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Centre Alexis Vautrin-CRAN, Unité Mixte de Recherche 7039 Centre National de la Recherche Scientifique-UHP-INPL Nancy-University, Vandoevre-lès-Nancy, France (L.T., N.T., F.G., M.B.-H.); Laboratoire de Mécanique et Ingénierie Cellulaire et Tissulaire, Unité Mixte de Recherche 7563 Centre National de la Recherche Scientifique-INPL, LEMTA et IFR 111 Centre National de la Recherche Scientifique-UHP-INPL-CHU (D.D.), Nancy-University, Vand
uvre-lès-Nancy, France; Laboratoire de Spectrométrie de Masse et de Chimie Laser, Université Paul Verlaine-Metz, Metz, France (M.D., B.M.); and DCPR-GRAPP, Groupe ENSIC-Unité Mixte de Recherche 7630 Centre National de la Recherche Scientifique-INPL, Nancy-University, Nancy, France (C.F.)
(Received November 8, 2006; accepted February 2, 2007)
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). This can be improved by using third-generation PS, which consist of PS to which a tumor-targeting moiety is attached (Sharman et al., 2004). Among these tumor-targeting molecules, peptides are receiving increasing interest in the field of PDT (Schneider et al., 2006). They not only exist in a natural form but also can be designed synthetically as novel molecules. In addition, the effective tissue penetration of short synthetic peptides in combination with their selective binding and internalizing capacity by cancer cells make peptides ideal candidates for the delivery of therapeutic agents such as PS (Schneider et al., 2006).
We recently reported the synthesis and in vitro efficacy of a new peptide-conjugated PS (referred to hereafter as TPC-Ahx-ATWLPPR) having affinity for endothelial cells of the tumor neovasculature by targeting the vascular endothelial growth factor (VEGF165) receptor neuropilin-1 (NRP-1) and not the type 2 VEGF receptor (VEGFR2/KDR), as previously thought, through its peptidic moiety (Tirand et al., 2006). TPC-Ahx-ATWLPPR displayed enhanced uptake and photodynamic properties in endothelial cells compared with its nonconjugated counterpart TPC. Destruction of the neovasculature of tumors may indirectly lead to tumor eradication, following deprivation of life-sustaining nutrients and oxygen (Folkman, 1995; Dougherty et al., 1998), and this antivascular effect is thought to play a major part in the destruction of some vascularized tumors by PDT (Ichikawa et al., 2005).
Despite their numerous advantages over other molecules (e.g., antibodies, proteins) as targeting agents, the main disadvantage of peptides is related to their natural structural conformation, which makes them extremely sensitive to endopeptidases and exopeptidases present in most tissues (Adessi and Soto, 2002). As a result, they often display low stability in biological fluids, which limits their use in vivo. Whereas the PS moiety of TPC-Ahx-ATWLPPR is responsible for the photocytotoxic activity of the conjugate, the peptidic part is responsible for its selectivity. Degradation of the peptidic moiety of the molecule would impair the selectivity of the conjugate and would both allow it to accumulate in normal tissues, where the activated PS could exert nondesirable photocytotoxicity, and decrease the amount of PS delivered to targeted diseased tissues. Therefore, to optimize the in vivo use of the new conjugate TPC-Ahx-ATWLPPR that proved very effective in vitro (Tirand et al., 2006), we sought to investigate its stability after i.v. injection to mice bearing NRP-1-expressing tumors. The metabolic products were identified and quantified by matrix-assisted laser desorption ionization/time of flight (MALDI-TOF) mass spectrometry and reverse-phase high-performance liquid chromatography (HPLC), respectively. Besides, to gain mechanistic insight in the degradation process observed in vivo, the in vitro stability of TPC-Ahx-ATWLPPR was studied in human and mouse plasma, as well as in human umbilical vein endothelial cells (HUVEC) that express NRP-1.
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). Stock solutions of 2 mM in dimethyl sulfoxide, aliquoted and kept in the dark at –20°C, were used.
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Cell Line and Culture. For in vitro experiments, NRP-1-expressing HUVEC, pooled from several donors, were used (Cambrex, Verviers, Belgium) and routinely grown in endothelial growth medium (EGM2), as described previously (Tirand et al., 2006).
