Multichannel MRI Labeling of Mammalian Cells by Switchable

Letter pubs.acs.org/NanoLett

Multichannel MRI Labeling of Mammalian Cells by Switchable Nanocarriers for Hyperpolarized Xenon Stefan Klippel,†,‡ Christian Freund,‡ and Leif Schröder*,† †

ERC Project BiosensorImaging, Leibniz-Institut für Molekulare Pharmakologie (FMP), 13125 Berlin, Germany Protein Biochemistry Group, Freie Universität Berlin, 14195 Berlin, Germany

‡

S Supporting Information *

ABSTRACT: We demonstrate a concept for multichannel MRI cell-labeling using encapsulated laser-polarized xenon. Conceptually different Xe trapping properties of two nanocarriers, namely macrocyclic cages as individual hosts or compartmentalization into nanodroplets, ensure a large chemical shift separation for Xe bound in either of the carriers even after cellular internalization. Two differently labeled mammalian cell populations were imaged by frequency selective saturation transfer resulting in a switchable “twocolor” xenon-MRI contrast at micro- to nanomolar Xe carrier concentrations. KEYWORDS: Multiplexing, cell-labeling, MRI, xenon, hyperpolarization, CEST

N

MR detection of laser-polarized 129 Xe has been investigated in many studies based on the ability of dissolved xenon to form inclusion complexes with supramolecular hosts,1−4 hydrophobic binding pockets of proteins,5,6 or bacterial gas vesicles.7 An important feature of these interactions is the unique chemical shift of encapsulated xenon, which depends on the molecular structure of the host system and the immediate environment. It has therefore been postulated to use different xenon hosts that confer individual chemical shifts to label and detect different molecular targets for the purpose of diagnostic imaging.8 This multiplexing approach is of great interest for analyzing the complexity of biological systems in vivo.9 Examples include tracking of cell fate in cell therapy10 and molecular profiling of tumors by targeted tracers.11 Optical methods generally have high sensitivity and allow for the multichannel detection of several tracers, but their use is constrained to small animal experiments due to limited penetration depth in tissue.12,13 MRI has the potential to overcome this limitation because it uses deep penetrating radio waves and also comes with a spectral dispersion for encoding multiple tracers along the chemical shift dimension, as demonstrated for chemical exchange saturation transfer (CEST)14 agents. These compounds have through their frequency selectivity a significantly higher specificity over conventional relaxation-based MRI contrast agents. However, the detection limit of 1H-CEST is still in the range of 10−3 M.7 Herein we demonstrate a very sensitive xenon-MRI approach that allows for the multiplexed localization of two individual nanoparticulate xenon hosts that are acting as cell tracers: cryptophane-A cages (CrA) and perfluoroctyl bromide (PFOB) © 2014 American Chemical Society

nanodroplets. Both hosts confer a unique chemical shift to xenon to enable a frequency-selective detection of two distinctly labeled mammalian cell populations resulting in a switchable “two-color” MRI contrast. The high sensitivity of the demonstrated approach is based on combining laser-induced xenon hyperpolarization and indirect cell tracer detection by a host-specific saturation transfer (Hyper-CEST).15 This study compares two conceptually different xenon nanocarriers: cryptophane-A as a host with a tailored cavity (cavity volume ca. 90 Å3, one xenon atom per host)16 and PFOB nanodroplets (carrier volume ca. 4 × 109 Å3) with the capacity to bind many xenon atoms. Also, Xe atoms encounter interactions with different local environments that have impact on the chemical shift. In CrA, it is surrounded by the aromatic rings of the well-fitted CTV caps whereas in PFOB it is immersed within the linear structures of the fluorocarbon chains. This was chosen to ensure a large chemical shift separation of Xe between both nanocarriers that will also be retained after further changes in the microenvironment such as cellular internalization. Well-separated resonance frequencies are ideal to exploit the large signal amplification inherent to the Hyper-CEST technique. With regard to single-type labeling, cryptophane-A has been used in vitro for MRI cell-tracking at low micromolar concentrations17 and for molecular imaging of cell surface receptors at nM concentrations,18 i.e., significantly lower than conventional Gd contrast agents.19 While inclusion complexes Received: July 3, 2014 Revised: August 28, 2014 Published: September 23, 2014 5721

