The association of silicon microparticles with endothelial cells in drug delivery to the vasculature

The association of silicon microparticles with endothelial cells in drug delivery to the vasculature

Biomaterials 30 (2009) 2440–2448 Contents lists available at ScienceDirect Biomaterials journal homepage: www.elsevier.com/locate/biomaterials The ...

2MB Sizes 3 Downloads 20 Views

Biomaterials 30 (2009) 2440–2448

Contents lists available at ScienceDirect

Biomaterials journal homepage: www.elsevier.com/locate/biomaterials

The association of silicon microparticles with endothelial cells in drug delivery to the vasculature Rita E. Serda a, *, Jianhua Gu a, Rohan C. Bhavane a, XueWu Liu a, Ciro Chiappini b, Paolo Decuzzi c, d, Mauro Ferrari a, e, f, * a

Brown Institute of Molecular Medicine, University of Texas Health Science Center, Nanomedicine Division, 1825 Pressler, Suite 537, Houston, TX 77030, USA University of Texas at Austin, Department of Biomedical Engineering, 1 University Station, C0400, Austin, TX 78712, USA Universita’ degli Studi Magna Græcia, Viale Europa, Italy d School of Health Information Sciences, University of Texas Health Science Center Houston, 7000 Fannin, Houston, TX 77030, USA e Rice University, Department of Bioengineering, Houston, TX 77005, USA f University of Texas MD Anderson Cancer Center, Department of Therapeutics, Unit 422, 1515 Holcombe Boulevard, Houston, TX 77030, USA b c

a r t i c l e i n f o

a b s t r a c t

Article history: Received 3 December 2008 Accepted 3 January 2009 Available online 12 February 2009

Endothelial targeting is an approach evolving for drug delivery to the vasculature of pathological lesions. Nano-porous silicon-based multi-functional particles are of particular interest, since they can be manufactured in essentially any size and shape, employing methods of photolithography, to optimize their ability to localize on target endothelia. In this study we tested the impact of surface charge, serum opsonization, and inflammation on the ability of vascular endothelial cells to associate with nano-porous silicon microparticles. Vascular endothelial cells were capable of rapidly internalizing both positive and negative silicon microparticles by an actin-dependent mechanism involving both phagocytosis and macropinocytosis. However, following serum opsonization, internalization was selective for APTES (originally positive) modified microparticles, despite the finding that all opsonized microparticles had a net negative charge. Conversely, macrophages displayed a preference for internalization of serum opsonized oxidized (originally negative) microparticles, supporting the choice of positive microparticles for endothelial targeting. The internalization of opsonized microparticles by endothelial cells was further enhanced by the presence of inflammatory cytokines. These findings suggest that it may be possible to bioengineer silicon microparticles to favor opsonization with proteins that enhance uptake by endothelial cells, without a concurrent enhanced uptake by macrophages. Published by Elsevier Ltd.

Keywords: Endothelia Phagocytosis Silicon Microparticles

1. Introduction Nano-structured delivery systems are emerging as powerful tools for the systemic delivery of therapeutic molecules and imaging agents for different biomedical applications, from cancer [1,2] to cardiovascular diseases [3]. These delivery systems can be loaded with a multitude of drug molecules and contrast agents to simultaneously provide therapeutic and imaging capabilities. Following intravenous injection, these particles are transported by the blood stream into different vascular districts. Targeting diseased vasculature is a strategy already demonstrated to be effective for the detection of

* Correspondence at: Mauro Ferrari and Rita E. Serda, Brown Institute of Molecular Medicine, Nanomedicine Division, University of Texas Health Science Center, 1825 Pressler Street, Suite 537, Houston, TX 77030, USA. Tel.: þ1 713 500 2309; fax: þ1 713 500 2313. E-mail addresses: [email protected] (R.E. Serda), [email protected] edu (M. Ferrari). 0142-9612/$ – see front matter Published by Elsevier Ltd. doi:10.1016/j.biomaterials.2009.01.019

atherosclerotic plaque and cardiovascular alterations [4,5], and it is becoming more prominent in cancer applications [6] following the pioneering work of Hood and co-workers [7] that introduced the notion of targeting via integrins such as aV-b3. Typically, delivery vectors are conjugated with ligand molecules that specifically recognize and interact with receptors uniquely or over-expressed at pathological lesions [8–10]. Targeting ligands include peptides identified from phage-displayed libraries [11], thioaptamers [12], and antibodies. Receptors for these ligands act as vascular docking sites for the circulating particles. The most prominent bottle-neck to systemic delivery of particles, regardless of whether they are designed for vascular endothelial or tumor cell surface targeting, is their uptake by professional phagocytes, either circulating as monocytes/macrophages or accumulating in specific organs, such as the liver, spleen and lungs. Attempts at bypassing phagocytic uptake by shielding with polymers, such as poly(ethylene glycol), successfully prolongs circulation time, but also limits attachment of targeting ligands and uptake by target populations [13].

