Interaction of Nanoparticles with Cells - DATAPDF.COM

Biomacromolecules 2009, 10, 2379–2400

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Interaction of Nanoparticles with Cells Volker Maila¨nder*,†,‡,§ and Katharina Landfester†,| Max Planck Institute for Polymer Research, Ackermannweg 10, 55128 Mainz, Germany, University Medicine of the Johannes Gutenberg University, III. Medical Clinic, Langenbeckstr. 1, 55131 Mainz, Germany, Institute for Clinical Transfusion Medicine and Immunogenetics Ulm, Department of Transfusion Medicine, University of Ulm, Helmholtzstr. 10, 89081 Ulm, Germany, and Institute of Organic Chemistry III-Macromolecular Chemistry and Organic Materials, University of Ulm, Albert-Einstein-Allee 11, 89081 Ulm, Germany Received March 4, 2009; Revised Manuscript Received May 20, 2009

Nanoparticles and their interaction with human cells have been a focus of many groups during the past decade. We discuss and review here the progress in the field of understanding and harnessing the interactions of polymeric nanoparticles synthesized by the miniemulsion process with different cell types. Nanotechnology and the hereby produced nanomaterials have promised to make use of specific properties of supramolecular assemblies and nanomaterials so that hitherto inaccessible effects can be exploited for new applications. Examples are superparamagnetism or the high surface area helpful for catalysis and adsorption. In biology and medicine, superparamagnetic iron oxide nanoparticles have been used for cell selection and as magnetic resonance imaging (MRI) contrast agents. Furthermore, uptake of nanoparticles into a wide variety of cells is an effect that seems to be specific for materials in the range of 50-200 nm. Surface modifications (positively or negatively charged side groups of the polymers, amino acids, or peptides/proteins) enhance this uptake. Knowledge about factors influencing cellular uptake, like size, surface properties, cell type, and endocytotic pathways, enables optimization of labeling and selection of cells and nanoparticles for applications in vitro and in vivo. For in vivo applications, we will focus on how nanoparticles can cross the blood-brain barrier.

1. Introduction In the past few years we and others have been investigating how cells derived from human origin interact with nanomaterials. This is especially interesting as cellular therapy offers great opportunities for regenerative medicine, especially repair of tissue function after organ damage. Here nanomaterials carrying specific molecules could influence the fate of differentiation of these cells or enable the detection of migration and homing of cells. Nanomaterials are characterized by their size, which is well below the micrometer range where we would locate cells. On the other side, they are larger than single molecules, small molecules, or even proteins, that is, they are in a range of several nanometers up to some hundred nanometers. These materials are referred to as nanomaterials. Nanotechnology and the hereby produced nanomaterials have promised to make use of specific properties of such supramolecular assemblies and materials so that hitherto inaccessible effects can be exploited for new applications. Many of these properties are not active for single molecules or assemblies in the micrometer range. Some effects that are only found in this size range are in the field of physics, for example, optical properties of nanoparticles (gold nanoparticles in different colors depending on their size, fluorescent “quantum dots”), superparamagnetism of small magnetic nanoparticles,1 or supercool* To whom correspondence should be addressed. Tel.: +49(0) 152 03125250. Fax: +49(0)6131 379-100. E-mail: volker.mailaender@ mpip-mainz.mpg.de. † Max Planck Institute for Polymer Research. ‡ University Medicine of the Johannes Gutenberg University. § Institute for Clinical Transfusion Medicine and Immunogenetics Ulm. | Institute of Organic Chemistry III-Macromolecular Chemistry and Organic Materials.

ing of fluids in confined geometries.2 Some of these effects are exploited in chemistry like the high surface area for catalysis3 and adsorption.4 In biology and medicine, superparamagnetic iron oxide nanoparticles have been used for cell selection5 and as magnetic resonance imaging (MRI) contrast agents. During the last years when we and others have been working in this field it became clear that in cell biology one of the effects that seems to be specific for materials in the range of 50-200 nm is the uptake of nanoparticles into a wide variety of cells.6,7 While labeling and selection of cells are possible applications, targeted nanoparticulate drug delivery is one of the most promising techniques for increasing the efficiency of drugs8,9 and also of influencing the differentiation of stem cells. With the incorporation or adsorption of drugs to a carrier system, the drug can be effectively protected from degradation or metabolization after its administration. Even new routes of administration, like, for example, oral application of insulin or heparin might be possible10,11 or transdermal delivery of drugs.12,13 Nanoparticles are also a promising tool for the delivery of drugs that were abandoned from further development because they had no access to their intracellular target or they were sensitive to extracellular degradation like enzymatic digestion of nucleic acids. Also a variety of small molecules is either not taken up by most cell types or may be even actively removed from the cytoplasm of the cell, for example, by multidrug resistance related proteins.14 While hematopoetic stem cell populations have been collected, processed, and manipulated for clinical reapplications in malignant and nonmalignant diseases for more than three decades, further applications have been under investigation in the last years. Regenerative therapy using cell preparations of different origin to repair damaged tissue is a rapidly evolving field in medicine. To understand and harness the possibilities

10.1021/bm900266r CCC: $40.75  2009 American Chemical Society Published on Web 07/28/2009

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Biomacromolecules, Vol. 10, No. 9, 2009

Figure 1. Schematic of the miniemulsion process.

of nanoparticles for (stem) cell therapy the interactions of nanoparticles with cells need to be investigated. This review will focus on the use of polymeric nanoparticles as synthesized by the miniemulsion process and on superparamagnetic iron oxide nanoparticles as they are used in medicine for MRI imaging as they are approved for use in humans and therefore would be readily available for clinical studies. It is the motivation of this review to demonstrate the influence of different polymeric materials on the interaction with cells. Here the influence of the polymer itself and other components, like surfactants, used for the preparation of nanoparticles are reviewed. In addition, the interaction behavior should be altered deliberately by using monomers that have substituted side groups. Hereby, surface functionalization can be achieved. The surface of the nanoparticles, which is the interaction partner with the cell surface and other cellular compartments and proteins, is altered intentionally, for example, charge, amino acids, and so on. We set out to describe and study the influence of these alterations on the amount of uptake. This is interesting as either a high amount uptake of nanoparticles is advantageous as in intracellular drug delivery or cell labeling or it should be avoided as in the extracellular delivery of a drug. Here also the influence of the density of side groups per square nanometer at least for some nanoparticles should be investigated, thereby establishing a kind of density-uptake relationship. Also the uptake kinetic is summarized here. Subcellular localization is to be determined, for example, to use nanoparticles for DNA delivery it would be desirable to bring nanoparticles into the nucleus. We also wanted to investigate and dissect the uptake mechanisms by which these nanoparticles are endocytosed by cells, and for this purpose, pharmacological inhibitors for defined endocytotic pathways are used. Besides the individual cell membrane, there are even more complex barriers as they are present in vivo, like the blood-brain barrier, which consists of several cell types controlling the translocation of substances and also nanoparticles into the brain. Furthermore, the usefulness of a variety of reporters is demonstrated here. The nanoparticles can either act as marker systems for future in vivo studies (like investigations on homing and trafficking of (stem) cells) or in a more complex form with encapsulated substances as possible drug delivery systems. 1.1. Nanoparticle Synthesis. The experimental nanoparticles were synthesized via the miniemulsion process, which enables the defined modification of the nanoparticles’ parameters with a complex structure. The principle of miniemulsion is described in detail in ref 2 and shown in Figure 1. Shortly, an oily phase and a water phase are mixed. Oil droplets are formed by stirring.

Maila¨nder and Landfester

These are in the micrometer range and their sizes differ in a wide range. To homogenize them, high shear stress is applied, for example, by ultrasonification or high pressure homogenizers. Hereby nanodroplets are formed and these are stabilized by a surfactant. If the oily phase consists of monomers, they can be polymerized; therefore, the geometry and size of the droplets is preserved. Also, substances in the nanoparticles are hereby enclosed. For cell experiments, many of the nanoparticles were dialyzed to get rid of the nonreacted monomers and most of the surfactant. By picking a different surfactant and changing the ratio of water/oil, one can also yield aqueous nanodroplets in an oily continuous phase (inverse miniemulsion). As can be seen from the principle of miniemulsion, a wide variety of polymers, enclosed substances (reporters, drugs etc.), surfactants, and geometries (nanoparticles, nanocapsules etc.) with different surface functionalities can be synthesized (see Figure 2 for illustration). 1.1.1. Nanoparticle CoVered in this ReView. Besides the commercially available nanoparticles of which we have used two superparamagnetic nanoparticles that are approved for use in humans (Feridex (marketed in Europe as Endorem) and Resovist), we have started to investigate the interaction of nanoparticles with cells by using specifically designed nanoparticles. Each set of these nanoparticles was designed to answer specific questions on the interaction of nanoparticles with cells. These nanoparticles should not be toxic for the cell tested. We started with polystyrene nanoparticles (see Table 1) because styrene is cheap, well-known, and the polymerization in miniemulsions yields polymeric nanoparticles with a narrow size distribution. These nanoparticles show long-term stability on the shelf and in cell experiments, as they cannot be degraded. In several series, an unmodified particle system was synthesized (see Table 1, P1, AMPM0, VHPM0, VHMPM0). The surface can be easily altered by addition of a comonomer that, for example, yields carboxylic groups on the surface when acrylic acid is used (“VHPM” and “VHMPM” nanoparticles). Further modifications like adding amino acids can be performed (Table 1, nanoparticle VH3KaLys). The carboxyl group is used as anchor for coupling reactions in these cases. A comonomer with amino groups has also been used for these studies (aminoethyl methacrylate hydrochloride ((AEMH), see Table 1, “HF” and “AMPM” nanoparticles). Different reporters can be enclosed in a (hydrophobic) polymer shell (see Table 1) of which a fluorescent dye (PMI), superparamagnetic iron oxide particles (SPIO), and gadolinium complexes (gadovist, magnevist) were studied. To include hydrophilic substances, nanocapsules were obtained in an inverse miniemulsion process (Table 1, “cap” nanocapsules). First, nanodroplets of 100-550 nm were formed. These are subsequently encapsulated by an interfacial polymerization in polymeric shells of polyurethane, polyurea, and cross-linked dextran. The shell thickness was adjusted by the monomer concentration, resulting in wall thicknesses of 10-50 nm. In further studies we speculated that the uptake of nanoparticles into cells could be facilitated if polymers with a high similarity to natural structures are used. The terpene structure is found in many substances in nature, like in essential oils and pheromones and is known as the monomer isoprene in polymer science (see Table 1, “mkpi” nanoparticles). The uptake of the nonfunctionalized polyisoprene nanoparticles, without any transfection agents, was analyzed, while the remaining double bonds

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Figure 2. “Chemical toolbox” from which nanoparticles can be built is illustrated.

in the polyisoprene latex can also be used to functionalize the polymeric particles.15,16 As unfunctionalized nanoparticles were taken up in high amounts as compared to polystyrene we synthesized composite nanoparticles of different amounts of polystyrene/polyisoprene in order to tune the amount of nanoparticles taken up (Table 1, “mksi” and see chapter 3.1.3). Poly(methyl methacrylate) (PMMA) was chosen as for some applications, like vaccination, these nanoparticles are proposed17 although they are not biodegradable. While the polymers discussed before are not biodegradable, poly(alkylcyanoacrylate) nanoparticles are biocompatible and biodegradable and can absorb or entrap bioactive compounds. Therefore, we investigated them as tools for applications where the nanoparticles are administered, for example, as drug carrier systems (Table 1). For poly(alkylcyanoacrylate) nanoparticles, also, penetration of the blood-brain barrier has been described. Another type of biodegradable nanoparticles was made from polyesters: poly(ε-caprolactone), poly(D,L-lactide), poly(D,Llactide-co-glycolide). These have been chosen as they show differences in their degradation time and hydrophobicity (Table 1, “PLLA”, “PCL”, “PLGA”, see chapter 3.1.6). As we speculated that the surface charge would influence the uptake of nanoparticles into cells, a series of fluorescent polystyrene latex particles with carboxyl and amino functionalities on their surface was synthesized by the miniemulsion technique. Furthermore amino acids like lysine (the monomer of poly-L-lysine, a transfection agent used here and by other groups) was coupled to the surface. For poly(n-butylcyanoacrylate) (PBCA) nanoparticles also poly(ethylene glycol) was coupled to decrease unspecific cellular uptake (Table 1, nanoparticles MP and MH). 1.2. Cell Types. Because the interaction of nanoparticles with cells can possibly depend on the cell type, but on the other hand, not all available cell lines or primary cell types can be screened, we selected four different cell types with which the nanoparticles could be investigated: a primary human stem cell type, that is, mesenchymal stem cells (MSC), and the three cell lines HeLa, Jurkat, and KG1a. These cell lines were chosen as they can serve as models for clinically interesting cellular targets. Mesenchymal stem cells (MSC) are an interesting cell population for regenerative medicine.18-20 MSCs are characterized by a high proliferation and differentiation potential making them the ideal candidate for cellular therapeutics in regenerative medicine.20 They were characterized by surface markers (CD 73, 90, 105 positive, while negative for a range of other markers like CD 34, 45) and by their differentiation potential (osteogenic,

chondrogenic, adipogenic). Therefore, many of the nanoparticles mentioned below were tested in MSCs. Malignant cell lines are easy to culture and tumors are interesting targets for in vivo applications of nanoparticles. Therefore, we chose HeLa as a model cell line. Jurkat cells served as model for T cells. Migration and homing of these cells for immune therapeutic application is studied by many groups. Finally, KG1a cells are a primitive cell line of the hematopoetic system. They are therefore used in many studies as a model for CD34+ human hematopoetic stem cells. Not all, but still many, nanoparticles were tested in Jurkat and KG1a cells reviewed here.