Animals and Tumor Model. Female athymic Foxn1 nude mice (nu/nu) were obtained from Harlan (Gannat, France), were 7 to 9 weeks old, and weighed 20 to 25 g. Animal procedures were performed according to institutional and national guidelines. The model of human malignant glioma was obtained using U87 cells, as described previously (Tirand et al., 2006).
In Vivo Metabolism of TPC-Ahx-ATWLPPR. When tumors reached a volume of 70 ± 30 mm3 (15 ± 5 days after tumor grafting), TPC-Ahx-ATWLPPR (2.8 mg/kg) dissolved in polyethylene glycol (PEG) 400/ethanol/water (30:20:50, v/v/v) was injected via the tail vein, and mice were kept in the dark. After a time ranging from 1 to 48 h, mice were anesthetized by i.p. injection of ketamine (Ketalar, Panpharma, Fougères, France) at 60 mg/kg and xylazine (Rompun, Bayer Pharma, Puteaux, France) at 8 mg/kg. Blood samples were collected in heparinized tubes (BD Vacutainer, Becton Dickinson, Plymouth, UK) by cardiac puncture and cooled on ice, and the plasma was separated by centrifugation (700g; 10 min) within 30 min of collection. Mice were sacrificed by cervical dislocation, and the tissues were carefully excised, rinsed with saline, and blotted dry. The tissues selected for dissection were tumor, liver, kidney, spleen, and skin (removed from the back of the mice). At least three animals were used per time point. All the samples were protected from light. Blood, tumor, and organ samples were kept at –80°C in polypropylene tubes until analysis by HPLC and/or MALDI-TOF mass spectrometry.
In Vitro Stability of TPC-Ahx-ATWLPPR. Human and mouse plasma were obtained from whole blood of human volunteers and nude mice, respectively. Mice and human blood samples were collected in heparinized tubes by heart puncture or using an indwelling i.v. cannula placed in the arm, respectively. After immediate centrifugation at 700g for 10 min at 4°C, the supernatant (plasma) was withdrawn and frozen at –80°C in aliquots until required. TPC-Ahx-ATWLPPR was added to prewarmed human or mouse plasma at a final concentration of 1 µM and incubated up to 24 h at 37°C. Samples were taken after 2, 6, and 24 h and kept at –80°C in polypropylene tubes until analysis by HPLC.
For in vitro cell experiments, HUVEC were grown in T-25 flasks (initial seeding concentration: 5.104 cells/ml) for 3 days. EGM2 medium (3 ml) containing 1 µM TPC-Ahx-ATWLPPR (with or without 30 mM ammonium chloride) (Merck Eurolab A.A., Fontenay-sous-Bois, France) was then added. At different times, ranging from 1 to 24 h, the EGM2 cell medium was collected. Subsequently, cells were rinsed twice with cold PBS, trypsinized, counted, centrifuged (220g; 5 min), and kept at –80°C in polypropylene tubes until analysis by HPLC and/or MALDI-TOF mass spectrometry. All the incubations were performed in triplicate.
Preparation of Samples for HPLC and MALDI-TOF Mass Spectrometry Analyses. Extraction of the PS from plasma, tumor, organs, cells, and cell medium was carried out as described previously (Barberi-Heyob et al., 2004; Tirand et al., 2006). To remove surface blood, tissue samples (tumor, organs) were rinsed in physiological saline, blotted dry, and weighed. They were crushed in 500 µl of Tris EDTA molybdate buffer (10 mM Tris, 1.66 mM EDTA, 5 mM molybdate, pH 7.4) using an Ultra-Turrax T25 device (IKA, Labortechnik, Janke and Kunkel, Staufen, Germany). Then, all the samples were spiked with 100 µl of 5,10,15,20-tetrakis(m-hydroxyphenyl)porphyrin (mTHPP) (500 ng/ml in methanol) as an internal standard of extraction. Extraction involved solvent precipitation using methanol combined with dimethyl sulfoxide (5:0.1, v/v). Samples were then vortexed, homogenized for 30 min, and sonicated for 10 min (Branson 1200, Roucaire Instruments Scientifiques, Les Ulis, France). Tissue and cellular debris were removed by centrifugation (2500g; 15 min). The PS-containing organic phase was then concentrated by evaporation at room temperature for at least 3 h with a Speedvac apparatus (Fisher Bioblock Scientific, Illkirch, France) and resuspended into 200 µl of methanol.
