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* Department of Medicine, Washington University in St. Louis, School of Medicine, St. Louis, Missouri, USA;
Departments of Pathology and Laboratory Medicine, Saint Louis University School of Medicine, St. Louis, Missouri, USA; and
Philips Medical Systems, Andover, Massachusetts, USA
1Correspondence: Washington University School of Medicine, Campus Box 8215, 660 South Euclid Ave., St. Louis, MO, USA 63110. E-mail: saw{at}howdy.wustl.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: endothelial progenitor cells • nanoparticles • contrast agent
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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2
3
4)
. Current labeling methods typically employ iron oxide nanoparticles to produce "negative" (i.e., dark) contrast effects in proton (1H) images based on susceptibility disturbances to the local magnetic fields surrounding targeted cells, or alternatively "positive" (i.e., bright) contrast based on off-resonance techniques (e.g., IRON) (5)
. In general, these particles are not readily internalized by stem cells (6)
and require adjunctive transfection techniques such as cationic agents (7)
or mechanical approaches such as electroporation (8
, 9)
. Although these methods can achieve rapid labeling and appear relatively nontoxic in acute settings, the opportunity for simultaneously tracking multiple cell types each bearing a unique MR signature is beyond the range of these agents. Such a goal could be important for identifying and tracking the myriad cell types that are involved cooperatively in many pathological processes and in regenerative therapeutics.
Accordingly, on the basis of prior work from our laboratory and others (10
11
12)
, we have developed a liquid perfluorocarbon nanoparticle that is endocytosed readily by stem/progenitor cells without the need for conjunctive transfection approaches, and that can employ multiple unique fluorocarbon labels for fluorine (19F) MR imaging and spectroscopy. A major advantage of the 19F molecular imaging approach previously demonstrated for thrombus detection is the lack of any measurable tissue background signal from endogenous fluorine (10)
, which creates superior contrast-to-noise features and permits definitive localization of the uniquely labeled cells, as compared to the iron oxide proton-based agents. Finally, 19F signals are intrinsically quantitative based on localized spectroscopy and have the potential to report the actual concentration of particles (and thus stem cells) in a given region. In this work, we demonstrate the use of these fluorine nanoparticles for sensitive detection of stem/progenitor cells in vivo after systemic injection and for rapid imaging in mouse skeletal muscle in situ at both 11.7T and 1.5T, which is the most common clinical imaging field strength.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Briefly, the emulsions comprised 20% (v/v) perfluorocarbon (PFC) either perfluorooctylbromide (PFOB) or perfluoro-15-crown-5 ether (CE), 1.5% (w/v) of a surfactant/lipid comixture, and 1.7% (w/v) glycerin in distilled, deionized water. Fluorescent nanoparticles additionally contained fluorescent-conjugated phospholipids either 2.05 mol% of nitrobenzoxadiazole (NBD) (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2–1,3-benzoxadiazol-4-yl)) or 0.135 mol% of rhodamine (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl)) (Avanti Polar Lipids, Inc., Alabaster, AL, USA) in the surfactant layer. The mixture of surfactant components, PFC, and water was blended and then emulsified at 20,000 PSI for 4 min in an ice bath (S110 Microfluidics emulsifier, Microfluidics, Newton, MA). Particle size analysis by laser light scattering (Malvern Instruments, Malvern, Worcestershire, UK) measured sizes of 224 nm (polydispersity=0.350) and 233 nm (polydispersity=0.173) for PFOB and CE formulations, respectively.
Isolation and culture of mononuclear cells
Human umbilical cord blood was obtained from the Cardinal Glennon Children’s Hospital (St. Louis, MO, USA) and was used in accordance with ethical guidelines and accepted human studies protocols at Washington University School of Medicine. Mononuclear cells (MNCs) were isolated by density gradient centrifugation with Ficoll-Paque Plus (Amersham Biosciences, Piscataway, NJ, USA). MNCs were plated at concentrations of 5 x 105 cells/cm2 on fibronectin-coated plates (RetroNectin; Takara, Otsu, Japan) and grown in modified endothelial growth media (Clonetics EGM-2+20% FBS; Cambrex, East Rutherford, NJ, USA) designed to promote differentiation along the endothelial lineage. At day 2, nonadherent cells were removed and transferred to fresh fibronectin-coated plates (14
, 15)
.