dx.doi.org/10.1021/nl502498w | Nano Lett. 2014, 14, 5721−5726

Nano Letters

Letter

Figure 1. Principle of multichannel Hyper-CEST detection for cell labeling. The MRI signal of hyperpolarized xenon in bulk solution acts as a sensing medium. It reports on the spatial distribution of two distinct cell-internalized xenon inclusion complexes (CrA and PFOB) both characterized by a unique xenon chemical shift. A reference scan visualizes the unaffected distribution of xenon in solution ((a) off-resonant saturation). The localization of cells either labeled with CrA ((b) on-resonant saturation for xenon@CrA in cells) or PFOB ((c) on-resonant saturation for xenon@ PFOB in cells) is encoded within the sensing medium by a frequency-specific saturation transfer acting on xenon atoms temporarily entrapped within the respective cell-internalized nanocarrier. Schematic spectra are shown for clarification as are participating xenon pools on top. Peak widths differ due to different exchange conditions.

solution signal of xenon. A first reference scan visualizes the unaffected distribution of the sensing medium (rf-saturation: off-resonant). The second scan encodes the localization of CrAlabeled cells by a frequency selective depolarization of xenon within cell-internalized CrA (rf-saturation: on-resonant for xenon@CrA in cells). The depolarization of this typically lowconcentrated spin ensemble is transferred into the abundant spin-pool of xenon in solution by efficient chemical exchange (ca. 30 Hz for cryptophane-A at room temperature).23 As the T1 relaxation time of xenon allows for saturation pulses of several seconds,24 the depolarization gets efficiently amplified within the solution pool where it is detected. As a consequence, the sensing medium gets depleted at the localization of CrAlabeled cells. The localization of PFOB-labeled cells is encoded by the same saturation transfer principle while the RF-pulse is placed on the resonance frequency of xenon within cellinternalized PFOB (rf-saturation: on-resonant for xenon@ PFOB in cells) in a third scan. Though CrA has a cavity that yields a high binding constant for Xe (∼103 M−1 in water),25 the PFOB nanocarriers are expected to yield high image contrast at even lower concentrations due to their ability to serve as host for many Xe atoms. Differences in Xe exchange dynamics and a certain droplet size distribution for this host manifests in different NMR spectral qualities (line broadening) that could potentially limit multichannel detection with this agent. Therefore, this study aims to demonstrate that these two xenon hosts with their different spectral properties can be distinguished from each other in cellular environment under feasible CEST saturation parameters. Monitoring the Cellular Uptake of Xenon Hosts. We started by investigating the Hyper-CEST properties of cell internalized PFOB nanodroplets. Different studies suggest that various mammalian cell types can be easily labeled with perfluorcarbon nanodroplets.22,26,27 We decided to label mouse

of xenon in PFOB nanodroplets have been detected with Hyper-CEST at extremely low concentrations, they have not been detected in a cellular environment and demonstration of their Xe MRI imaging capability has not been described to date.20 In order to build different Hyper-CEST contrast agents that can be used in combination, a relatively big chemical shift separation (ca. 30 ppm) is highly desirable. This is an important consideration due to the expected broadening of each CEST peak (as a consequence of relaxation effects, magnetic field inhomogeneity as well as the use of strong saturation pulses21) likely to occur in vivo. The chemical shift for xenon entrapped in PFOB nanodroplet solutions (reported as 120 ppm from Xe in the gas phase) is therefore suitable to be tested in combination with xenon inside cryptophane-A molecules (reported as 60 ppm from Xe in the gas phase) for multicolor cell-labeling. Moreover, PBOB nanodroplets have several favorable characteristics, notably their “straightforward, inexpensive production”, a known in vivo behavior due to their proven use as ultrasound and 19F-MRI contrast agents as well as the lowest detection level (subpicomolar for nonlocalized detection) reported for Hyper-CEST agents so far.20 These properties along with the demonstrated possibility to label different cell types such as stem- and progenitor cells while preserving viability22 also motivated this study. Principle of Multichannel Hyper-CEST MRI. The MRI localization of either CrA- or PFOB-labeled cells is achieved by a frequency selective depletion of the xenon signal in solution as conceptually illustrated in Figure 1. Through CEST, this solution signal thus acts sequentially as an indirect sensing medium for the xenon in each host agent. Both types of xenon hosts are characterized by a unique xenon chemical shift after cellular internalization. Selective MRI localization of both cell tracers is achieved within three subsequent measurements by varying the saturation-frequency offset while detecting the 5722