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

Phagocytosis of particulates, apoptotic bodies, and aged erythrocytes is predominately attributed to macrophages, neutrophils and dendritic cells. However, several studies have described phagocytosis of large objects by endothelial cells, including the uptake of latex particles, zymosan, apoptotic bodies and microorganisms [14–18]. In this study, the role of endothelial cells in cellular uptake of porous silicon microparticles is explored. The advantages of using porous silicon for drug delivery include biocompatibility, exquisite control over rates of degradation, and precise control over size and shape by employing methods of photolithography [2,19,20]. Precise control over particle geometry allows realization of the computational prediction that size and shape influence vascular navigation, avoidance of biological barriers, localization, and biological pathways [21–23]. The porous nature of the silicon microparticles transforms the conceptual idea of multi-functional particles into a reality by creating vectors capable of releasing successive stages of carries and drugs intended to conquer sequential biological barriers to reach the intended target [24]. In this study, the effect of surface chemistry, serum opsonization and inflammation on the interplay between silicon microparticles and endothelial cells is explored. 2. Materials and methods 2.1. Silicon particle fabrication Nano-porous hemispherical silicon microparticles were designed, engineered, and fabricated in the Microelectronics Research Center at The University of Texas at Austin. Two sizes of microparticles were generated, with mean diameters of 1.6  0.2 and 3.2  0.2 mm, and pore sizes ranging from 5 to 10 nm. Processing details were recently published by our laboratory [2]. Briefly, heavily doped pþþ type (100) silicon wafers with resistivity of 0.005 ohm cm (Silicon Quest, Inc., Santa Clara, CA) were used as the silicon source. A 100 nm layer of low-stress silicon nitride was deposited using a Low Pressure Chemical Vapor Deposition (LPCVD) system. Standard photolithography was used to pattern the microparticles over the wafer using a contact aligner (EVG 620 aligner) and AZ5209 photoresist. Nitride on particle patterns was selectively removed by CF4 based reactive ion etching (RIE). After the photoresist was stripped in piranha solution, the wafer was placed in a home-made Teflon cell for two-step electrochemical etching. First, the wafers were etched in a mixture of hydrofluoric acid (HF) and ethanol (1:1 v/v) by applying a current density of 6 mA/cm2 for 105 s for 3.2 mm particles or 40 s for 1.6 mm particles, respectively. Then a high porosity release layer was formed by changing the current density to 320 mA/cm2 for 6 s in a 2:5 v/v mixture of HF and ethanol. Finally, the nitride layer was removed in HF after etching, and microparticles were released by ultrasound in isopropyl alcohol (IPA) for 1 min. The IPA solution containing porous silicon microparticles was collected and stored at 4  C. The morphology of the microparticles was examined by SEM.

2.2. Oxidation of silicon microparticles Silicon microparticles in isopropyl alcohol (IPA) were dried in a glass beaker kept on a hot plate (110  C). The dried microparticles were then treated with piranha solution (1 volume H2O2 and 2 volumes of H2SO4). The suspension was heated to 110–120  C for 2 h with intermittent sonication to disperse the microparticles. The suspension was then washed in deionized (DI) water until the pH of the suspension was w5.5–6. 2.3. Surface modification of silicon microparticles with APTES The oxidized microparticles were washed in IPA 3–4 times. They were then suspended in IPA containing 0.5% (v/v) APTES (Sigma) for 2 h at room temperature. The APTES modified microparticles were washed and stored in IPA. APTES modification was evaluated by measuring the zeta potential and by colorimetric analysis of amine density. The latter was found to correlate with zeta potential measurements.

2.4. PEG conjugation APTES modified microparticles were reacted with 10 mM mPEG-SCM-5000 (methoxy poly-ethylene glycol succinimidyl carboxymethyl; purchased from Laysan Bio Inc.) in acetonitrile for 1.5 h. The microparticles were then washed in distilled water 4–6 times to remove any unreacted mPEG. Zeta potential measurements were used to indicate adequate surface coating.

2441

2.5. Z2 analysis Microparticles were counted in a Z2 CoulterÒ Particle Counter and Size Analyzer (Beckman Coulter). The aperture size used for microparticle analysis was 50 mm. For analysis, microparticles were suspended in the balanced electrolyte solution of the instrument (ISOTONÒ II Diluent) and counted.

2.6. Cell culture HUVEC, purchased from Lonza Walkersville, Inc. (Walkersville, Maryland), were cultured in EBMÒ-2 medium (CloneticsÒ, CC-3156). Cells were maintained at 37  C in a humidified 5% CO2 atmosphere, and detached using a 0.25 mg/ml trypsin/EDTA solution (CloneticsÒ) upon reaching 80–90% confluency. For serum-free experiments, the media used was EBMÒ-2 medium (CloneticsÒ) supplemented with only hydrocortisone and GA-1000, plus 0.2% BSA. HUVECs were discarded after 7–8 passages. J774A.1 macrophage cells were purchased from American Type Culture Collection (Manassas, VA). Growth medium was Dulbecco’s Modified Eagle’s Medium containing 10% FBS, 100 mg/ml streptomycin and 100 U/ml Penicillin (Invitrogen; Carlsbad, CA). Cells were collected by scrapping.

2.7. Confocal microscopy HUVECs were grown on No. 1.5 glass coverslips. When confluent, cells were incubated with microparticles (1:10; cell:microparticle) for 60 min in serum-free media at 37  C. Cells were then washed with PBS, fixed and premeabilized with 0.1% triton X-100. PBS containing 1% BSA was used as a blocking agent prior to incubation with 200 nM Alexa Fluor 555 conjugated phallodin (Invitrogen) and anti-tubulin FITC conjugated antibody (Abcam Inc., Cambridge, MA). Coverslips were then washed and mounted on glass slides using Vectashield mounting media (Vector Laboratories, Burlingame, CA) containing a 1000-fold dilution DRAQ5 (Biostatus Limited, UK). Detection of silicon microparticles was based on autofluorescence using a 633 excitation laser. Images were acquired using a Leica DM6000 upright confocal microscope equipped with a 63 oil immersion objective.

2.8. Flow cytometry HUVECs (1.5  105 cells/well) were seeded into 6 well plates and 24 h later the cells were incubated with silicon microparticles (10–20 microparticles/cell) in serum-free media, or as specified, for the indicated time. FluoresbriteÒ YG Microspheres (1 mm; Polysciences) were incubated with HUVECs at 37  C for 60 min at a ratio of 1:1000 (cell:particle). For opsonization experiments, microparticles were pre-incubated with 100 ml of 100% FBS or physiological concentrations of BSA (4 g/dL in PBS) or ImmunoPureÒ Human Whole Molecule IgG (500 mg/dL; Pierce, Rockford, IL) on ice for 60 min. Following incubation with microparticles, cells were washed with PBS, harvested by trypsinization (HUVEC) or scrapping (J774), and resuspended in PBS containing 1.0% BSA and 0.1% sodium azide (FACS wash buffer). Microparticle association with cells was determined by measuring side scatter using a Becton Dickinson FACSCalibur Flow equipped with a 488-nm argon laser and CellQuest software (Becton Dickinson; San Jose, CA). Data is presented as either percentage of cells with microparticles (percent of cells with high side scatter) or mean particle uptake per cell (mean side scatter). Side scatter due to cells in the absence of particles has been subtracted from the presented data. Comparisons of FITC Dextran (0.5 mg/ml) internalization by HUVECs are based on fluorescent intensity measurements following incubation of cells with microparticles at 37  C for 1 h. The following antibodies were used to measure expression of Fc gamma globulin receptors: Monoclonal anti-Human CD64 FITC Conjugate (clone 10.1, BD Pharmingen), Monoclonal anti-Human CD32 FITC Conjugate (clone 3D3; BD Pharmingen), and Monoclonal anti-Human CD16 FITC Conjugate (clone 3G8; Sigma). Expression of FcgR, as well as microparticle uptake, were also studied on HUVECs stimulated for 48 h with TNF-a (10 ng/ml) and IFN-g (100 U/ml). Flow cytometric analysis was performed on live cells, which were identified by their lack of propidium iodide staining. Antigen density calculations were based on a QuickCalÒ calibration curve generated with QuantumÔ Simply CellularÒ anti-Mouse IgG (Bangs Laboratories, Inc.; Fishers, IN).

