JACC: CARDIOVASCULAR INTERVENTIONS
© 2009 BY THE AMERICAN COLLEGE OF CARDIOLOGY FOUNDATION
PUBLISHED BY ELSEVIER INC.
VOL. 2, NO. 8, 2009
ISSN 1936-8798/09/$36.00
DOI: 10.1016/j.jcin.2009.05.014
Magnetic Tagging Increases Delivery of
Circulating Progenitors in Vascular Injury
Panagiotis G. Kyrtatos, BMEDSCI,*†§ Pauliina Lehtolainen, PHD,*
Manfred Junemann-Ramirez, MD,‡ Ana Garcia-Prieto, PHD,储 Anthony N. Price, PHD,*
John F. Martin, MD,‡ David G. Gadian, DPHIL,† Quentin A. Pankhurst, PHD,§储
Mark F. Lythgoe, PHD*†
London, United Kingdom
Objectives We sought to magnetically tag endothelial progenitor cells (EPCs) with a clinical agent
and target them to a site of arterial injury using a magnetic device positioned outside the body.
Background Circulating EPCs are involved in physiological processes such as vascular re-endothelialization and post-ischemic neovascularization. However, the success of cell therapies depends on
the ability to deliver the cells to the site of injury.
Methods Human EPCs were labeled with iron oxide superparamagnetic nanoparticles. Cell viability
and differentiation were tested using flow cytometry. Following finite element modeling computer
simulations and flow testing in vitro, angioplasty was performed on rat common carotid arteries to
denude the endothelium and EPCs were administered with and without the presence of an external
magnetic device for 12 min.
Results Computer simulations indicated successful external magnetic cell targeting from a vessel
with flow rate similar to a rat common carotid artery; correspondingly there was a 6-fold increase in
cell capture in an in vitro flow system. Targeting enhanced cell retention at the site of injury by
5-fold at 24 h after implantation in vivo.
Conclusions Using an externally applied magnetic device, we have been able to enhance EPC localization at a site of common carotid artery injury. This technology could be more widely adapted to
localize cells in other organs and may provide a useful tool for the systemic injection of cell
therapies. (J Am Coll Cardiol Intv 2009;2:794 – 802) © 2009 by the American College of Cardiology
Foundation
From the *Centre for Advanced Biomedical Imaging, University College London (UCL) Department of Medicine and UCL
Institute of Child Health, London, United Kingdom; †Radiology and Physics Unit, UCL Institute of Child Health, London,
United Kingdom; ‡Centre for Cardiovascular Biology and Medicine, UCL Department of Medicine, London, United Kingdom;
§Davy-Faraday Research Laboratories, The Royal Institution of Great Britain, London, United Kingdom; and 储London Centre
for Nanotechnology, London, United Kingdom. Supported by the UCL Institute of Child Health (Child Health Research Appeal
Trust), the British Heart Foundation, the Alexander S. Onassis Public Benefit Foundation, and the Biotechnology and Biological
Sciences Research Council. Panagiotis G. Kyrtatos and Dr. Lehtolainen contributed equally to this manuscript.
Manuscript received May 4, 2009, accepted May 20, 2009.
Kyrtatos et al.
Magnetic Tagging Increases Cell Delivery
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One of the current challenges in the biomedical sciences is
the localization of stem cells to sites of interest for the repair
of tissue damage. Cellular therapies are increasingly applied
in clinical trials, and in recent years the use of hematopoietic
progenitors as repairing modules in the compromised cardiovascular system has been the focus of considerable
attention (1–3). In the context of ischemic heart disease,
these efforts have led to modest success (4,5). However,
delivery to specific targets within the body generally remains
a difficult task, in part due to low uptake at the site of injury.
As such, cell therapies would greatly benefit from methodologies aimed at targeting and monitoring cell trafficking.
See page 803
Superparamagnetic iron oxide nanoparticles (SPIO) offer
attractive possibilities in biomedicine as they can be incorporated into cells affording a controllable means of “tagging” (6).