dx.doi.org/10.1021/nl502498w | Nano Lett. 2014, 14, 5721−5726

Nano Letters

Letter

Figure 2. Cellular uptake of xenon inclusion complexes can be monitored by Hyper-CEST-spectroscopy. The intensity of xenon in solution was observed following a 5 s, 8 μT cw saturation at varying frequency offsets. (a) The CEST response of Xe@PFOB shifts upfield after cellular internalization while the spectral resolution is improved. (b) Coincubation of cells with PFOB and CrA reveals a xenon chemical shift separation of approximately 40 ppm between both cell-internalized xenon hosts. Chemical shift changes related to cellular uptake of the respective MRI tracers are indicated by gray bars. Schematics are used according to Figure 1.

fibroblasts (L929) because CrA-based cell labeling and HyperCEST detection has been already described in detail for this cell type.17 The cells where incubated for 18 h with cell culture medium containing 1.6 nM PFOB nanodroplets (diameter: 200 nm). The labeling conditions have been optimized in order to achieve high nanodroplet uptake while preserving cellular viability. We also aimed for a CEST effect that should be comparable with the one of CrA-labeled cells. Efficient PFOB uptake under the given conditions was confirmed by incubating cells with fluorescence-labeled nanodroplets (fluorescence dye DiI, see Supporting Information) followed by microscopy. After labeling cells were washed and resuspended (10 million cells/ mL) in fresh medium, while viability was confirmed by Trypan blue staining. Direct NMR detection of the PFOB-related xenon NMR signal was not possible at nanomolar droplet concentration. This is explained by relative fast chemical exchange between xenon in solution and the nanodroplets causing extensive line broadening for this spin ensemble and making it disappear in the noise level. While this exchange regime hampers direct detection it is beneficial for saturation transfer based detection. Therefore, Hyper-CEST spectroscopy allows analyzing the xenon chemical shift signature of PFOB-labeled mammalian cells (Figure 2a). While detecting the abundant signal of hyperpolarized xenon in solution, an rf-pulse (saturation: 5 s with 8 μT) was used to selectively saturate the xenon chemical shift range between 5 and 250 ppm in 5 ppm steps. For the incubation medium (diluted by a factor of 2 without cells) a broad saturation response centered at 120 ppm was observed in addition to the CEST-response for direct saturation of xenon in solution (at 190 ppm). The additional CEST-peak represents xenon within PFOB-nanodroplets as described by Pines and co-workers.20 The sample containing PFOB-labeled cells in suspension shows a narrower PFOB-related CEST-peak centered around 110 ppm. The supernatant of the measured cell sample (cells were removed by sedimentation) shows no CEST-response at 110 ppm, thus confirming that the signal is cell-related. The change in line width as well as in resonance position of the CEST peak for xenon within cell internalized PFOB nanodroplets compared to PFOB in solution can be related to different aspects. (i) A slower chemical exchange of xenon due to the

