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Molecular Imaging |
1 From the Dept of Radiology (H.E.D.L., S.M., T.M.L., E.J.R.), Institute of Pathology (M.R., G.P., J.S.), IIIrd Clinic of Internal Medicine, Laboratory of Stem Cell Physiology (R.B., C.P., R.A.J.O.), and Dept of Gynecology (H.E.D.L., M.R., G.P., S.M., R.B., J.S., T.M.L., C.P., E.J.R., R.A.J.O.), Technical University, Munich, Germany; Dept of Gynecology and Obstetrics, NeuPerlach Hospital, Munich, Germany (G.D.); and Guerbet Group, Paris, France (C.C.). From the 2003 RSNA annual meeting. Received Aug 6, 2003; revision requested Oct 22; final revision received Apr 3, 2004; accepted May 3. Supported by German Research Foundation grants DA 529/1–1, the StemMat project (Bavarian Government of Health, Food, and Consumer Interests), and the Dr Ingrid Leonhard Lorenz Foundation of the Technical University of Munich. Address correspondence to H.E.D.L., Dept of Radiology, UCSF Medical Center, University of California San Francisco, 513 Parnassus Ave, San Francisco, CA 94143 (e-mail: daldrup@radiology.ucsf.edu).
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODS RESULTS DISCUSSIONREFERENCES |
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MATERIALS AND METHODS: This study was approved by the ethical committee, and all women had given consent to donate umbilical cord blood for research. Twenty athymic female Balb/c mice underwent MR imaging before and 1, 4, 24, and 48 hours after intravenous injection of (1–3) x 107 human hematopoietic progenitor cells labeled with the superparamagnetic iron oxide particles ferumoxides through simple incubation (n = 10) or P7228 through lipofection (n = 10). Fifteen female Balb/c control mice were examined after intravenous injection of the pure contrast agents (n = 6 for both probes) or nonlabeled cells (n = 3). Signal intensities of liver, spleen, and bone marrow on MR images obtained before and after injection were measured and compared for significant differences by using the t test. MR imaging data were compared with the results of immunostaining against human CD31+ cells and against the coating of the contrast agents; these results served as the standard of reference.
RESULTS: Ferumoxides was internalized into more mature CD34– cells but not into CD34+ stem cells, while P7228 liposomes were internalized into both CD34– and CD34+ cells. After injection of iron oxide–labeled hematopoietic cells, a significant decrease in MR signal intensity was observed in liver and spleen at 1, 4, 24, and 48 hours after injection (P < .05) and in the bone marrow at 24 and 48 hours after injection (P < .05). The signal intensity decrease in bone marrow was significantly stronger after injection of iron oxide–labeled cells compared to controls that received injections of the pure contrast agent (P < .05). Results of histopathologic examination confirmed homing of iron oxide–labeled human progenitor cells in the murine recipient organs.
CONCLUSION: The in vivo distribution of intravenously administered iron oxide–labeled hematopoietic progenitor cells can be monitored with 1.5-T MR imaging equipment.
© RSNA, 2005
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODS RESULTS DISCUSSIONREFERENCES |
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So that transplanted stem cells can be visualized and tracked with MR imaging, they must be labeled with MR imaging contrast agents, and the development of dedicated labeling techniques is currently being broadly investigated (6–15). Initial cell-labeling techniques were hampered by limited concentration of internalized contrast agent, which resulted in a limited sensitivity of MR to depict the labeled cells. To compensate for this limited sensitivity, experimental cell-tracking studies were performed by using MR imagers with very high magnetic field strengths of up to 14 T (10); alternatively, the contrast agent–labeled cells were injected directly into the organ of interest (8), in which the cells had a small distribution volume, ensuring that contrast agent concentration remained relatively high in the evaluated field of view. With clinical 1.5-T MR equipment and clinically applicable contrast agents, it has—to our knowledge—not been possible to trace the in vivo distribution of intravenously injected hematopoietic cells to more than one final target organ or to depict the migration of the transplanted cells to several subsequent target organs over time (6).