2.9. Inhibition studies A 5 mg/ml Cytochalasin B solution in 95% ethanol (Sigma–Aldrich; St. Louis, MO) was prepared and stored at 20  C. For inhibition studies, HUVEC cells, plated in 6 well plates (80–90% confluent), were incubated in media containing Cytochalasin B (2.5, 5.0 and 10.0 mg/ml) for 60 min at 37  C prior to addition of microparticles and during the 60 min incubation with microparticles (4  106 microparticles per well; ratio 1:20). Following incubation, the cells were washed with PBS, collected by trypsinization and resuspended in FACS wash buffer. Samples were analyzed by flow cytometric analysis.

2442

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

A

cells

15 min

2μm 30 min

60 min

5μm

B

5 min

0

60 min

µm

5

Fig. 1. Internalization of silicon microparticles by endothelial cells. (A) HUVECs, grown on silicon supports, were incubated with 3.2 mm oxidized silicon microparticles (10:1 particles per cell) for 15–60 min at 37  C in serum-free media, followed by fixation, dehydration, and imaging by SEM. (B) Confocal micrographs of HUVECs incubated with 3.2 mm oxidized silicon microparticles (10:1) for 5 and 60 min at 37  C. Following incubation, cells were fixed, permeabilized, and stained with Alexa Fluor 555 Phalloidin. Microparticle autofluoresence was imaged following excitation at 633 nm.

2.10. Scanning electron microscopy

2.11. Transmission electron microscopy

Microparticle binding and uptake by HUVEC and J774 cells were observed by scanning electron microscopy. Cells were plated in 24 well plates containing 5  7 mm Silicon Chip Specimen Supports (Ted Pella, Inc., Redding, CA) at 5  104 cells per well. When cells were confluent, media containing microparticles (1:20, cell:microparticles, 0.5 ml/well) was introduced and cells were incubated at 37  C for the specified amount of time. Samples were washed with PBS and fixed in 2.5% glutaraldehyde for 30 min (Sigma–Aldrich; St. Louis, MO). After washing in PBS, cells were dehydrated in ascending concentrations of ethanol (30%, 50%, 70%, 90%, 95%, and 100%) for 10 min each. HUVECs were then incubated in a 50% alcohol–hexamet hyldisilazane (HMDS; Sigma) solution for 10 min followed by incubation in pure HMDS for 5 min and overnight incubation in a desiccator. Specimens were mounted on SEM stubs (Ted Pella, Inc.) using conductive adhesive tape (12 mm OD PELCO Tabs, Ted Pella, Inc.). Samples were sputter coated with a 10 nm layer of gold using a Plasma Sciences CrC-150 Sputtering System (Torr International, Inc.). SEM images were acquired under high vacuum, at 20.00 kV, spot size 3.0–5.0, using an FEI Quanta 400 FEG ESEM equipped with an ETD (SE) detector.

HUVECs were grown to 80% confluency in a 6 well plate. Using 1 ml of serumfree media per well, 1.6 mm or 3.2 mm particles were introduced at a cell:microparticle ratio of 1:10 at 37  C for either 15 or 120 min. HUVECs were then washed and fixed in a solution of 2% paraformaldehyde (16% solution purchased from Electron Microscopy Sciences; Hatfield, PA) and 3% glutaraldehyde (25% solution from Sigma) in phosphate buffered saline pH 7.4 (2 ml/well; Sigma) for 1 h. After fixation, the samples were washed and treated with 0.1% Millipore-filtered cacodylate buffered tannic acid, post-fixed with 1% buffered osmium tetroxide for 30 min, and stained en bloc with 1% Millipore-filtered uranyl acetate. The samples were dehydrated in increasing concentrations of ethanol, infiltrated, and embedded in Poly-bed 812 medium. The samples were polymerized in a 60  C oven for 2 days. Ultrathin sections were cut using a Leica Ultracut microtome (Leica, Deerfield, IL), stained with uranyl acetate and lead citrate in a Leica EM Stainer, and examined in a JEM 1010 transmission electron microscope (JOEL, USA, Inc., Peabody, MA) at an accelerating voltage of 80 kV. Digital images were obtained using the AMT Imaging System (Advanced Microscopy Techniques Cory, Danvers, MA).