Control (blank) plasma, tumor, organs, cells, and cell medium (i.e., containing no exogenous PS) were used to determine whether endogenous constituents coeluted with the PS. Calibration samples, used to construct calibration curves, were prepared by mixing plasma, tumor, organs, cells, or cell medium with appropriate concentrations of PS. For these control and calibration samples, the PS extraction procedure was identical to that described above.
HPLC Analysis. The chromatographic system was composed of a programmable solvent module (System Gold 126, Beckman Coulter, Fullerton, CA), an autosampler injector (507e, System Gold, Beckman Coulter), and a scanning fluorescence detector (RF-10A XL, Shimadzu, Kyoto, Japan). Analyses were performed by reverse-phase HPLC on a C18 analytical column (250 x 4.6 mm i.d., S-5 µm, YMC, Interchim, Montluçon, France) under isocratic elution conditions with a mobile phase of methanol/H2O (95:5, v/v) and a flow rate of 1 ml/min. Fluorescence emission was detected at 652 nm, with an excitation wavelength set at 416 nm. These wavelengths correspond to the maximal intensities of the fluorescence emission and absorption spectra of TPC-Ahx-ATWLPPR, respectively (Tirand et al., 2006). All the chemicals were of analytical grade quality. The chromatograms were acquired and analyzed using GOLD Nouveau software version 1.6 (Beckman Coulter). Quantitations were based on peak areas and deduced from calibration curves.
MALDI-TOF Mass Spectrometry Analysis. One microliter of a 2,5-dihydroxybenzoic acid solution (150 mg/ml in water/acetonitrile, 1:1, v/v) was mixed with 1 µl of sample and spotted on the stainless steel MALDI targets. The solvent was evaporated before insertion in the source. Mass spectra were acquired over the range 0 to 2000 Da. Analyses were performed on a Bruker Reflex IV TOF mass spectrometer (Bruker-Daltonic, Bremen, Germany) equipped with the SCOUT 384 probe ion source. The system used a pulsed nitrogen laser (337 nm, model VSL-337ND, Laser Science Inc., Boston, MA) with a maximal energy of 400 µJ/pulse. The detector signals were amplified and transferred to the XACQ program on a SUN workstation (Sun Microsystems Inc., Palo Alto, CA). Spectra were processed with the XMass 5.1 program (Bruker-Daltonic). External calibration of MALDI mass spectra was carried out using sodic and potassic distribution of PEG 600 and PEG 1500 mixtures.
Intracellular Localization by Confocal Laser Scanning Microscopy. Exponentially growing HUVEC were plated at 103 cells/well in Labtek-II eight-chambered coverslips (Dutscher, Brumath, France). After a 48-h attachment and growth period at 37°C, the cells were incubated with TPC-Ahx-ATWLPPR (1 µM) for 24 h and specific fluorescent organelle markers (Molecular Probes-Europe, Leiden, The Netherlands) using the procedure adapted from the experimental protocol previously reported (Di Stasio et al., 2005). The endoplasmic reticulum was stained by incubating cells for 30 s at room temperature with 1 µg/ml 3,3'-dihexyloxacarbocyanine iodide. Mitochondria were identified after staining the cells for 30 min at 37°C with 0.5 µM MitoTracker Green (Molecular Probes, Eugene, OR). LysoTracker Green (Molecular Probes) was used at a final concentration of 0.2 µM for 30 min to identify lysosomes. To visualize the Golgi apparatus, cells were labeled with 2 µM N-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-pentanoyl) sphingosine (BODIPY FL C5-ceramide) for 20 min at 4°C and then further incubated in dye-free EGM2 medium at 37°C for 30 min. At the end of the double staining, the labeling solution was removed by gentle rinsing with EGM2 medium, and cells were imaged using a confocal laser scanning microscope (TCS SP2-AOBS, Leica Microsystem, Wetzlar, Germany) equipped with a 63x, numerical aperture 1.3 oil immersion objective (Leica). A pinhole of 60.85 µm was used, and each image recorded contained 512 x 512 pixels. An argon laser was used as excitation light at 488 nm for all the organelle probes and TPC-Ahx-ATWLPPR. Fluorescence of the organelle probes was detected on channel 1 with a 505- to 545-nm band-pass emission filter. Channel 2 was used to detect the red fluorescence of TPC-Ahx-ATWLPPR with a 640- to 660-nm band-pass emission filter. The fluorescence images were displayed in green and red "false" colors for organelle markers and PS, respectively.