Newer and optimized labeling techniques have improved the efficiency of stem cell labeling and allowed the labeling of the cells with contrast agents that have been approved by the U.S. Food and Drug Administration or are currently being investigated in clinical trials (14,15). Thus, the purpose of this study was to evaluate the use of clinical 1.5-T MR imaging equipment to depict the in vivo distribution of iron oxide–labeled human hematopoietic progenitor cells in athymic mice.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODS RESULTS DISCUSSIONREFERENCES |
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Labeling of ferumoxides by means of simple incubation.—Ferumoxides particles have a diameter of 120–180 nm, an R1 relaxivity of 40 L · mmol–1 · sec–1, and an R2 relaxivity of 160 L · mmol–1 · sec–1 at 37°C and 0.47 T (13,14). For this study, 5 x 106 progenitor cells in 1 mL of Delbecco’s modified Eagle’s medium were incubated with a ferumoxides dose of 250 µg of iron for 4 hours.
Liposome-assisted labeling of P7228 particles.—P7228 particles have a diameter of 20–50 nm, an R1 relaxivity of 22 L · mmol–1 · sec–1, and an R2 relaxivity of 80 L · mmol–1 · sec–1 at 37°C and 0.47 T (12,15). P7228 particles are negatively charged and thus can be entrapped with cationic liposomes (15,19). Of note, ferumoxides particles could also be entrapped with cationic liposomes, but this would occur with a much lower efficiency owing to the fact that ferumoxides particles have a neutral coating, whereas P7228 particles have an anionic coating.
For this study, 5 x 106 progenitor cells in 1 mL of Delbecco’s modified Eagle’s medium were incubated with 20 µL of liposome formulation (Lipofectin; Life Technologies, Gibco BRL, Karlsruhe, Germany) and a P7228 dose of 250 µg iron for 4 hours. The P7228-containing liposomes fuse with the cell plasma membrane and deliver the contrast agent to the cell cytosol (15,19). All applied incubation procedures, doses, and incubation times were chosen on the basis of results of previous optimization experiments (15).
After the labeling procedures, cells were washed at least three times with HF/2+ (Hank’s balanced salt solution; Gibco BRL) supplemented with 2% fetal calf serum (PAN Biosystems), penicillin-streptomycin (Gibco BRL), and 10 mmol/L HEPES buffer (Gibco BRL). They were then diluted in 300–400 µL of HF/2+ and filtered through a 30-µm Fillcon filter (BD Immunocytometry Systems, Erembodegem, Belgium). Filtering was performed directly before cell injection. The cell suspensions were slowly injected into one of the tail veins of the mouse over 1–2 minutes.
Spectrometry
Because the majority of hematopoietic stem cells express the surface mucin CD34, we determined the iron uptake by CD34+ stem cells and by the more mature CD34– cells. CD34– and CD34+ cells were separated by using a cell sorter (Miltenyi Biotech, Bergisch Gladbach, Germany) and positive selection. The iron concentration within nonlabeled cells, as well as that in iron oxide–labeled CD34– and CD34+ cells, was quantified with atomic absorption spectrometry by using a polarized Zeeman atomic absorption spectrometer (model Z-8200; Hitachi, Japan) (15). If necessary, the cell suspensions were diluted with 0.05 mol/L HCl. For quality control, Lyphochek (Bio-Rad Laboratories, Munich, Germany), a control serum that is used to obtain standard curves before each experiment, was used.
Colony Assay
Human cord blood mononuclear cells or CD34+ stem cells, labeled with ferumoxides or P7228, were seeded as 20 000 cells (mononuclear cells) or 100 cells (CD34+ cells) per 1.1 mL of semisolid medium (H4435; StemCell Technologies, Vancouver, Canada). The cells were incubated in 35-mm dishes at 37°C, 5% CO2, and greater than 95% humidity. After 14 days, erythroid colonies were determined as the number of burst-forming erythroid units (BFU-E) (ie, colonies of three or more small colonies of up to 200 erythroid cells), mixed colonies were scored as the number of granulocyte/erythroid/macrophage/megakaryocyte colony-forming units (CFU-GEMM) (ie, colonies comprising a substantial proportion of both erythroid and myeloid progeny), and myeloid colonies were scored as the number of granulocyte/macrophage colony-forming units (CFU-GM) (ie, colonies of more than 50 cells consisting only of myeloid cells).