Immunophenotyping by flow cytometry
Before and after incubation with fluorescently labeled nanoparticles, cells were removed with cell dissociation solution (Sigma, St. Louis, MO, USA) and analyzed for cell surface markers with an array of monoclonal antibodies in appropriate combinations, including CD34-PE (phycoerythrin)-Cy7, human CD45-APC (allophycocyanin)or FITC, human CD31-PE, Tie-2-PE or APC (Becton Dickinson, San Jose, CA, USA), or CD133-APC or PE (Miltenyi Biotechnology, Bergish, Gladbach, Germany). Immunostained samples were analyzed on a Coulter FC-500 flow cytometer with RXP analysis software (Beckman-Coulter, Miami, FL, USA).
Cell loading and assessment
After 7–14 days in culture, cells were incubated for 12 h with a 30-pM concentration of fluorescently labeled PFOB or CE nanoparticles. The nanoparticle concentration was estimated from nominal particle size determined by laser light scattering and the total volume of perfluorocarbon incorporated into the formulation (10)
. No conjunctive transfection methods were used to label the cells. After incubation, cell pellets were prepared by removing free nanoparticles with PBS washing, detaching adherent cells from the surface, and preserving samples with 2% paraformaldehyde fixation for 30 min. Cell number was determined using a hemocytometer.
For confocal imaging, fixed cells were placed in #1.5 glass-bottom culture dishes (Bioptechs Inc., Butler, PA, USA). Fluorescence imaging of cell sections was conducted with a confocal microscope (Zeiss Meta 510, Thornwood, NY, USA) using standard filter sets. The location of nanoparticles with respect to the cell was determined with simultaneous differential interference contrast (DIC) imaging.
Cell survival after loading was determined by trypan blue exclusion. Cells were removed with trypsin, resuspended in PBS, and diluted 1:1 with 0.4% trypan blue (Sigma, St. Louis, MO). For a positive control, cells were heated at 45°C for 15 min. The number of viable and nonviable cells was counted using a hemocytometer. The percentage of trypan blue-positive cells was used to calculate cell survival.
To characterize functional performance of cells before and after PFC nanoparticle loading, cells were incubated with acetylated LDL (15 µg/mL for 3 h) and stained after fixation with Ulex europaeus agglutinin(UEA)-1 (10 µg/mL for 1 h), both combined denote cells of endothelial lineage (16
, 17)
.
Intravenous injection and assessment
Tumor-bearing mice were utilized to determine whether labeled cells injected intravenously (i.v.) could be detected in tissues of interest and to confirm prolonged functionality of PFC-labeled cells, according to their ability to colocalize to vascular tissues. Before injection cells were colabeled with rhodamine-labeled CE nanoparticles and AlexaFluor 488-labeled acLDL (Invitrogen, Carlsbad, CA), as described previously. Six million labeled cells in a final volume of 50 µl (12x104 cells/µl) were injected i.v. by jugular vein catheterization into an athymic mouse bearing a 10-day-old implanted tumor (human MDA-435 breast adenocarcinoma). After allowing 5 days for migration, tissues were collected and frozen in optimal cutting temperature (OCT) medium for histological analysis. Frozen sections (Leica Microsystems, Inc., Bannockburn, IL) were stained with 4',6'-diamidino-2-phenylidole (DAPI) (Vector Laboratories, Burlingame, CA) and imaged at high power (x400) with a Nikon Eclipse microscope. All animal procedures were approved and in accordance with guidelines of the Animal Studies Committee at Washington University Medical School.