additional diffusion barrier represented by the cellular membrane, (ii) an increase of the PFOB droplet size after cellular internalization, or (iii) a concentration-dependent change within the peak shape.20,21 Complementary to the fluorescence data, cell uptake of the PFOB nanodroplets therefore can be monitored by a 10 ppm upfield shift of PFOBencapsulated xenon at 310 K. A similar effect is known for Xe in cell internalized CrA cages.28 Hence, both types of Xe nanocarriers are able to distinguish between extracellular and cell-associated environment, a capability that does not come with simple fluorescence reporters. For the cellular uptake of CrA, the comparable xenon chemical shift change occurs along the opposite direction (downfield). To illustrate both effects, we repeated the cell labeling experiment with the addition of CrA to the PFOBincubation medium (final CrA concentration of 50 μM according to labeling conditions optimized elsewhere17) (Figure 2b). This time the incubation medium (dilution by a factor of 2) showed an additional narrow CEST peak representing CrA-bound xenon at 60 ppm next to peak of xenon in PFOB at 120 ppm. The CEST peaks of both contrast agents are then shifted toward each other after cellular uptake resulting in a chemical shift separation of 40 ppm at physiological temperature (310 K). Selective MRI Localization of PFOB- and CrA- Labeled Cells. We performed an MRI experiment to demonstrate the multiplexing capability of both Hyper-CEST contrast agents. Two batches of mouse fibroblasts were labeled with either PFOB nanodroplets (0.25 nM, diameter: 380 nm, droplet concentration had changed due to Ostwald ripening) or cryptophane-A (50 μM) individually (both for 18 h incubation). After washing, both cell batches were resuspended in cell culture medium (10 million cells/mL). We decided to label two cell populations of the same cell type (L929) although the concept allows for the multiplexed detection of different cell types as explained later on. This ensures that the observed chemical shift difference (40 ppm) between the signals of both cell populations is just related to the type of cell internalized xenon host and not influenced by the cell type (different cell size, metabolism, Xe uptake). Hence, both nanocarriers are tested in this study for their ability to serve as selectively MRI-switchable building blocks for future functionalization. 5723

dx.doi.org/10.1021/nl502498w | Nano Lett. 2014, 14, 5721−5726

Nano Letters

Letter

Figure 3. PFOB- and CrA-labeled cells can be selectively localized by multichannel xenon-MRI as demonstrated for a two compartment phantom (outer compartment, PFOB-labeled cells; inner compartment, CrA-labeled cells). (a) The saturation frequency offsets used for Hyper-CEST imaging are indicated within the associated CEST-spectrum (8 s, 10 μT cw saturation) by gray dotted lines. Exponential Lorentzian fits are shown as solid lines. Schematics are used according to Figure 1. (b) 1H-MR-Images are shown in overlay with pseudocolored Hyper-CEST-effects derived for both contrast agent-specific saturation conditions. Cell labeling was confirmed by laser scanning microscopy with fluorescence labeled versions of both Hyper-CEST agents as depicted within the circular insets.

their viability.17 It therefore simulates in vivo-like conditions where xenon has to be transported to the region of interest via the bloodstream after inhalation or infusion. In addition, no cell fragmentation occurs thereby excluding their contributions of cell released xenon hosts to the observed effects. Two batches of mouse fibroblasts were labeled with either PFOB nanoemulsion (1.6 nM, diameter: 200 nm) or CrA (50 μM) individually (both 18 h incubation) and encapsulated within solid alginate beads (ca. 1 mm diameter). The bioreactor consists of two separated compartments that were either filled with PFOB- or CrA-labeled cells (10 million cells/compartment). The intracellular concentration of both contrast agents was estimated by measuring the fluorescence intensity in cell lysates. Therefore, cells have been labeled with fluorescence labeled versions of both tracers (CrA, CrA-FAM; PFOB, PFOB-DiI) under incubation conditions identical to the MRI experiment. Using some simplifications (see Supporting Informations) the intracellular concentration within the shown MRI experiment is 80 nM for PFOB and 40 μM for CrA. By using Hyper-CEST (saturation: 10 s with 10 μT; for image postprocessing, see Supporting Information Figure S2) we were able to identify the localization of both individually labeled cell populations within 2 CEST acquisitions (2 min) per contrast agent (Figure 4). It should be noticed that in this initial study we aimed for relative high intracellular concentrations of both cell tracers for multichannel detection (when compared to cell tracking with each of the tracers individually). This was motivated by expected spillover effects between the PFOB- and CrA-Hyper-CEST responses that had to be minimized by the use of weak saturation pulses, thereby requiring relative high cell tracer concentrations. Nevertheless, variations in cell numbers as well as cell tracer concentrations can be considered by adjusting the saturation parameters for each of the xenon hosts individually. Translation of the Multiplexing Concept to Preclinical Applications. So far, the principle of xenon-based HyperCEST contrast agents has not been demonstrated in live animals. Nevertheless various studies within animals and humans demonstrated the potential to deliver dissolved hpxenon into strongly vascularized organs such as the brain after