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

2443

3. Results 3.1. Silicon microparticle uptake by endothelial cells Nano-structured silicon microparticles were fabricated using a porosification process involving photolithography [25]. This process allows for accurate control over particle size and geometry, two parameters important for drug delivery. To examine the response of endothelial cells to silicon microparticles, two sizes of microparticles, 1.6  0.2 and 3.2  0.2 mm, and three types of surface modifications were explored. All experiments were performed with both sizes of microparticles in triplicate, and representative experiments are presented. Initial experiments explored the response of endothelial cells to oxidized silicon microparticles. Microparticles were introduced to Human Umbilical Vein Endothelial Cells (HUVECs) at a ratio of 20 particles to every cell. Scanning electron microscopy (SEM) images demonstrated that early contact (15 min at 37  C) of HUVECs with microparticles induced formation of pseudopodia that extended outward to engulf the microparticles (Fig. 1A). SEM analysis at 30–60 min showed an increasing number of microparticles becoming completely internalized by HUVECs with time. Confocal microscopy images, taken after 5 min of incubation of microparticles with HUVECs at 37  C, verified the formation of cellular membrane extensions reaching out to surround the microparticles (Fig. 1B). After 60 min, the internalized microparticles were located in the perinuclear region of the cell. Microparticles were visualized by autofluorescence (excitation at 633 nm) and the cell periphery (i.e. actin) was labeled with Alexa Fluor 555 conjugated phalloidin. Based on an MTT cell proliferation assay, at 12–72 h cell viability was similar for cells cultured in media alone versus media containing both 5 and 10 particles to every cell (Appendix S1). 3.2. Size effect on microparticle uptake Both phagocytosis and macropinocytosis are characterized by extensive membrane reorganization. Phagocytic engulfment is characterized by the formation of tight-fitting phagosomes, while macropinocytotic uptake involves the formation of ruffles that engulf both solid cargo and fluid [26]. In this study, HUVECs incubated with 1.6 mm silicon microparticles for 15 min at 37  C were engulfed by mechanisms phenotypically resembling both macropinocytosis and phagocytosis (Fig. 2A). Macropinocytosis of particles by endothelial cells is supported by Hartig et al. [27] who recently reported that nonspecifically bound nanoparticulate complexes are taken up by microvascular endothelial cells by macropinocytosis. Internalization of the larger 3.2 mm particles by HUVECs was more consistent with classical phagocytosis, with the hallmark presence of an actin cup holding a microparticle (Fig. 2B). Involvement of macropinocytosis in microparticle uptake was supported by internalization of fluorescein isothiocyanate dextran (FD) from the cell media during microparticle uptake. A greater amount of FD was internalized by HUVECs in the presence of 1.6 mm particles compared to 3.2 mm particles (Fig. 2C) supporting a greater role for macropinocytosis in the uptake of the smaller microparticles. 3.3. Effect of Cytochalasin B on silicon microparticle uptake Membrane reorganization occurring during phagocytosis and macropinocytosis involves reorganization of the actin cytoskeleton, which leads to changes in the shape of the cell membrane. Cytochalasin B dissociates actin-binding protein from actin filaments, weakening existing actin filaments, and blocking actin polymerization [28]. To confirm the involvement of actin

Fig. 2. Early uptake of oxidized silicon microparticles by endothelial cells. HUVECs were incubated with either 1.6 mm (A) or 3.2 mm (B) oxidized silicon microparticles (10:1 particles per cell) for 15 min at 37  C in serum-free media. HUVECs were washed, fixed and processed for TEM analysis. Top row: direct magnification 3000; others 25,000. (C) Internalization of FITC dextran, as measured by flow cytometry, was used as a metric to measure fluid uptake indicative of macropinocytosis. HUVECs were incubated with either 1.6 mm (red line) or 3.2 mm (purple line) oxidized silicon microparticles in the presence of FITC dextran for 60 min. Controls included cells incubated with FITC dextran (green peak) or media (blue peak). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

Cells containing internalized microparticles have an increase in granularity as demonstrated by an upward shift in orthogonal (90 ) laser light scatter (side scatter) based on flow cytometric analysis. As the ratio of microparticles to cells increases, there is a linear increase in the percentage of cells with a high side scatter phenotype (Fig. 3C). Based on gating similar to that shown in Fig. 3D, side scatter was used as a metric to measure microparticle uptake by HUVECs (i.e. percent cells with internalized microparticles). Dose dependent inhibition of microparticle uptake by HUVECs was evaluated by flow cytometry at concentrations of Cytochalasin B of 2.5 mg/ml and higher. Cytochalasin B inhibited microparticle uptake by 85–88% (Fig. 3E). 3.4. Effect of surface charge on microparticle uptake The effect of surface charge on microparticle uptake was evaluated by flow cytometry and SEM analysis. Silicon microparticles were oxidized with a piranha solution [30:70 (v/v) H2O2:H2SO4] to create negatively charged, hydroxylated microparticles. Complete surface oxidation was confirmed by energy dispersive spectrophotometer analysis (data not shown). Surface modification with

particle uptake 1 µm HUVEC 1X

y = 15.4x + 0.4 R2 = 0.9605

0

5 10 20 particle to cell ratio

800 1000

D

Side Scatter

cells

600

control 60 min

cells + Particles (MP)

0

200

400

600

800 1000

0

Forward Scatter

E

0 µm 25

% cells with particles

CB 60 min

0 µm 25

40

0

0

25 µm

800 1000

spot mag WD HFW 6/28/2007 HV 11:22:32 AM 20.00 kV 5.5 30.000 x 9.7 mm 4.97 µm

Side Scatter

B

5 µm HUVEC 1X

90 80 70 60 50 40 30 20 10 0

400

HV spot mag 7/8/2007 WD HFW 11:38:15 AM 20.00 kV 5.5 7.500 x 9.2 mm 19.9 µm

CB 30 min

200

CB 15 min

600

C A

400

polymerization in the uptake of silicon microparticles, HUVECs were pre-incubated with Cytochalasin B (2.5 mg/ml) for 60 min, then incubated with both 1.6 mm and 3.2 mm oxidized microparticles for 15–60 min at 37  C. SEM analysis at 15 min shows a lack of pseudopodia engaging microparticles which have settled on the cell surface (Fig. 3A). After 30 min, the HUVECs appear to have initiated actin cup formation, which is a dense actin network that forms beneath the microparticles [29]; however, uptake is frustrated by Cytochalasin B, preventing internalization of the microparticles. In the absence of Cytochalasin B, further actin polymerization would induce expansion of the actin cup which would completely surround the microparticle, creating a phagosome. Confocal images taken after 60 min of incubation show microparticle uptake by HUVECs in the absence of Cytochalasin B (control), however, in agreement with SEM images, uptake of microparticles is blocked by addition of Cytochalasin B (CB) (Fig. 3B). Filipin and chlorpromazine, inhibitors of caveolae and clathrin mediated endocytosis, failed to block the uptake of silicon microparticles by HUVECs as visualized by confocal microscopy (R.E. Serda, unpublished data).

