Dose-Dependent Contribution of CD34-Positive Cell Transplantation to Concurrent Vasculogenesis and Cardiomyogenesis for Functional Regenerative Recovery After Myocardial Infarction
Background— Multilineage developmental capacity of the CD34+ cells, especially into cardiomyocytes and smooth muscle cells (SMCs), is still controversial. In the present study we performed a series of experiments to prove our hypothesis that vasculogenesis and cardiomyogenesis after myocardial infarction (MI) may be dose-dependently enhanced after CD34+ cell transplantation.
Methods and Results— Peripheral blood CD34+ cells were isolated from total mononuclear cells of patients with limb ischemia by apheresis after 5-day administration of granulocyte colony-stimulating factor. PBS and 1×103 (low), 1×105 (mid), or 5×105 (high) CD34+ cells were intramyocardially transplanted after ligation of the left anterior descending coronary artery of nude rats. Functional assessments with the use of echocardiography and a microtip conductance catheter at day 28 revealed dose-dependent preservation of left ventricular function by CD34+ cell transplantation. Necropsy examination disclosed dose-dependent augmentation of capillary density and dose-dependent inhibition of left ventricular fibrosis. Immunohistochemistry for human-specific brain natriuretic peptide demonstrated that human cardiomyocytes were dose-dependently observed in ischemic myocardium at day 28 (high, 2480±149; mid, 1860±141; low, 423±9; PBS, 0±0/mm2; P<0.05 for high versus mid and mid versus low). Immunostaining for smooth muscle actin and human leukocyte antigen or Ulex europaeus lectin type 1 also revealed dose-dependent vasculogenesis by endothelial cell and SMC development after CD34+ cell transplantation. Reverse transcriptase–polymerase chain reaction indicated that human-specific gene expression of cardiomyocyte (brain natriuretic peptide, cardiac troponin-I, myosin heavy chain, and Nkx 2.5), SMC (smooth muscle actin and sm22α), and endothelial cell (CD31 and KDR) markers were dose-dependently augmented in MI tissue.
Conclusions— Human CD34+ cell transplantation may have significant and dose-dependent potential for vasculogenesis and cardiomyogenesis with functional recovery from MI.
Received February 8, 2005; revision received August 2, 2005; accepted September 13, 2005.
The various cell types that make up the blood are of mesodermal origin and emanate from a common pool of hematopoietic stem cells (HSCs). During embryogenesis, hematopoietic and endothelial lineage cells1,2 are derived from common progenitor cells (hemangioblasts).3–6 In adults, CD34+ cells had been considered to be HSCs and clinically applied to the field of hematology for HSC transplantation.7,8 Recently, human peripheral blood CD34+ cells were reported to be an endothelial progenitor cell (EPC)–enriched population as well as a HSC fraction.9 Thereafter, therapeutic application of CD34+ cells for vascular regeneration has been performed in many preclinical studies. In the case of immunodeficient rats with acute myocardial infarction (MI), transplanted human CD34+ cells or ex vivo expanded EPCs incorporated into the site of myocardial neovascularization, differentiated into mature endothelial cells (ECs) (vasculogenesis), augmented capillary density, inhibited myocardial fibrosis and apoptosis, and preserved left ventricular (LV) function.10–12 In our institution, a phase I/II clinical trial of CD34+ cell transplantation has already been started in patients with chronic critical limb ischemia.
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Recently, Orlic et al13 and Jackson et al14 reported translineage differentiation of adult mouse HSCs into cardiomyocytes when introduced into heart by intramyocardial injection or via the circulation. This is followed by demonstrating that mouse c-Kit-positive bone marrow cells differentiate into myocardium and blood vessels in vivo.15 Further reports16,17 presented development of human peripheral blood CD34+ cells into cardiomyocytes, smooth muscle cells (SMCs), and ECs in a mouse model of acute MI. On the other hand, other investigators indicated that transplanted HSCs were unable to transdifferentiate into cardiomyocytes after MI.18,19 Given these extensive and controversial investigations into the potency of so-called stem/progenitor cells derived from peripheral blood and bone marrow in experimental models, we conducted this study to investigate not only vasculogenesis but also cardiomyogenesis derived from human CD34+ cells, the most practical human cells for clinical medicine, in a dose-dependent manner.