These particles lead to a marked decrease in the magnetic
resonance imaging (MRI) parameter T2* and the possibility of
visualizing their localization noninvasively on T2*-weighted
MRI (7–9). Furthermore, the magnetic properties of SPIOs
allow them to be manipulated mechanically by a magnetic field
gradient. This “action at a distance,” combined with the
intrinsic penetrability of magnetic fields into human tissue,
opens up potential applications involving the transport of
magnetically tagged biological entities. Such tagging has to
date mainly been the focus of drug delivery systems (10 –12),
including drug targeting in humans (13). Recent efforts on
cell targeting in the arterial circulation have been limited to
animal models; to achieve cell capture, these have necessitated the use of large nonbiodegradable micron-sized beads
(14 –16) or nanoparticle composites fabricated in-house
(17), and in most cases, the additional introduction of
permanent intravascular metallic devices not currently approved for human use (15–17). It may also be possible to
attract gene-carrying cells to post-capillary venules of tumors (18). However, the targeted delivery of progenitor cells
using external magnetic devices and clinically approved
iron-bearing agents has not yet been accomplished.
Early endothelial progenitor cells (EPCs) derived from
CD34⫹/CD133⫹ cells (19,20) have been shown to be
involved in post-ischemic neovascularization (21) and reendothelialization, reducing neointima formation following
arterial injury (22). We chose to target EPCs to a site of rat
common carotid artery (CCA) injury, as localization of such
progenitors to sites of vascular catheterization may help
prevent post-angioplasty restenosis (23,24). Although various SPIO formulations have been used in the past, their future
use as clinical agents is not straightforward due to safety issues
raised by cell labeling and the use of transfection agents. To
obviate these concerns, we have chosen to label human EPCs
with the U.S. Food and Drug Administration (FDA)-
795
approved nano-sized SPIO compound Endorem (Guerbet,
Paris, France), the only agent to date that has been used to
monitor cells in humans using MRI (8).
Methods
Cells. Human mononuclear cells (MNCs) were collected by
leukapheresis from peripheral blood of donors stimulated
with granulocyte colony-stimulating factor (25). We magnetically isolated the CD133⫹ progenitor cells using antihuman CD133 (epitope 1) microbeads (Miltenyi Biotec,
Bergisch Gladbach, Germany). These beads are biodegradable and typically disintegrate after a few days in culture.
Then, EPCs were derived by culturing CD133⫹ cells for up
to 21 days on fibronectin-coated plates (BD, Franklin
Lakes, New Jersey) in growth medium consisting of 20%
fetal bovine serum/endothelial basal medium 2 (Lonza,
Basel, Switzerland) and supplemented with fibroblast, insulinlike, and endothelial growth factors; ascorbic acid; heparin; and
recombinant human vascular endothelial growth factor 165 (25
ng/ml) (R&D Systems, MinneAbbreviations
apolis, Minnesota) (26). Growth
and Acronyms
media were supplemented with
CCA ⴝ common carotid
recombinant human stem cell facartery
tor (100 ng/ml), rhFlt-3/Flk-2 liEPC ⴝ endothelial progenitor
gand (50 ng/ml), and recombicell
nant human interleukin 3 (10 ng/
FDA ⴝ Food and Drug
ml) (R&D Systems) for the first
Administration
72 h. MNCs were grown for 2 h
MNC ⴝ mononuclear cell
in serum-free Dulbecco’s modiMRI ⴝ magnetic resonance
fied eagle medium (Gibco, Inimaging/images
vitrogen, Carlsbad, California)
SPIO ⴝ superparamagnetic
followed by removal of nonadheriron oxide
ent cells and culture of the adherent fraction in 10% fetal bovine
serum/Dulbecco’s modified eagle medium. All blood samples
were obtained with written consent and were approved by local
ethics committees.
Labeling of cells with SPIO. At day 9 of culturing, the
CD133⫹ cells were labeled with the FDA-approved SPIO
Endorem (also known as Feridex in the U.S.) (Guerbet)
using 0.5 mg/ml SPIO in growth medium. Suspension
CD133⫹ cells were labeled for 24 h and adherent cells for
1 h. Additionally, adherent cells were labeled for 2 or 24 h
to increase the amount of iron in the cells. Mononuclear
cells were labeled with SPIO by overnight incubation.