2. Nanoparticles as Labeling Agents for Cellular Therapeutics To detect homing and migration of transplanted cells, techniques like bioluminescence,21,22 radioactive substrates,23 near-infrared fluorescence24,25 and labeling with magnetic resonance imaging (MRI) contrast agents26 are applied in small animal studies. Of these, only labeling with radioactive agents and MRI contrast agents are suitable for studies in humans as well. Since the early use of “dextran-magnetite” nanoparticles in the 1970s and the discovery that they have a strong influence on the T2 in MRI was discovered,27 it took another decade until they were used for the enhancement of liver metastasis28 and lymph nodes29 by the group of Weissleder et al. Similar nanoparticles also evolved as tools for selection of cells5 and they are the basis of the MACS technology that is successfully used for graft engineering in transfusion medicine. A further modification was done by Bulte and the group of Frank/Arbab, as they have been using these “dextran magnetite” nanoparticles and incubated them with cells to follow the fate of these nanoparticles in vivo.26,30,31 These last groups were mostly radiologist and therefore they were studying only a limited number of nanoparticles. These studies used the then marketed nanoparticles (e.g., Feridex) without further chemical modification. Therefore, in these studies no investigations into the basic questions of the interaction nanoparticles/cells were possible. Only by using cross-linked dextran-coated iron oxide nanoparticles (“CLIOs”32) were modifications of nanoparticles made possible, but without changing the basic materials or studying the influence of modifications (e.g., surface charge, side groups) in a dose-dependent manner. We also started our work on clinically approved nanoparticles. The results lead to more specific questions concerning the

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Table 1. Overview of Nanoparticlesa nanoparticle carboxyl (COO-) functionalized NPs with different densities

control amino (NH3+) functionalized NPs with different densities

amino (NH3+) functionalized NPs with different densities

carboxyl (COO-) functionalized NPs with different densities and superparamagnetic cores lysine NP

monomer comonomer

polyisoprene/ polystyrene NPs

polystyrene NPs for endocytosis studies

zeta-potentialb (mV)

101 327 119 110 222 329 313 405 n.d. 587 >1 µm 397 222 597 277 116 57 190 155 162 136 162 162 146

PLLAH-1 poly(L-lactide) PLLAL-15 PCLH-16 poly(ε-ca-prolactone) PMI PCLL-17 PLGA-18 poly(D,L-lactide-co-glycolide) mkpi033 mkpi067

121c 106c 165c 139c 76c 192 145

-42c -41c -54c -47c -51c -59.6 -63.8

mkpi076 mkpi073 mkpi075 mkpi071 mkpi072 mksi13 mksi12 mksi11 mksi10 mksi09 mksi15 mksi06 mksi05 mksi14 PS+ PS-

138 81 140 86 100 168 163 154 158 155 141 131 134 148 113 121

+81.4 +60.7 +66.8 +74.0 +42.8 -57.5 -64.6 -58.3 -67.1 -59.2 -65.3 -64.7 -74.6 -69.4 +59 -60

PMMA cap17 cap19 cap24 cap25 nanocapsules (NCs) from cap26 cap74 polyurethane (PU) cap35 cap37 cap36 cap38 cap16 NCs from polyurea cap 18 cap31 NCs from dextran cap 33 UP-1 UP-2 UP-3 UH PP NPs from poly(butylcyan-acrylate) PH MP MH T80

NPs from polyisoprene

size (nm)

-20 -50 -59 -62 -64 -51 -42 -59 n.d. n.d. n.d. n.d. n.d. n.d. -12 -8 -2 5 27 29 32 36 n.d. n.d. n.d. n.d. n.d. n.d. -47.5 ) VH3Ka after EDC coupling of lysine -41 n.d. n.d. n.d. n.d. n.d. n.d. n.d 20-36 n.d. n.d. n.d. n.d. n.d. n.d. -70 -47 n.d. -72 -57 -89 -47 -51 -5

VHPM0 VHPM1 VHPM2 VHPM3 VHPM5 VHPM10 VHPM15 VHPM20 P1 HF3 HF4 HF5 HF11 HF6 AMPM0 AMPM1 AMPM2 AMPM3 AMPM5 AMPM10 AMPM15 AMPM20 VHMPM0 VHMPM1 VHMPM2 VHMPM5 VHMPM10 VHMPM15 VH3Ka VH3KaLys

poly(methy methacrylate)

biodegradable NPs

reporter

styrene

101 99 98 styrene + acrylic acid (AA) 100 97 number in NP name 103 gives wt% of AA) 148 150 styrene PMI 168 169 206 337 styrene +AEMH PMI 489 1290 styrene 217 175 162 styrene + AEMH (number in 154 NP name gives wt% 160 of AA) 155 143 119 57 53 styrene + acrylic acid (AA) 45 (number in NP name PMI + SPIO 52 46 gives wt% of AA) 67 68 see VH3Ka 151 -10.9 methyl methacrylate TDC + hexane-1,6-diol

PMI

PMI magnevist

TDC +propane-1,3-diol magnevist TDC +glycerol

gadovist 178 magnevist

TDC + diethylenetriamine

magnevist

cross-linked dextran

magnevist

butylcyan-acrylate

isoprene

PMI

PMI

isoprene/styrene

PMI

styrene

PMI

paper (ref no.)

note control NP

COO- per nm2:

0.19 0.31 0.39 0.40 0.66 2.19 4.48

control NP NH3+ per nm2:

0.07 0.09 0.10 0.61 15.11

control NP +

2

NH3 per nm :

COO- per nm2:

shell thickness:

shell thickness: shell thickness: shell thickness: shell thickness:

0.10 0.17 0.26 0.50 0.94 1.16 1.35 0.100 0.107 0.117 0.378 0.803 1.463 0.437

16-26 23-33 25-50 10-30 10-30 20-40 13-24 15-25 19-35 18-30 23-40 10-20 20-45

no functionalization Phe functionalized PEG functionalized

86 86 86 86 86 86

78 78 78 78 78 78 78 78 78 78 78 78 78 78 83 83 83 83 83 83 83 150

polysorbate 80 as surfactant. For in vivo studies. NPs with Lutensol as surfactant showed a larger diameter and a higher zeta potential SDS as surfactant longer sonication compared to Mkpi033

125 125 125 125 125 87 87

different surfactants used (see P11 for details)

wt%PI/wt%PS

86 47, 47, 86 47, 47, 47, 47, 6 6 6 6 6 6 6 6 6 6 6 6 6 6 62 62 62 62 62 62 62 62

90:10 80:20 70:30 60:40 50:50 30:70 20:80 10:80 0:100

87 87 87 87 87 87 87 87 87 87 87 87 87 87 85 85

a n.a. ) not applicable; n.d. ) not done; TDC ) tolylene-2,4-diisocyanate; NP ) nanoparticle; NC ) nanocapsule. b If zeta potential was measured at more than one pH, then the one closer to pH 7 was chosen for this overview; also, figures after dialysis were preferred over the before dialysis figure. c Values with SDS are given.

characteristics of the interaction of nanoparticles with cells. To answer these questions, nanoparticles with different reporters were synthesized. Here the miniemulsion process allows one to encapsulate many of the available reporters, like fluorescent dyes, magnetite (iron oxide, Fe3O4), gadolinium complexes, or nuclides. To answer specific questions, miniemulsion nanopar-

ticles varying only a specific parameter (like surface charge) and having an easily detectable reporter can be synthesized. This makes studies on the interaction of nanoparticles with cells easier as fluorescence microscopy and flow cytometers can be used for these studies instead of using wet chemistry detection of iron or Prussian blue staining, with the last two techniques being

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Figure 3. MSC stained with Prussian blue. MSC were incubated with SPIOs with and without PLL as transfection agent and cellular labeling was visualized histochemically by the Prussian blue reaction. (a) MSC without SPIO or PLL (control); (b) PLL only; (c) Feri1x/PLL; (d) Feri1x/s; (e) Feri10x/s; (f) Reso1x/PLL; (g) Reso1x/s; (h) Reso10x/s 1x)25 µg Fe/mL, 10x)250 µg Fe/mL. Scale bars are 50 µm wide. (Published with kind permission from Springer Science+Business Media: Maila¨nder, V.; Lorenz, M. R.; Holzapfel, V.; Musyanovych, A.; Fuchs, K.; Wiesneth, M.; Walther, P.; Landfester, K.; Schrezenmeier, H. Carboxylated Superparamagnetic Iron Oxide Particles Label Cells Intracellularly Without Transfection Agents.Mol. Imaging Biol. 2008, 10, 138-146, Figure 1. Copyright 2008.)

less sensitive for detection of nanoparticles than techniques using a fluorescent readout. Even two reporters can be included in one particle like a fluorescent dye and a superparamagnetic particle. 2.1. Reporters for MRI. Magnetic resonance imaging (MRI) is based on local proton concentration and the relaxation properties of the protons, which are influenced by the local environment. To enhance anatomic details such as blood vessels or tumors, contrast agents, which locally alter the relaxation properties of the protons, are administered. Contrast agents, which are used in current clinical practice, fall into two main categories: Superparamagnetic compounds such as iron oxide have a strong effect on the local longitudinal (T1) and transversal (T2) relaxation properties of the protons, with a more pronounced effect on the T2 properties. These agents normally appear hypointense in the final image. The second class of contrast agents utilizes paramagnetic compounds such as lanthanides (like gadolinium), which mainly reduce the T1 relaxation property and result in a brighter signal, if a T1 sensitive MRI technique is used. 2.2. Commercially Available Iron Oxide (Magnetite) Nanoparticles as Reporters for In Vitro and In Vivo Applications. The superparamagnetic effect of iron oxide nanoparticles, which are typically around 10 nm in size, makes them not only interesting contrast agents in magnetic resonance tomography,33 but they are also used as nonviral vehicles for gene therapy,34,35 drug delivery,9,36,37 immunization,11,38-40 and detoxification.4 Most approaches for cell labeling for in vivo studies in animals utilizes superparamagnetic iron oxide particles (SPIOs).

SPIOs are commercially available and FDA-approved for use in humans.26,30 However, to achieve intracellular uptake of the nanoparticles, transfection agents are needed in most studies.26,30,41-44 These transfection agents like Superfect, DOTAP, Lipofectamin,41,42 poly-L-lysine (PLL),42,44 or protamine43 are mostly cationic, positively charged molecules. Higher concentrations of these agents are toxic45,46 and, with the exception of protamine, they are not approved for any indication in clinical use. Avoiding a transfection agent in the process of cell labeling seems to be favorable as this would simplify the approval of studies in humans. For our initial experiments we used the two available contrast agents Resovist (generic name: ferucarbotran) and Feridex (generic name: ferumoxides). The magnetite nanoparticle aggregates are stabilized in the case of Resovist by carboxydextran, in Feridex by nonfunctionalized dextran. By evaluating the standard method for cell labeling (SPIOs plus transfection agent26,30,42), a strong Prussian blue staining was observed if the cells were incubated with the SPIOs plus transfection agent (see Figure 3c,f), and the results were easily reproduced as reported in the literature. Surprisingly, especially in Resovist without a transfection agent, cells also showed a few bluish spots in their cytoplasm (see Figure 3g). Triggered by this observation, we increased the amount of Resovist and Feridex and showed that Resovist was taken up spontaneously, that is, without the need for a transfection agent (Figure 3h). These findings were just missed by others, as most other groups were satisfied by evaluating one dose of SPIOs. The results from light microscopy were confirmed by TEM studies (see Figure 2 in ref 47).

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One of the factors that determine the detection limit of cells labeled with SPIOs is the amount of SPIOs per cell and the number of cells per voxel.48,49 Therefore, the amount of iron in each cell by using a quantitative wet chemical assay (ferrocine-based assay) was determined. For SPIOs with transfection agents, we found about the same amount of iron in our cell preparations as has been reported by several other groups with a range of 10 to 25 pg Fe/cell for MSCs and HeLas when they used SPIOs with a transfection agent.26,42 Without transfection agent and at a low concentration of SPIOs (i.e., 25 µg Fe/mL in incubation media), the uptake of iron oxide was not detected above background level (see ref 47, Figure 3, columns “Reso1x/---”, “Feri1x/---”) as has been also observed by other groups. However, by increasing the amount of SPIOs during incubation, there was a sharp increase of calculated iron per cell (see ref 47, Figure 3). Other groups have also reported such high payloads of labeling agent per cell.44,50-52 Nevertheless, we were questioning these results as from a biological point of view it would have been anticipated that the quantity of intracellular storage volume or the uptake mechanism should be a limiting factor and therefore the curve should flatten with higher concentrations of SPIOs, that is, should show a plateau. This has been demonstrated for other substances like transferrin, which are endocytosed by cells.53 On the other hand, we have shown for experimental SPIO nanoparticles that extracellular aggregation in cell culture media occurs with the nanoparticles cross-linked by material that was not observed with nanoparticle preparations without serum containing media (see Figure 13), white arrowheads). These conglomerates cannot be easily washed away as they adhere to the surface of the cells, they are also not digested by trypsin, at least after a few minutes of incubation at the concentrations of trypsin used for detachment of cells from the cell culture surface. This may be due to the fact that large quantities of proteins need to be cleaved before these agglomerates disintegrate and the trypsin would need more time. As in the literature, no method for removing or quantifying the amount of coagulated nanoparticles was reported before in such a complex nanoparticle-cell culture system, flow cytometry, also known as fluorescence-activated cell sorting (FACS), as a tool for quantifying aggregation (see ref 6, Figure 6, and ref 47, Figure 4) was evaluated. When nanoparticles are not coagulated, they are usually not detected by FACS, as most users set a threshold on the forward or sideward scatter in order not to count cellular-derived particles and contaminations (see ref 6, Figure 4b, c). But if the coagulated material eventually forms large enough clusters to be detected in the forward FACS scatter plot these events can be quantified. The coagulates can be further distinguished from cells as the coagulated particles do not express specific cell surface markers like CD55 for HeLas or CD44 for MSC. This distinct population of coagulated particles was detected for higher concentrations of SPIOs without a transfection agent and for SPIOs with a transfection agent. To evaluate only the uptake of live cells and not dead or dying (i.e., apoptotic) cells, we used 7-AAD, which is actively pumped out from living cells (i.e., negative for 7-AAD), while apoptotic cells are stained dim and dead cells appear as 7-AAD bright positive. After sorting for live cells (CD55 pos or CD44 pos/7-AAD neg/high FCS) the quantitative analysis of the iron content per cell revealed distinctively lower amounts of iron per cell as the coagulated clumps of nanoparticles were excluded from the quantification (Figure 4).


The relevance of this observation is not only that the amount of iron uptake reported in the literature must be questioned, but that these conglomerates would also hamper further applications. They will be indistinguishable from the labeled cells once brought into the bloodstream together with the labeled cells, especially if they are taken up by macrophages. Therefore, it would be questionable for signals detected by MRI if these signals represent a migrating target cell, aggregates of the nanoparticles, or macrophages that have phagocytosed injected aggregates. As a result we feel that iron contents per cell should only be reported if the extracellular aggregates have been removed thoroughly. FACS sorting is a well established way of purifying a cell population based on fluorescent markers and FSC/SSC and yields pure cell populations.54 After having evaluated FACS sorting for removing the coagulated nanoparticles in these cell preparations, we focused on the newly established conditions for intracellular uptake with higher SPIO concentrations without transfection agents. We examined the differences between Feridex and Resovist. For HeLa cells, Resovist is a suitable agent for labeling at a high concentration without the need to use a transfection agent (Figure 4, right graph, light gray column “Reso10x/---”). In contrast, Feridex is not so effective, demonstrating a difference between the two SPIO preparations (light grey column “Feri10x/---”). For MSCs, Resovist at a high concentration is very nicely suited and Feridex yields at least a detectable uptake (Figure 4, left graph). With these higher concentrations of SPIOs, but also when using transfection agents, special care should be taken when reporting iron contents per cell as it is crucial to remove SPIO aggregates from the media and from the cell surface. Using a fluorescent activated cell sorter is an elegant method for small-scale purification of cell preparations, although other methods need to be implemented if clinical studies should be performed. As the differential behavior concerning the uptake into cells between the two nanoparticles Resovist and Feridex was established, we were looking for differences between these two commercially available nanoparticles. Both nanoparticle preparations have embedded superparamagnetic cores of 10-12 nm in a biocompatible polymer yielding particles of a hydrodynamic diameter of 70-150 nm. The most striking difference in these nanoparticles lies in the biopolymers that are used for synthesis. The polymer in the case of Resovist is carboxydextran. Under physiological conditions (pH 7.4), carboxydextran is mostly in the deprotonated form, that is, negatively charged (or anionic). In contrast, Feridex is prepared with dextran, which is not bearing a negative charge. Therefore, we hypothesized that the negatively charged carboxyl groups of the carboxydextran are responsible for the differences seen between Resovist and Feridex. Although most transfection agents are cationic, that is, positively charged, also negatively charged substances can exert such a function.55,56 These anionic transfection agents have been mostly used to transfect small molecules or DNA but not nanoparticles. From these experiments we hypothesized that the anionic charge in the form of carboxyl groups in the carboxydextran of Resovist are responsible for a more efficient uptake. Furthermore, we wanted to know if the function of the positively charged poly-L-lysine, which probably absorbs to the surface of the nanoparticles, can be mimicked by modification of the surface of the nanoparticles by embedding amino groups on the surface. Third, we wanted to know if we could mimic the effect of separately added transfection agent poly-L-lysine, which is then only physically absorbed on the nanoparticle surface