A double-compartment phantom was prepared by inserting a 5 mm NMR tube (inner compartment) into a 10 mm NMR tube (outer compartment). The outer compartment was loaded with PFOB-labeled cells in solution while the inner one was filled with CrA-labeled cells in solution. Using Hyper-CEST spectroscopy (sampling step, 5 ppm; saturation, 8 s with 10 μT) four different spin pools could be resolved: xenon in solution at 192 ppm, Xe@PFOB in cells at 110 ppm, Xe@CrA in cells at 70 ppm, and Xe@CrA in solution at 60 ppm (Figure 3a). The additional peak for Xe@CrA in solution is related to CrA released from the cells. This peak is not resolved within the Hyper-CEST spectrum of labeled cells in Figure 2b due to different saturation conditions (the resonance could be revealed by reducing the sampling step size, data not shown). The release of CrA is a consequence of the xenon delivery strategy that was used within this initial in vitro experiment: hp-xenon is directly dissolved within the cell solution by using a spectrometer triggered bubble dispenser. This bubbling procedure causes fragmentation of sensitive cell types correlated with CrA release. Nevertheless, this method allows performing Hyper-CEST spectroscopy quickly which is beneficial for an initial characterization of the applied labeling strategy. Figure 3b illustrates the frequency selective Hyper-CESTMRI localization of both differently labeled cell populations. The Hyper-CEST-effect when saturating at 107 ppm (Xe@ PFOB in cells) is clearly restricted to the outer compartment with a mean value of 70%. A saturation pulse centered @ 70 ppm (Xe@CrA in cells) reveals the localization of CrA-labeled cells restricted to the inner compartment also with a mean Hyper-CEST effect of 70%. The Hyper-CEST effect has been calculated based on the average of 20 on-resonant (saturation @ 70 ppm or @ 107 ppm) and 20 off-resonant (saturation @ 326 ppm) images acquired with a CEST-weighted RARE sequence29 (saturation for 8 s with 13 μT; for image postprocessing, see Supporting Information Figure S1). Live Cell Multichannel Cell-Tracking within a Perfused Bioreactor System. To work out the detection limit with respect to measurement time, we performed a multiplexing experiment within an MRI compatible bioreactor system. The setup enables the perfusion of immobilized cells with hyperpolarized xenon saturated medium without impairing 5724

dx.doi.org/10.1021/nl502498w | Nano Lett. 2014, 14, 5721−5726

Nano Letters

Letter

Although not proven so far, it is expected that the same holds true for cell internalized PFOB because the Xe chemical shift is dominated by the immediate fluorocarbon environment that shields it from cellular material. Further on, multichannel detection can be extended for in vivo labeling applications by injecting targeted versions of both Hyper-CEST agents into appropriate animal models. This is a valid extension because the individual resonance frequency of Xe in either cryptophane-A or PFOB in cellular environments is practically independent from the labeling strategy (ex vivo or in vivo). This holds potential for multiplexed molecular imaging applications of various pathologies such as tumor profiling. Indeed, the molecular specificity of cryptophane conjugates for certain disease markers has been already demonstrated by the coupling of targeting units such as antibodies,37 peptides,38−40 or other ligands23,28,41 leading to cell surface labeling or receptor mediated cellular internalization. Similar approaches have been carried out for PFOB by the incorporation of targetspecific phospholipids into the outer lipid monolayer surrounding the droplets.42,43 This work illustrates the feasibility to perform highly sensitive, multichannel detection of mammalian cells by Xe MRI using synthetic Xe nanocarriers as contrast agents. Cellular uptake of PFOB nanodroplets was characterized and monitored by Hyper-CEST spectroscopy and revealed improved spectral resolution upon partitioning through the membrane. The expected superior CEST imaging performance of PFOB compared to CrA was also confirmed. As for the above-mentioned gas vesicle approach, 7 functionalization is feasible for all these types of Xe carriers though it might require routes different from simple postinsertion as doable for PFOB nanodroplets. The gas vesicle concept also came with multiplexed detection of vesicles within different bacterial species. This illustrates that the multichannel idea and the use of Xe nanocarriers with a high Xe load (as opposed to 1:1 complex formation) can be achieved in different approaches and significantly improves the perspectives for Xe Hyper-CEST MRI toward a colorful future.