Animals and Animal Procedures
The study was approved by the institutional Committee on Animal Research. Forty-one female Balb/c mice were initially included in the study. Of these, six mice were excluded from evaluations because they died directly after intravenous cell injections (n = 4) or because of problems with the anesthesia (n = 2).
The study animals, which completed the whole study procedure, comprised 20 female nude Balb/c-AnNCrl mice that were 30 days old (Charles River Laboratories, Sulzfeld, Germany). These mice received an intravenous injection of ferumoxides-labeled cells (n = 10) or P7228-labeled cells (n = 10); the animals were randomized to the two contrast agent groups. In each group, 1 x 107 (n = 2), 2 x 107 (n = 2), or 3 x 107 (n = 6) labeled cells were injected into the tail vein. Control experiments were performed in 15 female Balb/c control mice that were 30 days old, which received intravenous injections of either 3 x 107 nonlabeled progenitor cells (n = 3) or the contrast agents alone at clinically applied doses (ferumoxides: 0.8 mg iron per kilogram, n = 6; and P7228: 2.6 mg iron per kilogram, n = 6).
All mice were anesthetized for MR imaging with an intraperitoneal administration of 0.5 mg/kg medetomidin (1 mg/mL, Domitor; Pfizer, New York), 5 mg/kg midazolam (5 mg/mL, Dormicum; Roche, Vienna, Austria), and 0.05 mg/kg fentanyl (0.05 mg/mL, Fentanyl; Janssen-Cilag, Germany). After completion of MR imaging, anesthesia was reversed with 2.5 mg/kg atipamezol (5 mg/mL; Antisedan, Pfizer), 0.5 mg/kg flumazenil (0.1 mg/mL, Anexate; Roche), and 1.2 mg/kg naloxon (0.4 mg/mL, Narcanti; Janssen-Cilag). After completion of imaging procedures, the animals were sacrificed with an intracardiac overdose of pentobarbital.
MR Imaging
MR imaging of the animals was performed in each group before and 1 (n = 3), 4 (all), 24 (all), and 48 (n = 3) hours after injection of labeled cells or contrast agent. Imaging was performed with a 1.5-T MR imaging unit (ACS NT; Philips, Best, the Netherlands) and a dedicated birdcage coil (Medical Advances, Milwaukee, Wis). Pulse sequences comprised (a) coronal T2-weighted two-dimensional turbo spin-echo (SE), 3700/90 (repetition time msec/echo time msec) sequences with a 90° flip angle and a section thickness of 600 µm and (b) coronal T2*-weighted three-dimensional fast field-echo, 32/14 sequences with a flip angle of 15° and an effective section thickness of 400 µm. MR images were acquired with a field of view of 100 x 80 mm, a 5122 pixel matrix, and an in-plane spatial resolution of 200 x 150 µm. Average signal intensities of liver, spleen, and bone marrow before and after cell injection were measured by one investigator (H.E.D.), who was blinded to the applied labeling procedure, using three or four operator-defined regions of interest per tissue. The size of the regions depended on the diameter of the anatomic region being investigated; a minimum of 20 pixels was required per region. Signal-to-noise ratios (SNRs) were calculated by dividing signal intensity data of the target organ by the image (background) noise (random fluctuations in signal intensity), which was measured in the background anterior to the depicted object (20).