The present results of human CD34-positive cell transplantation into an immunodeficient rat MI model demonstrated that the collaborative multilineage differentiation potential of CD34+ cells into not only ECs but cardiomyocytes and SMCs was enhanced by cell dose escalation and was conducive to heart regeneration in terms of functional and histological recovery through vasculogenesis and cardiomyogenesis.
Isolation of CD34+ Cells From Patients With Critical Limb Ischemia
Peripheral blood total mononuclear cells were obtained from 2 male patients aged 21 and 40 years with Buerger disease by apheresis after 5-day subcutaneous administration of granulocyte colony-stimulating factor (G-CSF) (10 μg/kg per day). Bone marrow–derived CD34+ cells were isolated from total mononuclear cells by the magnetic cell sorting system CliniMACS (Miltenyi Biotec Inc, Auburn, Calif).20 These patients received intramuscular transplantation of 105 CD34+ cells/kg according to the protocol of a phase I/II dose-escalation clinical trial. The CD34+ cell fraction had a purity of >99%, as determined by fluorescence-activated cell sorting (FACS) analysis with the use of a CD34-specific monoclonal antibody (Becton Dickinson, San Jose, Calif) (Figure 1a and 1b). Remaining CD34+ cells were used for the following experiments. Informed consent with regard to the cell therapy and experimental use of the remaining cells was obtained from the patients before the case registration. The clinical study protocol was approved by the institutional ethics committees of Kobe Institute of Biomedical Research and Innovation and Kobe City General Hospital.
Female athymic nude rats (F344/N Jcl rnu/rnu, CIEA Japan, Inc, Tokyo, Japan) aged 7 to 8 weeks and weighing 130 to 145 g were used in this study. The institutional animal care and use committees of RIKEN Center for Developmental Biology approved all animal procedures, including human cell transplantation.
Induction of MI and Cell Transplantation
Rats were anesthetized with ketamine and xylazine (60 and 10 mg/kg IP, respectively). MI was induced by ligating the left anterior descending coronary artery as described previously.12 In brief, after the fourth to fifth intercostal space was opened, the heart was exteriorized, and the pericardium was incised. Thereafter, the heart was held with forceps, and MI was induced by ligating the left anterior descending coronary artery near its origin with a 6-0 Proline suture. Twenty minutes after MI, rats received intramyocardial transplantation of 1×103 (low group), 1×105 (mid group), or 5×105 (high group) CD34+ cells resuspended with 120 μL of PBS or the same volume of PBS without cells (n=12 in each group when the first patient’s cells were used; n=4 in each group for the second patient’s cells). After the injection was completed, the thorax was closed.