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Figure 4. Quantitative analysis of iron content of cell preparations before sort (dark bars) and after (light gray bars) sorting the fraction of live cells (CD55 or CD44 pos/7-AAD neg/high FSC). (a) HeLa cells incubated with and without SPIOs or PLL and iron content calculated as pg Fe/cell. (b) MSC cells incubated with and without SPIOs or PLL and iron content calculated as pg Fe/cell. 1× ) 25 µg Fe/mL. Error bars are standard deviation of triplicates. (Reprinted with kind permission from Springer Science+Business Media: Maila¨nder, V.; Lorenz, M. R.; Holzapfel, V.; Musyanovych, A.; Fuchs, K.; Wiesneth, M.; Walther, P.; Landfester, K.; Schrezenmeier, H. Carboxylated Superparamagnetic Iron Oxide Particles Label Cells Intracellularly Without Transfection Agents.Mol. Imaging Biol. 2008, 10, 138-146, Figure 5. Copyright 2008.)

presumably by covalently binding the monomer lysine to the nanoparticles’ surface. By using functional comonomers in the miniemulsion polymerization process, such surface functionalizations can be achieved (see section 3.2.1). Fluorescent reporters can be embedded into the polymeric nanoparticles during the synthetic process and can therefore enable rapid testing and quantification (see section 2.5). 2.3. Experimental Polystyrene Nanoparticles with Iron Oxide (Magnetite) and a Fluorescent Dye as Reporter. To improve the properties of the magnetite nanoparticles with regard to aggregation or coagulation and also iron leakage, it was desirable that the magnetic nanoparticles can be efficiently encapsulated in a hydrophobic polymer shell in contrast to the biologically unstable dextran coating. This goal was achieved by Landfester et al. by using the miniemulsion process just a while ago.57,58 The encapsulation ensures that the shell is not washed off in hydrophilic media, which would result in sedimentation and aggregation of the magnetic core particles. At the same time, high magnetite contents and uniform distribution of magnetite in the polymer can be achieved.58 The polymer chosen for encapsulation in the studies discussed here is polystyrene because it is cheap, well-known, and can easily be functionalized by copolymerization, which allowed answering the main question of the previous studies, which is control and dependency of the amount of cell uptake by modification of the surface properties (see section 3.2). As polystyrene is not biodegradable, it can be used for long-term studies in animal models while studies in humans would be prohibited. Toxicity of polystyrene nanoparticles has been studied, and although some toxicity was seen with nanoparticles in macrophages,59,60 it appears to be less toxic than poly(methyl methacrylate) particles, which are under investigation for applications in humans.61 To have an easy way to follow the behavior of nanoparticles, dual-reporter nanoparticles, that is, nanoparticles that can be detected by different ways of detection, for example, by fluorescence and by MRI, were synthesized (see ref 62). A series of magnetic polystyrene particles encapsulating magnetite nanoparticles (10-12 nm) in a hydrophobic poly(styrene-co-

acrylic acid) shell that also had the fluorescent dye PMI encapsulated was synthesized by a three-step miniemulsion process (see ref 62, Table 1). A high amount of iron oxide (magnetite) was incorporated by this process (typically 30-40% (w/w)). This polymerization of the monomer styrene yielded nanoparticles in the range of 45 to 70 nm. These results have been confirmed by others.63,64 Furthermore, by copolymerization of styrene with the hydrophilic acrylic acid, the amount of carboxyl groups on the surface was varied, thereby enabling quite complex nanoparticles. Functionalization with acrylic acid also offers the opportunity for further functionalization with biomolecules like amino acids or antibodies for targeting the particles. The surface modification can be used for coupling lysine, glutamine or asparagine to the carboxylic groups by the 1-ethyl3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC) coupling.65 Antibodies can be bound to the surface to modify the particles in biomedically interesting ways. For biomedical evaluation, the nanoparticles were incubated with different cell types. The introduction of carboxyl groups on the particle’s surface enhanced the uptake of nanoparticles as demonstrated by the detection of the fluorescent signal by fluorescent activated cell sorter (FACS, see ref 62, Figure 6) and laser scanning microscopy (LSM, see Figure 5, left panel). While nanoparticles where readily detectable by FACS and laser scanning microscopy, we did not find uptake of more than 1 pg Fe/cell (not shown). The quantity of iron oxide in the cells that is required for most biomedical applications (like detection by magnetic resonance imaging) has to be significantly higher (around 10-20 pg Fe/cell) as shown by others.66 A further increase of uptake can be accomplished by transfection agents like poly-L-lysine or other positively charged polymers (see refs 30, 31, and 62, Figure 8, column 1xVH3ka+PLL). This enables detection by Prussian blue staining (Figure 5, right panel). While adding poly-L-lysine to the nanoparticles as a separate reagent as has been described in the literature before26,31,42,66 this functionality can also be engrafted into the surface of the nanoparticles by covalently coupling of lysine to carboxyl groups.

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Figure 5. Left panel: Laser scanning microscopy of HeLa cells. Cell membranes are stained with RH414 in red (contours) and the particles in green (dots in the cells). (A) Untreated control, (B) VHMPM2 (2% of acrylic acid), (C) VHMPM5 (5% of acrylic acid), (D) VHMPM10 (10% of acrylic acid). Right Panel: Prussian blue staining of VH3ka and VH3kaLys in HeLas. While there are no blue spots in A (control cells), there are bluish spots in VH3ka when PLL (B) was added and VH3kaLys (C) even without the addition of a transfection agent. Reprinted with permission from IOP Publishing from ref 62. Copyright 2006 IOP Publishing.

When evaluating these lysine-functionalized nanoparticles we found that the amount of iron that can be transfected with them was even higher than with the nanoparticles with a transfection agent only physically adsorbed (see ref 62, Figure 8). By Prussian blue staining the iron oxide could be detected (Figure 5, right panel). Furthermore, the subcellular localization of these nanoparticles was demonstrated to be clustered in endosomal compartments by TEM (see ref 62, Figure 10). While coagulation and clumping was a major concern with the commercially available dextran nanoparticles Resovist or Feridex we never observed coagulation or clumping with the carboxyl functionalized nanoparticles in the cell culture media. Notably even at the highest concentrations tested all particles which were not washed away were located inside the tested cell types and not only absorbed on the surface of the cells (see ref 62, Figure 7D). This makes this type of nanoparticle suitable for a wide range of biomedical applications such as use as contrast agent for magnetic resonance imaging, hyperthermia,67,68 and selection of cells. 2.4. Gadolinium as a Reporter in Nanoparticles. Commonly used paramagnetic substances that are active in T1 sequences as contrast agents for MRI are hydrophilic paramagnetic gadolinium complexes like Gd(DTPA) or Gd(DOTA). Because the gadolinium complex is still a small molecule, a high dose in terms of molar amount of the complex needs to be administered to achieve a significant contrast enhancement. This can easily lead to increased osmotic pressure, and as the osmotic pressure is proportional to the number of molecules in the bloodstream, a reduction of the number of infused molecules would be favorable. For the reduction of injected osmotic active compounds, several MRI active complexes can be combined into a single (supra-)molecular assembly like a nanoparticle. Additionally, this concept has the potential for functionalization of the nanocapsule’s surface ensuring specific enhancement of regions in which certain enzymes, antigens, or other target structures are located at high local concentrations. Several approaches of combining gadolinium molecules are described in the literature. Lipinski et al. used Gd-containing

immunomicelles in the range of 85 to 130 nm with approximately 4-9 Gd atoms per micelle.69 Because micelles show a limited stability after injection into the bloodstream due to dilution effects, gadolinium-rhodamine nanoparticles for cell labeling and MRI imaging were produced by mixing lipid monomers like phosphatidylcholine or Gd-lipid with diacetylene, a subsequent photolysis polymerization leads to an increased stability.70 Wooley et al synthesized gadolinium-labeled crosslinked nanoparticles with hydrodynamic diameters of about 40 nm using diblock copolymers (poly(acrylic acid) and poly(methacrylic acid)) micelles, which were then covalently crosslinked by amidation with ethyl amines. Gadolinium coordinated pentaacetic acids were then located at the surface of the micelles in a hydrated shell allowing a rapid water exchange and therefore high relaxivities.71 Yu et al. intensively investigated the synthesis of fibrin-targeted contrast agents for fibrin detection of human thrombus in vitro.72 As contrast agent, lipid-encapsulated perfluorocarbon nanoparticles with numerous Gd-DTPA complexes incorporated onto the outer surface were synthesized by other groups.72,73 On particles with 250 nm diameter, more than 90.000 Gd3+ per particle could be attached. A 20-40% decrease in T1 relaxation was demonstrated. The encapsulation of gadolinium complexes inside apoferritin spheres was presented by Aime et al. The technique allowed encapsulation of approximately 10 units of the hydrophilic gadolinium chelate complex per apoferritin sphere, which lead to a significant increase of the T1 relativity compared to free gadolinium chelate.74 A further approach was developed by Reynolds et al. with the synthesis of core-shell nanoparticles loaded with gadolinium salt in an intermediate layer. The nanoparticles were synthesized by a three-step approach. First, a polymeric core was synthesized with carboxylic acid groups as functional groups. In a second step the salt Gd(NO)3 was bound to the carboxylic groups. In the final step a shell is added by a second polymerization to cover the Gd layer. Due to the porous shell a rapid exchange of water and therefore a reduction in relaxation time could be determined.75

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Figure 6. Left: Plot of 1/T1 against the concentration of trivalent gadolinium ion to obtain the T1 relaxivity in PU nanocapsules cap19 and in the pure contrast agent Magnevist. Right: TEM images of PU polymer capsules (cap26). Published with permission from ref 78. Copyright 2007 Wiley-VCH.

To maintain relaxation properties after encapsulation of these lanthanides, ensuring sufficient water exchange between the encapsulated contrast agent and the surrounding 1H protons is the most important factor and concern after encapsulation. Therefore, the shell of the nanocapsule needs to be adjusted so that the gadolinium complex cannot diffuse out while the 1H protons can easily enter and leave the interior of the nanocapsule. Furthermore, the application of polymer nanocapsules as new targeted contrast agents would require stability of the capsules in the bloodstream to avoid leakage of the Gd complex into the systemic blood flow. Tiarks et al. synthesized in a one-step procedure hollow polymer nanocapsules of sizes between 80 and 400 nm with a hydrophobic core by using the direct aqueous miniemulsion technique.76 The inverse miniemulsion process allows the synthesis of nanocapsules with an aqueous core. Polyurea, polythiourea, or polyurethane shells are obtained by interfacial polyaddition at the droplet interface.77 For specific targeting of the contrast agent the polymer shells can be prefunctionalized for further coupling reactions with highly selective agents, e.g. peptides or antibodies. Nanocapsules containing the hydrophilic gadolinium complexes Magnevist and Gadovist (structural formulas, see ref 62, Figure 1) were obtained in an inverse miniemulsion process (see ref 78, Figure 2). First, nanodroplets of 100-550 nm were formed that are subsequently encapsulated by an interfacial polymerization in polymeric shells of polyurethane, polyurea, and cross-linked dextran. The shell thickness was adjusted by the monomer concentration. Hereby the shell can be made impermeable for the Gd complex. In contrast to other methods reported in the literature this system therefore shows a high degree of flexibility. Due to the porosity of the polymeric shell, a diffusion of water molecules through the capsule walls can be ensured, that is, encapsulation does not compromise the T1 relaxivity of the contrast compound in magnetic resonance imaging. Wall thicknesses of 20-40 nm for the nanocapsules were obtained (Figure 6). These nanoparticles were first tested for their relaxivity in cyclohexane as continuous phase, where they show low T1 relaxivity. This is easily understood when keeping in mind that, due to the limited water content inside the capsules and the

missing water molecules outside of the nanocapsules, the number of protons is by far not sufficient for local strong T1 shortening. However, after transferring the nanocapsules to water as continuous phase, the T1 relaxivity is not inferior when compared to nonencapsulated Magnevist solution (Figure 6). The unchanged T1 relaxivities prove a sufficient water/proton exchange through the polymeric shell. The T1 relaxivity was found to depend on the thickness of the polymer shell, which is affecting the efficiency of the water exchange. Because these nanoparticles should be brought into the bloodstream, measurements simulating physiological conditions (isotonic NaCl solution, human blood) were performed (see ref 62, Table 3). Blood cells and proteins could alter the exchange rate of water from the outside to the inside, for example, by coating the nanoparticles and thereby increasing the wall thickness or decreasing porosity. The results clearly show that in all cases the proton exchange was sufficiently high. The suggested approach appears not to be limited to the encapsulation of Magnevist as proven by the Gadovist example. This approach can likely act as a new basis for versatile contrast agents in MRI. The experiments clearly show the potential of using nanocapsules as new contrast agent material for MRI. A functionalization of the nanocapsules is now possible and will allow the specific targeting of defined molecular aims for detection of specific diseases. 2.5. Fluorescent Dyes as Markers in Nanoparticles. Polystyrene and the other polymers can hardly be identified once brought into a biological system and tagging additives are necessary to detect the nanoparticles. While the superparamagnetic iron oxide cores of 10-12 nm are easily identified by TEM or detected by MRI as shown before by others,31,66 synthesis of such complex compound nanoparticles and the subsequent quantification after incubation with cells is much more tedious. The easiest way also from the synthesis part of this work is to incorporate fluorescent dyes.79,80 There are different options for connecting a fluorescent dye to a polymeric particle, for example, physical adsorption onto the particle surface81 or copolymerization of a fluorescent monomer.82 Physical adsorp-

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tion causes problems such as instability in changing environments and is limited by the desorption processes of the adsorbed material. Another possibility is embedding a highly hydrophobic fluorescent dye into the polymeric matrix of the nanoparticle. We showed with the dual-reporter nanoparticles (see section 2.3) that the most versatile readout system is embedding a fluorescent dye into the nanoparticle as this system can be visualized by cLSM and uptake can be quantified by FACS on a single cell level or by fluorescence spectroscopy quite easily and without extensive preparation of the cell samples. In the miniemulsion process, the fluorescent dye is emulsified in the hydrophobic monomer (styrene) phase and it cannot diffuse through the water phase. In a subsequent polymerization process, the monomeric droplets are converted to polymer particles without changing the identity of the droplets,76 thereby enclosing the fluorescent dye. As fluorescent dye, N-(2,6-diisopropylphenyl)perylene-3,4dicarboximide (PMI) was chosen as it is stable during the particle preparation and polymerization process (for structure information on PMI see ref 83, Figure 1). It is also highly hydrophobic. When illuminated by the laser light, for example, in cLSM, it is more stable than many other fluorescent dyes. This enables long-term and repeated evaluation of nanoparticle uptake. Furthermore, excitation with an abundantly available argon laser emitting at 488 nm is possible, while emission is at 539 nm far enough shifted to work with a common setup in FACS and cLSM. Therefore, these nanoparticles are quite well suited for applications, as described also by others.82

