Cardiac repair with intramyocardial injection
of allogeneic mesenchymal stem cells
after myocardial infarction
Luciano C. Amado*†, Anastasios P. Saliaris*†, Karl H. Schuleri*†, Marcus St. John*, Jin-Sheng Xie*, Stephen Cattaneo‡,
Daniel J. Durand*†, Torin Fitton‡, Jin Qiang Kuang§¶, Garrick Stewart*, Stephanie Lehrke*†, William W. Baumgartner‡,
Bradley J. Martin§¶, Alan W. Heldman*†, and Joshua M. Hare*†?
*Department of Medicine, Cardiology Division, and‡Department of Surgery, Division of Cardiac Surgery, The Johns Hopkins Hospital, Blalock 618, 600 North
Wolfe Street, Baltimore, MD 21287;†Institute for Cell Engineering, Broadway Research Building, Suite 651, 733 North Broadway,The Johns Hopkins
University School of Medicine, Baltimore, MD 21205; and§Osiris Therapeutics, 2001 Aliceanna Street, Baltimore, MD 21231-3043
Communicated by Victor A. McKusick, The Johns Hopkins University School of Medicine, Baltimore, MD, June 2, 2005 (received for review April 13, 2005)
Although clinical trials of autologous whole bone marrow for
cardiac repair demonstrate promising results, many practical and
mechanistic issues regarding this therapy remain highly contro-
versial. Here, we report the results of a randomized study of
bone-marrow-derived mesenchymal stem cells, administered to
pigs, which offer several new insights regarding cellular cardio-
myoplasty. First, cells were safely injected by using a percutane-
ous-injection catheter 3 d after myocardial infarction. Second,
cellular transplantation resulted in long-term engraftment, pro-
found reduction in scar formation, and near-normalization of
cardiac function. Third, transplanted cells were preprepared from
an allogeneic donor and were not rejected, a major practical
advance for widespread application of this therapy. Together,
these findings demonstrate that the direct injection of cellular
grafts into damaged myocardium is safe and effective in the
periinfarct period. The direct delivery of cells to necrotic myocar-
dium offers a valuable alternative to intracoronary cell injections,
and the use of allogeneic mesenchymal stem cells provides a
valuable strategy for cardiac regenerative therapy that avoids the
need for preparing autologous cells from the recipient.
myocardium. Small clinical trials conducted in the perimyocardial
infarction (MI) period (1–5) with intracoronary infusion of autol-
ogous whole bone marrow preparations have suggested moderate
improvements in cardiac function (1–3). However, recent experi-
mental studies questioning the engraftment of hematopoietic stem
cells (6, 7) and clinical findings that bone marrow cells do not
engraft in the infarct zone or reduce infarct size have lead to
controversy regarding the mechanism behind these promising
Despite these concerns, there is increasing evidence that a
specific bone marrow constituent, the mesenchymal stem cell
(MSC), has cardiac reparative properties. MSCs engraft (9), have
and growth factors that stimulate endogenous repair mechanisms
(11, 12). Furthermore, MSCs have several properties that contrib-
ute to an ability to evade rejection (13–15). In this regard, they lack
cell-surface B-7 costimulatory molecules (16, 17) and may also
serve as an allogeneic graft, thereby avoiding the need for bone
marrow harvesting from prospective recipients, an extraordinary
therapeutic advantage for this cell type.
An additional consideration for the development of cellular
infusions of cells do not appear to infiltrate the MI zone or reduce
MI size (3). Accordingly, the direct delivery of cells into the area of
tissue necrosis may circumvent this potential limitation of cellular
derived cell-based therapies to repair or regenerate damaged
Here, to address the hypothesis that MSCs reduce MI size and
blinded, placebo-controlled trial of MSCs in pigs after MI. Addi-
tional studies were performed by using cardiac MRI. The aims of
this study were to (i) demonstrate that stem cells can be safely
administered directly to damaged myocardium via injection cath-
eter, (ii) test the efficacy of preprepared allogeneic MSCs as a
the mechanism of benefit is cellular engraftment, which results in
cardiac myocyte regeneration, reduced infarct size, and improved
Animal Model. AllanimalstudieswereapprovedbytheInstitutional
Animal Care and Use Committee and comply with the Guide for
the Care and Use of Laboratory Animals (National Institutes of
were conducted. First, we performed a randomized study of
animals (n ? 14) that underwent surgical induction of MI and
cardiac oxygen consumption. Second, a group of chronically in-
strumented animals (n ? 4) were submitted to MI by balloon
occlusion of the left anterior descending coronary artery (LAD),
after recovery from surgery, so that measures of normal cardiac
studied noninvasively with MRI after MI induced by balloon
occlusion of the LAD.