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In Vitro Stability in Plasma. To assess the involvement of the plasma compartment in the degradation process, we followed the in vitro stability of TPC-Ahx-ATWLPPR in plasma. A concentration of 1 µM was chosen because it represents a relevant in vivo concentration. Indeed, after injection of 2.8 mg/kg TPC-Ahx-ATWLPPR in glioma-bearing mice, the in vivo plasma concentrations of this peptide-PS conjugate were in the range of 200 to 6000 ng/ml (0.1–3.8 µM) from 1 to 48 h p.i. (Fig. 3C). TPC-Ahx-ATWLPPR was stable in vitro in human plasma up to 24 h at 37°C (Fig. 4). Likewise, no degradation could be observed in mouse plasma up to 24 h in the same conditions (not shown). This suggested that the degradation observed in mouse plasma in vivo did not involve plasma peptidases.
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In Vivo Biodistribution and Metabolism in Glioma-Bearing Nude Mice. To explore the biodistribution and stability of TPC-Ahx-ATWLPPR in tumor-bearing mice, PS concentrations were determined in the tumor and organs of mice (n = 3 or 4) at different times. Following i.v. injection into tumor-bearing mice, PS accumulated to the highest levels in liver and spleen (Fig. 5A). It should be noted that part of the PS quantities estimated in the liver, a highly vascularized organ, is caused by PS present in the blood circulation because our values have not been corrected for blood content of the tissue. Moreover, endogenous porphyrins (e.g., heme) are present at significant levels in the liver. These porphyrins could not be completely separated from our PS; indeed, part of their elution peaks superimposed to the ones of our PS (i.e., similar retention times). As a result, the real PS quantities in the liver are in fact lower than those presented in Fig. 5A but would remain much higher than the PS quantities in all the other organs considered. Total PS levels in the tumor were higher than in the skin at all the time points (Fig. 5A).
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Whereas TPC-Ahx-ATWLPPR was stable in vitro in plasma for at least 24 h, a degradation product could be observed in vivo in plasma (Fig. 3A2), as well as in other organs (Fig. 5A). In the organs of the reticuloendothelial system, TPC-Ahx-ATWLPPR was rapidly degraded, and the major part of the hydrolysis of the peptide bond(s) occurred in the liver (Fig. 5A). Indeed, the degradation product represented more than 85% of the total amount of PS present in the liver as early as 2 h p.i. TPC-Ahx-A was characterized as the main metabolic product from liver by MALDI-TOF analysis (Fig. 5B). On the contrary, the percentages of degradation measured at 2 and 4 h p.i. in the tumor were inferior to 25 and 45%, respectively (Fig. 5A). Likewise, the extent and rate of degradation in the skin were very low, with no degradation observed up to 4 h p.i. (Fig. 5A). Therefore, the degradation observed in vivo in the plasma likely resulted from a release from some organs (i.e., liver, spleen) where the degradation took place.
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Localization of TPC-Ahx-ATWLPPR. Intracellular localization in HUVEC following incubation with TPC-Ahx-ATWLPPR (1 µM, 24 h) was studied using fluorescent organelle probes and confocal fluorescence microscopy. Yellow color on merged images from intracellular localization of the PS (red) and the probe (green) indicates colocalization of both (Fig. 8). TPC-Ahx-ATWLPPR mostly localized into lysosomes (Fig. 8A) compared with Golgi apparatus, mitochondria, and endoplasmic reticulum (Fig. 8, B–D, respectively).