Labeled cells were washed twice in 15 ml phosphatebuffered saline.
Quantification of cellular SPIO uptake. We used a superconducting quantum interference device (27) to measure the
amount of Fe3O4 in the cells. To separate SPIO signal from
background signal coming from the cells and media, measurements were performed at –263oC. The samples were
saturated in an applied field of 2-T, which was subsequently
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Kyrtatos et al.
Magnetic Tagging Increases Cell Delivery
removed to leave the SPIO particles in a magnetized state.
Comparison of this remnant signal with that of an undiluted
SPIO sample of known Fe3O4 concentration allowed quantification of Fe3O4 per cell.
Magnetic actuation. Neodymium iron boron permanent
disk magnets (1.195-T, grade N35SH, Magnet Applications Ltd., Hertfordshire, United Kingdom) 22 mm in
diameter and 16 mm in height were used to test cell viability
and differentiation following magnetic attraction. The
SPIO-labeled EPCs were placed 1 mm from the magnet
surface (maximum: 19.8 T2/m). For the flow system and in
vivo magnetic targeting, a magnetic actuator was constructed using 5 10 ⫻ 10 ⫻ 25-mm3 NdFeB grade N35
blocks, attached together inside a machined aluminum
casing in a configuration of rotating magnetization (Halbach array), which concentrates the force on 1 side of the
actuator (28).
Viability assays. The CellTiter 96 AQueous One Solution
Assay (Promega, Madison, Wisconsin) was performed after
24 h of magnetic attraction of labeled CD133⫹ cells. The
tetrazolium compound in this assay undergoes reduction to
a colored formazan product by viable cells, primarily mediated by reduced nicotinamide adenine dinucleotide phosphate (29). Colorimetric quantification of viability of samples of 3 ⫻ 104 CD133⫹ cells was performed using a
spectrophotometric microplate reader (GENios, Tecan
Group Ltd., Männedorf, Switzerland) (490 nm, at 4 h).
Sample activity was normalized to the mean absorbance of
control (no magnet). Cell apoptosis was analyzed using an
annexin-V-fluorescein isothiocyanate/propidium iodide
flow cytometric assay (ApoTarget, Invitrogen).
Flow cytometry. Immunophenotypic characteristics of
CD133⫹ cells labeled with SPIO at day 10 were analyzed
with a fluorescence-activated cell sorter (FACSCalibur,
BD) at day 21 of cultivation. Cells pre-treated with crystallizable fragment receptor blocking reagent (Miltenyi Biotec)
were stained using the following antihuman antibodies:
phycoerythrin (PE)-CD304 BDCA-4/neuropilin-1 (30),
biotin-CD144/vascular endothelial cadherin (Bender MedSystems Inc., Burlingame, California) with streptavidinfluorescein isothiocyanate for endothelial differentiation;
PE-CD14 and CD11b (BD Biosciences, San Jose, California) for monocytic differentiation. Fluorochrome- and dosematched isotypes were used as controls. Cells were suspended in 20 mmol/l N-2-hydroxyethylpiperazine-N-2ethanesulfonic acid/0.5% bovine serum albumin/phosphatebuffered saline containing 10 ␮g/ml 7-aminoactinomycin D
(Sigma Aldrich, St. Louis, Missouri). The 7-aminoactinomycin D–positive, nonviable cells were excluded and 1 ⫻ 104
viable cells were scored per analysis (CellQuestPro, BD).
