In Vivo Monitoring of Inflammation After Cardiac and Cerebral Ischemia by Fluorine Magnetic Resonance Imaging
Background— In this study, we developed and validated a new approach for in vivo visualization of inflammatory processes by magnetic resonance imaging using biochemically inert nanoemulsions of perfluorocarbons (PFCs).
Methods and Results— Local inflammation was provoked in 2 separate murine models of acute cardiac and cerebral ischemia, followed by intravenous injection of PFCs. Simultaneous acquisition of morphologically matching proton (1H) and fluorine (19F) images enabled an exact anatomic localization of PFCs after application. Repetitive 1H/19F magnetic resonance imaging at 9.4 T revealed a time-dependent infiltration of injected PFCs into the border zone of infarcted areas in both injury models, and histology demonstrated a colocalization of PFCs with cells of the monocyte/macrophage system. We regularly found the accumulation of PFCs in lymph nodes. Using rhodamine-labeled PFCs, we identified circulating monocytes/macrophages as the main cell fraction taking up injected nanoparticles.
Conclusions— PFCs can serve as a “positive” contrast agent for the detection of inflammation by magnetic resonance imaging, permitting a spatial resolution close to the anatomic 1H image and an excellent degree of specificity resulting from the lack of any 19F background. Because PFCs are nontoxic, this approach may have a broad application in the imaging and diagnosis of numerous inflammatory disease states.
Received September 4, 2007; accepted April 28, 2008.
Inflammation is associated with a large number of human diseases such as atherosclerosis, glomerulonephritis, inflammatory bowel disease, transplant rejection, neurodegenerative brain diseases, brain and spinal cord trauma, myocarditis, and ischemic heart disease. Thus, the medical problem is vast and an exact diagnosis is often difficult. Accordingly, therapy frequently is limited to symptomatic treatment and the success of the prescribed therapy is difficult to assess. Although recent advances involve various imaging modalities such as positron emission tomography, computed tomography, magnetic resonance imaging (MRI), optical imaging, and ultrasound imaging,1–3 the visualization of inflammatory processes still poses a serious challenge, especially because in the initial phase the affected tissue does not exhibit specific physical properties that can be used to create contrast between inflamed and healthy regions.
Editorial p 109
Clinical Perspective p 148
Among the different noninvasive imaging modalities capable of whole-body imaging such as positron emission tomography and single-photon emission computed tomography, MRI provides superior resolution and the potential to generate the required contrast to noninflamed areas by gadolinium enhancement. However, this attempt relies on the transient accumulation of intravascularly applied gadolinium contrast agent in the interstitial space because of enhanced endothelial permeability,4,5 which is a rather nonspecific phenomenon found to be associated with a variety of diseases. A more defined approach to delineate inflammatory areas from surrounding tissue is the tagging of infiltrating, immunocompetent cells with contrast agents.6,7 Noninvasive visualization of immigrating cells by MRI has so far used predominantly superparamagnetic iron oxide particles, taking advantage of the high affinity of these species for the monocyte/macrophage system.8,9 Despite its excellent sensitivity, this attempt has the disadvantage that the particles are not detected directly. Local deposition results in regional magnetic field inhomogeneities and thus depletion of the MR signal. Consequently, anatomic proton (1H) MRIs often are difficult to interpret because it is not always clear whether dark areas are caused by these nanoparticles or by other inhomogeneities. At present, no method is available for a true positive MRI identification of infiltrating cells into inflamed tissue.