Surgical Preparation. Female Yorkshire pigs underwent surgical
instrumentation for subsequent noninvasive measurement of LV
pressure and dimension and myocardial oxygen consumption (19,
and great cardiac vein. Endocardial ultrasound crystals (Sonomet-
rics, Ontario, Canada) were inserted to measure short-axis dimen-
cava for graded preload reduction to assess LV-pressure–
dimension relations. A 4–5 mm flow probe (Transonics, Ithaca,
flow. A solid-state miniature pressure transducer (P22, Konigsberg
Instruments, Pasadena, CA) was placed in the LV apex for high-
fidelity recordings of LV pressure. Additional pacing leads were
Abbreviations: Ees, ventricular elastance, slope of the end-systolic pressure–dimension
relationship; LAD, left anterior descending coronary artery; LV, left ventricular; MI, myo-
cardial infarction; MSC, mesenchymal stem cell; MVO2, myocardial oxygen consumption
per cardiac cycle; SW, stroke work.
¶J.Q.K. and B.J.M. are employees of Osiris Therapeutics.
?To whom correspondence should be addressed. E-mail: email@example.com.
© 2005 by The National Academy of Sciences of the USA
August 9, 2005 ?
vol. 102 ?
During surgery, MI was induced by a 60-min occlusion of the
LAD, followed by reperfusion. Stainless steel surgical clips were
placed to delineate the injured myocardial area. At 3 d after MI,
animals were randomized to receive intramyocardial injections of
either allogeneic porcine MSCs (2.0 ? 108cells, n ? 7) or placebo
7) under fluoroscopy, with a helical-needle-tipped injection cath-
eter advanced to the left ventricle through a steerable guide
catheter (BioCardia, South San Francisco, CA). MSC injections
were guided by the presence and location of the surgical markers.
Normal hemodynamic values in the swine were obtained from four
additional animals after surgical preparation and instrumentation
by a 1-h occlusion of the LAD with a balloon angioplasty catheter
Hemodynamic and Energetic Measurements. Pressure–dimension
data were recorded at steady state and during transient inferior
vena cava occlusion. Myocardial contractility and?or work were in-
dexed by the maximal rate of isovolumetric contraction (?dP?dt),
stroke work (SW), and ventricular elastance, slope of the end-
systolic pressure–dimension relationship (Ees) (21). Preload was
evaluated as effective arterial elastance (22), the ratio of LV
end-systolic pressure to stroke dimension. Diastolic function was
indexed by LV end-diastolic pressure and the time constant of
ventricular relaxation (?; Glantz formula) (23). Hemodynamic
pressure–dimension data were digitized at 200 Hz and stored for
subsequent analysis on a personal computer by using custom
software. Myocardial oxygen consumption per cardiac cycle
(MVO2) was calculated from the arteriovenous difference of oxy-
blood, multiplied by LAD flow and divided by heart rate. Cardiac
mechanical efficiency was calculated as the SW?MVO2ratio (20).
MRI. To gain a better understanding of the effect of MSC therapy
in the heart, infarct size and global function were assessed by MRI.
MI was induced under fluoroscopic guidance: A catheter sheath (8
French) was placed in the right carotid artery, through which a
coronary angioplasty balloon (3.5 French, 20 mm) was advanced
into the proximal LAD. MI was created by inflating the balloon for
60 min, after which the balloon was deflated and the artery
reperfused. After reperfusion, the catheter sheath in the carotid
At 3 d after MI, animals were randomized to receive (n ? 6) or not
Scientific, Natick, MA). An additional group of animals (n ? 6)
received Feridex-labeled (Berlex Laboratories) MSCs, so as to
visualize the cells by MRI. Cell retention was estimated by quan-
tifying the presence and intensity of the hypoenhanced regions
caused by the presence of iron-oxide-labeled cells.
MRI images were acquired by using a 1.5-T MR scanner (CV?i,
GE Medical Systems, Waukesha, WI) at four time points after
injection, 2 d and 1, 4, and 8 weeks. To assess engraftment of the
MRI-labeled MSCs, high-resolution MRI images were obtained as
acquired, covering the entire left ventricle, by using fast-gradient
recoiled echo (24). Imaging parameters were as follows: repetition
time (TR) ? 8.7 ms; echo delay time (TE) ? 2.4 ms, flip angle ?
20°; 512 ? 512 matrix; 5-mm slice thickness?no gap; 32-kHz
bandwidth; 28-cm FOV and 3 number of acquisitions (NSA).
injection of contrast [Magnevist, Berlex Laboratories, gadolinium
diethylenetriamine-pentaacetate dimeglumine (Gd-DTPA); 0.2
mmol?kg of body weight] by using an ECG-gated, breath-hold,
interleaved, inversion recovery, fast-gradient recoiled echo pulse
sequence. A total of 8–10 contiguous short-axis slices were pre-
parameters were as follows: TR?TE?inversion recovery time
matrix; 8-mm slice thickness?no gap; 31.2 kHz; 28-cm FOV and 2
NSA. Inversion recovery time was adjusted as needed to null the
normal myocardium (25).
Global LV function was assessed by using a steady-state free-
precession pulse sequence (26). Short-axis cine images were ac-
quired at the same location as the DE-MRI images. Image param-
256 ? 160 matrix; 8-mm slice thickness?no gap; 125 kHz; 28-cm
FOV and 1 NSA. Images were analyzed by using a custom research
software package (CINE TOOL, GE Medical Systems). Infarct
areas were defined based on the full-width at half-maximum
Stem Cell Harvest and Isolation. Male swine MSCs were obtained,
isolated, and expanded as described in ref. 9. Briefly, bone marrow
At 5–7 d after plating, hematopoetic and other nonadherent cells
MSC population was expanded in culture. All used cells were
harvested when they reached 80–90% confluence at passage 3.