MRI. The SPIO-labeled CD133⫹ cells were embedded in
low melting point agarose (0.5%, 37°C) at concentrations of
745, 165, 49.5, 16.5, and 0 ⫻ 103 cells/ml in 250-␮l
Eppendorf tubes (Eppendorf North America, Westbury,
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AUGUST 2009:794 – 802
New York) and were cooled to 4°C. Suspension in agarose
allowed for homogeneous distribution of the cells. The
tubes were imaged in a 9.4-T horizontal bore Varian MRI
system (VNMRS, Varian Inc., Palo Alto, California) using
a 39-mm radiofrequency coil (RAPID Biomedical GmbH,
Rimpar, Germany). The T2*-weighted images were acquired using a spoiled gradient echo sequence (echo time
[TE] ⫽ 25 ms, repetition time [TR] ⫽ 200 ms, flip angle
[FA] ⫽ 30, 5122 matrix). The T2* maps were obtained
from 20 echo times (TE ⫽ 2.5 to 30 ms, TR ⫽ 200 ms,
FA ⫽ 30, 2562 matrix) and produced using the ImageJ MRI
Analysis Calculator plug-in (K. Schmidt, ImageJ, U.S.
National Institutes of Health, Bethesda, Maryland).
Computer simulations and in vitro flow system. Magnet
designs for acquiring optimal magnetic field strength to
capture intravascular SPIO-labeled cells were modeled using Opera-2d and -3d (Cobham Technical Services, Aurora, Illinois) and COMSOL Multiphysics 3.1 (COMSOL
Inc., Burlington, Massachusetts) software. Fluid flow rate,
cell size and Fe3O4 content, and distance of magnet from
the vessel were considered by solving for the Khan and
Richardson hydrodynamic drag force and the attractive
force from the magnet, in parabolic nonpulsatile laminar
flow. Simulated cells were 10 ␮m in diameter, 3.9 pg
Fe3O4/cell, within a circulating blood volume of 20 ml. Cell
capture was defined as approximation and contact with the
vessel wall within the 50-mm length of the array. The
Halbach magnet design described earlier was assessed in an
in vitro flow system by placing 1 ⫻ 106/ml SPIO-labeled
blood MNCs (3.5 pg Fe3O4/cell) in peristaltic pump-driven
flow (10 ml/min, mean velocity: 0.4 m/s) using biocompatible (ISO 10993) Tygon S-54 elastic microtubing (SaintGobain Performance Plastics, Courbevoie, France). The
magnet array was placed at distances of 1 and 5 mm from
the tubing. The mean of 5 hemocytometer cell counts was
calculated. Digital images were acquired using a standard
dissection microscope and a digital camera.
Vascular injury and in vivo magnetic targeting. Male
Sprague-Dawley rats (n ⫽ 13, 380 to 420 g) (Charles River
Laboratories International, Wilmington, Massachusetts)
were anaesthetized using midazolam (625 ␮g/100 g) and
fentanyl (40 ␮g/100 g) supported by halothane at 0.5% to
2% at 2 l/min oxygen. The left CCA was exposed at the
bifurcation to the external and internal carotid artery and a
2-F embolectomy catheter (Edwards Lifesciences Corp.,
Irvine, California) was advanced proximally through an
arteriotomy into the CCA. The catheter was inflated with
100 ␮l of air and rotated as it was being retracted (31). The
SPIO-labeled CD133⫹ cells were labeled with 5-␮mol/l
CellTracker Green 5-chloromethylfluorescein diacetate
(Invitrogen-Molecular Probes). This dye is activated by
intracellular esterases and only stains viable cells. We
administered 5 ⫻ 105 cells in the CCA, which was
temporarily clamped for 10 min with the magnet array
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797
placed at a 5-mm distance. The circulation was restored and
the magnet removed after a further 2 min of magnetic
actuation (n ⫽ 5). Two types of controls were used: 1) CCA
injury was performed and labeled cells were injected without
magnetic actuation (n ⫽ 6); and 2) magnetic attraction of
labeled cells was performed on uninjured vessels (n ⫽ 2).
Tissues were retrieved 24 h later and fixed with 1%
p-fluoro-phenylalanine/phosphate-buffered saline. All experimental procedures complied with the institutional
guidelines for animal experiments.
Confocal microscopy and quantitation of cell engraftment.
