged DNA through its N-terminal zinc finger motifs, which activates its catalytic C-terminal domain to hydrolyze NAD⁺ and produce linear and branched poly(ADP-ribose) (PAR) chains that can extend over hundreds of ADP-ribose units (1–5). The rapid binding of PARP1 and PARP2 to DNA is critical for the resealing of DNA single-strand breaks (SSB) during base excision repair (BER) and for the repair of topoisomerase I cleavage complexes (6–11). A large number of SSBs are generated endogenously as well as BER intermediates (12, 13). When replication forks encounter SSBs, they generate double-strand breaks (DSB) that need to be repaired by homologous recombination (7, 13–15). Accordingly, PARP inhibition results in the accumulation of recombinogenic substrates marked by RAD51 and γH2AX nuclear foci (16, 17).

Since the discovery of the synthetic lethality of PARP inhibitors in homologous recombination–deficient cells (15, 18–23), the mechanism by which PARP inhibitors exert their cytotoxicity has been dominantly interpreted as an accumulation of unrepaired SSBs resulting from catalytic PARP inhibition (7, 24). Hence, more highly efficacious PARP catalytic inhibitors with IC₅₀ (inhibitory concentration 50%) values reaching the low nanomolar range have been developed (4, 21, 24–26).

Recently, we showed that, in addition to catalytic inhibition, PARP inhibitors exert their cytotoxicity by trapping PARP1 and PARP2 on SSB sites (27). Such PARP–DNA complexes are more effective at killing cancer cells than unrepaired SSBs caused by the absence of PARP. Because the cytotoxicity is mediated by the presence of PARP1 and PARP2, PARP inhibitors have been proposed to act as “PARP poisons”. PARP trapping is also not merely interpreted as resulting from catalytic PARP inhibition, which prevents dissociation of PARP from DNA and is required for repair completion (28), because the potency to trap PARP–DNA complexes varies widely across the different PARP inhibitors and is not correlated with their PARP catalytic inhibition potency (27). Indeed, veliparib is a highly potent catalytic PARP inhibitor with relatively limited trapping of PARP–DNA complexes in comparison with niraparib and olaparib (27). Therefore, we have proposed that PARP inhibitors should be categorized according to PARP poisoning capability in addition to catalytic PARP inhibition.

In the present study, we examined the ability of three clinical inhibitors, olaparib (AstraZeneca), rucaparib (Clovis), and BMN 673 (BioMarin; Fig. 1A) in terms of catalytic PARP inhibition and PARP poisoning. We also evaluated potential off-target effects of these drugs. To do so, we took advantage of the fact that avian cells genetically lack PARP2 (29). PARP1^−/− avian B lymphoblast DT40 cells are equivalent to PARP1 and PARP2 double-knockout cells, and do not have detectable level of poly(ADP-ribosyl)ation (PARylation; refs. 27, 29). We also compared the cytotoxicity of the three PARP inhibitors as single agents in the BRCA-deficient DT40 cells, and in human prostate cancer and Ewing's sarcoma cells, which have been reported to be selectively sensitive to PARP inhibitors (30), in the NCI60 cell line panel, and in combination with the DNA alkylating agents, methyl methane sulfonate (MMS), and temozolomide.

The DT40 cell lines used in this study were obtained from the Laboratory of Radiation Genetics Graduate School of Medicine in Kyoto University (Kyoto, Japan) in 2011–2012. Human prostate cancer cells (DU145) and human breast cancer cells (MDA-MB231) were obtained from the National Cancer Institute Developmental Therapeutics Program (Frederick, USA) in 2011–2012. The human Ewing's sarcoma cell line (EW8) was a kind gift from Dr. Lee Helman, Pediatric Oncology Branch, National Cancer Institute (NCI), NIH (Bethesda, MD) obtained in 2012. We did not authenticate these cells in our laboratory. BMN 673 and LT674 were provided by Dr. Leonard E. Post (BioMarin Pharmaceutical Inc). Olaparib, rucaparib, and temozolomide were obtained from the Drug Synthesis and Chemistry Branch, Developmental Therapeutics Program, DCTD, NCI. Drug stock solutions were made in dimethyl sulfoxide (DMSO) at 10 mmol/L or 100 mmol/L. The stock solutions were stored at −20°C in the dark and diluted in culture medium immediately before use. Of note, 1% or 10% MMS was prepared fresh each time from 99% MMS (129925; Sigma-Aldrich) in PBS, and then diluted in culture medium to final concentration.