DMSO, 5% porcine serum albumin, and 85% Plasmalyte. Cells
were placed in cryo bags at a concentration of 5–10 million MSCs
per ml and frozen in a control-rate freezer to ?180°C until the day
of implantation. By using trypan blue staining, the viability of all
thawed MSC lots was verified to be ?85% before use in the study.
To ensure that the grafts were allogeneic, the MSC donor animals
were of a different strain (Durok–Landrace) than the recipients
Cell Engraftment. Cell engraftment was assessed in the preparation
of both surgical and nonsurgical (MRI) animals. First, in the
surgically instrumented animals, MSCs were labeled after thawing
with the cross-linkable membrane dye CM-DiI and the nuclear dye
DAPI (Molecular Probes) according the manufacturer’s protocol,
at a concentration of 2 ?g?ml cell suspension. Before injection, the
cells were thoroughly washed in serum-free DMEM, resuspended
in 10 ml of PBS, and held on ice at an approximate concentration
of 20 million cells per ml. Second, MRI studies were performed to
8-week study period. MSCs were magnetically labeled before
ml, Feridex, Berlex Laboratories) (24). The presence of labeled
MSCs within the myocardium was assessed by MRI at 2 d after
injection and, subsequently, at 1, 4, and 8 weeks. Feridex labeling
was also confirmed by Prussian blue stain immediately after
Immunohistochemistry. The animals were followed for an 8-week
period and were then humanely killed. Their hearts were analyzed
at both microscopic and gross levels. Myocardial tissue was pre-
pared for immunohistochemistry, as described in ref. 28. Tissue
samples were obtained from three specific areas: infarct zone,
infarct border zone, and remote tissue. Antibodies examined in-
cluded ?-actinin (Sigma), troponin-T (Sigma), tropomyosin
(Sigma), myosin heavy chain-MHC (Developmental Studies Hy-
(Affinity BioReagents, Golden, CO). Ki67 and c-kit stains (Ven-
tana Medical Systems, Tucson, AZ) were performed to assess the
presence of active dividing and progenitor cells, respectively.
Amado et al.
August 9, 2005 ?
vol. 102 ?
no. 32 ?
Infarct Size by Gross Pathology. Myocardial fibrosis was determined
purpose, hearts were excised and sectioned into 8-mm-thick short-
axis slices. Each slice was weighed and digitally photographed.
Analysis was performed in fresh, nonstained hearts. TTC (2,3,5-
triphenyltetrazolium chloride) staining was avoided, so as to pre-
serve the myocardial tissue for engraftment-histological analysis.
We have previously validated that quantification of infarct size is
setting of chronic scarred tissue (data not shown). Infarcted areas
and LV borders were manually traced for each slice by using a
custom research software package IMAGE ANALYSIS 4.0.2 beta ver-
sion (Scion, Frederick, MD). Infarct size was determined, in a
blinded fashion, as percentage of LV mass from the digital pictures
and normalized by the weight of the slice.
Statistical Analysis. All analyses were performed blind to the
randomization. All values are expressed as mean ? SEM. Hemo-
dynamic parameters at baseline were compared between groups by
using Student’s t test. Differences in the hemodynamic indices
repeated measurements ANOVA and between groups by using
two-way ANOVA with an interaction term. A level of P ? 0.05 was
considered statistically significant.
MSC Intramyocardial Injection. MSCs were injected via LV cathe-
terization under fluoroscopic guidance in both surgically instru-
mented and MRI animals. The surgically instrumented animals
(n ? 14) were randomized to receive intramyocardial injections of
(13.4 ? 1, ?0.5 ml each). Additional animals (n ? 18) underwent
serial MRI imaging. At 3 d after MI, no animal had died, exhibited
malignant arrhythmias, or demonstrated evidence of cardiac per-
of our delivery approach.
MSC Engraftment. Engraftment was monitored by both MRI and
histological evaluation. MRI imaging of pigs receiving Feridex-
labeled cells (n ? 6) demonstrated a gradual loss of the intensity of
the iron oxide label over the 2-month period, yet there was
significant retention of implanted MSCs, with 42.4 ? 15% of the
magnetic label detected at the end of 8 weeks (Fig. 1; P ? 0.05 vs.
2 d postinjection). Histologic evaluation of the infarct, border zone,
and remote myocardium was performed in the surgically instru-
mented animals that received Di-I- and DAPI-labeled cells. MSCs
labeled with these membrane and nuclear dyes, respectively, (n ?
7) were present throughout the infarct regions and border zones up
to 8 weeks after implantation and expressed muscle-specific pro-
teins not expressed by cultured MSCs (Fig. 2). MSCs were also
detected in vascular structures, where they expressed vascular
endothelium growth factor and Von Willebrand factor, suggesting
differentiation and?or incorporation into vascular smooth muscle
and?or endothelium (data not shown). Extracardiac MSC engraft-
ment, differentiation into ectopic tissues (adipose, bone, cartilage,
Impact of MSC Treatment on Infarct Size. Gross pathologic exami-
nation revealed that MSC treatment substantially decreased the
percentage of left ventricle occupied by fibrosis (Fig. 3A). In the
surgical group, placebo pigs exhibited scar tissue occupying 16 ?