Figure 4. Live cell multichannel xenon-MRI within a perfused bioreactor system. Alginate-immobilized cells, labeled with either PFOB (intracellular concentration, 80 nM) or CrA (intracellular concentration, 40 μM) has been perfused with xenon-saturated medium. Selective cell tracking could be achieved within 2 HyperCEST acquisitions per contrast agent. Cell labeling of alginate encapsulated cells was confirmed by laser scanning microscopy with fluorescence-labeled versions of both Hyper-CEST agents as depicted within the circular inlets. Schematics are used according to Figure 1.

xenon inhalation in sufficient amounts to map its distribution by MRI.30−32 The advantage of the Hyper-CEST approach is that as long as the distribution of free Xe is detectable, its switchable knockout will identify the presence of specific Xe nanocarriers. In this context, pharmacokinetic modeling suggested that the achieved xenon concentrations are sufficient to detect 400 pM concentrations of xenon vesicles with Hyper-CEST schemes that are adopted to in vivo requirements.7 Such gas vesicles (ca. 4.1−16.5 × 109 Å3) represent genetically encoded hosts instead of synthetic nanocarriers and might be somewhat superior in terms of their Xe cargo capacity compared to the PFOB nanodroplets (ca. 4.2 × 109 Å3 for 200 nm diameter) used in this study. However, application of the gas vesicle concept to live animals is more complex then administration of synthetic nanocarriers. Hence, detection of nanomolar PFOB concentration appears to be feasible in vivo. Sensitivity limitation originating from scaling the setup to whole mice or even larger organisms only affects the detection conditions for the pool of dissolved Xe that does not represent a technical limitation (studies mentioned above). Moreover, continuous infusion of dissolved hp-xenon is known as an alternative delivery strategy within mouse experiments.33,34 Therefore, the multichannel cell labeling concept demonstrated here provides interesting experimental opportunities for a desired in vivo translation of the Hyper-CEST principle. Another important aspect for the feasibility of in vivo experiments is the separate delivery of the (targeted) nanocarrier and the hyperpolarized nuclei. This allows convenient timing for increased uptake/labeling in target structures and improved wash-out in surrounding areas. A potential experiment would be the injection and tracking of two different cell types after their individual ex vivo labeling. This would allow studying cell fate with relevance for cell therapeutic applications. Multichannel detection of different cell types is possible because the chemical shift of cell-internalized cryptophane-A is practically cell-independent as demonstrated for fibroblasts,17 macrophages,35 and brain endothelial cells.36

■

ASSOCIATED CONTENT

S Supporting Information *

Materials and methods, including preparation of PFOB nanoemulsion, cell labeling and NMR sample preparation, fluorescence microscopy and quantification, as well as NMR experiments and MR-image postprocessing. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Homepage: https://www. fmp-berlin.de/schroeder. Notes

The authors declare no competing financial interest.

■

REFERENCES

(1) Bartik, K.; Luhmer, M.; Heyes, S. J.; Ottinger, R.; Reisse, J. J. Magn. Reson. B 1995, 109, 164−168. (2) Bartik, K.; Luhmer, M.; Dutasta, J.-P.; Collet, A.; Reisse, J. J. Am. Chem. Soc. 1998, 120, 784−791. (3) Kim, B. S.; Ko, Y. H.; Kim, Y.; Lee, H. J.; Selvapalam, N.; Lee, H. C.; Kim, K. Chem. Commun. (Cambridge, U.K.) 2008, 2756−2758. (4) Brotin, T.; Dutasta, J.-P. Chem. Rev. 2009, 109, 88−130.