To prepare whole-cell lysates, cells were lysed with CelLytic M lysis reagent (C2978; Sigma-Aldrich). After thorough mixing and incubation at 4°C for 30 minutes, lysates were centrifuged at 15,000 × rpm at 4°C for 10 minutes, and supernatants were collected. To prepare chromatin-bound subcellular fraction, 10 million DT40 cells with 10 mL medium in 15 mL tube or semiconfluent human cells with 10 mL medium in 10 cm dish were treated with indicated drugs for 30 minutes or 4 hours, respectively. Cells were collected and fractionated using a Subcellular Protein Fractionation Kit from Thermo Scientific (78840) following the manufacturer's instructions. Immunoblotting was carried out using standard procedures.

Rabbit polyclonal anti-PARP1 antibody (sc-7150) was purchased from Santa Cruz Biotechnology. Rabbit polyclonal anti-histone H3 antibody (07–690) was from Upstate Biotechnology. Rabbit polyclonal anti-PAR polymer antibody (#4336-BPC-100) was from Trevigen. Rabbit polyclonal anti-PARP2 antibody (ab93416) was from Abcam. Secondary antibodies were horseradish peroxidase (HRP)–conjugated antibodies to rabbit immunoglobulin G (IgG; GE Healthcare).

The validated chemiluminescent immunoassay for PAR using commercially available reagents has been described previously (31). The detailed laboratory procedures can be viewed at the website (32).

To measure the sensitivity of cells to drugs, cells were continuously exposed to various concentrations of the drugs for 72 hours in triplicate. For DT40 cells, 200 cells were seeded into 384-well white plates (#6007680 PerkinElmer Life Sciences) in 40 μL of medium per well. For human cells, 1,500 cells were seeded in 96-well while plates (#6005680 PerkinElmer Life Sciences) in 100 μL of medium per well. Cell survival was determined using the ATPlite 1step Kits (PerkinElmer). Briefly, 20 or 50 μL ATPlite solution for 384- or 96-well plate, respectively, was added to each well. After 5 minutes, luminescence was measured with an EnVision 2104 Multilabel Reader (PerkinElmer). The ATP concentration in untreated cells was defined as 100%. Viability (%) of treated cells was defined as ATP concentration of treated cells/ATP concentration of untreated cells × 100.

The cell viability assays across the NCI60 cell panel were carried out by the NCI/NIH Developmental Therapeutics Program (33) by using standard procedures (34–36).

The fluorescence anisotropy experiments were carried out with a 30-bp duplex labeled with 5′-Alexa Fluor 488. The deoxyoligonucleotide (sequence: 5′-Alexa Fluor 488-ACCCTGCTGTGGGCdUGGAGAACAAGGTGAT) was annealed to its cDNA strand in buffer containing 50 mmol/L potassium acetate, 20 mmol/L Tris acetate, 10 mmol/L magnesium acetate, and 1 mmol/L dithiothreitol, pH 7.9. All oligonucleotides were purchased from Integrated DNA Technologies. Uracil-DNA glycosylase and APE1 (New England Biolabs) were added to the annealed DNA sample and incubated at 37°C for 1 hour. The resulting DNA construct contains a DNA nick and a 5′-dRP at the nicked site. The completion of digestion is verified by denaturing PAGE. For fluorescence anisotropy measurements, 250 nmol/L recombinant PARP1 (a kind gift from Dr. Valerie Schreiber, University of Strasbourg, Strasbourg, France), 1 nmol/L DNA construct, and increasing concentration of PARP inhibitors were combined and incubated for 30 minutes in a buffer containing 50 mmol/L Tris–HCl (pH 8.0), 4 mmol/L MgCl₂, 100 mmol/L NaCl, and 100 ng/μL bovine serum albumin (BSA). Of note, 1 mmol/L NAD⁺ was added to the samples to initiate the experiment and the fluorescence anisotropy values were measured at indicated time using an EnVision 2104 Multilabel Reader equipped with a FITC FP Label (PerkinElmer). The control samples lack NAD⁺ or PARP inhibitor. The fluorescence anisotropy values reported here are average of three independent experiments.