4.6% of the left ventricle, and infarcts were transmural, beginning
subendocardially and extending ?50% of the LV-wall thickness, a
finding similar to that observed in humans surviving MI with
reperfusion of infarct-related arteries (29). However, in MSC-
treated pigs, only 3.4 ? 1.2% of the left ventricle was occupied by
scar tissue (P ? 0.008 vs. placebo, Fig. 3 A and B). Interestingly, in
the MSC-treated group, the infarct region was confined to the
midmyocardium, and viable tissue was clearly visible at both the
subendocardial and subepicardial zones (Fig. 3 C and D). The
subendocardial tissue rim is consistent with the growth of new
tissue, because myocardial infarction is known to develop in a
wave-front pattern, progressing from the subendocardium toward
the subepicardial layer (30). We further substantiated this obser-
rim on gross pathology in treated vs. control animals. Indeed, the
rim was substantially thicker in MSC-treated vs. nontreated ani-
mals (2.4 ? 0.2 and 1.5 ? 0.2 mm, respectively; n ? 6 each, Fig. 3E,
P ? 0.05).
To delineate mechanisms for the growth of this tissue rim, we
evaluated MSC-injected tissue 10 d after cell injection. Immuno-
histochemical analysis of MI and MI border zones in MSC-treated
animals revealed the presence of a band of myocytes positive for
Pseudolong-axis image obtained after MSC injection. Magnetically labeled
MSCs appear as hypoenhanced areas (arrows) visible at 2 d and at 1, 4, and 8
weeks after injection. (Right) Bar graph illustrating Feridex retention over 8
weeks. (B) Histologic evaluation of postinfarct myocardium. (Left) Hematox-
ylin?eosin (H&E) stains obtained from noninjected infarcted area, demon-
strating an abundance of inflammatory cells (filled arrows) surrounding the
necrotic region (?160 magnification). (Right) H&E and Prussian blue (PB)
Open arrows indicate Feridex (iron)-labeled MSCs.
Engraftment of allogeneic porcine MSCs assessed with MRI. (A)
(blue-staining nuclei and red-staining membranes, respectively) and fluorescent
MSCs in proximity to host myocardium. Several muscle-specific proteins are
detected by immunofluorescence, including ?-actinin (C), phospholamban (D),
of immunofluorescent antibodies and DiI.
Engraftment and differentiation of MSCs. DAPI- and Di-I-labeled MSCs
www.pnas.org?cgi?doi?10.1073?pnas.0504388102Amado et al.
c-kit- and Ki-67-staining cells 10 d after MI (Fig. 4). This band was
not present in control animals, although both groups contained
c-kit-positive hematopoeitic cells in capillaries. This finding is
consistent with both MSC differentiation and a paracrine effect,
stimulating endogenous cellular replication, both of which are
MRI. Delayed-enhancement MRI confirmed the decrease in MI
size observed at the time of postmortem analysis. MSC treatment
the course of the study (from 20.7 ? 3.5 to 9.9 ? 1.3% of the left
ventricle, at 3 d and at 8 weeks, respectively; P ? 0.05, n ? 6),
2F, P ? 0.05 vs. MSC group).
MSC Therapy Effect on Cardiac Function. We next examined whether
this new tissue formation translated into functional recovery and
measured the effect of MSC therapy on post-MI hemodynamics in
1 depicts indices of ventricular function before MI creation and at
3 d after MI in the two treatment groups (MSC or placebo
injections). As shown, there is a sizable and nearly identical
function (increased LVPED and ?) (P ? 0.05 vs. normal pre-MI
values), in both groups, indicative of equivalent degrees of injury.
In the placebo-treated group (n ? 7), impaired cardiac function
evident at 3 d after MI showed either no sign of recovery or a
contraction fell, and end-diastolic pressure rose, as did ? (Table 1
and Fig. 5). In marked contrast, animals receiving MSCs (n ? 7)
exhibited recovery to essentially normal levels of both systolic [Ees
(? fell to 34.2 ? 1.2 msec; Fig. 5). LV end-diastolic pressure did not
return to normal levels at 8 weeks, although a significant improve-
ment was observed (P ? 0.05 vs. placebo group; Table 1).
To further evaluate the effect of MSC therapy on global cardiac
function, MRI determination of LV ejection fraction (LVEF) was
performed. Immediately after MI, there was a marked deteriora-
example of MI scar formation in placebo-treated (Upper) and MSC-treated
(Lower) animals at 8 weeks after injury. MSC injection reduces the scar, which is
confined to the midmyocardium because of viable tissue in subendocardial and
in MSC-treated vs. placebo-treated pigs in a randomized study (*, P ? 0.008). (C)
MSC cardiomyoplasty augments development of new myocardium (Left). At 8
weeks after MI, the subendocardial rim is thicker in the MSC group (arrows).