200

2444

16 14 12 10 8 6 4 2 0

200

400

600

800 1000

Forward Scatter

Cytochalasin B (CB)

cells

MP

CB 2.5

CB 10

CB 25

Fig. 3. Cytochalasin B blocks microparticle uptake by HUVECs. (A) SEM images of HUVECs incubated with a mixture of 1.6 mm and 3.2 mm oxidized silicon microparticles at 37  C for 15 and 30 min in serum-free media containing 2.5 mg/ml Cytochalasin B. (B) Confocal micrographs of control or Cytochalasin B treated HUVECs incubated with 3.2 mm silicon microparticles for 60 min and stained with FITC conjugated anti-tubulin antibody, Alexa Fluor 555 Phalloidin (actin), and DRAQ5 (nuclear). (C) Flow cytometric analysis showing orthogonal 90 laser light scatter (side scatter) by HUVECs after incubation with increasing numbers of microparticles (0–40 particles per cell) for 60 min at 37  C. (D) Flow cytometry dot blots showing the increase in side scatter caused by cell uptake of microparticles. Control cells are shown on the left and HUVECs incubated with 10 microparticles per cell for 60 min (37  C) are to the right. The box designates cells with internalized microparticles (MP), which is illustrated in the graph below. (E) HUVECs, pretreated for 60 min with 2.5–25 mg/ml Cytochalasin B (CB), were incubated with silicon microparticles for an additional 60 min at 37  C. Cells were harvested by trypsinization and particle uptake (i.e. side light scatter) was measured as shown in ‘‘D’’.

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

3-aminopropyltriethoxysilane (APTES) yielded positively charged, amine modified microparticles. APTES modified microparticles were further conjugated with methoxy poly-ethylene glycol-5000 (mPEG-5000) to create shielded microparticles with a more neutral charge. In the presence of serum-free media, HUVECs nondiscriminately internalized both positive and negative microparticles (Fig. 4A). SEM images are of cells after 1 h at 37  C with microparticles. Since it is well established that PEG creates a shield around particles, causing them to avoid or delay phagocytosis by macrophages, we examined uptake of mPEG-5000 microparticles by endothelial cells. Surface modification of silicon microparticles with mPEG-5000 effectively suppressed internalization of microparticles by HUVECs at 1 h (Fig. 4A). 3.5. Effect of serum opsonization on microparticle uptake Microparticles were incubated with 100% fetal bovine serum (hereafter serum) for 60 min on ice, followed by incubation with

oxidized

A

Endothelial cells - serum-free

HUVECs in the continued presence of 5% serum. Based on zeta potential measurements, despite the original surface charge, all opsonized silicon microparticles had a net negative surface charge (Fig. 4B). Based on flow cytometric analysis, while internalization of oxidized microparticles by HUVECs was strongly inhibited (78% less uptake than non-opsonized oxidized microparticles), serum opsonization failed to have an impact on internalization of APTES microparticles by HUVECs. Opsonization of PEG modified microparticles lead to a 27% decrease in uptake by endothelial cells (Fig. 4C). To evaluate the generality of inhibition of uptake of negatively charged microparticles by endothelial cells following serum opsonization, negatively charged polystyrene microparticles (FluoresbriteÒ YG, spherical, 1 mm; Polysciences) were opsonized in serum, and then incubated with HUVECs in media containing 5% serum. In the presence of serum, there was a 33-fold decrease in uptake of negative polystyrene microparticles, supporting the conclusion that serum blockade of internalization of originally negatively charged microparticles is a global phenomenon regardless of the type of microparticle used (Appendix S2).

B

serum-free

serum

OXID

-43.48 ± 1.65

-32.21 ± 3.43

APTES

15.12 ± 4.33

-40.28 ± 2.78

PEG

-3.84 ± 1.99

-27.82 ± 1.71

particle uptake

C

70 60 50 40 30 20 10 0

Endothelial

D

50 40

serum free serum

** *

oxidized

particle uptake

APTES

2445

APTES

PEG

Macrophage

30

*

20 10

serum-free serum

0 APTES

PEG

oxidized

Fig. 4. Silicon microparticle surface charge and binding by serum opsonins directs uptake by endothelial cells. (A) HUVECs grown on silicon supports were incubated with nonserum opsonized negative (oxidized), positive (APTES), or mPEG-5000 modified microparticles for 60 min at 37  C. SEM images were taken at direct magnification 10,000 (left) and 20,000 (right). (B) Electrostatic (zeta) potential of microparticles before and after serum opsonization (60 min, 4  C). (C) Flow cytometric analysis of microparticle uptake by HUVECs (20:1; microparticles:cell) in serum-free or serum-containing media (*p < 0.0004; **p < 0.015). Particle uptake is the percent of cells with high side scatter. (D) Flow cytometry analysis of microparticle uptake by J774 macrophages in serum-free or serum-containing media (*p < 0.03).

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

A

% cells with particles

Similar to endothelial cells, J774 mouse macrophages were able to take up positive and negative silicon microparticles nondiscriminately in serum-free media (Fig. 4D). However, under normalized conditions with serum present, J774 cells were also responsive to signals provided by particle bound opsonins. In contrast to endothelial cells, macrophages favored uptake of opsonized, originally negatively charged, oxidized microparticles by 29% (p < 0.05). Opsonization of particulates by antibodies enhances phagocytosis by macrophages via binding to gamma immunoglobulin (IgG) receptors (FcgRs) on the cell surface [30]. However, FcgRs are reported to be either absent or expressed at low levels on unstimulated endothelial cells [31–33]. IgG opsonization would therefore be expected to inhibit uptake of particles by endothelial cells. Additionally, Arvidsson et al. [34] has shown rapid (1 min) binding of antibody [specific for high molecular weight kininogen (HMWK)] to oxidized silicon immersed in plasma. As expected, opsonization of silicon microparticles in ImmunoPure Human Whole Molecule IgG at 500 mg/dL IgG (normal range, 400–1800 mg/dL) inhibited the uptake of oxidized microparticles by HUVECs, both in antibody free media (88%) and in media supplemented with antibody (76%) (Fig. 5A). Consistent with published data, IgG opsonization enhanced microparticle uptake by macrophages (by 35% and 47% in the absence or presence of free IgG, respectively) (Fig. 5B). In contrast to negative microparticles, IgG opsonization of positive microparticles failed to have an impact on microparticle uptake by endothelial cells, suggesting either inadequate binding or an incorrect orientation by bound IgG (Appendix S3). Leroux et al. [35] reported that opsonization of PEG-coated nanoparticles by serum proteins enhanced their uptake by monocytes. They further showed that IgG present in the serum was responsible for the enhanced uptake of PEGylated particles. Here