We first tested the potency of BMN 673 to inhibit total cellular PARylation in comparison with olaparib and rucaparib. Western blot analyses against PAR using total cell lysates of drug-treated wild-type DT40 cells (27) showed that all three PARP inhibitors reduced total cellular PAR levels in a concentration-dependent manner at submicromolar concentrations, with BMN 673 producing full inhibition at 0.1 μmol/L (Fig. 1B). Figure 1C and Table 1 show a quantitative analysis using the clinically validated PAR ELISA assay (27, 37) with IC₅₀ (inhibitory concentration 50%) and IC₉₀ (inhibitory concentration 90%) for all three drugs in DT40 cells and human prostate DU145 cancer cells. In DT40 cells, BMN 673 was approximately 2-fold more potent than olaparib, and rucaparib was approximately 3-fold less potent than olaparib. In DU145 cells, the three drugs were comparable. We also compared BMN 673 with its isomer, LT674 (Supplementary Fig. S1). LT674 showed no detectable cellular PARylation inhibition by Western blotting and several orders of magnitude less reduction of PAR by ELISA. These results demonstrate that BMN 673, olaparib, and rucaparib are highly potent PARP inhibitors at low nanomolar concentrations, and they are indistinguishable above 0.1 μmol/L as PAR levels become flat, and close to zero under these conditions.

To determine the selective targeting of PARP1 by BMN 673, we examined its single-agent cytotoxicity in wild-type, PARP1^−/−, and BRCA2tr/− [BRCA2 truncated mutant that is deficient in homologous recombination (38)] DT40 cells (Fig. 2A; ref. 27). We measured ATP concentration to evaluate cellular viability across this study. Because alteration of NAD⁺ pool by catalytic PARP inhibition might affect the ATP pool, we checked ATP concentration in BMN 673 treated and untreated cells in the same conditions as Fig. 1B (Supplementary Fig. S2). We confirmed that catalytic PARP inhibition did not alter the total ATP concentration. BMN 673 showed single-agent cytotoxicity at nanomolar concentrations. Yet, PARP1^−/− cells were immune to BMN 673, indicating a PARP1 requirement for the cytotoxicity of BMN 673, and therefore the selective targeting of PARP1 with no off-target effect for BMN 673. Similar results were observed for olaparib (ref. 27; Fig. 2A, second panel from top). Rucaparib also showed PARP-dependent cytotoxicity at low micromolar concentrations (Fig. 2A, third panel from top).

The additional sensitivity of wild-type compared with PARP1^−/− cells is mediated by PARP1 (27, 39). The IC₉₀ of wild-type DT40 cells to BMN 673 was six to 10 times lower than for the other two PARP inhibitors. We also confirmed by flow cytometry analyses the higher cytotoxicity of BMN 673 compared with olaparib and rucaparib (Supplementary Fig. S3A). As expected (20, 27), homologous recombination–deficient BRCA2tr/− cells showed greater sensitivity than wild-type cells to all three drugs. The IC₉₀ of BRCA2tr/− cells to BMN 673 was 25 to 33 times lower than for olaparib and rucaparib (Fig. 2A, bottom). Moreover, LT674, the inactive stereoisomer of BMN 673, was markedly less cytotoxic (∼100-fold) even in the BRCA2-deficient cells (Supplementary Fig. S4).

We next examined the cytotoxicity of each drug in human cancer cells (Fig. 2B). Ewing's sarcoma cells have recently been identified as being selectively sensitive to olaparib (30), and EW8 is an Ewing's sarcoma cell line that carries the EWS-FLI1 translocation (40). DU145, a prostate cancer cell line without TMPRSS2-ERG translocation, is among the most sensitive NCI60 cell lines to olaparib and BMN 673. Both cell lines showed comparable sensitivity curves with the three PARP inhibitors (Fig. 2B). The IC₉₀ of BMN 673 was 10- and 5-fold lower than that of olaparib in DU145 and EW8 cells, respectively (Fig. 2B, bottom). Rucaparib was also markedly less cytotoxic than BMN 673 in EW8 and DU145 cells (Fig. 2B). Consistently, the flow cytometry analyses revealed higher cytotoxic effect in BMN 673 compared with olaparib and rucaparib (Supplementary Fig. S3B). These results indicate that BMN 673 produces greater PARP-mediated cytotoxicity than olaparib and rucaparib.