In this study, we demonstrate the feasibility and safety of imaging inflammation in mice with a “positive” contrast at high local resolution with fluorine MRI. The naturally occurring stable fluorine isotope 19F (100%) is MR active and exhibits a sensitivity close to the 1H nucleus.10,11 Because of the lack of any 19F background in the body, observed signals originating from injected 19F-containing compounds exhibit an excellent degree of specificity. The merging of recorded 19F images with simultaneously acquired, morphologically matching 1H images enables an exact anatomic localization of fluorinated substances as “hot spots.”12 In the present investigation, we used nanoparticles containing perfluorocarbons (PFCs), a family of compounds known to be biochemically inert. Some of the PFC members such as perfluorodecalin, perfluorotripropylamine, perfluorodichloroctane, and perfluorooctyl bromide (also known as perflubron) were already used in patients as artificial blood substitutes.13 However, we used perfluoro-15-crown-5 ether, a PFC in which all 20 fluorine nuclei are chemically and magnetically equivalent and thus exhibit superior properties for 19F MRI detection.14 In contrast to previous studies using 19F MRI of PFCs to track injected stem/progenitor cells after ex vivo loading,15,16 we applied emulsified PFCs systemically, resulting in an efficient and selective enrichment in circulating cells of the monocyte/macrophage system. This approach enabled us to monitor the infiltration of immunocompetent cells into inflammatory areas in an acceptable acquisition time with a spatial resolution close to the anatomic 1H image.
An expanded Methods section can be found in the online-only Data Supplement.
Preparation of the PFC Emulsion
Purified egg lecithin (E 80 S, 4% wt/wt, a generous gift from Lipoid, Ludwigshafen, Germany) was dispersed in isotonic phosphate buffer (10 mmol/L phosphate, 150 mmol/L NaCl, pH 7.4) by magnetic stirring at room temperature for 30 minutes. When lissamine rhodamine B (rhodamine dihexadecanoic phosphatidylethanolamine, Molecular Probes, Leiden, the Netherlands) was used as a fluorescent lipid marker, a lipid mixture of lecithin and rhodamine dihexadecanoic phosphatidylethanolamine (99.5/0.5 mol/mol) was dissolved in ethanol, and the solvent was subsequently removed under reduced pressure at 35°C, followed by evaporation under high vacuum. The resulting lipid film was hydrated with buffer by gentle mixing and stirring. After addition of the perfluoro-15-crown-5 ether (10% wt/wt, Fluorochem Ltd, Glossop, UK), the dispersion was pretreated with a high-performance disperser (T18 basic ULTRA TURRAX, IKA Werke GmbH & Co KG, Staufen, Germany) at 14 000 rpm for 2 minutes. The resulting crude emulsion was high-pressure homogenized (70 MPa, 10 cycles, APV Gaulin Micron Laboratory 40, APV, Unna, Germany). The formed nanoemulsion was filtered through a 0.22-μm sterile filter unit (Millex-GS, Millipore, Ireland) and stored until application at 6°C.
Animal experiments were performed in accordance with the national guidelines on animal care and were approved by the Bezirks-regierung Düsseldorf. The male mice (C57BL/6; 20 to 25 g body weight; 10 to 12 weeks of age) used in this study were bred at the Tierversuchsanlage of Heinrich-Heine-Universität (Düsseldorf, Germany). They were fed a standard chow diet and received tap water ad libitum. In total, 60 mice were investigated: blood analysis and controls with PFC and saline injections, n=30 and 10, respectively; myocardial infarction, n=12; and cerebral ischemia, n=8. Myocardial infarction was provoked by ligation of the left anterior descending coronary artery (LAD). In a separate experimental series, focal cerebral ischemia was induced by photothrombosis (see the online-only Data Supplement for a complete description of both injury models). A detailed schematic of the experimental protocols applied to the different groups is shown in online-only Data Supplement Figure I.
Mice were anesthetized with isoflurane (2.0%) with a home-built nose cone. A total volume of 100 μL (for fluorescence experiments) or up to 500 μL (for MRI) of the PFC emulsion was given intravenously through the tail vein at the time indicated in the different experiments.
Data were recorded on a Bruker DRX 9.4-T wide-bore (89-mm) nuclear MR spectrometer (Bruker, Rheinstetten, Germany) operating at frequencies of 400.13 MHz for 1H and 376.46 MHz for 19F measurements. A Bruker microimaging unit (Mini 0.5) equipped with an actively shielded 57-mm gradient set was used, and images were taken from a 30-mm birdcage resonator tunable to 1H and 19F. After acquisition of the morphological 1H images, the resonator was tuned to 19F, and anatomically matching 19F images were recorded. For superimposing the images of both nuclei, the “hot iron” color lookup table (ParaVision, Bruker) was applied to 19F images.