Spectroscopy and imaging at 11.7T
19F MR spectroscopy (MRS) of labeled cells was performed on a Varian 11.7T scanner using a custom-designed 0.5 cm 4-turn solenoid RF coil. Labeled cells were contained within a centrifuge tube and analyzed together with an internal standard of PFOB or CE nanoparticle emulsions provided by inserting the cell tube into a slightly larger tube containing the emulsion standard; all samples were in equilibrium with ambient air. 19F MRS (number of averages=64, 90° flip angle, acquisition time: 2 min) of cells was performed for quantitative evaluation of intracellular labeling of nanoparticles using a spin-echo sequence. To ensure minimal effects from pO2 during quantification, a 2 s TR (
3T1, where the measured 19F T1800 ms for intracellular CE nanoparticles) was used to ensure the recovery of 19F magnetization in each repetition. For quantification, an external standard was made containing a mixture of CE and PFOB emulsion in known amounts, where the ratio was determined between the CE peak and PFOB peak (–80 ppm) areas with similar T1 and T2 values (10)
. For cell pellets, a 5-µl internal standard was provided containing either 100% PFOB or 10% CE emulsion for CE or PFOB-labeled cells, respectively.
The detection limit of CE- and PFOB-labeled cells was determined using 19F spectroscopy and imaging. For sensitivity measurements, tubes containing between 2x103-32x103 CE labeled cells and 5x103-16x104 PFOB labeled cells were analyzed by 19F MRS, as described above. Cell pellets with 19F signal intensity greater than the mean+3SD of noise were considered detectable. For 19F imaging, 860,000 CE nanoparticle-labeled cells condensed into a pellet were imaged using a gradient echo sequence (image matrix: 64x64, FOV: 1x1 cm2, slice thickness: 2 mm, TR: 50 ms, TE: 3 ms, number of averages=64, 
7 min total acquisition time). Pixels within the pellet with 19F signal intensity greater than the mean+3SD of background noise were considered detectable. Since the 19F signal intensity is linearly related to the amount of 19F molecules (10)
, the 19F imaging detection limit of CE nanoparticle-labeled cells was calculated:
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4 min total acquisition time; and 1H imaging parameters—image matrix: 256x256, FOV 6x6 cm2, slice thickness: 2 mm, TR: 300 ms, TE: 3 ms, number of averages=2,2.5 min acquisition time). The spectra of CE and PFOB were used to define the offset frequency of RF output for 19F imaging. Prior to sacrifice of tumor-bearing mice, the liver was assessed with 19F spectroscopy to determine whether cells naturally localized to a functioning tissue could be detected. For in vivo imaging, 5 days after injection, the mouse was anesthetized with ketamine (87 mg/kg) and xylazine (12 mg/kg). A double-tuned surface coil (1.5 cm diameter) was placed over the right abdomen of the mouse. Initially, a 1H spin-echo scout image was acquired to confirm that the liver was located in the field-of-view of the surface coil. Subsequently, the coil was retuned for 19F spectroscopy (acquisition time: 2 min, TR: 1s, number-of-averages: 128).
Imaging at 1.5T
For in situ imaging, 4 million fixed and labeled cells suspended in 50 µl of PBS were injected into the right hind limb of a euthanized mouse and imaged with a 5-cm square surface coil tuned to 60.1 MHz to transmit and receive. A projection image of the cells was generated with a true steady-state free precession (i.e., "balanced" FFE) pulse sequence (TE 5 ms, TR 10 ms, 512 signal averages, 2.5x2.5 mm reconstructed in-plane resolution, 60 degree flip angle, 35-mm slice thickness, 7 min total scan time) (18)
. Matching 1H images were acquired for comparison using the quadrature body coil for transmission and 4-cm diameter surface coil for receipt (3D T1-weighted turbo spin echo sequence, TE 15 ms, TR 363 ms, 7 signal averages, 0.2x0.2 mm reconstructed in-plane resolution, 90 degree flip angle, 2-mm slice thickness, 10 slices, 11 min total scan time).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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A). Cell sectioning through the nucleus confirms that nanoparticles were located in the cell cytoplasm and not simply bound to the cell plasma membrane.