Hematoxylin?eosin (H&E) stain of the subendocardial rim demonstrates cardio-
myocytes in both control and MSC (?100 magnification). (D) Short-axis delayed-
At 1 week after MSC injection, there is anteroseptal myocardial necrosis and
extensive microvascular obstruction (white arrow in Left). MSC administration
leads to myocardial tissue regeneration (red arrows) and reduction of necrotic
myocardium (yellow arrows) 8 weeks later (Center). (Left) Corresponding gross
tissue with MSCs (*, P ? 0.05 vs. nontreated) after 8 weeks. (F) MI size assessed
from MRI delayed hyperenhancement, showing that, in nontreated animals,
infarct size does not decrease but is reduced ?50% with MSC treatment (*, P ?
0.05 vs. baseline images obtained 2 d after therapy; †, P ? 0.05 vs. nontreated).
in both groups and within capillaries (open arrow) in MSC but not in control
group. (B) Ki67-positive myocytes present within MSC-treated heart.
Myocardial regeneration in MSC-treated hearts. Immunohistochemical
Amado et al.
August 9, 2005 ?
vol. 102 ?
no. 32 ?
tion in LVEF, (25.3 ? 1.6 and 29.8 ? 1.9% for MSC- and
was minimal change in LVEF in the nontreated group (n ? 6),
LVEF increased from 25.3 ? 1.6 to 41.9 ? 0.7% in MSC-treated
which is published on the PNAS web site) during the 8-week
Post-MI heart failure is characterized by mechanoenergetic
uncoupling, a phenomenon in which MVO2 increases, despite
decreased cardiac work (33, 34). In both groups, cardiac energy
metabolism was impaired after MI (P ? 0.05 vs. normal pre-MI
values; Table 1 and Fig. 5). In placebo-treated animals, SW
decreased substantially during 4 weeks, accompanied by a para-
doxical gain in MVO2(Table 1), both factors together depressing
the SW?MVO2ratio (Fig. 5). Conversely, MSC therapy signifi-
cantly improved myocardial efficiency, increasing SW (from
374.4 ? 59.3 to 654.4 ? 129.3 mmHg?mm at 8 weeks; Table 1),
while simultaneously decreasing MVO2(from 10.3 ? 2 to 3.7 ? 1.8
J per beat), both toward normal (Table 1). Thus, MSC therapy
exerts favorable effects on the damaged heart that extends to
2.5 ? 0.6 at 3 d after MI to a normal level of 10 ? 5.6 (P ? 0.05
vs. placebo; Fig. 5) at 4 weeks. The restoration in mechanoener-
getics was the earliest observable benefit of MSC treatment,
preceding the changes in global cardiac function.
The present work addresses major issues regarding cardiac regen-
erative therapy. Here, we show that, in pigs, allogeneic MSCs can
be safely and effectively delivered via injection catheter to a region
of damaged myocardium, where they engraft and express proteins
normally restricted to cardiac myocytes, vascular endothelium, and
smooth muscle. Engrafted MSCs dramatically reduce the extent of
necrotic myocardium and promote the regeneration of new, con-
tractile myocardium along the subendocardial surface of the MI.
followed by near-normalization of systolic and diastolic cardiac
function and substantial increases in global cardiac performance.
Taken together, these findings offer therapeutic promise for the
application of cellular therapeutics to myocardial infarction. Im-
portantly MSCs have the potential to be administered as a prepre-
pared allogeneic graft, avoiding the need to perform bone marrow
harvests on recipient patients.
Both experimental (6, 35) and clinical (1, 3) studies have con-
tributed to the controversy surrounding the extent and mechanism
of cardiac repair resulting from bone marrow cellular therapy. In
experiments conducted on mice, bone marrow c-kit-positive cells
have the capacity to engraft in damaged myocardium and differ-
entiate into cardiac myocytes (35, 36). On the other hand, several
neither (6, 7). Several small clinical trials demonstrate that autol-
ogous whole bone marrow infused into the coronary artery mar-
ginally improves the ejection fraction (3). Importantly, whole bone
marrow cells are reported by positron emission tomography scan-
ning to be unable to infiltrate areas of myocyte necrosis (8).
Table 1. Hemodynamic measurements
3 d post-MI8 weeks post-MI
LV end-diastolic pressure, mm Hg
LV end-systolic pressure, mm Hg
Arterial elastance, mm Hg?mm
dP?dtmax, mm Hg?s
Ees, mm Hg?mm
SW, mm Hg?mm
MVO2, J per beat
8.4 ? 2.3
107.1 ? 4.2
14 ? 2
2,560 ? 266
16.3 ? 2.4
36.2 ? 1.8
771 ? 116.5
3.2 ? 0.9
9.1 ? 1.6
21.5 ? 5.8*
98.8 ? 6.6
20 ? 2.5
1,734 ? 218*
11.2 ? 0.5*
42 ? 1*
434.8 ? 59.9*
7.4 ? 1.4*
3.8 ? 0.8*
26 ? 3.6*
112.6 ? 5.6
24.8 ? 4.2
1,802 ? 80*
8.1 ? 3.1*
44.3 ? 2.3*
374.4 ? 59.3*
10.3 ? 2*
2.5 ? 0.6*
29.8 ? 7.6
117.7 ? 22.2
26.1 ? 8.7
1,720 ? 351
7.9 ? 1.2
52.6 ? 11.6
470.3 ? 86.9
12.9 ? 1.4‡
2.9 ? 0.1‡
20 ? 6.4†
113.8 ? 5.8
17.1 ? 3
2,465 ? 574†
17.1 ? 2†
34.2 ? 1.2†
654.4 ? 129.3†
3.7 ? 1.8‡
10 ? 5.6‡
MVO2indicates myocardial oxygen consumption and SW?MVO2myocardial efficiency.