Mice were anesthetized with 1.5% isoflurane and were kept at 37°C. For functional cardiac analysis, 1H images of murine hearts were acquired essentially as described17 with an ECG- and respiratory-triggered fast-gradient-echo cine sequence (field of view [FOV], 30×30 mm2; matrix, 128×128; slice thickness, 1 mm). Corresponding 19F images were recorded from the same FOV using a multislice rapid acquisition with relaxation enhancement (RARE) sequence: RARE factor, 64; matrix, 64×64; slice thickness, 2 mm; averages, 256; acquisition time, 19.12 minutes. For fusion with 19F images, additional 1H data sets with a slice thickness of 2 mm were recorded. Brain images were acquired using multislice RARE sequences for both nuclei from a reduced FOV of 20×20 mm2 but otherwise unaltered geometry (see the online-only Data Supplement for a more detailed description of MRI setup, acquisition parameters, and quantification procedures).
Blood was obtained from the vena cava inferior at various times after injection of the PFC emulsion as indicated in the different experiments. Determination of serum markers of liver function was performed by the Central Laboratory of the University Hospital Düsseldorf using clinical routine protocols. In separate experiments, mononuclear cells were isolated from the blood samples by centrifugation over Histopaque density gradient (2.5-mL layers of both 1083 and 1119 [Sigma, Taufkirchen, Germany], 25 minutes, 700g at room temperature). Thereafter, either the tube was immediately transferred into the nuclear MR spectrometer for MRI (see the online-only Data Supplement for details) or the mononuclear cells were collected from the interface of the layers and analyzed by fluorescence-activated cell sorter (see the next section).
In preceding experiments with the murine macrophage cell line RAW 246.7 loaded in vitro with rhodamine-labeled PFCs (online-only Data Supplement Figure II), we confirmed that fluorescence of rhodamine bound to the coat of the PFC particles is detectable by fluorescence-activated cell sorter analysis (data not shown). Freshly prepared peripheral blood mononuclear cells were stained for flow cytometric analysis according to standard procedures (see the online-only Data Supplement for details). Cells were analyzed on a FACScalibur flow cytometer (Becton Dickinson, Franklin Lakes, NJ), and samples were gated on live cells based on forward and side scattering and by exclusion of propidium iodide–positive cells. For each sample, at least 10 000 live events were acquired and analyzed with the CellQuestPro software (Becton Dickinson, Franklin Lakes, NJ).
To avoid a dissociation of rhodamine label and markers of the initial PFC carrier as a result of downstream processes after infiltration, all organs analyzed by immunohistochemistry were excised 1 day after PFC injection. Slides were air dried, and red fluorescence images were recorded without further processing because of water solubility of rhodamine-labeled PFCs and the impossibility of adequate histological fixing of the nanoparticles. The sections selected for photographs were related to anatomic landmarks to ensure retrieval of the same area after immunohistochemistry. After processing for immunofluorescence of CD11b (see the online-only Data Supplement for a detailed description of protocols applied to heart and brain slices), cardiac and cerebral sections were again microscoped, making use of the anatomic landmarks defined in the previous session. Slides were viewed with an Olympus BX50 fluorescence microscope (Olympus, Hamburg, Germany) equipped with standard filter sets and using objectives without (before immunostaining) and with (after mounting) cover glass correction. We deliberately refrained from merging images taken before and after immunostaining because an exact overlay was hampered by unavoidable minute alterations of the dried histological slices during immunohistochemical incubation steps and subsequent mounting.
The authors had full access to and take full responsibility for the integrity of the data. All authors have read and agree to the manuscript as written.
PFC Infiltration Into the Heart After Infarction Assessed by In Vivo 19F MRI
Cardiac infarction was induced by ligation of the LAD, a procedure well known to be associated with an acute inflammatory response. Two hours after ligation, 500 μL of 10% perfluoro-15-crown-5 ether emulsion (average size, ≈130 nm; ζ potential, −31.3±1.5 mV) was applied via the tail vein (see the Methods section in the online-only Data Supplement for details on the PFC emulsion).