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Immunophenotyping of progenitor cells
To characterize the cell population derived from the combined influence of the endothelial conditioning media and elapsed time, we probed cells with monoclonal antibodies for characteristic stem and progenitor markers and analyzed expression with flow cytometry (19
20
21)
. The total population expressed hematopoietic, progenitor, and endothelial markers, including CD34, CD133, Tie-2, and CD31 (see Table 1
for averages and Supplementary Fig. 1 for representative dotplots). A significant part of the population (57±7) coexpressed the hematopoietic marker CD34 and the progenitor marker CD133, and a smaller population (3%) was CD34–CD133+. Coexpression of CD133 and the endothelial marker Tie-2 occurred in 63 ± 10% of cells with a small population (4%) of CD133+Tie-2– cells. Probing for the hematopoietic/leukocyte marker CD45 revealed a small portion (up to 5%) of CD133+CD45– cells, although CD45 was manifest in 78 ± 2% of the overall population. The similar population size (all 70%) indicates significant overlap between expression of CD34, CD133, Tie-2, CD31, and CD45, in which the presence of negative populations denotes the specificity of staining.
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To quantify the labeling efficiency and define the population of cells loaded with nanoparticles, we immunolabeled cells after nanoparticle incubation. Approximately 60 ± 5% of cells in the population contained nanoparticles and also expressed CD34, CD133, CD31, Tie-2, and CD45 (Table 1)
. Although we observed significant overlap between nanoparticle labeling and expression of CD133 and Tie-2, a small portion of CD133+ (2%) and Tie-2+ (2%) cells remained unlabeled. Incubation with nanoparticles did not alter the overall expression of probed markers within the population (Supplemental Fig. 1).
Lack of bioeffects of PFC nanoparticle loading on cell viability and function in vitro and in vivo
To confirm the safety of the labeling procedure, cell survival after exposure to either PFOB or CE nanoparticles was quantified after 12 h of incubation by trypan blue exclusion. We found high cell survival (90%) after 12 h with no significant difference from controls (Fig. 1B
). For positive controls, cells exposed to high temperature manifested a substantial loss of cell viability (>35% death). Accordingly, neither PFOB nor CE nanoparticles exerted any untoward effect on cell viability.
To evaluate cell function after nanoparticle loading, we incubated cells for 12 h with nonfluorescent nanoparticles and subsequently incubated with DiI-acetylated-LDL (acLDL), which is endocytosed independently through the cell’s surface scavenger receptors (17)
. We subsequently stained cells with FITC-labeled UEA-1 a lectin that binds the sugar fucose (16)
, which is used to denote cells of endothelial lineage (double positive acLDL+/fucose+) (15
, 19
, 22)
. Analysis of function with flow cytometry revealed a large population (84±5) that could bind and internalize acLDL (Fig. 1C
), despite being loaded previously with nanoparticles. In addition, acLDL+ cells stained positive for varied levels of UEA-1 (67±1), indicating that the cell population exhibited functional and structural features similar to cells of endothelial lineage. Unlabeled cells displayed similar staining characteristics, indicating no untoward effect of nanoparticles on cell function (data not shown).
An athymic mouse tumor model (MDA-435 line) was used to elucidate the in vivo functionality of PFC labeled stem/progenitor cells. The functionality of similar cells has previously been demonstrated in athymic mouse models where human cells were able to home, integrate, and function in vasculature at ischemic and tumor sites (23
24
25)
. Stem/progenitor cells colabeled with CE nanoparticles and AlexaFluor488-acLDL were i.v. injected 10 days after tumor implantation. After allowing an additional 5 days for cell migration (15 days old tumor), the location of transplanted cells was investigated in the vasculature along the tumor periphery with confocal microscopy. We observed that PFC labeled cells successfully homed to the tumor vasculature (Fig. 1D
), where cells within the vessel contained both CE nanoparticles (red) and acLDL (green), indicating the presence of intact cells (see Supplemental Video 1 for 3D projection). The ability of PFC-labeled cells to home to neovasculature indicates that these cells maintain preserved in vivo functionality.