5725

dx.doi.org/10.1021/nl502498w | Nano Lett. 2014, 14, 5721−5726

Nano Letters

Letter

(5) McKim, S.; Hinton, J. F. Biochim. Biophys. Acta, Biomembr. 1994, 1193, 186−198. (6) Bowers, C. R.; Storhaug, V.; Webster, C. E.; Bharatam, J.; Cottone, A.; Gianna, R.; Betsey, K.; Gaffney, B. J. J. Am. Chem. Soc. 1999, 121, 9370−9377. (7) Shapiro, M. G.; Ramirez, R. M.; Sperling, L. J.; Sun, G.; Sun, J.; Pines, A.; Schaffer, D. V.; Bajaj, V. S. Nat. Chem. 2014, 6, 629−634. (8) Spence, M. M.; Rubin, S. M.; Dimitrov, I. E.; Ruiz, E. J.; Wemmer, D. E.; Pines, A.; Yao, S. Q.; Tian, F.; Schultz, P. G. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 10654−10657. (9) Kobayashi, H.; Longmire, M. R.; Ogawa, M.; Choyke, P. L.; Kawamoto, S. Lancet Oncol. 2010, 11, 589−595. (10) De Vries, I. J. M.; Lesterhuis, W. J.; Barentsz, J. O.; Verdijk, P.; van Krieken, J. H.; Boerman, O. C.; Oyen, W. J. G.; Bonenkamp, J. J.; Boezeman, J. B.; Adema, G. J.; Bulte, J. W. M.; Scheenen, T. W. J.; Punt, C. J. A.; Heerschap, A.; Figdor, C. G. Nat. Biotechnol. 2005, 23, 1407−1413. (11) Hussain, T.; Nguyen, Q. T. Adv. Drug Delivery Rev. 2014, 66, 90−100. (12) Bentolila, L. A.; Ebenstein, Y.; Weiss, S. J. Nucl. Med. 2009, 50, 493−496. (13) Dinish, U. S.; Balasundaram, G.; Chang, Y.-T.; Olivo, M. Sci. Rep. 2014, 4, 4075. (14) Liu, G.; Moake, M.; Har-el, Y.; Long, C. M.; Chan, K. W. Y.; Cardona, A.; Jamil, M.; Walczak, P.; Gilad, A. A.; Sgouros, G.; van Zijl, P. C. M.; Bulte, J. W. M.; McMahon, M. T. Magn. Reson. Med. 2012, 67, 1106−1113. (15) Schröder, L.; Lowery, T. J.; Hilty, C.; Wemmer, D. E.; Pines, A. Science 2006, 314, 446−449. (16) Taratula, O.; Hill, P. A.; Khan, N. S.; Carroll, P. J.; Dmochowski, I. J. Nat. Commun. 2010, 1, 148. (17) Klippel, S.; Döpfert, J.; Jayapaul, J.; Kunth, M.; Rossella, F.; Schnurr, M.; Witte, C.; Freund, C.; Schröder, L. Angew. Chem., Int. Ed. 2014, 53, 493−496. (18) Rose, H. M.; Witte, C.; Rossella, F.; Klippel, S.; Freund, C.; Schröder, L. Proc. Natl. Acad. Sci. U.S.A. 2014, 111, 11697−11702. (19) Crich, S. G.; Biancone, L.; Cantaluppi, V.; Duò, D.; Esposito, G.; Russo, S.; Camussi, G.; Aime, S. Magn. Reson. Med. 2004, 51, 938− 944. (20) Stevens, T. K.; Ramirez, R. M.; Pines, A. J. Am. Chem. Soc. 2013, 135, 9576−9579. (21) Zaiss, M.; Schnurr, M.; Bachert, P. J. Chem. Phys. 2012, 136, 144106−144106−10. (22) Partlow, K. C.; Chen, J.; Brant, J. A.; Neubauer, A. M.; Meyerrose, T. E.; Creer, M. H.; Nolta, J. A.; Caruthers, S. D.; Lanza, G. M.; Wickline, S. A. FASEB J. 2007, 21, 1647−1654. (23) Spence, M. M.; Ruiz, E. J.; Rubin, S. M.; Lowery, T. J.; Winssinger, N.; Schultz, P. G.; Wemmer, D. E.; Pines, A. J. Am. Chem. Soc. 2004, 126, 15287−15294. (24) Spence, M. M.; Rubin, S. M.; Dimitrov, I. E.; Ruiz, E. J.; Wemmer, D. E.; Pines, A.; Yao, S. Q.; Tian, F.; Schultz, P. G. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 10654−10657. (25) Schröder, L. Phys. Med. 2013, 29, 3−16. (26) Ahrens, E. T.; Zhong, J. NMR Biomed. 2013, 26, 860−871. (27) Janjic, J. M.; Ahrens, E. T. Wiley Interdiscip. Rev.: Nanomed. Nanobiotechnol. 2009, 1, 492−501. (28) Boutin, C.; Stopin, A.; Lenda, F.; Brotin, T.; Dutasta, J.-P.; Jamin, N.; Sanson, A.; Boulard, Y.; Leteurtre, F.; Huber, G.; BogaertBuchmann, A.; Tassali, N.; Desvaux, H.; Carrière, M.; Berthault, P. Bioorg. Med. Chem. 2011, 19, 4135−4143. (29) Hennig, J.; Nauerth, A.; Friedburg, H. Magn. Reson. Med. Off. J. Soc. Magn. Reson. Med. Soc. Magn. Reson. Med. 1986, 3, 823−833. (30) Swanson, S. D.; Rosen, M. S.; Agranoff, B. W.; Coulter, K. P.; Welsh, R. C.; Chupp, T. E. Magn. Reson. Med. Off. J. Soc. Magn. Reson. Med. Soc. Magn. Reson. Med. 1997, 38, 695−698. (31) Zhou, X.; Sun, Y.; Mazzanti, M.; Henninger, N.; Mansour, J.; Fisher, M.; Albert, M. NMR Biomed. 2011, 24, 170−175. (32) Kilian, W.; Seifert, F.; Rinneberg, H. Magn. Reson. Med. 2004, 51, 843−847.