3. Uptake of Polymeric Nanoparticles into Cells The interaction with the cell and the uptake process of the polymeric nanoparticles into cells can be influenced by several factors like (i) structure and morphology of the polymer in the nanoparticles themselves. Hydrophilic and hydrophobic polymeric surface properties of nanoparticles are known to influence cell adhesion in the uptake process. Some groups have reported that with increasing hydrophobicity of the polymer the attachment on cells and subsequent internalization is enhanced;84 (ii) surface groups that are covalently bound to the nanoparticles by copolymerization; and (iii)an amphiphilic polymer that is physically adsorbed on the nanoparticles’ surface (as transfection agent). 3.1. Influence of Polymer on Uptake. Because the polymer itself is an important part of the nanoparticles, one should expect that the uptake behavior of nanoparticles could be altered by different polymers. This can only be studied when the surface charge and other parameters (e.g., size) are kept stabile. 3.1.1. Polystyrene. Styrene can be easily emulsified and polymerized in miniemulsions and there are many modifications possible like copolymerization with a monomer that yields surface functionalization (see refs 6, 62, and 85). Polystyrene does not degrade significantly in the context of cellular environment. This is a wanted property for long-term experiments in cell cultures and animals but is undesired for use in humans as accumulation of a substance and the negative side effects thereof are always of concern in human studies. Unmodified polystyrene nanoparticles were only taken up to a low extent by the cell lines tested (see ref 86, Figure 11, column AMPM0). The modifications and the results of the cell experiments are described in detail in section 3.2. 3.1.2. Polyisoprene. The uptake behavior of nanoparticles into cells could be facilitated if polymers with a high similarity to natural structures are used. The terpene structure is found in

Figure 7. Upper chart: FACS analysis of Jurkat cells incubated with different concentrations of dialyzed mkpi033 particles overnight for 24 h. Saturation of the curves is nearly complete after incubation of 2400 mg of solid content of the particles per mL of medium. Lower chart: FACS measurements of the fluorescence intensity of Jurkat cells after incubation of fluorescent PI nanoparticles for different time periods (0, 10, 30 min; 1, 2, 4, 16, and 48 h). In Jurkat cells, halfmaximum uptake is achieved within 1 h. Published with permission from ref 87. Copyright 2008 Wiley-VCH.

many substances in nature, like in essential oils and pheromones, and is known as the monomer isoprene in polymer science. Therefore, we speculated that polyisoprene (PI) nanoparticles are well suited for the interaction with the lipophobic part of the cell membrane and for cell uptake. The remaining double bonds in the PI latex can also be used to functionalize the polymeric particles.15,16 Before we set out to investigate this there was no literature on polyisoprene nanoparticles available. Fluorescent polyisoprene nanoparticles were synthesized by the miniemulsion technique as marker particles for cells. The uptake of the nonfunctionalized polyisoprene nanoparticles, without any transfection agents, was analyzed by experiments with an adherent (HeLa) and also a suspension (Jurkat) cell line. 3.1.2.1. Concentration Dependence of Particles on Uptake. For the analysis of the concentration dependence, two cell lines, HeLa and Jurkat, were incubated with increasing concentrations of PI nanoparticles (ranging from 37.5 to 4800 µg) in the culture medium for 24 h without the use of any transfection agents. Both cell lines (Jurkat Figure 7, upper chart; for HeLa, see ref 87, Figure 4A) show a steadily and rapid increase in the lower

NPs and Stem Cells

particle concentration range, while higher concentrations result in a flattening of the curve in both cell lines. This indicates that there is a maximum capacity of the amount of particles that can be taken up by these cell lines. This state of saturation might be due to a depletion of endocytotic mechanisms or saturation of the storage capacity of the cells. As a particle concentration of 75 µg · mL-1 in the culture medium was optimal for polystyrene nanoparticles in prior experiments and since this is for PI particles well below the saturation level, a particle concentration of 75 µg · mL-1 was used for further experiments. 3.1.2.2. Kinetics of the Uptake. For the evaluation of uptake kinetics, Jurkat and HeLa cells were incubated with a concentration of 75 µg · mL-1 of PI nanoparticles in the culture medium up to 48 h (see Figure 7 and in ref 87, Figure 5). The graphs for both cell lines reveal an exceedingly steep initial slope during the first hour. For polyisoprene nanoparticles, the uptake kinetics show that particle internalization starts during the first minutes of incubation and is finished after 48 h of incubation. This hastily uptake kinetic then levels off and results in saturation. Setting the 48 h value as maximum, the half-maximum value is reached after 1 h for HeLa cells and for Jurkat the half-maximum time is even below 1 h. Unfunctionalized PI nanoparticles show a far more rapid particle uptake during the first 4 h of incubation as compared with the uptake kinetics of PS particles with the transfection agent poly-L-lysin (PLL) or amino-functionalized PS nanoparticles (see ref 87, Figure 6A, PS nanoparticles with PLL; and see ref 87, Figure 6B, 0.21 NH2 groups per nm2). Even after 24 h of incubation with the functionalized PS nanoparticles the graph shows no saturation and the half-maximum value is well above 6 h. As the uptake kinetics of PI and PS nanoparticles are so different two distinct uptake mechanisms are likely. A tentative explanation for the rather high initial uptake of the PI particles might be a receptor involvement, although no specific receptor has been named for such a process of nanoparticle uptake. Receptor mediated internalization processes are usually much faster than constitutional uptake.7,88-90 3.1.2.3. TEM Studies of Intracellular Distribution of PI Nanoparticles. Intracellular distribution of nanoparticles was investigated by transmission electron microscopy (TEM). Like other types of nanoparticles that we and other groups have evaluated, polyisoprene particles are localized in endosomes. Polystyrene nanoparticles or SPIOs were always found in clusters of several nanoparticles (Figure 13). In contrast, polyisoprene particles are found in endosomes as single events. Intracellular PI nanoparticles were visible as black spots representing single nanoparticles (see ref 87, Figures 12 and 13). The particles are equally distributed through the whole cell body (see ref 87, Figure 12A, overview of the cell). After 4 and 24 h they are mainly localized as single PI nanoparticles in endosomes (see ref 87, Figure 12C, D, 24 h incubation; see ref 87, Figure 13, 4 h incubation). No change of this distribution pattern and localization is observed after 24 h of incubation, which is in line with the observations made by FACS. Only if PLL as a transfection agent is used, particles are found in clusters inside the cell, indicating a change of uptake mechanism or intracellular trafficking, like fusion of endosomes (see ref 87, Figure 14). However, the use of PLL also leads disadvantageously to an aggregation of the particles outside of the cells as we and others41 have reported. In conclusion, polyisoprene nanoparticles are internalized by different cell lines that are relevant for biomedical applications


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Figure 8. Uptake in HeLa cells after up to 24 h incubation (normalized on the maximal fluorescence intensity).

and especially Jurkat cells as a model for T cells show a fair amount of uptake in comparison to polystyrene nanoparticles or Feridex or Resovist. Therefore, they can be used to label these cells efficiently if a marker is incorporated in the particles. As polyisoprene is not or is hardly biodegradable, the particles should be suited for long-term applications in animal studies. 3.1.3. Nanoparticles Composed of Different Amounts of Polystyrene and Polyisoprene. Because (unfunctionalized) polystyrene particles show a lower uptake rate than PI nanoparticles, we attempted to tune the uptake rates by the amount of polystyrene in polyisoprene/polystyrene copolymer particles.87 As can be seen in ref 87, Figure 19, there is a trend toward higher uptake when the amount of PI is increased, thereby confirming the results obtained from pure PS or PI particles. 3.1.4. PMMA Nanoparticles. For some applications, like vaccination, particles of nonbiodegradable polymers like poly(methyl methacrylate) (PMMA) are proposed.17 As this material was synthesized for another project, we investigated it in cell culture experiments. As can be seen in Figure 8, PMMA uptake kinetic is comparable to polystyrene and poly-L-lactide, therefore broadening the choice of polymers that can be used. 3.1.5. PBCA Nanoparticles. While the polymers discussed before are not biodegradable, poly(alkylcyanoacrylate) nanoparticles are biocompatible and biodegradable and can absorb or entrap bioactive compounds, making them ideal tools for applications where the nanoparticles are administered repeatedly, for example, as drug carrier systems. A large number of different compounds has been used as “payload”, for example, inorganic crystallites (magnetite91), various drugs (methotrexate,92 doxorubicin,93-95) and even oligopeptides (dalargin96,97) or proteins (insulin10,98,99). Couvreur100 used a HCl solution containing a polymeric, nonionic surfactant to which the alkylcyanoacrylate monomer is added dropwise. The described particles show a broad distribution of sizes ranging from under 100 nm to more than 1 µm. The particle size, the stability of the dispersion, and the molar masses of the polymer depend largely on the pH of the continuous phase98,101-103 and on the type and concentration of the surfactant.104,105 Despite the extensive application of the emulsion polymerization with nonionic or polymeric surfactants for the preparation of poly(alkylcyanoacrylate) nanoparticles, there are several limitations. Especially the low polymer content of the dispersions of about 1 wt % and the high amount of surfactant

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Figure 9. CLSM analysis of MSCs incubated for 4 h with (A) negative control; (B) unfunctionalized particles; (C) Phe-functionalized particles; (D) MePEG-functionalized particles. (Published with permission from ref 83. Copyright 2007 Wiley-VCH).

compared to the monomer with a ratio surfactant to monomer of 1:1 or even more (see, e.g., refs 96 and 106) should be mentioned here. Additionally, the stabilizer present in the commercially available monomer causes severe problems. Up to now modification of the PBCA particle surface is achieved by the choice of surfactant, which is physically adsorbed, like, for example, polysorbates96,97 or chemically bonded via, for example, a hydroxyl group of dextran104,107 or poly(ethylene glycol) (PEG)108 to the particles’ surface. These modifications with polysorbates allow the particles to permeate through the blood-brain barrier (see section 5), while PEGylated particles show long persistence in the circulatory system. Applying the miniemulsion technique fluorescent dye labeled unfunctionalized and functionalized PBCA nanoparticles were prepared. The molar mass distribution of the polymer is dependent on the acid used as continuous phase and the applied initiator solution (see ref 83, Figure 2). The choice of the initiator determines the particles’ surface functionalization. Unfunctionalized particles are obtained after initiation with NaOH solution, while amines, amino acids, or poly(ethylene glycol)s used as initiators lead to surface functionalized particles. For the PBCA particles, the amount of uptake was nearly linear when 0.75 to 75 µg/mL of the polymeric nanoparticles was added (see ref 83, Figure 6), thus ensuring that uptake mechanisms were not saturated. We found that the molar mass of the polymer determines the onset and extent of apoptosis (see section 4), while the total uptake does not depend on the molar mass. Different uptake kinetics are obtained with HeLa and Jurkat cells after incubation with the same particle batch (see ref 83, Figure 8). The intracellular particle distribution, visualized by confocal laser scanning microscopy, does not show significant differences for either of the cell lines or particle batches (see Figure 9 for MSC and ref 83, Figures 9 and 10). 3.1.6. Polyester Nanoparticles: Poly(ε-caprolactone), Poly(D,Llactide), Poly(D,L-lactide-co-glycolide). Besides the already mentioned poly(alkylcyanoacrylate)s (PACA) like PBCA, (bio)degradable polymers like polyesters (e.g., poly(ε-capro-


lactone), or poly(D,L-lactide)) are the first choice for sustained release application.11,99,109-113 These polyesters are not only biodegradable, they are also biocompatible and possess a low toxicity in vivo (for in vitro effects of PBCA, see section 5). Therefore, they are used for the controlled release of pharmacologically active substances.114-117 As for other nanoparticles (e.g., polystyrene) mentioned before, the monomer was emulsified and then the monomer in these nanodroplets was polymerized. In contrast, polymerization of L-lactide is hard to achieve in miniemulsions as the chemical reaction (polycondensation or ring-opening) cannot be done easily in water or in close contact with water. The process here is based on the emulsification of an organic phase containing dissolved preformed polymer within an aqueous solution containing surfactant. This also leads to monodisperse nanodroplets in the size range of 50-500 nm.76 In the second stage, the organic solvent is removed from the system, which results in the precipitation of the polymer. Additionally, the process allows the entrapment of hydrophobic components (emulsion/ solvent evaporation process). Hereby nanoparticles were prepared from a series of biodegradable polymers: poly(L-lactide) (PLLA), poly(ε-caprolactone) (PCL), and poly(D,L-lactide-coglycolide) (PLGA) with incorporated fluorescent molecules. These particles can then be used as platform particles for the encapsulation of various hydrophobic materials, for example, drugs. 3.1.6.1. Polymer Degradation. The mechanism of the aliphatic polyesters degradation is hydrolysis of backbone ester groups. Based on the physicochemical characteristics of the polymers, the most hydrophobic and crystalline PCL and PLLA nanoparticles should degrade slower than PLGA.118-123 Taking into account that the ultrasonication step (US) during the miniemulsion formation induced the polymer degradation,124 the weight average molecular weight of the final particles prepared without ultrasonication was also studied (see ref 125, Figure 3). The degradation of polymers with a high initial molecular weight is more affected by the ultrasonication compared to the polymers with shorter chain length. With time the molecular weight decreased rapidly as a result of the random hydrolytic cleavage of ester bonds. The degradation rate of the amorphous PLGA particles was slightly faster compared to the other polymers, which is due to the higher permeability of the material for water molecules. Over a period of 5 months, the molecular weights of PLLA, PCL, and PLGA nanoparticles prepared with SDS decreased by 40, 50, and 47%, respectively. The loss in molecular weight of the same particles but stabilized with Lutensol AT50 were 20, 30, and 27%, respectively {see ref 125, Figure 3). The observed results confirmed that the presence of ionic groups on the particle surface (by SDS) as ionic groups accelerate the degradation rate of the polymer due to the better water access.126 Interestingly, the particle size did not change significantly. These results are in agreement with the recent findings of Vert et al.122 3.1.6.2. Cellular Uptake of Biodegradable Nanoparticles: Flow Cytometry. For Jurkat and HeLa cells, the uptake patterns of the various particle variants were similar (see ref 125, Figure 4). The uptake of poly(L-lactide) (PLLA) and poly(εcaprolactone) (PCL) particles was hardly influenced by the molecular weight of the polymer (compare samples H (stands for high molecular weight) and L (low molecular weight)). Even so, a consistent correlation between ζ-potential or particle size and cellular uptake could not be detected. Within this series of nanoparticles the PLGA nanoparticles show a higher amount of uptake in the cells.

NPs and Stem Cells

Figure 10. CLSM images of Jurkat (A-D) and HeLa (E-H) cells. A and E are negative controls, the other images were taken after a 24 h incubation with the respective types of polymer particles stabilized with Lutensol AT50. The particles contain PMI as green fluorescent dye and the cell membranes are stained red with RH414. This dye also stains the cytoplasm of dead cells as visible in A and B. Published with permission from ref 125. Copyright 2008 Wiley-VCH.