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Discussion |
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). In this work, we first investigated the in vivo stability of TPC-Ahx-ATWLPPR following i.v. injection into tumor-bearing mice. A significant degradation could be observed in plasma from 4 h p.i. We identified the main metabolic product as being TPC-Ahx-A. This metabolite would very probably have lost any selectivity for angiogenic endothelial cells that line the neovessels of the tumor because the arginyl residue at the C terminus of the ATWLPPR peptide has been shown to be essential for affinity for NRP-1 (Tournaire et al., 2001). Therefore, for future in vivo assessments of photodynamic activity of TPC-Ahx-ATWLPPR, the drug-light interval (DLI, i.e., the time interval between PS injection and its subsequent activation by light) should preferably, according to this study, be inferior or equal to 4 h. Indeed, after that time, more than 20% of the PS present in plasma is TPC-Ahx-A, which does not possess any specificity for endothelial cells lining the tumor vessels, contrary to TPC-Ahx-ATWLPPR. Using a DLI longer than 4 h would certainly allow PS accumulation in nontargeted tissues (e.g., skin), with potential skin photosensitivity problems, but would probably not improve much its uptake in target tissues (i.e., endothelium of tumor vessels).
The PS (TPC-Ahx-ATWLPPR and TPC-Ahx-A) accumulated at much higher levels in the liver than in the tumor. This has also been observed for other hydrophobic PS, including Foscan (GmbH, Ingelheim, Germany), with tumor to liver ratios of PS concentrations as low as 0.03 (Jones et al., 2003), but only PS present in the tumor will be activated because of localized light irradiation. It should be noted that PS present at the time of irradiation in plasma and endothelial cells lining the tumor vessels, and not only in tumor cells, also play a major part in tumor eradication by PDT by inducing vascular damage and occlusion of tumor vessels, which leads to tumor deprivation in nutrients and oxygen (Cramers, 2003).
A variety of proteases are present in human plasma, including both exopeptidases and endopeptidases (McDonald and Barrette, 1980, 1986). Therefore, we assessed the in vitro stability of TPC-Ahx-ATWLPPR in mouse and human plasma and did not observe any degradation during at least 24 h at 37°C, thus discarding any role of the plasma peptidases in the degradation phenomenon. The conjugated PS accumulated to high levels into organs of the reticuloendothelial system (i.e., liver, spleen), in agreement with its hydrophobic character, like other lipophilic PS (e.g., Foscan) (Jori and Fabris, 1998; Jones et al., 2003). Indeed, on administration into the bloodstream, most PS associate with various serum proteins, including both high- and low-density lipoproteins (LDL) and albumin (Hopkinson et al., 1999). LDL are believed to be important in the transportation of hydrophobic PS. 
2-Macroglobulin/LDL receptors are widely found in cells of various types, with LDL uptake most pronounced in adrenals, liver, and spleen (Rosenkranz et al., 2000).
The conjugated PS was rapidly degraded in organs of the reticuloendothelial system, as soon as 1 h p.i. Some degradation also occurred quite rapidly in the tumor, less than in metabolic organs but more than in plasma. All these results strongly suggested that the degradation product TPC-Ahx-A identified in plasma resulted from the degradation of the peptide in organs of the reticuloendothelial system, mainly liver, and subsequent release into the bloodstream.
To gain mechanistic insight into the degradation phenomenon, we undertook in vitro studies in HUVEC. TPC-Ahx-ATWLPPR was not degraded into the cell culture medium, thus allowing it to accumulate at high levels into cells through active targeting of NRP-1, as confirmed by the high depletion rate of the PS observed in the medium. On the contrary, following internalization by HUVEC, the peptide moiety of TPC-Ahx-ATWLPPR was progressively degraded, mostly into TPC-Ahx-A, as observed in vivo, but also to a lesser extent into TPC-Ahx-AT and TPC-Ahx-ATWLPP. Degradation of the peptidic moiety inside cells is not detrimental to our strategy because the peptide has already played its targeting role, and that, at this stage, only the PS moiety is involved in the photocytotoxic effect.