Histopathologic and Immunohistochemical Evaluations
Samples of labeled and nonlabeled CD34– and CD34+ cells were stained with Prussian blue (Merck, Haar, Germany) to identify intracytoplasmic iron oxide contrast agent particles. Immunohistochemistry was performed on paraffin tissue sections (1.5 µm), stained with monoclonal antibodies against CD31+ human cells (DAKO, Hamburg, Germany) and monoclonal antibodies against the coating of the ferumoxides P7228 contrast agent (StemCell Technologies) and visualized by using horseradish peroxidase– or alkaline phosphatase–conjugated secondary antibodies (LSAB kits; DAKO). The tissue sections were counterstained with Mayer hematoxylin solution. Colocalization of human CD31+ cells and dextran-coated iron oxide particles was demonstrated on deparaffinized sections by using CD31 and secondary fluorescent antibodies TRITC (tetramethyl rhodamin) and fluorescein-isothiocyanate–conjugated antidextran antibodies (StemCell Technologies).
Statistical Analysis
Initial injections of iron oxide–labeled cells were considered pilot experiments to determine the minimal required cell number that produced visible changes in MR signal intensity. For this pilot study, increasing quantities of cells—1 x 107, 2 x 107, and 3 x 107 cells—were injected into two animals each in each contrast agent group. The initial number of cells was chosen on the basis of results from previous studies (15). Results of the pilot study revealed that only injections of 3 x 107 cells caused visible signal intensity changes in all organs. In these subgroups, we increased the number of examined animals to six. This was the minimal number of animals, determined with power analysis, that was needed to prove the statistical significance of differences in MR signal intensity before and after cell injection. The power analysis assumed a nondirectional statistic testing of MR data for no difference versus difference.
The MR signal intensities before and after cell injections, quantified as SNR data, were presented as means and standard errors of the mean. To compare differences in these quantitative MR data before and after injection of 3 x 107 cells, a two-tailed paired Student t test was used. Differences in SNR data at different time points before and after injection in the same animals were tested for significance with an analysis of variance for repeated measurements. Differences in colony assays without and with iron oxide labeling were compared by using an unpaired t test. Statistical significance was assigned if P < .05. All statistical computations were processed by using Statview 4.1 software (Abacus, Berkeley, Calif).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODS RESULTS DISCUSSIONREFERENCES |
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODS RESULTS DISCUSSIONREFERENCES |
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Previous experimental approaches to label hematopoietic cells—such as use of lymphocytes (6) and progenitor cells (10)—with iron oxide contrast agents were not sensitive enough to provide an in vivo MR depiction of the cells after they were intravenously injected. Rather, postmortem excised tissue samples had to be evaluated in order to prove homing of the transplanted hematopoietic cells to recipient organs (6,10). Several studies focused on the design of new, optimized probes, which maximized labeling efficiency and—consequently—MR detectability of transplanted cells (8,13). However, a drawback of such new probes is that they are not readily available for other researchers or for clinical applications. Other investigators focused on labeling probes that are commercially available and may be clinically applicable (11,12,14). With our study, we intended to combine optimized labeling protocols (15) and comercially available probes. Of note, labeling techniques with radiotracers might also be clinically available and would provide a much higher sensitivity for cell detection than that achieved with our MR method (22). However, although the sensitivity of scintigraphy is high, its spatial resolution is limited, its temporal resolution for serial follow-up studies would be limited by radioactive decay of the tracer, and radiotoxic cell damage could occur (22).
An advantage of labeling with ferumoxides is that ferumoxides is approved by the U.S. Food and Drug Administration as a liver contrast agent and, thus, may be available for clinical applications. Likewise, we labeled hematopoietic cells with ferumoxides by means of simple incubation, because this method would be one most likely to be approved for clinical applications. As shown by our data, ferumoxides labeled CD34– hematopoietic cells efficiently, but the CD34+ stem cells remained unlabeled. Since the seeding efficiency of sorted CD34+ stem cells is nearly identical to that of the total population of hematopoietic donor cells (23) (ie, the CD34+ and CD34– cells distribute with comparable relative quantitative proportions and within the same time interval to liver, spleen, and bone marrow) this may not be a disadvantage. On the other hand, since the CD34+ stem cells are not labeled, any impairment of CD34+ stem cells by the labeling procedure is excluded.