*P ? 0.05 vs. normal (pre-MI).
†P ? 0.05 vs. placebo (2-way ANOVA).
‡Listed values were measured at 4 weeks.
NS for differences between groups at day 3 after MI). (Top) Ventricular
elastance (Ees) declines in placebo-treated pigs but increases dramatically in
the MSC group, with complete restoration of Ees to normal levels at 8 weeks
(P ? NS vs. normal). (Middle) Isovolemic ventricular relaxation (?) returns to
normal in MSC-treated pigs but remains impaired in placebo-treated pigs.
(Bottom) The impact of MSC therapy on myocardial mechanoenergetics
MVO2increases, leading to a reduced SW?MVO2ratio. In MSC-treated pigs,
SW increases and MVO2decreases, resulting in augmented SW?MVO2and
restoration of mechanoenergetic coupling toward normal [P ? 0.05 vs. D3,
‡, P ? 0.05 between groups by two-way ANOVA].
www.pnas.org?cgi?doi?10.1073?pnas.0504388102 Amado et al.
Based on concerns regarding the ability of bone marrow prep- Download full-text
arations to enter the region of damaged myocardium (8), we
predicted that direct injection would enhance cell engraftment.
Indeed, catheter-based cell injection was safe and highly effective,
producing areas of regenerated myocardium and diminishing the
amount of fibrotic tissue. Functionally, this therapy essentially
normalized systolic and diastolic cardiac function. This approach
represents a potential advance over the majority of clinical trials,
which have used intracoronary techniques for cell delivery, have
variable success in reducing infarct size (1–4, 37). The relative
contribution to the degree of recovery we report here of cell type
vs. the delivery method remains unclear. It should be mentioned
that previous reports suggest that intracoronary infusion of MSCs
to cause microvascular obstruction (38). On the other hand, there
is a possibility that intravenously administered MSCs may track to
the heart in response to injury signals (39).
This study demonstrates the potential therapeutic effects of
from bone marrow and, because of their multilineage potential (9,
10) and immunological advantages (16–18), represent ideal candi-
dates for allogeneic cell therapy. Our findings are supported by
earlier studies in rats, in which MSCs, genetically modified to
survive in ischemic myocardium, decreased infarct size (40). The
present results indicate that unmodified allogeneic cells are thera-
for the application of this therapy. Indeed, clinical trials for allo-
geneic MSC therapy have received FDA approval.
The mechanism by which cellular therapy limits the extent of
damaged myocardium after ischemic insult remains highly contro-
versial, and three general mechanisms are proposed: transdiffer-
entiation (35, 40), cell fusion (41), and paracrine signaling (42). It
may be argued, however, that the net result of effective cellular
therapy is a reduction in size of the MI and restoration of myocar-
dial contraction. Here, we describe evidence of cardiac myocyte
regeneration. Our results are consistent with the presence of both
mechanisms, transdifferentiation of transplanted MSCs into the
cardiac myocyte lineage and increased endogenous repair mecha-
and paracrine effects are operative in cellular regenerative strate-
provides a plausible target for paracrine signaling due to injected
MSCs (44–47). Our findings that MSCs express VEGF, which is
linked to both neoangiogenesis (48) and stem cell homing and
migration (11, 49), also support a contribution of paracrine stim-
ulation of endogenous repair mechanisms.
cardiomyoplasty. We did not observe this complication, rather,
noting a trend toward decreased sudden death in MSC-treated
animals (L.C.A. and J.M.H., unpublished observations), suggesting
that the implantation of MSCs into infarcted myocardium is safe
and, perhaps, even protects from postinfarct arrythmias.
In summary, allogeneic MSCs injected into regions of damaged
myocardium 3 d after MI engraft, stimulate cardiac regeneration,
and profoundly decrease myocardial infarct size. As a result, there
is an early restoration of cardiac energy metabolism, followed by
in both systolic and diastolic cardiac function. The successful
application of allogeneic cells injected intramyocardially offers
insights for practical application of this therapy in future clinical
This work was supported by The Johns Hopkins University School of
Medicine Institute for Cell Engineering, Osiris Therapeutics, The
Donald W. Reynolds Foundation, and National Institutes of Health
Grants R21 HL-72185 and R01 NIA AG025017.