To test whether the three PARP inhibitors have other cellular target(s) beside PARP1 and PARP2, the drugs were tested at high concentrations in PARP1^−/− DT40 cells (Fig. 2C). Unlike BMN 673 and olaparib, 50 μmol/L rucaparib showed marked PARP1/2-independent cytotoxicity. The off-target effect of rucaparib (with respect to PARP1/2) was also demonstrated in the human MDA-MB231 breast cancer cell line (Fig. 2D), one of the NCI60 cell lines (34, 41), that was insensitive to 100 μmol/L olaparib or BMN 673 (Fig. 3). Rucaparib decreased MDA-MB231 cells viabilities down to 50% at 50 μmol/L, whereas olaparib and BMN 673 showed no apparent effect (Fig. 2D). Moreover the sensitivity data across NCI60 showed that the cytotoxicity profile of rucaparib is not correlated with olaparib and BMN 673, whereas olaparib and BMN 673 are significantly correlated with each other, as expected for drugs with similar targets (ref. 41; Fig. 3; Table 2). Notably, the IC₅₀ of BMN 673 was overall lower than that of olaparib in the olaparib-sensitive cell lines, which is consistent with the fact that BMN 673 has higher PARP-mediated cytotoxicity than olaparib.

Trapping PARP1 and PARP2 on damaged DNA has recently been proposed as a mechanism accounting for the cytotoxicity of olaparib, niraparib, and to a lesser extent veliparib (27). Using a cellular assay to measure PARP trapping on damaged DNA (27), we examined chromatin-bound PARP1 and PARP2 (Fig. 4). Wild-type DT40 and prostate cancer DU145 cells were treated with different concentrations of PARP inhibitors in the presence of 0.01% MMS to produce base damage that recruits PARP1/2 (Fig. 4). DT40 cells only have PARP1 (no PARP2; Fig. 4A), whereas DU145 have both PARP1 and PARP2 (Fig. 4B; ref. 27). PARP1 and PARP2 were not detectable in chromatin-bound fractions without drug exposure (Fig. 4A and B, lanes 1 and 8). Although olaparib and rucaparib induced similar amounts of PARP–DNA complexes (Fig. 4A and B, lanes 2–7), 0.1 μmol/L BMN 673 induced equivalent levels of PARP–DNA complexes as 10 μmol/L olaparib (Fig. 4A and B, lanes 11 and 12), indicating that BMN 673 is approximately 100-fold more potent than olaparib and rucaparib at trapping PARP1 and PARP2.

To further investigate the differential trapping of PARP–DNA complexes by BMN 673 at the molecular level, we expanded PARP1–DNA binding using fluorescence anisotropy (27). A nicked oligonucleotide duplex DNA with a single 5′-dRP end at the break site was used as a fluorescent substrate (Fig. 5A). Its anisotropy was enhanced upon PARP1 binding to the damaged DNA site. PARylation following addition of NAD⁺ reduced the fluorescence anisotropy signal by freeing the DNA. Figure 5B shows that both PARP inhibitors enhanced the fluorescence anisotropy signal, which reflects the stabilization of PARP1–DNA complexes. BMN 673 was approximately 40-fold more potent than olaparib. Time-course experiments following NAD⁺ addition also showed that BMN 673 slowed the dissociation of PARP1–DNA complexes more efficiently than olaparib (Fig. 5C). Together, these results demonstrate that BMN 673 is markedly more effective at trapping PARP than olaparib and rucaparib.

The lack of detectable off-target effects of BMN 673 (with respect to PARP1/2; see Figs. 1 and 2 and Supplementary Fig. S4), and its high potency to trap PARP–DNA complexes (Figs. 4 and 5), led us to test the combinations of BMN 673 with alkylating agents in comparison with olaparib and rucaparib. As expected from the well-established role of PARylation for SSB repair, PARP1^−/− cells were hypersensitive to MMS (compare open and closed circles in Fig. 6A and B). Consistent with the recently established role of PARP–DNA complexes in the cytotoxicity of PARP inhibitors (27, 39), each drug had no impact on MMS sensitivity in PARP1^−/− cells (Fig. 6B), confirming the lack of off-target effects of the three PARP inhibitors tested up to 1 μmol/L concentration.