Sensitive 11.7T spectroscopy and imaging of labeled cells
For 11.7T spectroscopy and imaging, cells were loaded with CE or PFOB nanoparticles, preserved with paraformaldehyde fixation, and condensed into a pellet by centrifugation. To quantify the amount of perfluorocarbon incorporated into the cells, known amounts of PFOB or CE nanoparticles were utilized as standards for CE- or PFOB-loaded cells, respectively. MR spectroscopy indicates that the spectrum of CE comprises a single 19F peak (Fig. 2
A, arrow) (26)
, whereas the spectrum of PFOB is characterized by multiple peaks that span 60 ppm in the frequency domain (Fig. 2A
, all other peaks). The spectra were readily detectable (2 min acquisition time) with little background noise; in fact no 19F signal was detected from control cells (data not shown). We observed a narrower line width from intracellular PFC compared to the external standard, which could be attributed to a difference in volume between pelleted cells and the external standard. The ratio between the PFOB peaks at –80 and –62 ppm varies among acquired spectra. Contributing factors include saturation effects, RF offset, or sweep width of RF excitation pulse. To minimize inaccuracies, we utilized the peaks with the most similar T1 and T2 values (i.e., the PFOB peak at –80 ppm and the single CE peak), which were measured to have a 1:7 relative signal. Quantification of the amount of perfluorocarbon in the cell pellets after 12 h of incubation indicated fluorine levels of up to 3 pmol of perfluorocarbon per cell (Fig. 2B
), where long TR times were used to minimize effect of pO2. We observed higher cellular levels of PFOB vs. CE nanoparticles (Fig. 2B
: 2.90±0.048 vs. 0.515±0.009 PFC (pmol)/cell, respectively). These levels of PFC per cell were sufficient to permit highly sensitive detection with 19F MR spectroscopy of labeled cells, resulting in distinct 19F signals generating a signal-to-noise ratio (SNR) 
3 for as few as 2000 CE-labeled and 10,000 PFOB-labeled cells (data not shown). To demonstrate the sensitivity of imaging labeled cells, 19F MRI of a cell pellet containing 860,000 CE nanoparticle-labeled cells showed that as few as 6100 cells in a single voxel could be detected within 7 min (Supplemental Fig. 2).
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In situ localization of labeled cells with 11.7T and 1.5T
To determine the feasibility of detecting cells at specific tissue sites after local delivery, 1 million cells (2x104/µl) loaded with CE or PFOB nanoparticles were injected into mouse thigh skeletal muscle (Fig. 3
A). At 11.7T, the CE or PFOB nanoparticle-labeled cells were imaged (4 min scan time) by selecting the frequency of the RF excitation pulse based on the PFOB (Fig. 3B
) or CE spectrum (Fig. 3C
). Specifically, for 19F MR imaging of PFOB-labeled cells, the center frequency of the PFOB peak (at approximately –120 ppm as shown in Fig. 2A
) was selected as the RF output frequency; whereas CE imaging was performed on the single peak. Overlaying the fluorine images atop a conventional matched proton image reveals the 19F PFOB and CE signals are located in the left and right mouse thigh, respectively (Fig. 2D
).
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For imaging at 1.5 T, a similar in situ injection of 4 million CE-labeled cells (8x104/µl) produced a strong fluorine signal (Fig. 3E
) in 7 min. When overlaid atop a proton image, the fluorine signal is located in the mouse thigh just caudal to the bright signal from the gut (Fig. 3F
). These images suggest that nanoparticle internalization by cells yielded sufficient local concentrations of 19F for imaging at either 11.7T or 1.5T in short 
7 min image acquisition times. In addition, all acquired images evidenced essentially no fluorine background signal and little noise, which permits definitive assignment of the fluorine signal to the labeled cells.
In vivo detection of labeled cells with 11.7T spectroscopy
To investigate the feasibility of detecting cells dispersed within tissue after systemic delivery, 6 million cells (12x104/µl) colabeled with CE nanoparticles and acetylated LDL were injected i.v. Histological analysis revealed colabeled cells were present in the liver (Fig. 4
A), where overlay of signal due to acLDL (green), CE nanoparticles (red), and nuclei (blue) confirm the presence of intact cells (indicated by arrows). Localized MR spectroscopy of the liver reveals a 19F peak located at the chemical shift of CE nanoparticles (SNR=4.5), confirming the presence of labeled cells and demonstrating sensitivity of detection (Fig. 4B
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, 18)
. Ahrens et al. recently followed this work with a novel application of similar CE nanoparticles for labeling and tracking dendritic cells in vivo (12)
. Ahrens et al. imaged at 11.7 T for 3 h to acquire full image data sets for local injections of 4–8 million cells labeled with conjunctive approaches, such as cationic lipids to facilitate ingestion. We now extend these results to rapid imaging of labeled stem/progenitor cells at both research and clinical field strengths with multiple unique perfluorocarbon labels, and we illustrate spectroscopic methods for quantification of the cellular uptake and content of nanoparticles.