1. Schachinger, V., Assmus, B., Britten, M. B., Honold, J., Lehmann, R., Teupe, C., Abolmaali,
N. D., Vogl, T. J., Hofmann, W. K., Martin, H., et al. (2004) J. Am. Coll. Cardiol. 44, 1690–1699.
2. Britten, M. B., Abolmaali, N. D., Assmus, B., Lehmann, R., Honold, J., Schmitt, J., Vogl,
T. J., Martin, H., Schachinger, V., Dimmeler, S., et al. (2003) Circulation 108, 2212–2218.
3. Wollert, K. C., Meyer, G. P., Lotz, J., Ringes-Lichtenberg, S., Lippolt, P., Breidenbach, C.,
Fichtner, S., Korte, T., Hornig, B., Messinger, D., et al. (2004) Lancet 364, 141–148.
4. Assmus, B., Schachinger, V., Teupe, C., Britten, M., Lehmann, R., Dobert, N., Grunwald,
F., Aicher, A., Urbich, C., Martin, H., et al. (2002) Circulation 106, 3009–3017.
5. Strauer, B. E., Brehm, M., Zeus, T., Kostering, M., Hernandez, A., Sorg, R. V., Kogler, G.
& Wernet, P. (2002) Circulation 106, 1913–1918.
6. Murry, C. E., Soonpaa, M. H., Reinecke, H., Nakajima, H., Nakajima, H. O., Rubart, M.,
Pasumarthi, K. B., Ismail, V. J., Bartelmez, S. H., Poppa, V., et al. (2004) Nature 428, 664–668.
7. Balsam, L. B., Wagers, A. J., Christensen, J. L., Kofidis, T., Weissman, I. L. & Robbins, R. C.
(2004) Nature 428, 668–673.
8. Hofmann, M., Wollert, K. C., Meyer, G. P., Menke, A., Arseniev, L., Hertenstein, B.,
Ganser, A., Knapp, W. H. & Drexler, H. (2005) Circulation 111, 2198–2202.
9. Pittenger, M. F., Mackay, A. M., Beck, S. C., Jaiswal, R. K., Douglas, R., Mosca, J. D.,
Moorman, M. A., Simonetti, D. W., Craig, S. & Marshak, D. R. (1999) Science 284, 143–147.
10. Makino, S., Fukuda, K., Miyoshi, S., Konishi, F., Kodama, H., Pan, J., Sano, M., Takahashi,
T., Hori, S., Abe, H., et al. (1999) J. Clin. Invest. 103, 697–705.
11. Tang, Y. L., Zhao, Q., Zhang, Y. C., Cheng, L., Liu, M., Shi, J., Yang, Y. Z., Pan, C., Ge,
J. & Phillips, M. I. (2004) Regul. Pept. 117, 3–10.
12. Kucia, M., Dawn, B., Hunt, G., Guo, Y., Wysoczynski, M., Majka, M., Ratajczak, J.,
Rezzoug, F., Ildstad, S. T., Bolli, R., et al. (2004) Circ. Res. 95, 1191–1199.
13. Bartholomew, A., Sturgeon, C., Siatskas, M., Ferrer, K., McIntosh, K., Patil, S., Hardy, W.,
Devine, S., Ucker, D., Deans, R., et al. (2002) Exp. Hematol. 30, 42–48.
14. Le Blanc, K., Tammik, L., Sundberg, B., Haynesworth, S. E. & Ringden, O. (2003) Scand.
J. Immunol. 57, 11–20.
15. Tse, W. T., Pendleton, J. D., Beyer, W. M., Egalka, M. C. & Guinan, E. C. (2003)
Transplantation 75, 389–397.
16. Majumdar, M. K., Keane-Moore, M., Buyaner, D., Hardy, W. B., Moorman, M. A.,
McIntosh, K. R. & Mosca, J. D. (2003) J. Biomed. Sci. 10, 228–241.
18. Di Nicola, M., Carlo-Stella, C., Magni, M., Milanesi, M., Longoni, P. D., Matteucci, P.,
Grisanti, S. & Gianni, A. M. (2002) Blood 99, 3838–3843.
19. Saavedra, W. F., Paolocci, N., St. John, M. E., Skaf, M. W., Stewart, G. C., Xie, J. S.,
Harrison, R. W., Zeichner, J., Mudrick, D., Marban, E., et al. (2002) Circ. Res. 90, 297–304.
20. Ekelund, U. E., Harrison, R. W., Shokek, O., Thakkar, R. N., Tunin, R. S., Senzaki, H., Kass,
D. A., Marban, E. & Hare, J. M. (1999) Circ. Res. 85, 437–445.
21. Hare, J. M., Kass, D. A. & Stamler, J. S. (1999) Nat. Med. 5, 1241–1242.
22. Kelly, R. P., Ting, C.-T., Yang, T.-M., Liu, C.-P., Maughan, W. L., Chang, M.-S. & Kass,
D. A. (1992) Circulation 86, 513–521.
23. Gilbert, J. C. & Glantz, S. A. (1989) Circ. Res. 64, 827–852.
24. Kraitchman, D. L., Heldman, A. W., Atalar, E., Amado, L. C., Martin, B. J., Pittenger, M. F.,
Hare, J. M. & Bulte, J. W. (2003) Circulation 107, 2290–2293.