On the other hand, the three drugs produced supra-additive effects in wild-type DT40 cells treated with MMS in a concentration-dependent manner (Fig. 6A). Notably, the MMS sensitivity of wild-type cells treated with 0.1 μmol/L olaparib, 0.1 μmol/L rucaparib, or 0.001 μmol/L BMN 673 was greater than that of PARP1^−/− cells (Fig. 6A and B). These results indicate that BMN 673 induces PARP-mediated cytotoxicity approximately 100 times more efficiently than olaparib or rucaparib and that its cytotoxicity is mediated not only by PARP catalytic inhibition but also by trapping PARP–DNA complexes.

Human prostate cancer DU145 and Ewing's sarcoma EW8 cells also showed supra-sensitization to MMS at submicromolar concentration for all three PARP inhibitors (Fig. 6C, middle and bottom). However, BMN 673 was clearly more effective than olaparib or rucaparib (Fig. 6C, middle and bottom). Notably, breast cancer MDA-MB231 cells, a cell line tolerant to PARP inhibitors (see above), were not markedly sensitized to MMS even with 1 μmol/L of each PARP inhibitor (Fig. 6C, top).

To extend these findings to a clinically relevant combination, we tested temozolomide, an alkylating agent known to act synergistically with other PARP inhibitors (9, 42, 43). Figure 6D shows that all three PARP inhibitors enhanced the cytotoxicity of temozolomide, with BMN 673 being markedly more potent than olaparib and rucaparib. These results demonstrate that BMN 673 is the most potent drug among the three PARP inhibitors tested in combination with temozolomide.

In this study, we report that BMN 673 is the most potent PARP-trapping drug tested to date. It is approximately 2 orders of magnitude more potent than olaparib both in prostate cancer DU145 and lymphoma DT40 cells for both PARP1 and PARP2 (see Fig. 4). We also show that olaparib and rucaparib trap PARP1 and PARP2 with comparable efficiency. The present results complement our recent study (27) revealing PARP–DNA complex trapping, and comparing olaparib with niraparib and veliparib. Veliparib differed from the two other drugs by its much weaker ability to trap PARP–DNA complexes despite its remarkable activity as a PARP catalytic inhibitor (27).

Our data indicate that BMN 673 is only slightly more potent than olaparib and rucaparib at inhibiting PARP catalytic activity. The differential potencies of the drugs at trapping PARP versus inhibiting PARP catalytic activity may possibly be interpreted as resulting from an allosteric effect of the drugs (27). As shown in Fig. 1A, the chemical structure of BMN 673 is rigid, whereas olaparib and rucaparib are flexible. This might explain their weaker impact on PARP trapping. The binding of PARP inhibitor to the catalytic pocket of PARP1 and PARP2 may enhance the binding between DNA and the DNA-binding domains of PARP, which would be the converse allosteric effect produced by the binding of PARP to DNA, which induces conformational distortions that stimulate the catalytic domain (44). Notably, LT674, the inactive enantiomer of BMN 673, is markedly less active than BMN 673 both at PARP-mediated cytotoxicity and at inhibiting its catalytic activity. This difference between the enantiomers reflects the optimal structure of BMN 673 for PARP binding and the inability of LT674 to fit in the nicotinamide-binding pocket (45). We believe that BMN 673 can now be viewed not only as a valuable anticancer agent but also as a molecular tool to elucidate PARP allosteric regulation. For the comprehensive understanding of the mechanism of differential PARP trapping, further studies such as co-crystal structure analysis will be required.