Endothelial cells

20 15 10 5 0 serum-free

receptors per cell

C

3.6. Phagocytosis following cytokine stimulation of HUVECs Since it has been reported that activation of endothelial cells by pro-inflammatory cytokines enhances expression of cell adhesion molecules [36,37], as well as FcgRs [31], microparticle uptake following 48 h stimulation with TNF-a (10 ng/ml) and IFN-g (100 U/ ml) was examined by flow cytometric analysis. The data is expressed in terms of mean side scatter to give a measure of the relative number of particles per cell (scatter due to control cells has been subtracted). Internalization of serum opsonized silicon microparticles by HUVECs was enhanced for all groups of microparticles; however, a clear preference for opsonized APTES microparticles continued to exist. Following cytokine stimulation, 1.9-fold more APTES microparticles were engulfed compared to oxidized microparticles (Fig. 6A). J774 macrophages continued to show a clear preference for internalization of opsonized oxidized microparticles (1.6-fold greater uptake than positive microparticles). Cytokine stimulation did not affect the number of microparticles

B

30 25

we show that serum opsonization of PEGylated microparticles decreased their uptake by endothelial cells (Fig. 4C), consistent with blocked uptake of IgG-opsonized negative microparticles by endothelial cells. Analysis of FcgR expression on cells by flow cytometry confirmed high levels of FcgRII expression by macrophages. On the other hand, endothelial cells lacked expression of both FcgRI and FcgRIII, and expressed low levels of FcgRII (Fig. 5C). Based on a calibration curve generated using QuantumÔ Simply CellularÒ microspheres, HUVECs express approximately 10,000 FcgRII per cell, compared to approximately 320,000 FcgRII per macrophage. Treatment of cells with TNFa (10 ng/ml) and IFN-g (100 U/ml) for 48 h increased expression of FcgRII on HUVECs to approximately 35,500 receptors per cell (Fig. 5D).

IgG opson IgG opson + IgG media

% cells with particles

2446

30

Macrophages

25 20 15 10 5 0

serum-free IgG opson IgG opson + IgG media

D

500000

FcγγRII

IgG 400000 300000 200000

FcγRI

HUVEC

FcγRII Stim. HUVEC

FcγRIII

9,173 35,542

J774

324,343

Stim. J774

313,018

100000 0 HUVEC

stimulated HUVEC

J774

stimulated J774

Fig. 5. IgG opsonization blocks uptake of negative microparticles by HUVECs while stimulating uptake by J774 macrophages. Flow cytometry analysis of non-serum opsonized versus IgG-opsonized oxidized microparticle uptake by HUVECs (A) and J774 (B) cells. Oxidized microparticles were pre-opsonized with pure IgG for 60 min on ice, then incubated with cells for 60 min in serum-free media at 37  C (10 particles per cell). Cells were trypsinized following incubation and side scatter (percent cells with high SSC) was measured by flow cytometry. (C–D) Basal and stimulated [TNF-a (10 ng/ml) and IFN-g (100 U/ml) for 48 h] HUVECs and J774 cells were labeled with fluorescein isothiocyanate (FITC)-conjugated antibodies specific for FCgRI, FCgRII, and FCgRIII. Surface expression was determined by flow cytometric analysis. Quantitative expression of receptors was determined using a QuickCalÒ calibration curve. The data shown is the number of receptors expressed per cell.

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

engulfed by macrophages (Fig. 6B). PEGylation had a major impact on microparticle uptake by macrophages, with 2.8-fold less uptake compared to negative, oxidized microparticles. A novel finding was that opsonization of negative microparticles inhibited their uptake by endothelial cells to an extent equal to that of PEGylation, under both basal and inflammatory conditions (Fig. 6A). SEM images taken after 15 min at 37  C show the membranes extending outward to wrap around serum opsonized APTES microparticles by HUVECs (Fig. 6C) and serum opsonized oxidized microparticles by J774 cells (Fig. 6D). 4. Discussion In this study we found that cationic silicon microparticles were preferentially internalized by endothelial cells via phagocytosis and macropinocytosis. These observations are consistent with recent findings on liposomes, which suggest that cationic particles have a propensity for localizing in tumor vessels due to selective adherence to endothelial cells [38]. Several in vivo studies suggest that the pattern of opsonins adsorbed to the surface of particulates determines the population of phagocytic cells responsible for their clearance [39]. Current studies are underway to determine which serum constituents bind to positive silicon microparticles and act as opsonins for the uptake of microparticles by endothelial cells. Antibodies, which are opsonins with respect to particle uptake by macrophages, were dys-opsonins with respect to the uptake of microparticles by endothelial cells. We have shown that antibody opsonization selectively impacted the uptake of oxidized, anionic silicon microparticles. Thus, irrespective of size, cationic particles, ranging from nano to micron size, are targets for binding and uptake by vascular endothelial cells.