In contrast, the surfactant located on the surface of the nanoparticles influenced the cellular uptake with the exception of PCL particle uptake in Jurkat cells. In most cases anionically stabilized SDS particle variants were taken up to a greater extent than Lutensol AT50 analogues (nonionically stabilized). This effect was most pronounced for PLLA particle uptake (see ref 125, Figure 4), whereas it was much weaker for PLGA and PCL particle uptake.6,86 3.1.6.3. Cellular Uptake of Biodegradable Nanoparticles: cLSM. Intracellular location of the nanoparticles was confirmed by confocal laser scanning microscopy (cLSM, see Figure 10). In these images of Jurkat and HeLa cells, the respective types of polymer particles stabilized with Lutensol AT50 are shown after 24 h of incubation. Images after uptake of SDS variants gave similar results. Due to the different PMI contents of the


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particles (see ref 125, Table 4), they may not be compared by fluorescence microscopy with each other. The staining patterns are similar in Jurkat and HeLa cells. It is noticeable that the uptake of poly(ε-caprolactone) (e.g., PCLH-16) and poly(D,Llactide-co-glycolide) (e.g., PLGA-18) particles results in a diffuse background staining with partly bright spots, whereas background staining is very weak after the poly(L-lactide) (e.g., PLLAL-15) particle uptake (see Figure 10 and ref 125, Figure 8). In Figure 11 TEM images of HeLa cells after a 24 h incubation showed poly(L-lactide) particles as bright spots in TEM in the vicinity of cells and intracellularly. Filopod-like membrane extensions seemed to enclose the particles. In most cases, multiple particles were localized in endosomes of varying sizes, which is comparable to SPIOs or polystyrene but in contrast to polyisoprene. Some individual particles seemed to have escaped from those compartments (arrows in Figure 11). Win and Feng127 reported the nuclear uptake of vitamin E TPGS-coated poly(D,L-lactide-co-glycolide) (PLGA) particles of ∼200 nm size into Caco-2 cells. This is contradictory to our results with PLGA and also with all other nanoparticles tested, where we never found particles located in the nucleus, neither during cLSM nor during TEM imaging. The uptake kinetics were studied for poly(L-lactide) (PLLAL15 Lut) and poly(ε-caprolactone) (PCLH-16 Lut), both formulated with the nonionic (PEG-containing) surfactant Lutensol AT50. As demonstrated in ref 125, Figure 7, the initial uptake rates for both particles are rather high during the first several hours of incubation and reach a plateau. Uptake of PCLH-16 Lut occurs faster than uptake of PLLAL-15 Lut. For comparison, we synthesized polystyrene nanoparticles by a miniemulsion copolymerization process, which were functionalized with amino groups and stabilized by a cationic surfactant (NSC particles, Dz ) 113 nm, 5760 amino groups per particle).128 Recently, the uptake of similar amino-functionalized polystyrene particles which were stabilized by nonionic Lutensol AT50 surfactant was investigated in HeLa and Jurkat cell lines under similar conditions.6,86 While the half-maximum amount for PCLH-16 Lut and PLLAH-1 Lut is reached within less than 2 h and less than 4 h, respectively, the amino-functionalized polystyrene particles showed a more steady increase of cellular uptake with no plateau within 24 h. From these preliminary data, it can again be hypothesized that different endocytotic mechanisms are involved in the uptake of the different particle materials. Already after 15 min of incubation of poly(ε-caprolactone) particles, a small but detectable amount of particles was localized inside HeLa cells, as detected by cLSM, thereby confirming the results of the FACS studies (see ref 125, Figure 8). A more diffuse staining of cytoplasm is noted after 24 h compared with a more clustered one after 2 h. This effect might be caused by distribution events like endosomal escape or by degradation of particles. 3.2. Influence of Transfection Agents: Surface Modifications of Nanoparticles by Covalently Linked Groups. 3.2.1. Functionalization of Nanoparticle Surfaces by Carboxylic and Amino Side Groups. Understanding the interactions of nanoparticles with cells is crucial for improving their interaction in vivo and in vitro. Especially the uptake into cells and the degradation in intracellular compartments is of high importance; either internalization is the aspired goal, for example, for transfection or labeling, or it should be avoided, for example, for blood pool contrast agents or nanoparticles for slow release of extracellular components. Here we focus on ways to increase the rate of intracellular uptake of polymeric particles. Since cell membranes are

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Figure 11. TEM images of HeLa cells incubated 24 h with PLLAH-1 Lut particles. Arrows indicate particles that seem to have escaped from endosomal compartments. (Published with permission from ref 125. Copyright 2008 Wiley-VCH).

negatively charged, positively charged nanoparticles were expected to be taken up even more efficiently than negatively charged nanoparticles. To this end positively charged (cationic) transfection agents129 were used by many groups.31 However, these are toxic and not approved for clinical applications. Hence, applications in human trials and therapeutic interventions cannot be performed. As a series of nanoparticles with different amounts of surface charge produced under the same conditions was not available, previous attempts to study the influence of surface charge of nanoparticles on cellular uptake were hampered. 3.2.1.1. Copolymerization of Styrene and Monomers with Charged Side Groups and Characterization of Uptake into Cells. To investigate the influence of surface charge on the uptake of nanoparticles into cells a series of fluorescent polystyrene latex particles with carboxyl and amino functionalities on their surface was synthesized by the miniemulsion technique. Also, here the fluorescent dye N-(2,6-diisopropylphenyl)perylene-3,4-dicarboximide (PMI) was incorporated as reporter into the copolymer nanoparticles formulated from styrene and acrylic acid (to obtain negatively charged surfaces containing carboxylic groups) or styrene and aminoethyl methacrylate hydrochloride (to obtain positively charged surfaces containing amino groups). This allowed the quantitative readout of particle uptake, as the amount of PMI per g of polymeric dispersion was determined. The miniemulsion allows for the specific modification of the surface by inserting functional groups, which play an important role in further applications. This also enables in a second step to couple biomolecules covalently to the nanoparticles (like amino acids peptides, proteins, DNA, etc.), which proved to be more efficient and stable than physical adsorption.130,131 The particle size depended on the amount and nature of the functional comonomer and was in the range of 100-175 nm. All latexes were characterized by transmission electron microscopy (TEM), dynamic light scattering, UV-vis spectroscopy, and ζ-potential measurements. The amount of surface functional groups was determined by electrolyte titration. Cell uptake was visualized using fluorescence microscopy. The correlation of the uptake of nanoparticles with the surface charge was determined by FACS measurements. There were slight differences observed between poly(St-coAA) and poly(St-co-AEMH) particles. First, the size of the nanoparticles was influenced by the comonomer in a relatively

narrow size range. Increasing the acrylic acid fraction gave larger particles (see ref 86, Table 2). On the contrary, with an increase in the amount of AEMH, smaller particles were obtained. Composite particles with a size range of 100-150 nm for AA as a comonomer and of 120-175 nm for AEMH were synthesized. These size variations should not alter cellular uptake as only particles much smaller (below 50 nm) or much larger (above 300 nm) are reported to be taken up in a different manner in the literature (see also section 6). Therefore, this observation should be irrelevant in the following cell experiments. Furthermore, we were interested in differences in the interaction of submicrometer particles with a variety of cell lines, as these are important for in vitro and in vivo applications. Altered uptake behavior in nonphagocytotic HeLa, Jurkat and mesenchymal stem cells was observed after surface modification of polystyrene nanoparticles with carboxy or amino groups. For the cell types studied, with the exception of MSCs, a clear correlation of surface charge and fluorescence intensity could be shown (see Figure 12 and see ref 6, Figures 1, 2, and 8). By using surface functionalized polymeric nanoparticles with a range of carboxyl group densities, a density-uptake relationship was established in addition. We showed that for an efficient uptake of the nanoparticles a negatively charged surface was sufficient and no transfection agents were needed also comparing them to uncharged particles into HeLa cells and MSCs (see ref 6, Figure 1). The density of the carboxyl groups showed an optimum for particles around 0.5 COOH groups per nm2 and a decrease in uptake if the density of the carboxyl groups was increased (see ref 86, Figure 10). It was demonstrated that intracellular uptake is not only dependent on the charge of the polymeric model nanoparticle, but also on the type of target cell. We showed that for MSCs, the presence of carboxyl groups increases the cellular uptake in comparison to a nonfunctionalized particle. The density of carboxyl groups is less relevant for MSCs as compared to HeLa cells, and a small quantity is already sufficient to induce cell uptake (see ref 47, Figure 6). This explains why we detected differences in the uptake of Resovist made of carboxydextran and Feridex, as Feridex is made of dextran with citric acid (see section 2.2). This addition of citric acid may account for a few carboxyl group on the surface of the Feridex particles that may be sufficient for intracellular uptake in MSCs, but may result in inferior uptake rates in HeLa cells as a higher density of carboxyl groups is needed for an effective intracellular uptake in HeLa cells. Moreover, Feridex did not yield high amounts of iron oxide in

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Figure 12. Left: FACS measurements of HeLa cells incubated for 24 h with fluorescent poly(St-co-AA) nanoparticles. Each bar shows the average normalized fluorescence intensity (nFL). Each latex was tested in triplicate. Right: FACS measurements of HeLa cells incubated for 24 h with fluorescent amino functionalized nanoparticles. Each latex was tested in triplicate. (Published with permission from ref 86. Copyright 2005 Wiley-VCH).

Figure 13. (A,B) TEM analysis of MSCs. (A) Incubation for 24 h with particle HF3. The particles (arrows) can be found in groups in intracellular compartments that resemble endosomes. (B) MSCs incubated with HF5 for 24 h. Particles are attached to the cell membrane (black arrows). Electron dense material can be seen between the attached particles (white arrowheads). (C,D) cLSM of KG1a cells. (C) HF4 and (D) HF5 (green channel), where the green signal is located in clusters at the cell membrane (arrow). (Reprinted with permission from Elsevier. Lorenz, M.R.; Holzapfel, V.; Musyanovych, A.; Nothelfer, K.; Walther, P.; Frank, H.; Landfester, K.; Schrezenmeier, H.; Maila¨nder, V. Uptake of functionalized, fluorescent-labeled polymeric particles in different cell lines and stem cells. Biomaterials 2006, 27, 2820-2828. Copyright 2006 (http://www.sciencedirect.com/science/journal/01429612).

HeLa cells, and the uptake of the experimental nanoparticles was also less pronounced in HeLa cells. 3.2.1.2. Comparison of Amino and Carboxy Functionalized Nanoparticles. To compare different experiments, an unfunctionalized nanoparticle (termed VHPM1 or P1) was synthesized. By normalizing the uptake of this nanoparticle, we were able to compare different experiments. For HeLa cells, for example, we found that, while carboxy functionalized PS nanoparticles of a size of 100 nm only show a slight increase of total particle uptake after 24 h (about 6× better than P1 (see ref 47, Figure 6)) compared to unfunctionalized PS particles of the same size, amino functionalized particles show a more than 40-fold increase in uptake (see ref 47, Figure 1). The different uptake behaviors for surface modified nanoparticles (either covalently attached or physically absorbed moieties) can be explained for positively charged nanoparticles at least in part by the fact that the cell membrane itself is negatively charged. Therefore, positively charged nanoparticles should adhere to cell membranes easily. Why negatively charged nanoparticles show also an enhanced uptake is not so easily explained, but here interactions with surface molecules, for example, proteins, may be responsible for adhesion and uptake. Also, phosphatidylserine in the cell membrane has been linked to fusion events of cationic agents with cell membranes.132

3.2.1.3. Subcellular Distribution of Charged Nanoparticles: TEM, cLSM. Confocal laser scanning microscopy (cLSM) and transmission electron microscopy (TEM) revealed differences in subcellular localization of the nanoparticles. In MSCs and HeLas, nanoparticles were mostly located inside of cellular compartments resembling endosomes (Figure 13A, B), while cytoplasmatic localization was not observed. Nanoparticles were also not detected in mitochondria, the cell nucleus or the Golgi apparatus. On the other hand, in Jurkat and KG1a, nanoparticles were predominantly located in clusters on or near the cell surface, as shown by cLSM studies (Figure 13C,D). Scanning electron microscopy showed microvilli to be involved in this process of adherence of nanoparticles to KG1a and Jurkats (see ref 6, Figure 10). The attachment of the particles to the cell membrane as the first step seems to be mostly affected by the surface charge of the particles, while the differences in the intracellular localization between various cell lines can be explained by different endocytotic/pinocytotic properties of the cell lines HeLa, MSC, KG1a, and Jurkat. Even by cLSM an increase in uptake with increasing surface charges was detected. While in carboxylic functionalized nanoparticles no aggregation was observed, a tendency to aggregation in the medium and on the cell surface was

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observed with very high amounts of amino groups (see ref 6, Figure 3D, E). 3.2.2. Functionalization of PBCA Nanoparticles. While with other routes of PBCA synthesis surface functionalization is hard to achieve, by applying the miniemulsion technique it was possible to prepare stable dispersions of fluorescence dye labeled PBCA nanoparticles of a size smaller than 250 nm with different surface functionalities. This is done by using the functional group as the starter of the polymer chain. For example, phenylalanine can be used for functionalization without using an EDC coupling reaction as done for coupling lysine to the polystyrene nanoparticles (see section 2.3). It could be shown that phenylalanine particles are internalized by HeLa, Jurkat, and mesenchymal stem cells effectively (see ref 83, Figures 8-11). Cellular uptake kinetics for HeLa and Jurkat cells are shown in ref 83 Figure 8. This means that the cellular uptake kinetics and the targeting abilities of PBCA nanoparticles can be influenced by basic modifications of the particles’ surface properties and size without applying further steps during or after the nanoparticle preparation. All modifications can easily be accomplished by the application of the miniemulsion technique. 3.2.2.1. Functionalization of PBCA Nanoparticles with PEG. Despite the protection of the drug in nanoparticles, not all of it reaches the site where it is required. Several reasons can interfere with the effective delivery of the drug to the desired destination when the nanocarrier is injected into an animal or human being. The drug may loose its protection due to a rapid degradation process of the carrier system, the carrier system shows unspecific affinity to any type of tissue, the particles might be opsonized by absorption of serum proteins and phagocytosed by macrophages, or they may be simply excreted from the organism. Targeting or avoidance of specific cell types on one hand and simultaneous protection of the particles on the other hand should overcome these obstacles. The most common approach to protect particles from opsonization is the formation of a hydrophilic corona, e.g. from PEG or carbohydrates, around the particles.133-138 This shell minimizes unspecific adsorption of proteins on the particle surface, which renders the particles “invisible” for macrophages (“stealth” effect). Therefore, the particles remain unaffected in the circulatory system, thereby increasing the chance that the site of disease is reached by the particles and as, for example, metastases of a tumor show an enhanced permeability with a decreased rate of clearance, more nanoparticles are trapped at the site of the disease (“passive targeting”).139-141 For PBCA nanoparticles, we were able to show that uptake into Jurkat cells can be minimized by covalently bonding PEG on the surface of such nanoparticles. Interestingly, this was not observed for all cell types, that is, we did not find any effect for HeLa cells (see ref 83, Figure 8, last row). This indicates that surface modification by PEG is important for modulation of uptake into Jurkat and that the effect of PEG functionalization may depend on the cell type. As mentioned before, PEG is known to reduce phagocytosis of polymeric particles due to altered adsorption of proteins, which are responsible for cellular recognition and uptake processes.