After surgery and application of the contrast agent, all animals (n=6) were imaged 5 times within 7 days. The infarcted area was localized by acquisition of fast-gradient-echo 1H cine movies via akinesis of the affected region within the left ventricle. Subsequently, anatomically matching 19F images were recorded for tracking of the injected PFCs. A typical example of consecutively recorded 1H and 19F images obtained 4 days after ligation of the LAD is illustrated in Figure 1A. The end-diastolic 1H image (Figure 1A, left) clearly shows the presence of ventricular dilatation and wall thinning within the infarcted area, and in the corresponding 19F image (Figure 1A, middle), a signal pattern matched the shape of the free left ventricular wall. Merging of these images (Figure 1A, right) confirms the localization of PFCs within the anterior, lateral, and posterior walls. In all animals studied, 19F signal also was detected in the adjacent chest tissue, where thoracotomy for LAD ligation was performed. Note that no background signal from other tissue is present. Repetitive measurements from day 1 after LAD ligation revealed a time-dependent accumulation of PFCs within the infarcted region as shown in a representative example in Figure 1B. End-diastolic 1H images acquired 1, 3, and 6 days after induction of myocardial infarction show the progressive left ventricular dilatation as a consequence of the insult. Merging with the matching 19F images (red) demonstrates the successive infiltration of PFCs into the affected area of the heart and the region of the chest injured by surgery. Detected 19F signals were restricted to the area near the infarcted region of the heart; at no time were infiltrating PFCs observed within the septum (see online-only Data Supplement Table I for individual data of all animals studied).
Although strong PFC signals were found in ex vivo 19F images of blood components (see below), in vivo signals from PFCs in the circulation were not detectable at all (eg, no signal within ventricular chambers; Figure 1). Even when 19F images were acquired immediately after injection, no 19F signal from the streaming blood could be observed because the pulse sequence used for 19F MRI (RARE) results in a signal void of flowing blood particles. Therefore, detected signals can be attributed unequivocally to accumulated PFCs in the tissue without contamination from 19F signals of circulating PFCs.
Uptake and Transport of PFCs by Cells of the Monocyte/Macrophage System
To characterize the mode by which PFCs can enter the injured heart tissue, murine blood samples were investigated ex vivo by 19F MRI after intravenous application of the emulsion. 19F images acquired after density gradient centrifugation of blood collected at different points after injection revealed a time-dependent accumulation of the 19F signal within the layer of the mononuclear cells (Figure 2). However, 3 days after injection, the PFCs were completely cleared from the bloodstream and were no longer detectable by 19F MRI.
To further specify the cell population containing the PFCs, experiments were performed using rhodamine-labeled PFCs. These experiments enabled us to trace the fluorescence label not only within the mononuclear blood cells by flow cytometry but also within the inflamed region by means of fluorescence microscopy of tissue sections.
After tail vein injection of fluorescently labeled PFCs and subsequent collection of blood samples, we analyzed the layer of mononuclear cells containing the PFCs as assessed by ex vivo 19F MRI (Figure 2). As shown in Figure 3A, 2 hours after injection of rhodamine-labeled PFCs, almost a fifth of the mononuclear cells were found to be positive for rhodamine, with the large majority of the labeled cells (≈80%) exhibiting the monocyte/macrophage marker CD11b (Figure 3B, top). Approximately half of this cell type was detected to be loaded with PFC particles (Figure 3C). The remaining rhodamine-positive cells were observed to be B cells (B220; Figure 3B, middle), with a marginal amount of T cells (<2%; CD3; Figure 3B, bottom). Control experiments in vitro with a murine macrophage cell line confirmed that the labeled PFCs are avidly taken up by macrophages (online-only Data Supplement Figure II).
The fate of rhodamine-labeled PFCs in cardiac tissue was investigated by histology. Microscopic survey images obtained from the same mouse shown in Figure 1A are displayed in Figure 4A. Micrographs show a pattern of rhodamine fluorescence that is similar to the signal distribution in the corresponding 19F MRI acquired immediately before organ excision (Figure 1A, right). The main fluorescence signals were located exclusively within the injured area. No rhodamine fluorescence was observed in the septum and necrotic areas, as confirmed by staining with triphenyltetrazolium chloride (data not shown).