A

*

We observed that microparticle internalization by endothelial cells was enhanced by pro-inflammatory cytokine stimulation, supporting superior uptake of silicon microparticles at sites of chronic inflammation. At sites of pathological inflammation, upregulation of adhesion molecules on endothelial cells leads to recruitment of leukocytes [40]. As just presented, the altered recognition properties of endothelial cells also inspire enhanced binding to porous silicon microparticles. Silicon microparticles either remained attached to the endothelial membrane or were internalized via phagocytosis. The array of opsonins having the most dramatic influence on the interaction between endothelia and silicon microparticles were those binding to the surface of positively charged microparticles. Future studies aimed at optimizing the adherence of microparticles to inflamed endothelium will simultaneously incorporate targeting ligands selective for enriched adhesion molecules and maintenance of an overall positive charge to favorably influence binding by endothelial specific opsonins. Another potential targeting strategy that emerges from this study takes into account microparticles that bind to the endothelial cell surface, but have not been internalized. Relying on enhanced binding of favorable opsonized cationic microparticles at inflammatory lesions, subsequent transmigration could be supported by attaching targeting molecules to the surface of silicon microparticles that are involved in secondary steps of transmigration. For example, initiation of transmigration involves PECAM, with completion depending on CD99 [41,42]. Attachment of CD99 to the microparticle surface would thus favor migration across the endothelial barrier. Once across, the release of secondary particles, such as iron oxide nanoparticles and liposomes, from the pores of the silicon microparticles would home to the tumor cell itself via cancer specific targeting ligands. With this logic, one may envision

C

Endothelial

250

2447

control

200

stimulated 150 100

Particle uptake

50 0 oxidized

aptes

1 micron

peg

D

B

Macrophage

250

control

200

stimulated 150 100 50 0 oxidized

aptes

peg

2 micron

Fig. 6. Inflammatory cytokines stimulate silicon microparticle uptake by endothelial cells. (A, B) Flow cytometry analysis of serum opsonized, silicon microparticle uptake by HUVECs (A) and J774 cells (B) in cells under basal conditions (control) or following stimulation with TNF-a and IFN-g for 48 h (*p < 0.045). Cells were incubated with 20 microparticles (negative, positive, or neutral PEG) per cell for 60 min at 37  C. Microparticle uptake is presented as mean side scatter, that is, average uptake per cell. (C, D) SEM images of silicon microparticle uptake by HUVEC (C) and J774 (D) cells (30 min, 37  C) in the presence of serum.

2448

R.E. Serda et al. / Biomaterials 30 (2009) 2440–2448

transforming biological barriers into stepping stones leading to the targeted tumor lesion. 5. Conclusions In summary, cationic silicon microparticles bind serum opsonins that favor uptake by endothelia, with enhanced uptake by cytokine inflamed endothelia. Strategies for the attachment of targeting ligands to the microparticle surface should incorporate methods that preserve or impart an overall positive charge to the microparticle surface. The optimal design of delivery vehicles must take into account the final presentation of the opsonized particle. Acknowledgements We wish to thank Angelo Benedetto and the Rice Shared Resource Facility for training in scanning electron microscopy and Kenneth Dunner Jr. at MD Anderson Cancer Research Center for sample processing and image analysis by transmission electron microscopy. We also thank Dr. Fredika Robertson for advice on the technical writing of the manuscript and Matt Landry for aid in assembling high resolution figures for publication. This research was supported by grants DODW81XWH-07-1-0596, DODW81XWH-04-2-0035 and DODW81XWH-07-2-0101; NASA Grant #SA23-06-017; and State of Texas, Emerging Technology Fund. Appendix. Supplementary data Supplementary data associated with this article can be found in the online version, at doi:10.1016/j.biomaterials.2009.01.019. References [1] Ferrari M. Cancer nanotechnology: opportunities and challenges. Nat Rev 2005;5:161–71. [2] Tasciotti E, Liu X, Bhavane R, Plant K, Leonard A, Price B, et al. Mesoporous silicon particles as a multistage delivery system for imaging and therapeutic applications. Nat Nanotechnol 2008;3:151–7. [3] Cyrus T, Lanza GM, Wickline SA. Molecular imaging by cardiovascular MR. J Cardiovasc Magn Reson 2007;9:827–43. [4] Lanza G, Winter P, Cyrus T, Caruthers S, Marsh J, Hughes M, et al. Nanomedicine opportunities in cardiology. Ann N Y Acad Sci 2006;1080:451–65. [5] Kang HW, Torres D, Wald L, Weissleder R, Bogdanov Jr AA. Targeted imaging of human endothelial-specific marker in a model of adoptive cell transfer. Lab Invest 2006;86:599–609. [6] Hu G, Lijowski M, Zhang H, Partlow KC, Caruthers SD, Kiefer G, et al. Imaging of Vx-2 rabbit tumors with alpha(nu)beta3-integrin-targeted 111In nanoparticles. Int J Cancer 2007;120:1951–7. [7] Hood JD, Bednarski M, Frausto R, Guccione S, Reisfeld RA, Xiang R, et al. Tumor regression by targeted gene delivery to the neovasculature. Science 2002;296:2404–7. [8] Rossin R, Muro S, Welch MJ, Muzykantov VR, Schuster DP. In vivo imaging of 64Cu-labeled polymer nanoparticles targeted to the lung endothelium. J Nucl Med 2008;49:103–11. [9] Ding B, Dziubla T, Shuvaev VV, Muro S, Muzykantov VR. Advanced drug delivery systems that target the vascular endothelium. Adv Drug Deliv Syst 2006;6:98–112. [10] Koning GA, Schiffelers RM, Wauben MHM, Kok RJ, Mastrobattista E, Molema G, et al. Targeting of angiogenic endothelial cells at sites of inflammation by dexamethasone phosphate-containing RGD peptide liposomes inhibits experimental arthritis. Arthritis Rheum 2006;54:1198–208. [11] Arap W, Haedicke W, Bernasconi M, Kain R, Rajotte D, Krajewski S, et al. Targeting the prostate for destruction through a vascular address. Proc Natl Acad Sci U S A 2002;99:1527–31. [12] Bassett SE, Fennewald SM, King DJ, Li X, Herzog NK, Shope R, et al. Combinatorial selection and edited combinatorial selection of phosphorothioate aptamers targeting human nuclear factor-kappaB RelA/p50 and RelA/RelA. Biochemistry 2004;43:9105–15. [13] Gabizon A, Shmeeda H, Horowitz AT, Zalipsky S. Tumor cell targeting of liposome-entrapped drugs with phospholipid-anchored folic acid–PEG conjugates. Adv Drug Deliv Rev 2004;56:1177–92.