The distribution of a PS within cells depends on the route by which it enters, as well as on its physicochemical properties, e.g., its hydrophobicity/hydrophilicity; the type, number, and arrangement of its charged groups; the presence of a central atom in the tetrapyrrole structure; its aggregation state, etc. (Rosenkranz et al., 2000). For example, LDL-bound PS can enter target cells via receptor-mediated endocytosis. Chlorin e6 covalently bound to LDL has been detected in enzymatically active lysosomes (Schmidt et al., 1992). We showed that TPC-Ahx-ATWLPPR also mostly localized into lysosomes, in agreement with the fact that TPC-Ahx-ATWLPPR targets NRP-1 and should be incorporated into cells by a receptor-mediated endocytosis mechanism. Lysosomes are known to contain a large variety of hydrolytic enzymes, which degrade proteins and other substances taken up by endocytosis. The main proteolytic activities experienced by endocytosed peptides and proteins in the endosomal/lysosomal pathway belong to a family of papain-like proteases called cathepsins (Roberts, 2005). The pH rate profiles indicate that the low pH of the lysosomes (approximately 5.5) is optimal for the majority of cathepsins. Treatment with the lysosomotropic weak base ammonium chloride significantly decreased the extent of intracellular degradation.
Because some peptides can be significantly stabilized by glycosylation or the addition of D-amino acids at the N terminus (Powell et al., 1993), the relatively slow rate of degradation observed in plasma may be, at least in part, caused by the steric protection afforded by the PS moiety at the N terminus, which may prevent the peptide from being degraded by aminopeptidases. Moreover, no oxidation into a porphyrin, which would present less interesting photophysical properties than the corresponding chlorin, could be observed on the PS moiety in any experiment, as assessed by MALDI-TOF analysis, contrary to what has been observed with other PS (Laville et al., 2004).
To achieve higher selective delivery to the tumor neovessels, the in vivo stability of the peptide moiety of the conjugate may be increased through formulation and/or chemical modification. The first strategy encompasses the use of pegylated liposomes, which will impact on the pharmacokinetics of the PS and which may limit its accumulation in the reticuloendothelial system. For example, Ichikawa et al. (2005) encapsulated benzoporphyrin derivative monoacid ring A in pegylated liposomes modified with the H-Ala-Pro-Arg-Pro-Gly-OH pentapeptide, which had earlier been isolated as a peptide specific to angiogenic endothelial cells. Pegylation of liposomes aims to avoid opsonization in the bloodstream, which is a prerequisite for the clearance of the liposomes by the reticuloendothelial system, such as liver and spleen (Ichikawa et al., 2005). The second method implies the generation of modified peptides with improved stability properties, through (e.g., cyclization) the use of D-amino acids and reduced peptide bonds, etc. (Adessi and Soto, 2002), for which the knowledge of the site of degradation on the peptides that were determined in this study is essential.
The ATWLPPR peptide has already been used in several in vitro and in vivo studies (Janssen et al., 2003; Rodrigues et al., 2003; Perret et al., 2004; Renno et al., 2004), but, to the best of our knowledge, none has reported on its instability. The present study draws attention to this potential problem with peptides, especially in the case of targeting strategies, and provides useful information for the choice of the DLI for in vivo assessments of photodynamic activity and for the future design of more stable molecules.
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This work was presented in part at the 97th Annual Meeting of the American Association for Cancer Research, Washington, DC, April 1–5, 2006.
Article, publication date, and citation information can be found at http://dmd.aspetjournals.org.
ABBREVIATIONS: PDT, photodynamic therapy; PS, photosensitizer(s); TPC, 5-(4-carboxyphenyl)-10,15,20-triphenylchlorin; Ahx, 6-aminohexanoic acid; ATWLPPR, H-Ala-Thr-Trp-Leu-Pro-Pro-Arg-OH; VEGF, vascular endothelial growth factor; NRP-1, neuropilin-1; MALDI-TOF, matrix-assisted laser desorption ionization/time of flight; HPLC, high-performance liquid chromatography; HUVEC, human umbilical vein endothelial cell(s); Fmoc, 9-fluorenyl-methoxy-carbonyl; PBS, phosphate-buffered saline; EGM, endothelial growth medium; PEG, polyethylene glycol; mTHPP, 5,10,15,20-tetrakis(m-hydroxyphenyl)porphyrin; p.i., postinjection; DLI, drug-light interval; LDL, low-density lipoprotein(s).
Address correspondence to: Muriel Barberi-Heyob, Laboratoire de Recherche, Centre Alexis Vautrin, Avenue de Bourgogne, F-54511 Vandoeuvre-les-Nancy Cedex, France. E-mail: m.barberi{at}nancy.fnclcc.fr
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3).