Flow Cytometry Studies and Monoclonal Antibodies
Regular flow cytometric profiles were analyzed with a FACSCalibur analyzer and CELLQuest software (Becton Dickinson Immunocytometry Systems, Mountain View, Calif). The instrument was aligned and calibrated daily with the use of a 4-color mixture of CaliBRITE beads (BD Biosciences, San Jose, Calif) with FACSComp software (BD Biosciences). Dead cells were excluded from the plots on the basis of propidium iodide (PI) staining (Sigma Co, St Louis, Mo). CD34+ cells were washed twice with Hanks’ balanced salt solution (HBSS) containing 3.0% heat-activated fetal calf serum (FCS), incubated with 10 μL of FcR blocking regent to increase the specificity of monoclonal antibodies (Milteyi Biotic) for 20 minutes at 4°C, and incubated with the monoclonal antibodies for 30 minutes at 4°C. The stained cells were washed 3 times with PBS containing 3.0% FCS, resuspended in 0.5 mL of HBSS/3% FCS/PI, and analyzed by FACSCalibur flow cytometer (Becton Dickinson, Franklin Lakes, NJ). The following monoclonal antibodies were used to characterize the CD34+ cell population: CD34-FITC (clone My10, BD), CD34-PE (clone 581, Pharmingen, San Diego, Calif), CD45-FITC (clone HI30, Pharmingen), CD31-FITC (clone WM59, Pharmingen), KDR (Sigma), AC133-APC (clone AC133, Pharmingen), VE-cadherin-APC (HyCult Biotechnology, Uden, The Netherlands), IgG1-FITC isotype controls (Pharmingen), IgG1-PE isotype controls (Pharmingen), IgG1-APC isotype controls (Pharmingen), IgG2a-APC isotype controls (Pharmingen), and PI (Sigma Chemical Co, St Louis, Mo).
Physiological Assessment of LV Function Using Echocardiography and Microtip Conductance Catheter
Transthoracic echocardiography (SONOS 5500, Philips Medical Systems) was performed to evaluate LV function immediately before and 5 and 28 days after MI. Under general anesthesia with ketamine and xylazine, LV end-diastolic and end-systolic dimensions (LVEDD and LVESD, respectively) and fractional shortening (FS) were measured at the midpapillary muscle level. Regional wall motion score (RWMS) was evaluated per published criteria.21 All procedures and analyses were performed by an experienced researcher who was blinded to treatment.
Immediately after the final echocardiography on day 28, the rats underwent cardiac catheterization for more invasive and precise assessment of global LV function.22 A 2.0F micromanometer-tipped conductance catheter (SPR 838, Millar Instruments Inc, Houston, Tex) was inserted via the right carotid artery into the LV cavity. LV pressure and its derivative (LV dP/dt) were continuously monitored with a multiple recording system (Pressure-Volume Conductance System ARIA and Pressure-Volume Analysis Using P-V Analysis Software [Millar Instruments Inc, Houston, Tex] and Power Laboratory DAQ System [ADInstrument, Australia]).
Heart rate, LV end-diastolic pressure (LVEDP), LV ejection fraction (EF), and the maximum and minimum LV dP/dt (+dP/dt and −dP/dt, respectively) were recorded continuously for 20 minutes. All data were acquired under stable hemodynamic conditions. All procedures and analyses were performed by an experienced researcher who was blinded to treatment.
All rats were killed 28 days after transplantation with an overdose of ketamine and xylazine. At necropsy, hearts were sliced in a broadleaf fashion into 4 transverse sections from apex to base, embedded in OCT compound, snap-frozen in liquid nitrogen, and stored at −80°C for Masson trichrome staining, immunohistochemistry, and fluorescence in situ hybridization (FISH). Rat hearts in OCT blocks were sectioned, and 5-μm serial sections were collected on slides followed by fixation with 4.0% paraformaldehyde at 4°C for 5 minutes and stained immediately. Total RNA was isolated by selective dissection of peri-infarct area in LV myocardium for reverse transcriptase–polymerase chain reaction (RT-PCR).
Morphometric Evaluation of Capillary Density and Infarct Size
Histochemical staining with isolectin B4 (Vector Laboratories, Burlingame, Calif) was performed, and capillary density was morphometrically evaluated by histological examination of 5 randomly selected fields of tissue sections recovered from segments of LV myocardium subserved by the occluded left anterior descending coronary artery. Capillaries were recognized as tubular structures positive for isolectin B4. To elucidate the severity of myocardial fibrosis, Masson trichrome staining was performed on frozen sections from each tissue block, and the stained sections were used to measure the average ratio of fibrosis area to entire LV area (percent fibrosis area). All morphometric studies were performed by 2 examiners who were blinded to treatment.