[14] Steffan AM, Gendrault JL, McCuskey RS, McCuskey PA, Kirn A. Phagocytosis, an unrecognized property of murine endothelial liver cells. Hepatology 1986;6:830–6. [15] Langeggen H, Namork E, Johnson E, Hetland G. HUVEC take up opsonized zymosan particles and secrete cytokines IL-6 and IL-8 in vitro. FEMS Immunol Med Microbiol 2003;36:55–61. [16] Dini L, Lentini A, Diez GD, Rocha M, Falasca L, Serafino L, et al. Phagocytosis of apoptotic bodies by liver endothelial cells. J Cell Sci 1995;108:967–73. [17] Kirsh T, Woywodt A, Beese M, Wyss K, Park J-K, Erdbruegger U, et al. Engulfment of apoptotic cells by microvascular endothelial cells induces proinflammatory responses. Blood 2007;109:2854–62. [18] Bengualid V, Hatcher VB, Diamond B, Blumberg EA, Lowy FD. Staphylococcus aureus infection of human endothelial cells potentiates Fc receptor expression. J Immunol 1990;145:4279–83. [19] Canham LT. Bioactive silicon structure fabrication through nanoetching techniques. Adv Mater 1995;7:1033–7. [20] Godin B, Gu J, Serda RE, Ferrati S, Liu X, Chiappini C, et al. Multistage mesoporous silicon-based nanocarriers: biocompatibility and controlled degradation in physiological fluids. New York City, NY: 35th Annual Meeting and Exposition of the Controlled Release Society; 2008. p. 575. [21] Ferrari M. Nanogeometry beyond drug delivery. Nat Nanotechnol 2008;3:131–2. [22] Decuzzi P, Lee S, Bhushan B, Ferrari M. A theoretical model for the margination of particles within blood vessels. Ann Biomed Eng 2005;33:179–90. [23] Decuzzi P, Lee S, Decuzzi P, Ferrari M. Adhesion of microfabricated particles on vascular endothelium: a parametric analysis. Ann Biomed Eng 2004;32:793–802. [24] Sanhai WR, Sakamoto JH, Canady R, Ferrari M. Seven challenges for nanomedicine. Nat Nanotechnol 2008;3:242–4. [25] Cohen MH, Melnik K, Boiarski AA, Ferrari M, Martin FJ. Microfabrication of silicon-based nanoporous particulates for medical applications. Biomed Microdevices 2003;5:253–9. [26] Krysko DV, Brouckaert G, Kalai M, Vandenabeele P, D’Herde K. Mechanisms of internalization of apoptotic and necrotic L929 cells by a macrophage cell line studied by electron microscopy. J Morphol 2003;258:336–45. [27] Hartig SM, Greene RR, Carlesso G, Higginbotham JN, Khan WN, Prokop A, et al. Kinetic analysis of nanoparticulate polyelectrolyte complex interactions with endothelial cells. Biomaterials 2007;28:3843–55. [28] Hartwig JH, Stossel TP. Interactions of actin, myosin, and an actin-binding protein of rabbit pulmonary macrophages. III. Effects of cytochalasin B. J Cell Biol 1976;71:295–303. [29] Stockem W, Hoffmann HU, Gruber B. Dynamics of the cytoskeleton in Amoeba proteus. I. Redistribution of microinjected fluorescein-labeled actin during locomotion, immobilization and phagocytosis. Cell Tissue Res 1983;232:79–96. [30] Coleman DL. Regulation of macrophage phagocytosis. Eur J Clin Microbiol 1986;5:1–5. [31] Pan LF, Kreisle RA, Shi YD. Detection of Fcgamma receptors on human endothelial cells stimulated with cytokines tumour necrosis factor-alpha (TNF-alpha) and interferon-gamma (IFN-gamma). Clin Exp Immunol 1998;112:533–8. [32] Alberto MF, Bermejo EI, Lazzari MA. Receptor expression for IgG constant fraction in human umbilical vein endothelial cells. Thromb Res 2000;97:505–11. [33] Vielma S, Virella G, Gorod A, Lopes-Virella M. Chlamydophila pneumoniae infection of human aortic endothelial cells induces the expression of FC gamma receptor II (FcgammaRII). Clin Immunol 2002;104:265–73. [34] Arvidsson S, Askendal A, Tengvall P. Blood plasma contact activation on silicon, titanium and aluminium. Biomaterials 2007;28:1346–54. [35] Leroux JC, De JF, Anner B, Doelker E, Gurny R. An investigation on the role of plasma and serum opsonins on the internalization of biodegradable poly(D,Llactic acid) nanoparticles by human monocytes. Life Sci 1995;57:695–703. [36] Klein CL, Bittinger F, Kohler H, Wagner M, Otto M, Hermanns I, et al. Comparative studies on vascular endothelium in vitro. 3. Effects of cytokines on the expression of E-selectin, ICAM-1 and VCAM-1 by cultured human endothelial cells obtained from different passages. Pathobiology 1995;63: 83–92. [37] Scholz D, Devaux B, Hirche A, Potzsch B, Kropp B, Schaper W, et al. Expression of adhesion molecules is specific and time-dependent in cytokine-stimulated endothelial cells in culture. Cell Tissue Res 1996;284:415–23. [38] Campbell RB, Fukumura D, Brown EB, Mazzola LM, Izumi Y, Jain RK, et al. Cationic charge determines the distribution of liposomes between the vascular and extravascular compartments of tumors. Cancer Res 2002;62:6831–6. [39] Borchard G, Kreuter J. The role of serum complement on the organ distribution of intravenously administered poly(methyl methacrylate) nanoparticles: effects of pre-coating with plasma and with serum complement. Pharm Res 1996;13:1055–8. [40] Sakhalkar HS, Dalal MK, Salem AK, Ansari R, Fu J, Kiani MD, et al. Leukocyteinspired biodegradable particles that selectively and avidly adhere to inflamed endothelium in vitro and in vivo. Proc Natl Acad Sci U S A 2003;100:15895–900. [41] Muller WA, Weigl SA, Deng X, Phillips DM. PECAM-1 is required for transendothelial migration of leukocytes. J Exp Med 1993;178:449–60. [42] Schenkel AR, Mamdouh Z, Chen X, Liebman RM, Muller WA. CD99 plays a major role in migration of monocytes through endothelial junctions. Nat Immunol 2002;3:116–8.