P7228 is a second-generation iron oxide contrast agent for MR imaging (15), is currently being evaluated in phase I and II clinical trials (24,25), and is not yet approved for routine clinical imaging applications. As shown by our data, liposome-assisted P7228 labeling affects all hematopoietic donor cells, including the CD34+ stem cells. Indeed, we have shown that this labeling procedure interferes neither with hematopoietic colony formation nor with differentiation of neural and embryonic stem cells (26). Thus, in the long term, the liposome-assisted labeling appears to be better suited for MR tracking studies of purified stem and progenitor cells, because it can depict all cell populations.
We did not find any impairment of our progenitor cells by ferumoxides or P7228 treatment. Indeed, in cultures of both ferumoxides-treated cells and P7228-treated cells, detection of colonies with an erythroid component appeared to be enhanced. Previous studies showed that iron uptake corelates with the number of surface receptors for transferrin, CD71 (27). Cord blood erythroid progenitors express higher levels of CD71 than do myeloid progenitors (28), which would explain why colonies with an erythroid component (BFU-E and CFU-GEMM) are even enhanced by treatment with ferumoxides or P7228.
A crucial goal in the development of new stem cell therapies is to achieve and prove the homing of the transplanted cells to the particular tissue where they should exert their therapeutic activity (2). This is of special importance if the cells are administered systemically (eg, after intravenous injection, when the cells pass several intermediate organs after reaching their final destination) rather than directly into the target organ. Data in our xenotransplant model showed that the in vivo distribution of systemically administered hematopoietic progenitor cells to intermediate organs (liver and spleen) and the desired final target organ (bone marrow) can be traced with MR imaging. The observed anatomic and temporal in vivo cell distribution corresponded in general to the distribution of human hematopoietic cells in immunodeficient NOD/SCID mice in previous studies (23,29,30). These cells disappeared rapidly from the circulation and accumulated in spleen and marrow (29,30). In bone marrow, an increasing accumulation of human cells was observed up to 24 hours after injection (22,23). This initial cell accretion was followed by a cell persistence in marrow (29,30).
Our MR studies were directed to depict early processes of cell transplantation with histopathologic correlation. In this context, it is important to distinguish between homing and engraftment of the cells. We evaluated the homing of intravenously injected hematopoietic cells, which is characterized by the in vivo distribution of transplanted cells to specific target organs before they start to divide; this usually occurs within 1 day after transplantation (31). Homing of hematopoietic progenitor and stem cells provides important information about the in vivo allocation of the originally transplanted cells without interference from subsequent dividing progeny. The majority of transplanted hematopoietic stem cells home specifically to the bone marrow (31,32). Although many investigators studied the homing process of hematopoietic cells in myeloablated animals, it has also been shown that homing in nonirradiated animals proceeds as efficiently as in irradiated animals (33). It even appeared that hematopoietic stem cells may home more efficiently in nonirradiated animals because of the preservation of the endothelial cell integrity within the target organs (34). Therefore, we investigated homing of human hematopoietic cells in nonirradiated nude mice with use of MR imaging within 24 hours of transplantation.
By contrast, we did not assess cell engraftment, which is characterized by subsequent proliferation and separation of progenitor cells and which usually occurs over several days or weeks after transplantation. It would be illogical to perform MR imaging after 1 week in this context, because at 1 week the initially labeled and injected, original progenitor and stem cells would no longer be present but would have divided and differentiated to more mature progeny. Our in vitro data did not show any impairment of the cell biology as a result of the labeling process. However, follow-up studies would be valuable to prove, in vivo, that the cell engraftment is not impaired by iron oxide depositions within the cytoplasm of the transplanted cells.
Our study design had some limitations. With regard to a potential clinical application, we standardized our procedures according to the injected cell number, not according to the applied contrast agent dose. In a clinical situation, 2 x 108 to 2 x 109 mononuclear hematopoietic cells per kilogram body weight are infused. Accordingly, we injected (1.0–3.0) x 107 cells in a 20-g mouse, which corresponds to 5 x 108 to 1.5 x 109 cells per kilogram. However, cell labeling with P7228 (uptake, 1.5 pg of iron per cell ± 0.7) was less efficient than cell labeling with ferumoxides (uptake, 2.2 pg of iron per cell ± 0.8). This lower efficiency of the P7228 labeling method, along with a lower R2 relaxivity of P7228 compared with that of ferumoxides, explains the lower MR contrast effect observed after injection of P7228-labeled cells as opposed to ferumoxides-labeled cells. We did not increase the number of injected P7228-labeled cells to improve the contrast. Increasing the number of cells injected would also not be done in a clinical situation, where the radiologist would have to tailor his or her labeling technique with respect to the required number of cells, and not vice versa.