To detect transplanted human cells in rat ischemic myocardium, immunohistochemistry was performed with following human-specific antibodies: human leukocyte antigen (HLA)–ABC (BD Pharmingen) and human nuclei antibody (HNA) (Chemicon International, Inc, Temecula, Calif) to detect various kinds of human cells, human-specific brain natriuretic peptide (hBNP),23 which was kindly given by Dr Hiroshi Itoh of Kyoto University Kyoto, Japan, to detect human cardiomyocytes, and human-specific Ulex europaeus lectin type 1 (UEA-1) (Vector Laboratories, Inc)24 for human ECs. Staining specificity of hBNP, HLA-ABC, HNA, and UEA-1 against human cells and lack of cross-reactivity to rat cells were confirmed by histochemical staining with the use of human and rat heart samples (data not shown). Double immunohistochemistry with HNA and cardiac troponin I (cTn-1) (Chemicon International, Inc) was performed to detect double-positive cells as human cardiomyocytes in rat myocardium. Double immunohistochemistry with HLA-ABC and smooth muscle actin (SMA) was performed to detect double-positive cells as human SMCs in rat myocardium. Similarly, double immunohistochemistry with hUEA-1 and SMA and for von Willebrand factor (vWF) (Chemicon International, Inc) and HNA was performed to detect UEA-1–positive cells and double-positive cells of vWF and HNA as human ECs in ischemic myocardium. The secondary antibodies for each immunostaining are as follows: FITC goat anti-mouse IgG (The Jackson Laboratory, Bar Harbor, Me) for hBNP staining, Alexa Flour 594–conjugated goat anti-mouse IgG1 (Molecular Probes) for HLA-ABC staining, Alexa Flour 488–conjugated goat anti-mouse IgG2a (Molecular Probes, Carlsbad, Calif) for cTn-I staining, Alexa Flour 488–conjugated goat anti-mouse IgG2a (Molecular Probes) for SMA, Cy3-conjugated streptavidin (Jackson ImmunoResearch, West Grove, Pa) for hUEA-1, Alexa Flour 488– and 594–conjugated goat anti-mouse IgG1 for HNA, and Alexa Flour 594–conjugated goat anti-rabbit IgG for vWF. DAPI solution was applied for 5 minutes for nuclear staining. Number of human cardiomyocytes, total (both human and rat) cardiomyocytes in ischemic myocardium detected as hBNP-positive cells, number of human SMCs as double-positive cells for HLA and SMA, and number of human ECs as hUEA-1–positive cells 28 days after MI were morphometrically quantified with the use of 5 randomly selected fields (from peri-infarction area to fibrosis area) of tissue sections.
Fluorescence In Situ Hybridization
To identify whether cardiac repair occurred through cell fusion in rat ischemic myocardium, FISH was performed with human Y chromosome painting probe and rat genome probe (nick translation methods) in MI tissue. Tissue sections were fixed immediately with 4.0% paraformaldehyde at 4°C for 20 minutes and predenatured, dehydrated, and denatured according to the manufacturer’s protocol. Sections were hybridized with a Cy-3–conjugated DNA probe for human Y chromosomes and a biotin-conjugated probe for rat genome overnight at 37°C. After posthybridization wash, TexRed-conjugated streptavidin was applied, and slides were counterstained with DAPI and examined.