Arteries were opened longitudinally and counterstained
with 4=,6-diamidino-2-phenylindole (Vector Labs, Burlingame, California). Total arterial surface confocal microscope images were acquired using a Leica TCS SP2 confocal
microscope. Reflectance scanning (32) was used to detect
intracellular SPIO. Image processing was performed using
the Volocity imaging software (Improvision, Waltham,
Massachusetts). Background correction was performed using the “rolling ball” algorithm implemented in ImageJ
software. Signal outliers were removed by application of a
despeckle median filter (W.S. Rasband, ImageJ, U.S. National Institutes of Health). To analyze cell engraftment,
x-y-z scans were rendered into a 3-dimensional vector
graphic format; this enabled quantification the total volume
of CellTracker-labeled cells per artery. Cell counts were
calculated from the total volume based on a 10-␮m cell
diameter and are presented per square millimeter of arterial
surface area.
Statistical analyses. We used SPSS version 11.5.1 (SPSS
Inc., Chicago, Illinois) and Prism version 4.02 (Graphpad
Software Inc., San Diego, California). Data were inspected
visually and with the Shapiro-Wilk test for small samples to
assess normality. The homogeneity of variance assumption
was evaluated with the Levene test. Mean values are quoted
with the standard error of the mean, medians are quoted
with the interquartile range (IQR). The p values reported
are 2-tailed and were considered statistically significant at
the 5% level (p ⬍ 0.05). In the apoptosis assays, equivalence
testing was considered appropriate because the null hypothesis was nondifference; 2 1-sided t tests were used with a 3%
equivalence limit at the 0.05 level (Schuirmann test). A
maximum acceptable difference of 3% was set a priori based
on 24-h serum-deprived positive control cells (3% mean
increase in early apoptosis, 16% increase in late apoptosis).
Other statistical tests are specified in context.
Results
Cell labeling and biological assays. To demonstrate that the
magnetic force following Endorem labeling was large
enough to attract cells to the vessel wall through a liquid, we
recorded the motion of labeled MNCs in real time as a
Figure 1. Magnetic Attraction Causes Translocation of SPIO-Labeled MNC
The estimated force is 19 pN per cell. (A) Video frames for t ⫽ 0 (application of magnet), 5, 10, and 15 s after exposure to the magnet. Relative
position of the magnet is displayed. (B) Count of cells in field of view (20⫻
objective). The cells transiently acquire a magnetic dipole moment and
experience intercell attraction, entering the field of view in aggregates
(increase in count rate between 5 and 10 s). The cell count rate subsequently decreases as the suspension volume closest to the magnet is
depleted of cells. MNC ⫽ mononuclear cell; SPIO ⫽ superparamagnetic
iron oxide.
magnetic force was applied (Fig. 1). Rapid movement
toward the magnet was observed as the cells responded to
the applied force.
The CD133⫹-derived EPC viability was normal following labeling and magnetic attraction (Fig. 2A) and no
increase in apoptosis was observed (Fig. 2B) when the
amount of SPIO in the cells was controlled. Increased
labeling resulted in a decrease in viability following magnetic attraction (Fig. 2A). This may be explained by
force-induced mechanical damage of intracellular structures
containing the iron nanoparticles (33). As shown in Figure
2C, there was normal expression of endothelial (vascular
endothelial cadherin, CD304/NP-1) and monocytic
(CD14, CD11b) markers of differentiation, 10 days after
labeling. Using in vitro MRI, cells were visible on T2*weighted images at dilutions as low as 25 cells/mm2 and
caused a marked reduction in T2* values from 8 cells/mm2
(Fig. 2D). Using a superconducting quantum interference
device, we calculated that labeled cells contained on average
3.9 pg Fe3O4 per cell. Given the magnetic field applied in
these experiments and the iron content of the cells, it is
possible to estimate a “cell-safe” magnetic force for EPCs
above which there is a risk of compromising viability; this
was 9.3 pN per cell.
External magnet design. An important consideration in our
experimental model is that the rat CCA is a nonsuperficial
structure approximately 5-mm from the skin in which there
are large hydrodynamic forces acting on the cells, with peak
systolic blood flow rate up to 10 ml/min (34,35). However,
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Figure 2. Labeling of CD133ⴙ Cells With SPIO
(A) Viability assay following 24 h of magnetic attraction (n ⫽ 4 per group, analysis of variance with Tamhane T2 post hoc test). There was no notable effect on
viability when the standard labeling method was used (adherent cells labeled for 1 h) when compared with the viability of the nonmagnet control group.