The nanomolar cytotoxicity of BMN 673 is notably greater than that of rucaparib or olaparib (≥10-fold in lymphoma DT40 and prostate cancer DU145 and ≥ 5-fold in Ewing's sarcoma EW8 cells; see Fig. 2). The potency of BMN 673 as a cytotoxic agent was just reported independently (46). However, these studies did not examine the molecular mechanism of action of BMN 673, especially with respect to PARP trapping. The potency of BMN 673 observed across the NCI60 cell line panel (see Fig. 3) showed significant correlation between BMN 673 and olaparib (Table 2). The greater cytotoxic potency of BMN 673 over olaparib and rucaparib can be related to the trapping of PARP–DNA complexes because knocking out PARP1 in lymphoma DT40 cells, which by itself is well-tolerated despite the fact that DT40 cells also lack PARP2 (29), conferred extreme resistance to BMN 673 (see Fig. 2). Moreover, the greater cytotoxicity of BMN 673 compared with olaparib is correlated with the greater potency of BMN 673 at trapping PARP (see Figs. 4 and 5), while both drugs are equally effective at inhibiting PARP catalytic activity (see Fig. 2). The BMN 673 findings reinforce our proposal (27) that PARP inhibitors should be categorized and evaluated on the basis of both PARP inhibition and PARP trapping.

Our study shows that rucaparib exhibits off-target effect with respect to PARP1 and PARP2 (Fig. 2C), which fits with a previous report showing that rucaparib has more promiscuous inhibitory activity (extending to PARP1-4 and tankyrases) than olaparib (specific to PARP1-4; ref. 47). The NCI60 data also revealed the differences between rucaparib and BMN 673 or olaparib, and the general cytotoxicity of rucaparib irrespective of cell lines and tissue origin (Fig. 3). We speculate that the inhibition of tankyrases may contribute to the broader cytotoxicity of rucaparib as tankyrase-1 RNA interference (RNAi) results in mitotic arrest (48).

Our results provide relevant information for the clinical use of PARP inhibitors. As single agent in BRCA-deficient cells, we found that BMN 673 demonstrates ∼30-fold greater potency in isogenic BRCA2-deficient lymphoma DT40 cells (see Fig. 2). Consistent results have just been reported independently using other cellular systems (45). BMN 673 is also significantly more potent than olaparib in combination with temozolomide or MMS (see Fig. 6; ref. 45), which is consistent with the enhanced trapping of PARP by BMN 673 and olaparib in the presence of MMS (see Fig. 4). Despite the fact that BMN 673 is a highly potent drug by inducing PARP–DNA complexes, it is surprising that half of the NCI60 cell lines are resistant even at 100 μmol/L BMN 673 (see Fig. 3). Further studies are warranted to elucidate why some cell lines are tolerant or selectively sensitive to PARP trapping. One possibility is that sensitive cell lines are deficient in postreplication repair, Fanconi anemia pathway, ATM, homologous recombination (27), or PTEN (49). It will also be important to determine whether the resistant cells exhibit preferential homologous recombination by 53BP1 inactivation (50, 51).

No potential conflicts of interest were disclosed.

Conception and design: J. Murai, S. Takeda, Y. Pommier

Development of methodology: J. Murai, S.-Y.N. Huang, S. Takeda, Y. Pommier

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): J. Murai, S.-Y.N. Huang, A. Renaud, J. Morris, J.H. Doroshow, Y. Pommier

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): J. Murai, S.-Y.N. Huang, Y. Zhang, J. Ji, Y. Pommier

Writing, review, and/or revision of the manuscript: J. Murai, Y. Zhang, J. Ji, B. Teicher, J.H. Doroshow, Y. Pommier

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): Y. Zhang, Y. Pommier

Study supervision: J. Ji, Y. Pommier

The authors thank the Intramural Program of the National Cancer Institute, Center for Cancer Research (CCR), by the Developmental Therapeutics Program (DTP), Division of Cancer Treatment and Diagnosis (DCTD) for sharing the NCI60 data and drugs. The authors also thank CRADA with BioMarin Pharmaceuticals Inc. for providing BMN 673 and LT674 compounds.

J. Murai was a recipient of fellowships from the John Mung Program (Kyoto University) and the Kyoto University Foundation. Y. Pommier and J. Murai were supported by the Intramural Program of the National Cancer Institute, Center for Cancer Research (Z01 BC 006150). J. Murai and S. Takeda were supported by JSPS Core-to-Core Program. J. Murai was supported by JSPS KAKENHI Grant Number 25740016. J. Ji was supported by the federal fund from the National Cancer Institute, National Institutes of Health (Contract No. HHSN261200800001E).

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