Immunostaining of tissue sections for the monocyte/macrophage marker CD11b with FITC revealed some colocalization of fluorescence patterns for cells of the monocyte/macrophage system (green) and for rhodamine-labeled PFCs (red), as shown in Figure 4B. It should be noted, however, that technical reasons precluded a precise merge of the differently labeled sections. Because of the water solubility of the rhodamine-labeled PFCs, red fluorescence images had to be taken before immunohistochemistry for CD11b and required careful selection of anatomic landmarks to ensure retrieval of the same area.
PFC Infiltration Into the Brain After Focal Cerebral Ischemia
In another set of experiments, focal cerebral ischemia was chosen as an additional model of acute inflammation. After ischemia was induced by photothrombosis, all animals (n=4) were imaged at regular intervals up to 4 weeks after surgery. In RARE 1H images, the ischemic region appeared initially as a bright area (Figure 5A, top left), and the corresponding 19F images clearly show infiltration of PFCs into the border zone of the infarct, which was detected at the earliest at day 4 after photothrombosis. 19F signal also was transiently observed supracranially at the location of skin incision (Figure 5A, bottom left). Characteristic 1H and 19F images (Figure 5A) acquired from an individual mouse 7, 9, 12, and 19 days after focal cerebral ischemia was induced definitely show movement of the PFCs with the rim of the shrinking infarct over time (see online-only Data Supplement Table II for individual data of all animals studied).
To support the notion that PFCs were carried into the ischemic region by monocytes/macrophages, experiments with rhodamine-labeled PFCs were again conducted (n=4). Microscopic survey images after FITC immunostaining for CD11b exhibited a pattern of green fluorescence comparable to that observed for the 19F signal in the preceding MR experiment (Figure 5B). Furthermore, comparison of red and green fluorescence at large magnification indicated colocalization of PFCs and CD11b-positive cells (online-only Data Supplement Figure III).
Detection Threshold and Absolute Quantification
The sensitivity of our present approach can be estimated from Figure 2 by correlating the number of cells contained in the layer of the mononuclear cells with the signal-to-noise ratio in the corresponding areas of 19F images. Two days after PFC injection, the mean signal-to-noise ratio within this layer was determined to be 24 at a voxel size of 0.44 μL (FOV, 30×30 mm2; matrix, 64×64; slice thickness, 2 mm). The mononuclear cell layer contained 1.16×106 cells distributed vertically over ≈1 mm and horizontally over the inner diameter of the tube (14 mm as derived from axial 1H images), which results in a cell number of ≈3300 per 19F MR voxel within this layer. Assuming a minimal signal-to-noise ratio of 3 as the detection threshold, as little as ≈400 cells are expected to be visible by MRI under these conditions. Taking into account that only a fraction of the mononuclear cells are loaded with PFCs (Figure 3), the detection limit may be even lower.
A similar conclusion was reached in a separate set of experiments in which RAW 264.7 macrophages were incubated ex vivo with PFCs under in vivo–like conditions and analyzed by 19F MRI after immobilization in agarose (for details, see the Methods section of the online-only Data Supplement). Stepwise dilution of PFC-loaded macrophages revealed that <200 cells were detectable within a voxel of 0.44 μL (online-only Data Supplement Figure IV). By calibration of the absolute 19F signal intensities with PFC concentration standards (R2=0.99892; online-only Data Supplement Figure V), the average PFC loading per cell was calculated to be 0.73±0.19 pmol (n=8). Assuming a similar uptake of PFCs in vivo, the number of PFC-containing cells within ischemic areas can be quantified by interpolation from 19F signal intensities of the affected regions (online-only Data Supplement Tables III and IV).