(33) Duhamel, G.; Choquet, P.; Grillon, E.; Lamalle, L.; Leviel, J.-L.; Ziegler, A.; Constantinesco, A. Magn. Reson. Med. 2001, 46, 208−212. (34) Cleveland, Z. I.; Möller, H. E.; Hedlund, L. W.; Nouls, J. C.; Freeman, M. S.; Qi, Y.; Driehuys, B. PLoS One 2012, 7, e31306. (35) Rossella, F.; Rose, H. M.; Witte, C.; Jayapaul, J.; Schröder, L. ChemPlusChem 2014, DOI: 10.1002/cplu.201402179. (36) Schnurr, M.; Sydow, K.; Rose, H. M.; Dathe, M.; Schröder, L. Adv. Healthcare Mater. 2014, DOI: 10.1002/adhm.201400224. (37) Palaniappan, K. K.; Ramirez, R. M.; Bajaj, V. S.; Wemmer, D. E.; Pines, A.; Francis, M. B. Angew. Chem., Int. Ed. 2013, 52, 4849−4853. (38) Seward, G. K.; Wei, Q.; Dmochowski, I. J. Bioconjugate Chem. 2008, 19, 2129−2135. (39) Schlundt, A.; Kilian, W.; Beyermann, M.; Sticht, J.; Günther, S.; Höpner, S.; Falk, K.; Roetzschke, O.; Mitschang, L.; Freund, C. Angew. Chem., Int. Ed. 2009, 48, 4142−4145. (40) Wei, Q.; Seward, G. K.; Hill, P. A.; Patton, B.; Dimitrov, I. E.; Kuzma, N. N.; Dmochowski, I. J. J. Am. Chem. Soc. 2006, 128, 13274− 13283. (41) Chambers, J. M.; Hill, P. A.; Aaron, J. A.; Han, Z.; Christianson, D. W.; Kuzma, N. N.; Dmochowski, I. J. J. Am. Chem. Soc. 2009, 131, 563−569. (42) Lanza, G. M.; Wallace, K. D.; Scott, M. J.; Cacheris, W. P.; Abendschein, D. R.; Christy, D. H.; Sharkey, A. M.; Miller, J. G.; Gaffney, P. J.; Wickline, S. A. Circulation 1996, 94, 3334−3340. (43) Giraudeau, C.; Geffroy, F.; Mériaux, S.; Boumezbeur, F.; Robert, P.; Port, M.; Robic, C.; Bihan, D. L.; Lethimonnier, F.; Valette, J. Angiogenesis 2013, 16, 171−179.

5726

dx.doi.org/10.1021/nl502498w | Nano Lett. 2014, 14, 5721−5726