RT-PCR Analysis of CD34+ Cells and Ischemic Heart Tissue
Total RNA was obtained from freshly isolated peripheral blood CD34+ cells of a patient and tissues of rat ischemic LV at day 28 with the use of TRIzol (Life Technologies, St Paul, Minn) according to the manufacturer’s instructions. The first-strand cDNA was synthesized with the use of the RNA LA PCR Kit version 1.1 (Takara, Otsu, Japan), amplified by Taq DNA polymerase (Advantage-GC cDNA PCR Kit, Clontech and AmpliTaq Gold DNA polymerase, Applied Biosystems). PCR was performed with a PCR thermocycler (MJ Research PTC-225). The human GAPDH (hGAPDH), total (human and rat) GAPDH, human CD34 (hCD34), hBNP, human cardiac troponin-I (hcTn-I), human myosin heavy chain-β (hMHC-β), human KDR (hKDR), and human Nkx 2.5 (hNkx 2.5) were amplified by Taq DNA polymerase (AmpliTaq Gold DNA polymerase, Applied Biosystems) under the following conditions: 35 cycles of 30 seconds of initial denaturation at 94°C, annealing at 56°C for 1 minute, and 30 seconds of extension at 72°C according to the manufacturer’s instructions. Human myosin heavy chain-α (hMHC-α), human SMA (hSMA), human sm22α (hsm22α), and human CD31 (hCD31) were amplified by Taq DNA polymerase (Advantage-GC cDNA PCR Kit, Clontech, Mountain View, Calif) under the following conditions: 37 cycles of 30 seconds of initial denaturation at 94°C, annealing at 68°C for 3 minutes, and 7 minutes of elongation at 64°C according to the manufacturer’s instructions. Subsequently, PCR products were visualized in 1.5% ethidium bromide–stained agarose gels. Human heart RNA distributed from Clontech (premium RNA) was used as positive control. To quantify human-specific cardiomyogenic and vasculogenic gene expression in rat ischemic myocardium after human CD34+ cell transplantation, we measured the band intensity of the RT-PCR image. Each gel was photographed onto positive/negative Polaroid film under ultraviolet illumination. The negative images were then captured with the use of an image scanner. After the images were recorded in a computer, the band intensities were processed with the NIH Image program (version 1.62) as described previously.25 These band intensities were used to calculate the ratio of human-specific marker (hGAPDH, hBNP, hcTn-I, hMHC-α, hMHC-β, hNkx 2.5, hSMA, hsm22α, hKDR, and hCD31) expression to total GAPDH expression.
To avoid interspecies cross-reactivity of the primer pairs between human and rat genes, we designed the following human-specific primers using Oligo software (Takara). All primer pairs did not show cross-reactivity to rat genes (data not shown). Primer pairs were as follows: hBNP primer sequence (146 bp): sense GCA AAA TGG TCC TCT ACA CC; antisense CAG GAC TTC CTC TTA ATG CC; hcTn-I primer sequence (218 bp): sense AAT TGC AGC TGA AGA CTC TG; antisense GAC TTT TGC CTC TAT GTC GT; hMHC-β primer sequence (214 bp): sense GCT AAA GGT CAA GGC CTA CA; antisense GCA GAT CAA GAT CTG GCA AA; hNkx 2.5 primer sequence (205 bp): sense GAG AGT TTG TGG CGG CGA TT; antisense CGA CGG CGA GAT AGC AAA GG; hMHC-α primer sequence (413 bp): sense GTC ATT GCT GAA ACC GAG AAT G; antisense GCA AAG TAC TGG ATG ACA CGC T; hSMA primer sequence (485 bp): sense TCT GGA CGC ACA ACT GGC ATC GT; antisense TAC ATA GTG GTG CCC CCT GAT AG; hsm22-α primer sequence (468 bp): sense CGG CTG GTG GAG TGG ATC ATA; antisense CCC TCT GTT GCT GCC CAT CTG A; hCD31 primer sequence (469 bp): sense AGG TCA AGC AGC ATC GTG GTC AAC AT; antisense TTG TCT TTG AAT ACC GCA G; hCD34 primer sequence (380 bp): sense AAT GAG GCC ACA ACA AAC ATC ACA; antisense CTG TCC TTC TTA AAC TCC GCA CAG C; hGAPDH primer sequence (596 bp): sense CTG ATG CCC CCA TGT TCG TC; antisense CAC CCT GTT GCT GTA GCC AAA TTC G; total GAPDH primer sequence (320 bp): sense GTG CCA GCC TCA TGT TCG TC; antisense CGC CAG TGT ACT CCA CGA CAT TTC G; hKDR primer sequence (468 bp): sense CAA ATG TGA AGC GGT CAA CAA AGT T; antisense ATG CTT TCC CCA ATA CTT GTC GTC T.