Control Experiments After PFC Injection
Without further intervention, at no time were 19F signals observed within the heart or the brain. However, 19F images showed a distinct signal in the spleen 1 day after injection of the PFC emulsion and a weaker signal in the liver that increased up to days 2 to 3, reaching an intensity similar to the signal from the spleen (online-only Data Supplement Figure VI). Interestingly, at the same time, additional signals regularly appeared in lymph nodes in the area of the upper thorax and the head and became clearly visible, as shown in Figure 6. The signals in the liver persisted for several months, but no adverse effects of the PFCs were observed in these animals, and serum markers of liver function were comparable to those of saline-treated animals (eg, the ratio of glutamic oxaloacetic transaminase to glutamic pyruvic transaminase was 2.53±1.01 [PFC, n=8] versus 2.26±0.57 [saline, n=7]).
The present study describes a novel approach for visualizing local inflammatory processes by 19F MRI using in vivo tagging of circulating monocytes/macrophages with biochemically inert PFCs. Our results show that intravenous application of emulsified PFCs after local inflammation is provoked by acute cardiac or cerebral ischemia results in the accumulation of 19F-labeled cells within injured areas. Detection of infiltrating monocytes/macrophages by 19F MRI at a field strength of 9.4 T is feasible in the mouse at an acceptable acquisition time (20 minutes) with a resolution close to the anatomic 1H image. Therefore, PFCs can serve as a “positive” contrast agent for inflammatory processes (Figure 7), exhibiting a high degree of specificity because of the lack of any 19F background.
Compared with previous 1H MRI approaches for visualizing the infiltration of immunocompetent cells into inflamed areas by use of superparamagnetic iron oxide particles, the method presented here has the advantage of a direct positive detection of the tagging agent and therefore has the potential to work also in tissues that generally appear very dark in 1H MRI such as the lungs. Although techniques have recently been described to image superparamagnetic iron oxide particles with a bright contrast,18 the physical basis of detection is still the disturbance of the regional magnetic field by these particles. Therefore, it often remains difficult to unequivocally assign alterations in local contrast to accumulating superparamagnetic iron oxide particles. Furthermore, iron-based contrast agents are readily metabolized, whereas the fluorinated crown ether used in this study is biologically inert and cannot easily be degraded. The reason is the very stable C-F bond and the dense electron cloud of the fluorine atom, which results in a protective sheath.19 Experimentally, this provides the unique possibility for specifically and permanently labeling circulating monocytes/macrophages and following their fate within the body. It is of note that an absolute quantification of the observed signals is feasible (online-only Data Supplement Figures IV and V and Tables I through IV), which can be translated into the number of infiltrating immunocompetent cells.
Recent 19F MRI tracking studies of cells loaded ex vivo with PFCs and subsequently injected into mice either required long acquisition times (up to 3 hours)15 or were limited in spatial resolution (voxel size, 26 μL16 compared with 0.2 to 0.4 μL in the present work). In the latter investigation, the limit of detection was reported to be ≈6000 labeled cells. The substantial higher sensitivity observed in our study is most likely due to the fact that the monocyte/macrophage system in vivo more effectively takes up the injected PFCs compared with stem/progenitor cells incubated ex vivo. Labeling of ≈50% of the total monocyte/macrophage cell population (Figure 3C) raises a question about function and integrity of the loaded cells. Previous studies revealed that perfluoro-15-crown-5 ether labeling had no significant effect on cell proliferation, function, or maturation.15,16 It seems likely that this also applies to the monocyte/macrophage system because both the time course of accumulation and the localization of PFC-containing monocytes/macrophages within ischemic areas are in good agreement with previous data on myocardial20,21 and cerebral infarction,6,9 suggesting unaltered infiltration kinetics and distribution of loaded cells. Furthermore, we did not observe any adverse effects on the animals after PFC injection, and no changes were noted in the release of liver enzymes, although this organ is a major site of PFC accumulation.
An interesting observation of this study was that lymph nodes are clearly delineated in 19F images. Although the bulk of PFCs were found in CD11b-positive cells, it should be noted that ≈20% of the injected particles were taken up by B cells (Figure 3B). However, it is difficult to decide whether the labeling of lymph nodes is due to trapping of labeled B cells or to the accumulation of PFCs in resident macrophages. Therefore, we cannot exclude the possibility that local PFC deposition also may occur via an alternative pathway; nanoparticles carried by the lymphatic flow to the sites of inflammation could have been taken up by immunocompetent cells already present at the sites of injury before PFC injection.