4. Toxicity of the Nanoparticles Measured by 7-AAD As it was only a side aspect of this work and to give a rough estimation of the cytotoxicity of these nanoparticles, the studied cells were incubated for 4-24 h with all nanoparticles and


Figure 14. Toxicity of the nanoparticles for Jurkat cells. (Published with permission from ref 87. Copyright 2008 Wiley-VCH).

cytotoxicity was evaluated by 7-AAD. Untreated cells served as negative control. We chose 7-AAD as it can distinguish between live, apoptotic, and dead cells. FACS analysis was performed to measure the percentage of live, apoptotic, and dead cells (for example, see Figure 14). As we had used SPIOs (Feridex and Resovist) in 10× higher concentrations than was given in the literature, intensive cytotoxicity studies were done. Even these concentrations of SPIOs showed no cytotoxic effect (see ref 47, Supporting Information, Figure 2). For polystyrene nanoparticles, independent of their surface modification also no cytotoxicity was observed when cells were incubated for 24 h with these nanoparticles. Also, none of the particles composed of poly(L-lactide) (PLLA), poly(ε-caprolactone) (PCL), and poly(D,L-lactide-coglycolide) (PLGA) were cytotoxic, as quantified via 7-AADstaining, which showed HeLa cell vitalities of more than 98% (see ref 125 and unpublished data). For PBCA, the degradation products are n-butanol and poly(cyanoacrylicacid)andtheyareknowntobecytotoxic.110,142,143 Furthermore, the liberation rate of the degradation products can be expected to be dependent on the chain length of the polymer, because a shorter chain of partially hydrolyzed PBCA is more hydrophilic in total and more soluble in the aqueous medium than a longer one and can therefore be degraded more easily. PBCA nanoparticles containing polymers with different chain lengths were synthesized and we showed that cytotoxicity in vitro is dependent on the molar mass distribution of the polymer (see ref 83, Figure 7). Although we did not evaluate toxic effects systematically in all organs, we did not observe any disruption in brain histology during our in vivo experiments with PBCA nanoparticles (see section 5). Another source of cytotoxicity besides the polymer or its degradation product could be the surfactant as they can influence cell permeability. Interestingly no cytotoxicity was observed for the SDS stabilized PI nanoparticles for HeLa and Jurkats. Therefore, we evaluated with PI nanoparticles the influence of other surfactants. Two cationic surfactants were used: cetyl pyridium chloride and hexadecyl trimethyl ammonium-4-toluene sulfonate. At low concentrations of these surfactants (see Figure 14 mkpi075, respectively, mkpi076), no significant increase or only a slight increase in cytotoxicity was observed.

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Figure 15. Cryosections of the rat brains. The left image (transmitted) shows the optical transmission image of the section (40×); nano: fluorescence created by the PMI-labeled nanoparticles; Willebrand: endothelial cells stained with fluorescent antibody (von Willebrand factor primary and anti-IgG secondary antibody with fluorescent label); Merge 40× and 63×: image merged from the green and red channel. The scale bars represent 100 µm. (Published with permission from ref 150. Copyright 2008 Wiley-VCH).

Five times higher concentrations of these surfactants resulted in a high cytotoxicity rate (see Figure 14, mkpi072, respectively, mkpi073) with high numbers in the apoptotic and dead cell populations. The results for HeLa cells were similar, but the increase of cell toxicity showed up only in the apoptotic cell region.

5. Crossing the Blood-Brain Barrier Even after 30 years of intense research, the transport of drugs to the central nervous system (CNS) is a major challenge in pharmaceutical science. On their way from the bloodstream into the brain, most of the drugs encounter a nearly insurmountable obstacle, the blood-brain barrier (BBB). The BBB is formed by a dense layer of endothelial cells facing the blood flow linked by tight junctions. A basal membrane is interposed and on the other side the endothelial cells are lined by astrocytes and neurons. Small hydrophilic compounds with a molar mass lower than 150 g · mol-1 and hydrophobic compounds with a mass lower than 400 g · mol-1 can pass the cellular barrier by passive diffusion and thereby enter the brain. Compounds that do not meet these requirements are excluded from this path and remain in the bloodstream. Substances internalized by the endothelial cells are effectively removed from the cells by the p-glycoprotein efflux pump system if they are not recognized as necessary for the brain. For specific substances or systems essential for the brain metabolism, like amino acids, glucose, or low density lipoprotein (LDL) particles, specific receptors on the endothelial cells allow the barrier to be permeated.144 A more general approach proposes nanoparticulate (colloidal) systems like lipid or polymeric nanoparticles, liposomes, or micelles as drug carriers.110,145-147 Regarding polymeric nanoparticles, the surface characteristics, presumably in combination with the polymeric matrix, are responsible for the passage to the brain. Particularly PBCA based particles coated with polysorbates like Tween 80 or poloxamers like Pluronic F68 have been reported to successfully pass the BBB.148,149 The (bio)degradability of PBCA in the organism makes this polymeric matrix perfectly suitable for sustained release applications.

When fluorescent polysorbate 80 coated PBCA nanoparticles were used in an in vivo study, direct evidence for the presence of nanoparticles entering the brain and the retina of rats was found. The nanoparticles, prepared with a miniemulsion process, were in situ labeled with a fluorescent dye and coated with polysorbate 80. Human brain microvascular endothelial cells (BMEC) cells were used as in vitro model for the BBB. In the in vitro model (transwell system) the cells showed significant uptake of the particles (see ref 150, Figure 3), but no transcytosis could be observed in vitro (see ref 150, Figure 4). After the application of the particles in two concentrations to the animals, cryosections of the brains and retinas were prepared. Regarding the sections of the rat receiving the lower dose, colocalization of the applied fluorescent particles and the stained endothelial cells could be detected in the brain and the retina as well, indicating particle internalization in the endothelial cells (see Figure 15 and ref 150, Figure 7). On contrast in the higher dosage, the particles could be detected within the brain and the retina, with few colocalized signals, suggesting a passage through the blood-brain and blood-retina barrier. This difference between in vitro and in vivo data may be explained by the fact that BMEC in the transwell system may have changed their behavior due to the immortalization or that the cells have not yet established their ability for transcytosis of nanoparticles as this is a directed process for which further signals, for example, from the endoglial cells or the basal membrane may be important. Both other components of the BBB were not included in the present model. Like found by other groups, results from transwell experiments are not always in line with in vivo data. In vitro analysis may require a more complex BBB model and this points to the necessity to investigate questions concerning the BBB by in vivo models as the more relevant system.

6. Endocytosis As has been shown before, nanoparticles are taken up by a variety of cells. One of the pressing questions in the field is the route of uptake of these nanoparticles into cells. Insights into

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Figure 16. Mechanisms of endocytosis. The three main pathways are macropinocytosis, lipid raft-dependent mechanisms, and the clathrindependent pathway. Factors like actin, dynamin and cholesterol participate in different uptake mechanisms. Reprinted with permission from Elsevier: Kirkham, M.; Parton, R. G. Clathrin-independent endocytosis: New insights into caveolae and noncaveolar lipid raft carriers. Biochimica et Biophysica Acta 2005, 1745, 273-286. Copyright 2005 (http:// www.sciencedirect.com/science/journal/01674889).

the routes of uptake should allow us to manipulate the uptake of these nanomaterials into cells and may even enable targeting of the cargo to specific subcellular compartments. For small molecules and macromolecules like proteins, routes of uptake into cells have been described.151,152 Particles in the range of a few to several hundred nanometers are much larger and conditions and mechanisms of uptake involving several possible mechanisms like pinocytosis, nonspecific endocytosis, receptor-mediated endocytosis, and, for larger particles, phagocytosis have not been studied thoroughly.153 The exact mechanism of receptor-mediated endocytosis, for instance, is not completely understood, although some receptors for receptormediated endocytosis like opsonin, lectin, and scavenger receptors are known.154 Mechanisms of endocytosis have been studied extensively for biologically active molecules and also for viruses.155-158 The first route to be characterized were clathrin-coated pits, a specialized protein assembly of the triskelion clathrin forming a net-like basket.159 A second class of vesicles is coated with the protein caveolin and is therefore called caveolae. They are derived from lipid rafts.160 Lipid rafts are rigid membrane microdomains enriched with phospholipids, sphingolipids, and cholesterol.161 Uncoated vesicles can also be formed from these lipid rafts or by macropinocytosis and they can be bigger than coated ones and therefore may allow endocytosis of larger particles (>150 nm). Different receptors are known to be involved in endocytosis, some of them concentrating in clathrincoated pits or lipid raft regions. An overview of the endocytotic mechanisms is given in Figure 16, for a review see Kirkham et al.162 Model substances and inhibitors have been used to characterize routes of uptake,7,163 but many of the components like various receptors, dynamin, or cytoskeletal components are crucial in different pathways.162 Therefore, endocytotic pathways should be thought of as a network of different properties that can be combined with each other. For example, the small GTPase dynamin that pinches off the endocytotic sack from the cell membrane is essential during the internalization of clathrin or caveolin-coated vesicles, and therefore, inhibition of dynamin will affect both pathways (see Figure 16). On the other hand, even dynamin knockout mice survive because alternative molecular machinery provides ways of endocytosis.164 Furthermore, different cell types may have


different uptake mechanisms and may upregulate less prominent ones upon blocking of their standard pathway. As has been shown previously, surface charge is an important factor for endocytotic uptake into different cell lines (see section 3.2.1). Therefore, we expected that also uptake mechanisms may differ depending on the surface properties. In our studies, we evaluated which endocytotic mechanisms are involved in the uptake of well-defined positively and negatively charged 100 nm fluorescent polystyrene nanoparticles into HeLa cells. We demonstrated that, independent of the particle charge (cationic or anionic), endocytosis is energy dependent and highly reliant on dynamin and F-actin, as shown by inhibition of uptake in the presence of dynasore and cytochalasin D, respectively (see Figure 17). Because genistein inhibited the particle uptake, tyrosine specific protein kinases, located in lipid rafts, have to be involved in the uptake (see ref 85, Figure 8). These factors suggest a dynamin- and lipid raft-dependent uptake mechanism. However, a cholesterol sequestration or depletion by filipin or methyl-β-cyclodextrine, respectively, did not hinder the particle uptake (see ref 85, Figure 8). The particle surface charge impacts further uptake mechanisms: Macropinocytosis seems to be an important mechanism for positively charged nanoparticles as is demonstrated by the strong inhibition of positively charged nanoparticles by 5-(Nethyl-N-isopropyl)amiloride (EIPA), as shown in ref 85, Figure 9. The microtubule network and cyclooxygenases are also involved in uptake of positively charged particles as uptake was hindered in the presence of nocodazole and indomethacin, respectively (Figure 17, left panel and in ref 85, Figures 5 and 8). Clathrincoated pits only play a minor role in the uptake of positively charged nanoparticles, having no effect in endocytosis of negatively charged ones. About 20% of the endocytosis of positively charged particles is inhibited by chlorpromazine (see ref 85, Figure 6). This is plausible in the light of the TEM studies where several nanoparticles seemed to be engulfed by a structure (Figure 17, right) that would be most easily identified as macropinocytic and taking into account that the rigid clathrin structure cannot be dilated significantly.7 The question about the role of lipid rafts could not be unambiguously answered by these studies as the uptake was largely independent of cholesterol depletion/sequestration which should affect lipid rafts very effectively while inhibition of lipid raft-associated proteins showed a significant effect. Therefore, further studies (e.g., with CAV-1 antibodies) are warranted. It can be concluded that, in dependence on the surface charge of the nanoparticle, differences in uptake and intracellular trafficking of the endosomes may occur. Interestingly, negatively charged nanoparticles were less inhibited by a dynamin inhibitor pointing toward the possibility that a hitherto unidentified dynamin-independent process may contribute to the uptake of negatively charged nanoparticles.

7. Clinical Relevance and Applications of Nanoparticles Polymeric nanoparticles composed of dextran and an iron oxide core were the first to be used for diagnostic purposes in vivo in humans.28 Their application in MRI as a contrast agent exploited the superparamangnetic effect. This effect is a good example how materials (here iron oxide) in the nanometer range show hitherto unknown properties. Uptake by liver macrophages (Kupffer cells) is the main mechanism by which SPIOs act as contrast agents. In further investigations it became clear that the unspecific uptake of materials in the nanometer range165

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Figure 17. Left: Uptake of PS+ or PS- particles into HeLa cells under different inhibitory conditions. HeLa cells were preincubated for 30 min with the respective drugs, and particle uptake within 1 h was determined by flow cytometry. Mean values and standard deviations of two independent triplicate experiments are given. After incubation with filipin or MBCD, the fluorescence signal was never significantly lower than the respective positive control. Right: HeLa cells were incubated with positively charged polystyrene nanoparticles. HeLa cell during endocytosis of nanoparticles (Published with permission from ref 85. Copyright 2008 Wiley-VCH).

Figure 18. Scheme of possible applications of nanoparticles and nanocapsules.

hinders the further development of these nanoparticles when the liver or spleen should not be the target. Here the development of pegylated nanoparticles offers advantages (see ref 133 and also section 3.2.2.1) as they remain in the bloodstream for longer times. This gives the chance that the nanoparticles can reach other target organs in the body like a tumor139 or even enter the brain.140 Still the specific targeting of the nanoparticles is challenging as, for example, antibodies are more or less specific for a particular organ, but to reach the antigen on the cells they would not need to leave the bloodstream and to encounter the target antigens. This is usually prevented by the endothelial cells. The leakiness of the endothelial layers in tumors has been utilized as an unspecific targeting mechanism (“enhanced permeability and retention” effect, EPR139). The further development of nanoparticles as diagnostic or therapeutic tools depends on fulfilling the promise of specific delivery of nanoparticles inside the body. Only then applications as diagnostic (like iron oxide nanoparticles or gadolinium nanoparticles) or therapeutic tools (e.g., drug loaded nanoparticles) can be implemented in preclinical and clinical trials. Here the synthesis of nanoparticles by the miniemulsion process offers

great advantages as a wide variety of nanoparticles can be produced by this platform technology (see sections above). Surface modifications like altering the charge, attached proteins will play an important role here, but also the polymeric substance itself seems to have an influence on the field of application (see PBCA nanoparticles in section 5). The clinical application will also determine whether the nanoparticles should be biodegradable or not, like the polystyrene or polyisoprene nanoparticles, and also the time frame during which they should decompose can be adjusted by the use of the polymeric substance (see section 3.1.6). This is highly important for drug delivery, a broad field of application for nanoparticulate systems. Hyperthermia by using iron oxide nanoparticles is an interesting new therapeutic option but needs to be optimized so that only the tumor is specifically targeted by the nanoparticles. But also using nanoparticles as diagnostic tools like iron oxide nanoparticles or gadolinium containing nanocapsules as combination of marker substance and specific targeting agent like an antibody, a protein or an aptamer will provide new tools for detection of pathological processes.

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Further modifications like using pH reactive or enzyme degradable polymers will make the nanoparticles even more versatile and will improve the deposition in the desired target region of the body, like a tissue or tumor that (over)expresses a specific enzyme/protein or has a lower pH.