In this study, we observed that at least two distinct types of PFC-based nanoparticles were readily internalized by stem/progenitor cells without the need for conjunctive transfection strategies. Such methods used in conjunction with iron oxide particles can potentially lead to significant losses in cell viability (27)
. We achieved intracellular PFC levels of up to 3 pmol in 12 h without the aid of any additional agents (Fig. 2B
) while preserving viability (Fig. 1B
) and in vivo functionality (Fig. 1D
). The resultant intracellular fluorine levels were readily detectible at both 11.7 T and 1.5 T, considering the relatively low number of cells utilized and the reasonably short imaging times (under 7 min).
The usefulness of a tracking method depends on the sensitivity of the procedure. Because most cell tracking methods are not quantitative, the exact number of cells that accumulate at therapeutic sites is not well defined. Aisher et al. estimated that 3 x 104 endothelial progenitor cells (EPCs) labeled with 111In-oxine localize to infarcted rat heart muscle (28)
. In our study, we observed the ability to detect as few as 2000 CE-labeled and 10,000 PFOB-labeled cells with 19F MR spectroscopy, and 6000 CE-labeled cells with 19F MRI in vitro. Although this study was not designed to elucidate the minimum number of cells that could be detected under diverse experimental conditions, we demonstrate the feasibility of utilizing this method for multiple applications in the future. Given that clinical field strengths continue to advance beyond even 7 T, we feel the sensitivity should continue to improve over time.
Other laboratories have utilized special cell culture conditions to generate EPCs from cord blood (15
, 22
, 29
, 30)
. We noted that our cells expressed characteristic markers of EPCs such as CD34, CD133, CD31, and Tie-2, yet still expressed CD45 (Table 1)
. In vitro functional tests showed these cells readily internalized acLDL and stained positive for UEA-1, which are the functional compositional characteristics of endothelial lineage (15
, 19
, 22)
. The distribution of UEA-1 expression levels observed within acLDL+ cells (Fig. 1C
) suggests that these are endothelial progenitor cells similar to those created by others under similar culture conditions (22)
. We surmise that adjustments in culture conditions or more prolonged exposure to culture media could have been sufficient for cells to down-regulate CD45 expression. However, for the purpose of our study, we chose to generalize our cells as stem/progenitor cells. Our experience reveals the difficulty in precisely delineating these cells according to in vitro characterization of marker expression and/or functional type (31)
. Clearly, the most robust approach for defining stem/progenitor cells is by their function in vivo, such as homing to tumor vasculature (Fig. 1D
). The rapid, unequivocal, multiple labeling, and quantitative imaging methods developed in this study, such as labeling cells with multiple distinguishable PFCs (Fig. 1A
) and acquiring individual PFC images (Fig. 3, B, C
), should help to address this issue and aid in functional determination of individual and multiple cell types in the future.
The current study presents the first evidence confirming the feasibility for 19F imaging of both PFOB and CE nanoparticle-labeled stem/progenitor cells. PFC-labeled cells were detectable rapidly not only in vitro, but also in liver tissue in vivo with 11.7T spectroscopy and at potential target sites in situ using both experimental (11.7 T) and clinical (1.5 T) field strengths. The ability to quantify fluorine levels present and to differentiate distinct PFC signals with MR spectroscopy could provide additional advantages such as the type and number of cells accumulating at target sites. A variety of perfluorocarbons might serve as core materials for uniquely labeled nanoparticles (e.g., PFOB, CE, perfluorodecalin, perfluoro-1,8-dicholorooctane, tris(trifluoromethyl) benzene), which could be simultaneously tracked in different stem cells (32
, 33)
. In conclusion, PFC nanoparticles provide an unequivocal and unique MR signature for stem/progenitor cells, do not require conjunctive procedures to achieve cellular uptake, enable spatial cell localization with 19F MR imaging, and provide opportunity for quantification of stem cell concentration via 19F MR spectroscopy.
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ACKNOWLEDGMENTS |
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Received for publication June 30, 2006. Accepted for publication December 25, 2006.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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