25. Simonetti, O. P., Kim, R. J., Fieno, D. S., Hillenbrand, H. B., Wu, E., Bundy, J. M., Finn,
J. P. & Judd, R. M. (2001) Radiology 218, 215–223.
26. Slavin, G. S. & Saranathan, M. (2002) Magn. Reson. Med. 48, 934–941.
27. Amado, L. C., Gerber, B. L., Gupta, S. N., Rettmann, D. W., Szarf, G., Schock, R., Nasir,
K., Kraitchman, D. L. & Lima, J. A. (2004) J. Am. Coll. Cardiol. 44, 2383–2389.
28. Shake, J. G., Gruber, P. J., Baumgartner, W. A., Senechal, G., Meyers, J., Redmond, J. M.,
Pittenger, M. F. & Martin, B. J. (2002) Ann. Thorac. Surg. 73, 1919–1925.
29. Christian, T. F., Schwartz, R. S. & Gibbons, R. J. (1992) Circulation 86, 81–90.
30. Garcia-Dorado, D., Theroux, P., Desco, M., Solares, J., Elizaga, J., Fernandez-Aviles, F.,
Alonso, J. & Soriano, J. (1989) Am. J. Physiol. 256, H1266–H1273.
31. Dimmeler, S., Zeiher, A. M. & Schneider, M. D. (2005) J. Clin. Invest. 115, 572–583.
32. Pittenger, M. F. & Martin, B. J. (2004) Circ. Res. 95, 9–20.
33. Trines, S. A., Slager, C. J., Onderwater, T. A., Lamers, J. M., Verdouw, P. D. & Krams, R.
(2001) Cardiovasc. Res. 51, 122–130.
34. Hayashi, Y., Takeuchi, M., Takaoka, H., Hata, K., Mori, M. & Yokoyama, M. (1996)
Circulation 93, 932–939.
35. Kajstura, J., Rota, M., Whang, B., Cascapera, S., Hosoda, T., Bearzi, C., Nurzynska, D.,
Kasahara, H., Zias, E., Bonafe, M., et al. (2004) Circ. Res. 96, 127–137.
36. Orlic, D., Kajstura, J., Chimenti, S., Jakoniuk, I., Anderson, S. M., Li, B., Pickel, J., McKay,
R., Nadal-Ginard, B., Bodine, D. M., et al. (2001) Nature 410, 701–705.
W. (2004) Circ. Res. 94, 230–238.
38. Vulliet, P. R., Greeley, M., Halloran, S. M., MacDonald, K. A. & Kittleson, M. D. (2004)
Lancet 363, 783–784.
39. Nagaya, N., Fujii, T., Iwase, T., Ohgushi, H., Itoh, T., Uematsu, M., Yamagishi, M., Mori,
H., Kangawa, K. & Kitamura, S. (2004) Am. J. Physiol. 287, H2670–H2676.
40. Mangi, A. A., Noiseux, N., Kong, D., He, H., Rezvani, M., Ingwall, J. S. & Dzau, V. J. (2003)
Nat. Med. 9, 1195–1201.
41. Nygren, J. M., Jovinge, S., Breitbach, M., Sawen, P., Roll, W., Hescheler, J., Taneera, J.,
Fleischmann, B. K. & Jacobsen, S. E. (2004) Nat. Med. 10, 494–501.
42. Fraidenraich, D., Stillwell, E., Romero, E., Wilkes, D., Manova, K., Basson, C. T. & Benezra,
R. (2004) Science 306, 247–252.
43. Yoon, Y. S., Wecker, A., Heyd, L., Park, J. S., Tkebuchava, T., Kusano, K., Hanley, A.,
Scadova, H., Qin, G., Cha, D. H., et al. (2005) J. Clin. Invest. 115, 326–338.
44. Beltrami, A. P., Barlucchi, L., Torella, D., Baker, M., Limana, F., Chimenti, S., Kasahara,
H., Rota, M., Musso, E., Urbanek, K., et al. (2003) Cell 114, 763–776.
45. Oh, H., Bradfute, S. B., Gallardo, T. D., Nakamura, T., Gaussin, V., Mishina, Y., Pocius,
J., Michael, L. H., Behringer, R. R., Garry, D. J., et al. (2003) Proc. Natl. Acad. Sci. USA
46. Laugwitz, K. L., Moretti, A., Lam, J., Gruber, P., Chen, Y., Woodard, S., Lin, L. Z., Cai,
C. L., Lu, M. M., Reth, M., et al. (2005) Nature 433, 647–653.
47. Messina, E., De Angelis, L., Frati, G., Morrone, S., Chimenti, S., Fiordaliso, F., Salio, M.,
Battaglia, M., Latronico, M. V., Coletta, M., et al. (2004) Circ. Res. 95, 911–921.
48. Kocher, A. A., Schuster, M. D., Szabolcs, M. J., Takuma, S., Burkhoff, D., Wang, J., Homma,
S., Edwards, N. M. & Itescu, S. (2001) Nat. Med. 7, 430–436.
49. Eriksson, U. & Alitalo, K. (2002) Nat. Med. 8, 775–777.
Amado et al.
August 9, 2005 ?
vol. 102 ?
no. 32 ?