8. Summary Starting from experiments with commercially available superparamagnetic iron oxide nanoparticles intended here for labeling of (stem) cell preparations we stumbled on questions of how these nanomaterials interact with cells, that is, what are the crucial parameters on the side of the nanoparticles and the cells so that these are endocytosed. We investigated how the polymeric material influences cellular uptake. We were, for example, able to show that uptake kinetics is much faster for polyisoprene nanoparticles as compared to polystyrene. Nanoparticles can be made from a range of polymers used for technical applications as well as biocompatible and biodegradable ones. When two monomers are copolymerized (like polyisoprene and polystyrene), cell uptake can be tuned by changing the relative amount of the monomers. Also, modifications like a positive or negative charge on the surface of the nanoparticles enhances cellular uptake, we showed that this effect can be “titrated” by using a series of nanoparticles with a range of densities of effective side groups. Cellular uptake can be enhanced by transfection agents. Furthermore, uptake can also decreased by PEG if uptake is not favored for an application. To detect the nanoparticles different reporters were encapsulated like superparamagnetic iron oxide and gadolinium for MRI applications and fluorescence for in vitro studies. Also, more than one of these reporters can be included in these nanoparticles, thereby yielding dual-reporter nanoparticles. Subcellular distribution as studied by TEM found the nanoparticles mostly in endosomes. Nanoparticles were not found inside the cell nucleus. Therefore, any application like gene delivery would have to ensure that the DNA is released from the type of nanoparticles studied here to get into the cell nucleus. With all this instrumentarium of nanoparticles at hand, we were investigating that nanoparticles can be useful for crossing physiological barriers like the blood-brain barrier and what are the mechanisms of cellular endocytosis involved in uptake of nanoparticles. Concerning the cytotoxicity of nanoparticles we demonstrated that cytotoxicity can depend on the polymeric material like in PBCA but also on the type of surfactant. Performing enzymatic reaction like PCR in miniemulsion droplets is a very promising approach for designing nanocompartments that can perform more interesting tasks than can be done by the other nanoparticles mentioned (for illustration see Figure 18).

9. Outlook Future directions for this work will be in the field of “smart” nanoparticles, that is, the particles exert an effect, for example, as nanoreactors or they react to their surroundings as, for example, they only light up in a certain environment. This could be achieved by nanoparticles with pH dependent polymers or dyes or enzymatic degradation of nanocapsules therefore releasing their payload at specific sites. Using the synthesized nanoparticles for in vivo applications was not a goal of this work but is a long-term goal. For this purpose optimized nanoparticles will be used in animal models, for example, lysine


modified superparamagnetic iron oxide nanoparticles for detection in MRI scanners. The inclusion of other reporters will be investigated in future projects, for example, radionuclides, quenched fluorochromes, or bioluminescence (e.g., with luciferase). Also using the nanoparticles for in vivo imaging in an animal MRI scanner will be the next step for the superparamagnetic iron oxide nanoparticles. The possibilities of delivering drugs and gene therapeutics will be explored. For this purpose, we need to learn more about the intracellular trafficking and fate of these nanoparticles after the endocytotic process. Also, more studies will be done to evaluate toxicity of nanoparticles on cells, for example, by annexin, caspase activation. Also, the influence, for example, on release of cytokines and growth factors and metabolic functions will be evaluated. With all these nanoparticles at hand more in vivo experiments will be performed to determine organ distribution and in vivo effects of nanoparticles. Especially detecting homing and trafficking of (stem) cell preparations will be a major field of investigation in the future but also delivering, for example, cytotoxic drugs in vitro and in vivo. Here first experiments with paclitaxel and doxorubicin in nanoparticles and nanocapsules have been done. Acknowledgment. This work could not have been done without the support, feedback, and guidance of Prof. Hubert Schrezenmeier and Dr. Markus Wiesneth.

References and Notes (1) Jun, Y. W.; Jang, J. T.; Cheon, J. AdV. Exp. Med. Biol. 2007, 620, 85–106. (2) Landfester, K. Annu. ReV. Mater. Res. 2006, 36, 231–279. (3) Narayanan, R.; El-Sayed, M. A. J. Phys. Chem. B 2005, 109, 4357– 60. (4) Fallon, M. S.; Varshney, M.; Dennis, D. M.; Chauhan, A. J. Pharmacokinet. Pharmacodyn. 2004, 31, 381–400. (5) Miltenyi, S. Oncologist 1997, 2, 410–413. (6) Lorenz, M. R.; Holzapfel, V.; Musyanovych, A.; Nothelfer, K.; Walther, P.; Frank, H.; Landfester, K.; Schrezenmeier, H.; Mailander, V. Biomaterials 2006, 27, 2820–8. (7) Rejman, J.; Oberle, V.; Zuhorn, I. S.; Hoekstra, D. Biochem. J. 2004, 377, 159–169. (8) Bender, A. R.; von Briesen, H.; Kreuter, J.; Duncan, I. B.; Ru¨bsamenWaigmann, H. Antimicrob. Agents Chemother. 1996, 40, 1467–1471. (9) Wang, G.; Uludag, H. Expert Opin. Drug DeliVery 2008, 5, 499– 515. (10) Sullivan, C. O.; Birkinshaw, C. Biomaterials 2004, 25, 4375–4382. (11) Jiao, Y.; Ubrich, N.; Marchand-Arvier, M.; Vigneron, C.; Hoffman, M.; Lecompte, T.; Maincent, P. Circulation 2002, 105, 230–235. (12) Alvarez-Roman, R.; Naik, A.; Kalia, Y. N.; Guy, R. H.; Fessi, H. Pharm. Res. 2004, 21, 1818–25. (13) Alvarez-Roman, R.; Naik, A.; Kalia, Y. N.; Guy, R. H.; Fessi, H. J. Controlled Release 2004, 99, 53–62. (14) Kourti, M.; Vavatsi, N.; Gombakis, N.; Sidi, V.; Tzimagiorgis, G.; Papageorgiou, T.; Koliouskas, D.; Athanassiadou, F. Int. J. Hematol. 2007, 86, 166–73. (15) Derouet, D.; Mulder-Houdayer, S.; Brosse, J.-C. J. Appl. Polym. Sci. 2005, 95, 39–52. (16) Phinyocheep, P.; Phetphaisit, C. W.; Derouet, D.; Campistron, I.; Brosse, J. C. J. Appl. Polym. Sci. 2005, 95, 6–15. (17) Kreuter, J. J. Controlled Release 1991, 16, 169–176. (18) Zimmet, J.; Hare, J. Basic Res. Cardiol. 2005, 100, 471–81. (19) Kemp, K.; Hows, J.; Donaldson, C. Leuk. Lymphoma 2005, 46, 1531– 44. (20) Granero-Molto, F.; Weis, J. A.; Longobardi, L.; Spagnoli, A. Expert Opin. Biol. Ther. 2008, 8, 255–68. (21) Thorne, S. H.; Negrin, R. S.; Contag, C. H. Science 2006, 311, 1780– 4. (22) Contag, C. H. Neuroimaging Clin. N. Am. 2006, 16, 633–54. (23) Hofmann, M.; Wollert, K. C.; Meyer, G. P.; Menke, A.; Arseniev, L.; Hertenstein, B.; Ganser, A.; Knapp, W. H.; Drexler, H. Circulation. 2005, 111, 2198–202, Epub 2005 Apr 25.

NPs and Stem Cells (24) Michalet, X.; Pinaud, F. F.; Bentolila, L. A.; Tsay, J. M.; Doose, S.; Li, J. J.; Sundaresan, G.; Wu, A. M.; Gambhir, S. S.; Weiss, S. Science 2005, 307, 538–44. (25) Scarff, M.; Arnold, S. A.; Harvey, L. M.; McNeil, B. Crit. ReV. Biotechnol. 2006, 26, 17–39. (26) Frank, J. A.; Zywicke, H.; Jordan, E. K.; Mitchell, J.; Lewis, B. K.; Miller, B.; Bryant, L. H., Jr.; Bulte, J. W. Acad. Radiol. 2002, 9, S484-7. (27) Ohgushi, M.; Nagayama, K.; Wada, A. J. Magn. Reson. 1978, 29, 599–601. (28) Stark, D. D.; Weissleder, R.; Elizondo, G.; Hahn, P. F.; Saini, S.; Todd, L. E.; Wittenberg, J.; Ferrucci, J. T. Radiology 1988, 168, 297–301. (29) Weissleder, R.; Elizondo, G.; Wittenberg, J.; Lee, A. S.; Josephson, L.; Brady, T. J. Radiology 1990, 175, 494–8. (30) Kraitchman, D. L.; Heldman, A. W.; Atalar, E.; Amado, L. C.; Martin, B. J.; Pittenger, M. F.; Hare, J. M.; Bulte, J. W. Circulation 2003, 107, 2290–3, Epub 2003 May 5. (31) Arbab, A. S.; Bashaw, L. A.; Miller, B. R.; Jordan, E. K.; Bulte, J. W.; Frank, J. A. Transplantation 2003, 76, 1123–30. (32) Wunderbaldinger, P.; Josephson, L.; Weissleder, R. Acad. Radiol. 2002, 9, S304-6. (33) Lewin, M.; Carlesso, N.; Tung, C. H.; Tang, X. W.; Cory, D.; Scadden, D. T.; Weissleder, R. Nat. Biotechnol. 2000, 18, 410–4. (34) Cohen, H.; Levy, R. J.; Gao, J.; Fishbein, I.; Kousaev, V.; Sosnowski, S.; Slomkowski, S.; Golomb, G. Gene Ther. 2000, 7, 1896–905. (35) Panyam, J.; Labhasetwar, V. AdV. Drug DeliVery ReV. 2003, 55, 329– 47. (36) Gupta, A. K.; Curtis, A. S. J. Mater. Sci.: Mater. Med. 2004, 15, 493–6. (37) Chawla, J. S.; Amiji, M. M. Int. J. Pharm. 2002, 249, 127–38. (38) Elamanchili, P.; Diwan, M.; Cao, M.; Samuel, J. Vaccine 2004, 22, 2406–12. (39) Russell-Jones, G. J. J. Controlled Release 2000, 65, 49–54. (40) Damge, C.; Michel, C.; Aprahamian, M.; Couvreur, P. Diabetes 1988, 37, 246–251. (41) Montet-Abou, K.; Montet, X.; Weissleder, R.; Josephson, L. Mol. Imaging 2005, 4, 165–71. (42) Arbab, A. S.; Yocum, G. T.; Wilson, L. B.; Parwana, A.; Jordan, E. K.; Kalish, H.; Frank, J. A. Mol. Imaging 2004, 3, 24–32. (43) Arbab, A. S.; Yocum, G. T.; Kalish, H.; Jordan, E. K.; Anderson, S. A.; Khakoo, A. Y.; Read, E. J.; Frank, J. A. Blood 2004, 104, 1217–23, Epub 2004 Apr 20. (44) Ju, S.; Teng, G.; Zhang, Y.; Ma, M.; Chen, F.; Ni, Y. Magn. Reson. Imaging 2006, 24, 611–7, Epub 2006 Feb 17. (45) Hunter, A. C. AdV. Drug DeliVery ReV. 2006, 58, 1523–31. (46) Symonds, P.; Murray, J. C.; Hunter, A. C.; Debska, G.; Szewczyk, A.; Moghimi, S. M. FEBS Lett. 2005, 579, 6191–8. (47) Mailander, V.; Lorenz, M. R.; Holzapfel, V.; Musyanovych, A.; Fuchs, K.; Wiesneth, M.; Walther, P.; Landfester, K.; Schrezenmeier, H. Mol. Imaging Biol. 2008, 10, 138–46. (48) Heyn, C.; Bowen, C. V.; Rutt, B. K.; Foster, P. J. Magn. Reson. Med. 2005, 53, 312–20. (49) Dahnke, H.; Schaeffter, T. Magn. Reson. Med. 2005, 53, 1202–6. (50) Sun, R.; Dittrich, J.; Le-Huu, M.; Mueller, M. M.; Bedke, J.; Kartenbeck, J.; Lehmann, W. D.; Krueger, R.; Bock, M.; Huss, R.; Seliger, C.; Grone, H. J.; Misselwitz, B.; Semmler, W.; Kiessling, F. InVest. Radiol. 2005, 40, 504–13. (51) Metz, S.; Bonaterra, G.; Rudelius, M.; Settles, M.; Rummeny, E. J.; Daldrup-Link, H. E. Eur. Radiol. 2004, 14, 1851–8, Epub 2004 Jul 13. (52) Leuschner, C.; Kumar, C. S.; Hansel, W.; Soboyejo, W.; Zhou, J.; Hormes, J. Breast Cancer Res. Treat. 2006, 99, 163–76, Epub 2006 Jun 3. (53) Warren, R. A.; Green, F. A.; Enns, C. A. J. Biol. Chem. 1997, 272, 2116–21. (54) Eisenstein, M. Nature 2006, 441, 1179–85. (55) Fattal, E.; Couvreur, P.; Dubernet, C. AdV. Drug DeliVery ReV. 2004, 56, 931–46. (56) Freulon, I.; Roche, A. C.; Monsigny, M.; Mayer, R. Biochem. J. 2001, 354, 671–9. (57) Landfester, K.; Ramirez, L. J. Phys.: Condens. Matter 2003, 204, 22–31. (58) Ramirez, L.; Landfester, K. Macromol. Chem. Phys. 2003, 206, 2440– 2449. (59) Hamoir, J.; Nemmar, A.; Halloy, D.; Wirth, D.; Vincke, G.; Vanderplasschen, A.; Nemery, B.; Gustin, P. Toxicol. Appl. Pharmacol. 2003, 190, 278–85.