The results were statistically analyzed with the use of a software package (Statview 5.0, Abacus Concepts Inc, Berkeley, Calif). All values were expressed as mean±SE. Paired t tests were performed for comparison of data before and after treatment. The comparisons among 4 groups were made with 1-way ANOVAs. Post hoc analysis was performed by Fisher protected least significant difference test. Differences of P<0.05 were considered statistically significant.
Characterization of Freshly Isolated Peripheral Blood CD34+ Cells
The CD34+ cell fraction had a purity of >99%, as determined by FACS analysis with the use of anti-CD34, anti-CD45, anti-CD31, anti-AC133, anti-KDR, and anti–VE-cadherin monoclonal antibodies (Figure 1a and 1b). The RT-PCR analysis revealed the human-specific gene expression of CD34 and SMA but not of cardiomyocyte markers (hBNP, hcTn-I, hMHC-α, hMHC-β, and hNkx 2.5), hKDR, and hsm22α (Figure 1c).
Transplanted CD34+ Cells Dose-Dependently Preserve LV Function After MI
There were no significant differences in preoperative echocardiographic parameters, LVEDD, LVESD, FS, and RWMS among high, mid, low, and PBS groups. Echocardiography on day 5 revealed that the functional parameters were also similar in all groups (data not shown). Echocardiography performed 4 weeks after cell transplantation demonstrated that LVEDD was significantly smaller in the high group than in the low and PBS groups (P<0.05 versus low and P<0.01 versus PBS). LVEDD was also significantly smaller in the mid group than in the PBS group (P<0.01). However, LVEDD was similar in the high and mid groups and in the low and PBS groups (Figure 2a and 2b). LVESD 4 weeks after MI was significantly smaller in the high group than in the mid group and in the mid group than in the low group (high group, 0.564±0.01; mid group, 0.607±0.013; low group, 0.705±0.025; PBS group, 0.711±0.011 cm; P<0.01 for high versus mid and mid versus low) (Figure 2b). FS was significantly greater in the high group than in the mid group and in the mid group than in the low group (high group, 31.4±0.43%; mid group, 27.7±0.45%; low group, 20.2±0.58%; PBS group, 20.0±0.54%; P<0.01 for high versus mid and mid versus low) (Figure 2a and 2b). RWMS was significantly better preserved in the high group than in the mid group and in the mid group than in the low group (high group, 20.8±0.46; mid group, 23.4±0.15; low group, 27.2±0.2; PBS group, 26.8±0.24; P<0.01 for high versus mid and mid versus low). LVEDD, LVESD, FS, and RWMS 4 weeks after transplantation in the low group were not significantly different from those in the PBS group (Figure 2a and 2b).
Invasive hemodynamic study performed 4 weeks after transplantation revealed that heart rate and diastolic blood pressure were similar in each group. Systolic blood pressure was significantly greater in the high and mid groups than in the PBS group (P<0.05) (Table). The +dP/dt, absolute value of −dP/dt, and EF were significantly greater in the high group than in the mid group and in the mid group than in the low group (+dP/dt: high group, 11 131±106; mid group, 9772±111; low group, 7734±160; PBS group, 7322±233 mm Hg/s; P<0.01 for high versus mid and mid versus low) (Figure 2c) (−dP/dt: high group, −6403±209; mid group, −5753±170; low group, −4413±230; PBS group, −4415±212 mm Hg/s; P<0.01 for high versus mid and mid versus low) (Figure 2c) (EF: high group, 56.8±2.3%; mid group, 47.5±1.4%; low group, 34.2±2.1%; PBS group, 36.9±1.4%; P<0.01 for high versus mid and mid versus low) (Figure 2c). In addition, LVEDP 4 weeks after ischemia was significantly smaller in the high group than in the mid group and in the mid group than in the low group (high group, 7.8±0.6; mid group, 9.8±0.6; low group, 13.0±0.7; PBS group, 13.3±1.6 mm Hg; P<0.01 for high versus mid and mid versus low) (Figure 2c). The +dP/dt, −dP/dt, EF, and LVEDP 4 weeks after transplantation in the low group were not significantly different from those in the PBS group.