Increased adherent cell labeling for 2 and 24 h resulted in a 20% and 75% decrease in viability after magnetic attraction, respectively. (B) Mean difference in
apoptosis rates compared with apoptosis rates in the control group, at specified times after labeling and magnetic attraction. All groups passed equivalence testing with a 3% apoptosis limit. (C) Surface marker expression (percentage of live cells) at 10 days after labeling. No differences were observed on the expression
of differentiation markers (CS vs. LS and CA vs. LA) for vascular endothelial cadherin (VE-Cad, n ⫽ 6), CD304 (NP1, n ⫽ 6), CD14 (n ⫽ 6), or CD11b (n ⫽ 3). (D)
In vitro MRI; T2*-weighted images (i) and T2* maps (ii) with equivalent average T2* values plotted against cell concentration (iii). CA ⫽ control nonlabeled
adherent fraction; CI ⫽ confidence intervals; CS ⫽ control nonlabeled suspension fraction; EA ⫽ early apoptosis; LA ⫽ late apoptosis (B) or SPIO-labeled adherent fraction (C); LS ⫽ SPIO-labeled suspension fraction; MRI ⫽ magnetic resonance imaging; N ⫽ necrosis; SEM ⫽ standard error of the mean; T2* ⫽ MRI parameter, sensitive to SPIO presence; other abbreviations as in Figure 1.
there is a rapid decline of magnetic force at increased target
tissue distances, which poses a challenge to magnetic targeting using external devices in vivo (6,11). To overcome
this issue, we chose a permanent magnetic actuator design
that increases the force magnitude at large distances (Halbach array). With this actuator placed at a 5-mm vertical
distance from the target tissue, a computed maximum 8.7
pN force was applied on each cell, which is below the 9.3
pN limit.
Computer simulations. We proceeded by modeling the distribution of magnetic forces at distances of 1 and 5 mm
from the magnetic actuator (Fig. 3A) and simulated the
hydrodynamic and magnetic forces on intraluminal
CD133⫹ cells using finite element modeling. With the
vessel placed 1 mm away from the actuator, 11% of flowing
cells made contact with the endothelial surface at each pass
through the vessel. With the actuator at 5 mm, 2.3% of cells
made contact. If the cells adhered at the contact site,
approximately 50% of cells would be extracted from the
circulation within 15 min at 1 mm, or 60 min at 5 mm.
In vitro flow system. We investigated the targeting ability of
the actuator in vitro at the 2 distances using a 10-ml/min
flow system. At 1 mm, cells accumulated above the points of
predicted maximum force within 1 min after initiation of
flow (Fig. 3B). These cell aggregates dissociated a few
seconds after removal of the actuator. Targeting for 15 min
at 1 mm resulted in 252-fold increase in capture (mean ⫽ 41
⫻ 103 ⫾ 6 ⫻ 103 cells) versus control (mean ⫽ 163 ⫾ 41
cells) specimens. At 5 mm, there was a 6-fold increase in
capture (mean ⫽ 1,630 ⫾ 276 cells) versus nontargeted
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Figure 3. Computer Models and In Vitro Flow System
(A) Magnet array in aluminum casing; 5 bar magnets with rotating magnetization (top, digital image) and corresponding computer model (bottom), with color-coding of magnetic force magnitude, magnetic field lines displaying 1-sided flux, and 1 or 5 mm distance used in experiments (dotted lines). (B) In vitro cell
capture (top, digital images) at 1 min of 10 ml/min flow and corresponding force distribution along horizontal line 1 mm away from array (red plot), showing
predicted peaks of force magnitude at magnet junctions and cell aggregates within the vessel at these points (arrows and close up). Graph (right) displays
252-fold increase in cell capture after 15 min of magnetic targeting at 1 mm; n ⫽ 3, p ⫽ 0.003 (2-sample t test). (C) Force along the 5 mm horizontal line (left,
orange plot) and in vitro magnetic cell capture at 5 mm after 1 h of 10 ml/min flow (right); n ⫽ 4, p ⫽ 0.017 (2-sample t test with Welch correction).
control (mean ⫽ 281 ⫾ 21 cells) specimens after 1 h of flow
(Fig. 3C).