Likewise, in accordance with the usual clinical situation, we injected both CD34– cells (more than 99% of the injected cells) and CD34+ (about 0.1%–0.5%) mononuclear hematopoietic cells. For the ferumoxides studies, we did not isolate the nonlabeled CD34– cells from the whole cell population prior to injection, because we wanted to avoid deviations from current clinical protocols. For research purposes, it would be interesting to compare the in vivo behavior of the CD34+ cells with that of the CD34– cells. This would be possible only with the P7228 liposome–labeling technique, because only this technique allows labeling of both cell populations. However, such a study would necessitate a different setup compared with the current study, with cell separation taking place prior to injection and, ideally, injection of equal numbers of cells from each cell population. A study with this focus is currently being pursued by our group.
Our labeling procedure proved to be stable without release of contrast agent from the cells within the time of observation, up to 48 hours after injection. As shown by our histopathologic data, each iron oxide–positive cell within the recipient murine tissues was a transplanted human cell. Further follow-up studies are warranted to investigate the long-term pharmacokinetics of the internalized iron oxide, which would be expected to parallel the well-documented pharmacokinetics of the same iron oxide compounds after intravenous administration (ie, the dextran or semisynthetic coating would be slowly degraded and excreted by the kidneys, and the iron oxide would be incorporated into the body’s iron metabolism (eg, used for heme synthesis or stored in the liver) (16,17,24).
In summary, intravenously injected iron oxide–labeled human hematopoietic progenitor cells can be traced noninvasively in vivo with submillimeter anatomic resolution by using 1.5-T MR imaging equipment. This cell-tracking technique with clinically applicable MR imaging equipment and contrast agents may provide biological insights relevant for the development of new cell-based therapies and may be generally suited to monitor stem cell homing and engraftment in patients.
Potential clinical applications comprise MR imaging–based cell tracking studies of autologous and allogeneic bone marrow transplantation, particularly investigations of the homing specificity of various stem cell subtypes or genetically engineered stem cells, assessment of therapy effects on stem cell differentiation outcomes, and in vivo investigations of reasons for graft failure in certain populations (eg, patients with chronic myelogic leukemia) (1–3). Further applications could include in vivo monitoring of new cell-based therapies, such as homing of mesenchymal stem cells in injured myocardium (12), homing of neurologic stem cells in impaired brain tissue (8), and accumulation of transplanted natural killer cells in tumors. Finally, this method not only may be applicable to monitor therapy effects, but may be used diagnostically as an MR-based alternative to leukocyte scintigraphy (22): Iron oxide–labeled leukocytes may be used to specifically detect and characterize inflammatory tissue with improved anatomic resolution and without radiation exposure of the patient.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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See also Science to Practice in this issue
Author contributions: Guarantors of integrity of entire study, H.E.D., M.R., E.J.R., R.A.J.O.; study concepts, H.E.D., M.R., J.S., T.M.L., C.P., E.J.R., R.A.J.O.; study design, H.E.D., M.R., E.J.R., R.A.J.O.; literature research, H.E.D., M.R., S.M., R.A.J.O.; experimental studies, H.E.D., M.R., G.P., S.M., R.A.J.O.; data acquisition, H.E.D., M.R., G.P., S.M., R.A.J.O.; data analysis/interpretation, H.E.D., M.R., G.P., S.M., C.C., R.A.J.O.; statistical analysis, H.E.D., T.M.L., R.A.J.O.; manuscript preparation, definition of intellectual content, editing, and final version approval, all authors; manuscript revision/review, H.E.D., M.R., R.A.J.O.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODS RESULTS DISCUSSIONREFERENCES |
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