PFCs such as perflubron have been evaluated clinically as an artificial blood substitute. In these early studies, it was observed that perflubron is phagocytized by the reticuloendothelial system.22,23 In principle, perflubron should thus work as well as perfluoro-15-crown-5 ether, used in the present study, for 19F imaging of inflammatory processes. Perflubron has the additional advantage that it is readily cleared from the body through exhalation by the lungs within 1 week.24 Viewed from the MRI side, perflubron has a lower MRI sensitivity caused by signal splitting resulting from magnetically different 19F nuclei. However, this problem can be overcome by dedicated detection methods,25 the incorporation of gadolinium into the PFC droplets,26 or the preparation of emulsions with a higher PFC content. Furthermore, it should be noted that the voxel size in cardiac MR diagnostics at 3 T is in the range of 2 to 30 μL, whereas it was only 0.2 to 0.4 μL in our study at 9.4 T, which translates into a substantial sensitivity increase in the clinical setting.
We thank Jutta Ziemann, Barbara Emde, and Sabine Hamm for excellent technical assistance, as well as Andreas Neub (Freiburg) for the cryoTEM studies.
Sources of Funding
This study was supported by Sonderforschungsbereich 612, subproject Z2 (Drs Flögel and Schrader), Deutsche Forschungsgemeinschaft grants SCHR154/9 (Drs Flögel, Jacoby, and Schrader) and JA690/5–2 (Dr Jander), and National Institutes of Health grant P01 HL073361 (Drs Flögel and Schrader).
Tawakol A, Migrino RQ, Bashian GG, Bedri S, Vermylen D, Cury RC, Yates D, LaMuraglia GM, Furie K, Houser S, Gewirtz H, Muller JE, Brady TJ, Fischman AJ. In vivo 18F-fluorodeoxyglucose positron emission tomography imaging provides a noninvasive measure of carotid plaque inflammation in patients. J Am Coll Cardiol. 2006; 48: 1818–1824.
Jander S, Schroeter M, Saleh A. Imaging inflammation in acute brain ischemia. Stroke. 2007; 38: 642–645.
Holland GN, Bottomley PA, Hinshaw WS. 19F magnetic resonance imaging. J Magn Reson. 1977; 28: 133–136.
Partlow KC, Chen J, Brant JA, Neubauer AM, Meyerrose TE, Creer MH, Nolta JA, Caruthers SD, Lanza GM, Wickline SA. 19F magnetic resonance imaging for stem/progenitor cell tracking with multiple unique perfluorocarbon nanobeacons. FASEB J. 2007; 21: 1647–1654.
Currently, neither a clinically useful method to assess local inflammatory processes associated with the risk of plaque rupture nor a robust imaging method that provides information about local activity of inflammation (which plays a crucial role in various cardiovascular disease states such as ischemia/reperfusion, myocarditis, transplant rejection, or stroke) is available. In the present study, we demonstrate in murine models of myocardial and cerebral ischemia that nanoemulsions of perfluorocarbons can be used to precisely visualize localized inflammatory processes as hot spots by simultaneous acquisition of morphologically matching proton (1H) and fluorine (19F) magnetic resonance images. Injected perfluorocarbons are phagocytized primarily by monocytes/macrophages, resulting in 19F magnetic resonance imaging intensity signals along the border of infarcted areas as a result of progressive infiltration of the labeled immunocompetent cells. Because of the lack of any 19F background in the body, observed signals are robust and exhibit an excellent degree of specificity. Perfluorocarbons are biologically inert and have been shown to be nontoxic in humans. Thus, 19F MRI has the potential to be clinically applicable as a new diagnostic modality not only for acute but also for chronic inflammatory processes such as plaques in atherosclerosis.
The online-only Data Supplement, which consists of Methods, tables, and figures, can be found with this article at http://circ. ahajournals.org/cgi/content/full/CIRCULATIONAHA.107.737890/DC1.