External targeting in a small animal model. To investigate
the in vivo feasibility of magnetic targeting using external
magnets, we attempted to increase SPIO-CD133 cell engraftment to a site of rat CCA injury by placing the actuator
adjacent to the ventral aspect of the animal’s neck. Images in
Figures 4A and 4B display representative (median) confocal
scans of the arterial surface 24 h after cell delivery; there is
a notable increase in cell retention following magnetic
targeting. High power magnification (Fig. 4D) confirmed
the presence of nucleated cells (using 4=,6-diamidino-2phenylindole) and the presence of intracellular clusters of
SPIO nanoparticles in CD133⫹ cells was confirmed using
reflectance confocal microscopy (Fig. 4E). Application of an
external magnetic force using the actuator during and after
cell delivery increased CD133⫹ cell engraftment to the
injured vascular surface by a factor of 5.4 (median ⫽ 134,
IQR: 47 to 362 cells/mm2) compared with cell delivery
without the magnet (median ⫽ 25, IQR: 17 to 42 cells/
mm2). In the absence of vascular injury (control studies), no
cells engrafted to the endothelial surface when the magnetic
force was applied (cell count ⫽ 0).
Discussion
The concept of magnetic tagging and targeting could play
an important role for future advances in delivery and
noninvasive monitoring of cell-based therapeutic interventions. This is the first attempt to use a clinically approved
agent to externally target EPCs in a high-flow system such
as the arterial vasculature. We chose to use EPCs for this
study as these may hold high therapeutic value for the
future. Endothelial damage is an initiating event in many
cardiovascular disorders. Transplantation of these early
progenitor cells, which resemble hemangioblasts in their
properties of marker expression, proliferation, migration,
and differentiation, may expand the therapeutic arsenal to
repair vessels or endothelium and improve perfusion of
ischemic tissues by providing new progenitor cells capable of
responding to angiogenic stimuli, and have recently been
used for cardiac neovascularization in clinical trials (36 – 40).
In the majority of these and other studies on cardiac
regeneration, cells were administered via intracoronary injection with “flow-stop” periods of several minutes to
increase engraftment; yet it has been shown that despite this
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Figure 4. Magnetic Targeting Following Balloon Angioplasty
(A) Sample 2-dimensional tile, total surface area scanning of the luminal side of a rat common carotid artery following injury and cell delivery without magnetic
targeting; 18 cells/mm2. (B) Cell delivery with targeting; 142 cells/mm2. Square box included for orientation. (C) A 3-dimensional reconstruction used for quantitation. Objects meeting signal intensity and volume threshold criteria are shown in pink. (D) High power magnification of specimen showing CellTracker-labeled
cells (green) adhering to the injured vascular intima. Cell nuclei are labeled with 4,6-diamino-2-phenylindole (blue). (E) Reflectance confocal image of a CD133⫹
cell showing SPIO-nanoparticles (yellow). (F) Magnetic targeting increased adhesion of SPIO-labeled, CD133⫹-derived to the injured vascular intima (n ⫽ 5) versus to the nontargeted control group (n ⫽ 6); p ⫽ 0.017 (Wilcoxon-Mann-Whitney exact test). Medians are highlighted in red. Boxes represent the quartiles and
whiskers extend to the last value within 1.5 interquartile range from the quartiles. Cross (ⴙ) represents a value at 581 cells/mm2. Abbreviation as in Figure 1.
maneuver, only 1% to 3% of the cells remain on site (1,41),
and there is a pressing need for targeting strategies (42).