2399

(60) Olivier, V.; Duval, J. L.; Hindie, M.; Pouletaut, P.; Nagel, M. D. Cell Biol. Toxicol. 2003, 19, 145–59. (61) Horowitz, S. M.; Gautsch, T. L.; Frondoza, C. G.; Riley, L., Jr. J. Orthop. Res 1991, 9, 406–13. (62) Holzapel, V.; Lorenz, M.; Weiss, C. K.; Schrezenmeier, H.; Landfester, K.; Maila¨nder, V. J. Phys.: Condens. Matter 2006, 18, 2581– 2594. (63) Liu, X.; Guan, Y.; Ma, Z.; Liu, H. Langmuir 2004, 20, 10278–82. (64) Qian, Z.; Zhang, Z.; Chen, Y. J. Colloid Interface Sci. 2008, 327, 354–61. (65) Hua, X. F.; Liu, T. C.; Cao, Y. C.; Liu, B.; Wang, H. Q.; Wang, J. H.; Huang, Z. L.; Zhao, Y. D. Anal. Bioanal. Chem. 2006, 386, 1665–71. (66) Frank, J. A.; Anderson, S. A.; Kalsih, H.; Jordon, E. K.; Lewis, B. K.; Yocum, G. T.; Arbab, A. S. Cytotherapy 2004, 6, 621–5. (67) Kikumori, T.; Kobayashi, T.; Sawaki, M.; Imai, T. Breast Cancer Res. Treat. 2009, 113, 435-41. (68) Ito, A.; Kuga, Y.; Honda, H.; Kikkawa, H.; Horiuchi, A.; Watanabe, Y.; Kobayashi, T. Cancer Lett. 2004, 212, 167–75. (69) Lipinski, M. J.; Amirbekian, V.; Frias, J. C.; Aguinaldo, J. G. S.; Mani, V.; Briley-Saebo, K. C.; Fuster, V.; Fallon, J. T.; Fisher, E. A.; Fayad, Z. A. Magn. Reson. Med. 2006, 56, 601–610. (70) Vuu, K.; Xie, J. W.; McDonald, M. A.; Bernardo, M.; Hunter, F.; Zhang, Y. T.; Li, K.; Bednarski, M.; Guccione, S. Bioconjugate Chem. 2005, 16, 995–999. (71) Turner, J. L.; Pan, D. P. J.; Plummer, R.; Chen, Z. Y.; Whittaker, A. K.; Wooley, K. L. AdV. Funct. Mater. 2005, 15, 1248–1254. (72) Yu, X.; Song, S. K.; Chen, J.; Scott, M. J.; Fuhrhop, R. J.; Hall, C. S.; Gaffney, P. J.; Wickline, S. A.; Lanza, G. M. Magn. Reson. Med. 2000, 44, 867–72. (73) Morawski, A. M.; Winter, P. M.; Crowder, K. C.; Caruthers, S. D.; Fuhrhop, R. W.; Scott, M. J.; Robertson, J. D.; Abendschein, D. R.; Lanza, G. M.; Wickline, S. A. Magn. Reson. Med. 2004, 51, 480– 486. (74) Aime, S.; Frullano, L.; Crich, S. G. Angew. Chem., Int. Ed. 2002, 41, 1017. (75) Reynolds, C. H.; Annan, N.; Beshah, K.; Huber, J. H.; Shaber, S. H.; Lenkinski, R. E.; Wortman, J. A. J. Am. Chem. Soc. 2000, 122, 8940– 8945. (76) Landfester, K. Macromol. Rapid Commun. 2001, 22, 896–936. (77) Crespy, D.; Stark, M.; Hoffmann-Richter, C.; Ziener, U.; Landfester, K. Macromolecules 2007, 40, 3122–3135. (78) Jagielski, N.; Sharma, S.; Hombach, V.; Mailander, V.; Rasche, V.; Landfester, K. Macromol. Chem. Phys. 2007, 208, 2229–2241. (79) Taylor, J. R.; Fang, M. M.; Nie, S. M. Anal. Chem. 2000, 72, 1979– 1986. (80) Bhalgat, M. K.; Haugland, R. P.; Pollack, J. S.; Swan, S.; Haugland, R. P. J. Immunol. Methods 1998, 219, 57–68. (81) Charreyre, M. T.; Zhang, P.; Winnik, M. A.; Pichot, C.; Graillat, C. J. Colloid Interface Sci. 1995, 170, 374–382. (82) Tronc, F.; Li, M.; Lu, J. P.; Winnik, M. A.; Kaul, B. L.; Graciet, J. C. J. Polym. Sci., Part A: Polym. Chem. 2003, 41, 766–778. (83) Weiss, C. K.; Lorenz, M. R.; Landfester, K.; Maila¨nder, V. Macromol. Biosci. 2007, 7, 883–896. (84) Hu, Y.; Xie, J.; Tong, Y. W.; Wang, C. H. J. Controlled Release 2006, 6, 6. (85) Dausend, J.; Musyanovych, A.; Dass, M.; Walther, P.; Schrezenmeier, H.; Landfester, K.; Mailander, V. Macromol. Biosci. 2008, 8, 1135– 43. (86) Holzapfel, V.; Musyanovych, A.; Landfester, K.; Lorenz, M. R.; Mailander, V. Macromol. Chem. Phys. 2005, 205, 2440–9. (87) Lorenz, M. R.; Kohnle, M. V.; Dass, M.; Walther, P.; Hocherl, A.; Ziener, U.; Landfester, K.; Mailander, V. Macromol. Biosci. 2008, 8, 711–27. (88) Falcone, S.; Cocucci, E.; Podini, P.; Kirchhausen, T.; Clementi, E.; Meldolesi, J. J. Cell Sci. 2006, 119, 4758–4769. (89) Lai, S. K.; Hida, K.; Man, S. T.; Chen, C.; Machamer, C.; Schroer, T. A.; Hanes, J. Biomaterials 2007, 28, 2876–2884. (90) Nabi, I. R.; Le, P. U. J. Cell Biol. 2003, 161, 673–677. (91) Arias, J. L.; Gallardo, V.; Go´mez-Lopera, S. A.; Plaza, R. C.; Delgado, A. V. J. Controlled Release 2001, 77, 309–321. (92) Reddy, L. H.; Murthy, R. R. Acta Pharm. 2004, 54, 103–118. (93) Kattan, J.; Droz, J.-P.; Couvreur, P.; Marino, J.-P.; Boutan-Laroze, A.; Rougier, P.; Brault, P.; Vranckx, H.; Grognet, J.-M.; Morge, X.; Sancho-Garnier, H. InVest. New Drugs 1992, 10, 191–199. (94) Steiniger, S. C. J.; Kreuter, J.; Khalansky, A. S.; Skidan, I. N.; Bobruskin, A. I.; Smirnova, Z. S.; Severin, S. E.; Uhl, R.; Kock, M.; Geiger, K. D.; Gelperina, S. E. Int. J. Cancer 2004, 109, 759– 767.

2400


(95) Gulyaev, A. E.; Gelperina, S. E.; Skidan, I. N.; Antropov, A. S.; Kivman, G. Y.; Kreuter, J. Pharm. Res. 1999, 16, 1564–1569. (96) Alyautdin, R.; Gothier, D.; Petrov, V.; Kharkevich, D.; Kreuter, J. Eur. J. Pharm. Biopharm. 1995, 41, 44–48. (97) Olivier, J.-C.; Fenart, L.; Chauvet, R.; Pariat, C.; Cecchelli, R.; Couet, W. Pharm. Res. 1999, 16, 1836–1842. (98) Behan, N.; Birkinshaw, C. Macromol. Rapid Commun. 2001, 22, 41– 43. (99) Couvreur, P. Crit. ReV. Ther. Drug Carrier Syst. 1988, 5, 1–17. (100) Couvreur, P.; Kante, B.; Roland, M.; Speiser, P. J. Pharm. Sci. 1979, 68, 1521–1524. (101) Lescure, F.; Zimmer, C.; Roy, D.; Couvreur, P. J. Colloid Interface Sci. 1992, 154, 77–86. (102) El-Egakey, M. A.; Bentele, V.; Kreuter, J. Int. J. Pharm. 1983, 13, 349–352. (103) Douglas, S. J.; Illum, L.; Davis, S. S.; Kreuter, J. J. Colloid Interface Sci. 1984, 101, 149–158. (104) Douglas, S. J.; Illum, L.; Davis, S. S. J. Colloid Interface Sci. 1985, 103, 154–163. (105) Vasnick, L.; Couvreur, P.; Christiaens-Leyh, D.; Roland, M. Pharm. Res. 1985, 36–41. (106) Seijo, B.; Fattal, E.; Roblot-Treupel, L.; Couvreur, P. Int. J. Pharm. 1990, 62, 1–7. (107) Chauvierre, C.; Vauthier, C.; Labarre, D.; Hommel, H. Colloid Polym. Sci. 2004, 282, 1016–1025. (108) Peracchia, M. T.; Vauthier, C.; Passirani, C.; Couvreur, P.; Labarre, D. Life Sci. 1997, 61, 749–761. (109) Go¨pferich, A. Eur. J. Pharm. Biopharm. 1996, 42, 1–11. (110) Vauthier, C.; Dubernet, C.; Fattal, E.; Pinto-Alphandary, H.; Couvreur, P. AdV. Drug DeliVery ReV. 2003, 55, 519–548. (111) Kreuter, J. AdV. Drug DeliVery ReV. 2001, 47, 65–81. (112) Gref, R.; Minamitake, Y.; Peracchia, M. T.; Trubetskoy, V.; Torchilin, V.; Langer, R. Science 1994, 263, 1600–1603. (113) Gurny, R.; Peppas, N. A.; Harrington, D. D.; Banker, G. S. Drug DeV. Ind. Pharm. 1981, 7, 1–25. (114) Duvvuri, S.; Gaurav Janoria, K.; Mitra, A. K. Pharm. Res. 2006, 23, 215–223. (115) Lai, M. K.; Tsiang, R. C. C. J. Microencapsulation 2004, 21, 307– 316. (116) Mehta, R. C.; Jeyanthi, R.; Calis, S.; Thanoo, B. C.; Burton, K. W.; DeLuca, P. P. J. Controlled Release 1994, 29, 375–384. (117) Benny, O.; Duvshani-Eshet, M.; Cargioli, T.; Bello, L.; Bikfalvi, A.; Carroll, R. S.; Machluf, M. Clin. Cancer Res. 2005, 11, 768–776. (118) Tracy, M. A.; Ward, K. L.; Firouzabadian, L.; Wang, Y.; Dong, N.; Qian, R.; Zhang, Y. Biomaterials 1999, 20, 1057–1062. (119) Lemoine, D.; Francois, C.; Kedzierewicz, F.; Preat, V.; Hoffman, M.; Maincent, P. Biomaterials 1996, 17, 2191–2197. (120) Park, T. G. J. Controlled Release 1994, 30, 161–173. (121) Park, T. G. Biomaterials 1995, 16, 1123–1130. (122) Li, S.; Vert, M. Macromolecules 1994, 27, 3107–3110. (123) Makino, K.; Arakawa, M.; Kondo, T. Chem. Pharm. Bull. 1985, 33, 1195–1201. (124) Reich, G. Eur. J. Pharm. Biopharm 1998, 45, 165–171. (125) Musyanovych, A.; Schmitz-Wienke, J.; Mailander, V.; Walther, P.; Landfester, K. Macromol. Biosci. 2008, 8, 127–39. (126) Coffin, M. D.; McGinity, J. W. Pharm. Res. 1992, 9, 200–205. (127) Win, K. Y.; Feng, S.-S. Biomaterials 2005, 26, 2713–2722. (128) Musyanovych, A.; Rossmanith, R.; Tontsch, C.; Landfester, K. Langmuir 2007, 23, 5367–5376. (129) Gershon, H.; Ghirlando, R.; Guttman, S. B.; Minsky, A. Biochemistry 1993, 32, 7143–51. (130) Gibanel, S.; Heroguez, V.; Gnanou, Y.; Aramendia, E.; Bucsi, A.; Forcada, J. Polym. AdV. Technol. 2001, 12, 494–499. (131) Ortegavinuesa, J. L.; Hidalgoalvarez, R. Biotechnol. Bioeng. 1995, 47, 633–639.

Maila¨nder and Landfester (132) Stebelska, K.; Dubielecka, P. M.; Sikorski, A. F. J. Membr. Biol. 2005, 206, 203–14. (133) Vonarbourg, A.; Passirani, C.; Saulnier, P.; Benoit, J.-P. Biomaterials 2006, 27, 4356–4373. (134) Torchilin, V. P.; Trubetskoy, V. S. AdV. Drug DeliVery ReV. 1995, 16, 141–155. (135) Peracchia, M. T.; Vauthier, C.; Desmae¨le, D.; Gulik, A.; Dedieu, J.-C.; Demoy, M.; d’Angelo, J.; Couvreur, P. Pharm. Res. 1998, 15, 550-6. (136) Schauer, R. Trends Biochem. Sci. 1985, 10, 357–360. (137) Passirani, C.; Barratt, G.; Devissaguet, J. P.; Labarre, D. Pharm. Res. 1998, 15, 1046–1050. (138) Lemarchand, C.; Gref, R.; Passirani, C.; Garcion, E.; Petri, B.; Mu¨ller, R.; Costantini, D.; Couvreur, P. Biomaterials 2006, 27, 108–118. (139) Brigger, I.; Dubernet, C.; Couvreur, P. AdV. Drug DeliVery ReV. 2002, 54, 631–651. (140) Brigger, I.; Morizet, J.; Aubert, G.; Chacun, H.; Terrier-Lacombe, M.-J.; Couvreur, P.; Vassal, G. J Pharmacol. Exp. Ther. 2002, 303, 928–936. (141) Ranney, D. F. Biochem. Pharmacol. 2000, 59, 105–114. (142) Lherm, C.; Muller, R. H.; Puisieux, F.; Couvreur, P. Int. J. Pharm. 1992, 84, 13–22. (143) Kante, B.; Couvreur, P.; Duboiskrack, G.; Demeester, C.; Guiot, P.; Roland, M.; Mercier, M.; Speiser, P. J. Pharm. Sci. 1982, 71, 786– 790. (144) Begley, D. J. J. Pharm. Pharmacol. 1996, 48, 136–146. (145) Beduneau, A.; Saulnier, P.; Benoit, J. P. Biomaterials 2007, 28, 4947– 4967. (146) Jones, A. R.; Shusta, E. V. Pharm. Res. 2007, 24, 1759–1771. (147) Garcia-Garcia, E.; Andrieux, K.; Gil, S.; Couvreur, P. Int. J. Pharm. 2005, 298, 274–292. (148) Ambruosi, A.; Gelperina, S.; Khalansky, A.; Tanski, S.; Theisen, A.; Kreuter, J. J. Microencapsulation 2006, 23, 582–592. (149) Kreuter, J. J. Nanosci. Nanotechnol. 2004, 4, 484–488. (150) Weiss, C. K.; Kohnle, M. V.; Landfester, K.; Hauk, T.; Fischer, D.; Schmitz-Wienke, J.; Mailander, V. ChemMedChem 2008, 3, 1395– 403. (151) Fischer, R.; Waizenegger, T.; Kohler, K.; Brock, R. Biochim. Biophys. Acta 2002, 1564, 365–74. (152) Jensen, K. D.; Nori, A.; Tijerina, M.; Kopeckova, P.; Kopecek, J. J. Controlled Release 2003, 87, 89–105. (153) Pratten, M. K.; Lloyd, J. B. Biochim. Biophys. Acta 1986, 881, 307– 13. (154) Schulze, E.; Ferrucci, J. T., Jr.; Poss, K.; Lapointe, L.; Bogdanova, A.; Weissleder, R. InVest. Radiol. 1995, 30, 604–10. (155) Sieczkarski, S. B.; Whittaker, G. R. J. Virol. 2002, 76, 10455–10464. (156) Szymkiewicz, I.; Shupliakov, O.; Dikic, I. Biochem. J. 2004, 383, 1–11. (157) Stuart, A. D.; Brown, T. D. J. Virol. 2006, 80, 7500–9. (158) Llorente, A.; van Deurs, B.; Garred, O.; Eker, P.; Sandvig, K. J. Cell Sci. 2000, 113 (Pt 7), 1213–21. (159) Alberts, B.; Bray, D.; Lewis, M.; Raff, K.; Roberts, K.; Watson, J. D. Molecular biology of the cell; Garland Publishing, Inc: New York, 1989. (160) Anderson, R. G. Annu. ReV. Biochem. 1998, 67, 199–225. (161) Nichols, B. J. Cell Sci. 2003, 116, 4707–14. (162) Kirkham, M.; Parton, R. G. Biochim. Biophys. Acta 2005, 1746, 349– 63. (163) Qaddoumi, M. G.; Gukasyan, H. J.; Davda, J.; Labhasetwar, V.; Kim, K. J.; Lee, V. H. Mol. Vision 2003, 9, 559–68. (164) Klingauf, J. Neuron 2007, 54, 857–858. (165) Pouliquen, D.; Le Jeune, J. J.; Perdrisot, R.; Ermias, A.; Jallet, P. Magn. Reson. Imaging 1991, 9, 275–83.

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