We have based our cell targeting technique on a biodegradable iron oxide nanoparticle preparation, which should
facilitate transition to human studies, in contrast to other
nonbiodegradable agents used in cell tracking (43,44) and
targeting applications (14 –16). Previous investigations have
indicated that application of a magnetic force on labeled
cells can distort intracellular membranes (33). We noted
that the application of a magnetic force for 24 h had the
potential to compromise cell viability, an effect that was
dependent on the extent of iron loading. Although we did
not evaluate this in the current study, we anticipate that
higher magnetic forces will be tolerable if applied for shorter
lengths of time. By controlling iron uptake and the specific
design of magnetic devices, we have been able to safely
attract cells without affecting cell viability, apoptosis, or
differentiation in culture.
Most importantly, our data suggest that by tagging
human EPCs with an FDA-approved agent, it is possible to
safely target them in static and flowing conditions in vitro
and in vivo. We have assembled and tested a specialized
external magnetic actuator using an in vitro flow system.
Using this actuator, we have also been able to enhance cell
localization following common carotid artery injury in an
animal model; the increase in engraftment was 5-fold higher
than in control (nontargeted) cells. In this study, a flow
interruption period was implemented, which is similar to
those employed in clinical trials and is a favorable condition
for cell enrichment. Our computer simulations and in vitro
data suggest that in the presence of high flow rates at the
target area, very few cells are captured under nontargeted
conditions, which allows up to 252-fold enhancement in
vitro using magnetic attraction, depending on the distance
of the target tissue. This highlights the potential for
extensive targeting in the cardiovascular system, even in
areas of high blood velocity. Nevertheless, further in vivo
studies are warranted to assess EPC engraftment in a
variety of flowing conditions following peripheral intravenous injection. Our results also emphasize a key issue
regarding external targeting in vivo; invariably, cells
flowing in vessels closer to the magnetic source than the
target tissue will also be targeted (6). However, such cells
should be released following removal of the magnetic
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source in the absence of an underlying biological mechanism to keep them engrafted, as observed both in our in
vitro flow experiments and our noninjured control subjects in vivo.
In the injured arteries, we were not able to provide a
circumferential external magnetic force; previously this
problem has been partially circumvented by axial rotation of
the animal (14). Despite this fact, we did not observe any
consistent preferential targeting of the cells in the arterial
specimens. This may be due to a combination of factors.
The broad distribution of magnetic targeting forces due to
the large size of the magnetic actuator compared with the
size of the artery (at least 100o arc) is likely to disperse the
targeted cells. Small within-vessel variations in the extent of
damage, unrelated of the direction of the magnetic force,
could further facilitate nondirectional engraftment. In addition, the flow during the 24-h recirculation period may
enable cells to migrate away from the targeted regions to
areas of more pronounced damage. Indeed, in a recent
study, despite the presence of an intravascular permanently
magnetized device for 24 h, iron-labeled cells were present
in the damaged areas adjacent to the device, as well as on
it (15).
The magnetic control of cells inside the vasculature is a
fascinating and promising concept. In this preliminary
report, we have outlined the feasibility of external targeting
in the arterial circulation using a clinical-grade agent,
bringing this technology a step closer to intervention in
humans. Long-term studies will be necessary to investigate
the retention of the cells at the site of engraftment,
confirming a functionally beneficial increase in healing
following magnetic targeting. Ultimately, we envisage concurrent MR imaging and MR-based magnetic guidance of
labeled cells in the body, possibly in a manner similar to
magnetic guidance of intravascular catheters (45) and magnetic beads (46). This approach could augment localization
and simultaneous monitoring of cells in other organ systems
such as the heart or the brain and may prove complementary
to the systemic injection of cell therapies, thus expanding
the horizon of cardiovascular interventions and the future of
stem cell therapeutic strategies.
Reprint requests and correspondence: Dr. Mark Lythgoe, Centre for Advanced Biomedical Imaging, University College London,
72 Huntley Street, WC1E 6DD London, United Kingdom.
E-mail: mlythgoe@ich.ucl.ac.uk.
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Key Words: endothelial cells 䡲 local delivery 䡲 magnet 䡲
targeting 䡲 angioplasty.
APPENDIX
For more details regarding the methods and calculations used in this study,
please see the online version of this article.