Thus, transplantation of high and mid doses of CD34+ cells, but not the low dose, significantly preserved global and regional LV function after MI. The functional effect of CD34+ cells was dose-dependently observed.
Morphometric Evaluation of Capillary Density and Infarct Size
Myocardial neovascularization assessed by capillary density on day 28 was dose-dependently enhanced in rats receiving CD34+ cell transplantation (high group, 714.3±25.0; mid group, 535.8±31.0; low group, 320.9±36.0; PBS group, 291.3±19.0/mm2; P<0.01 for high versus mid and mid versus low). Capillary density in the low group was similar to that in the PBS group (Figure 3a and 3b). LV remodeling evaluated by the percent fibrosis area was dose-dependently inhibited in rats receiving CD34+ cell transplantation (high group, 16.0±2.6%; mid group, 22.4±1.9%; low group, 30.7±3.9%; PBS, 31.5±0.7%; P<0.01 for high versus mid and mid versus low groups). Percent fibrosis area was similar in the low and PBS groups (Figure 3c and 3d).
Thus, transplantation of high and mid CD34+ cells, but not the low dose, significantly preserved LV structural integrity after MI. The histological efficacy of CD34+ cells was dose-dependently observed.
Transplanted hCD34+ Cells Dose-Dependently Differentiate Into Cardiomyocytes
Differentiated human cardiomyocytes derived from the transplanted CD34+ cells were mainly identified in the rat peri-infarct myocardium by double staining both for cTn-I and HNA (Figure 4a to 4⇓e) and for hBNP and HNA (Figure 4f to 4⇓j). These findings suggest that transplanted CD34+ cells have potency of differentiation into cardiomyocytes. Double immunohistochemistry with hBNP and HNA also revealed specificity of the hBNP antibody for human cells in rat myocardium. Dose-dependent distribution of human cardiomyocytes in rat myocardium was observed both in samples stained with cTn-I and HNA (Figure 4⇓k) and in samples stained with hBNP (Figure 4⇓l). In fact, the numbers of hBNP-positive cardiomyocytes were dose-dependently observed in ischemic myocardium at day 28 (high group, 2480±149; mid group, 1860±141; low group, 423±9; PBS group, 0±0/mm2; P<0.05 for high versus mid and mid versus low groups). Total (both human and rat) cardiomyocytes were also dose-dependently observed in ischemic myocardium at day 28 (high group, 13 102±298; mid group, 11 936±238; low group, 6564±369; PBS group, 5707±300/mm2; P<0.05 for high versus mid and mid versus low groups) (Figure 4l and 4⇓m). Similar dose-dependent cardiomyogenesis was observed when CD34+ cells from another patient were used (Figure I in the online-only Data Supplement). These findings strongly suggest that transplanted CD34+ cells may have dose-dependent potency of differentiation into cardiomyocytes.
Transplanted hCD34+ Cells Dose-Dependently Differentiate Into SMCs
Human SMCs derived from the transplanted CD34+ cells were mainly identified in the vasculatures within the peri-infarct area by double staining for SMA and HLA-ABC. Identified human SMCs were dose-dependently observed after CD34+ cell transplantation (high group, 895±95; mid group, 180±11; low group, 0±0; PBS group, 0±0/mm2; P<0.01 for high versus mid and mid versus low groups). In contrast, differentiated human SMCs were not identified in PBS and low groups (Figure 5a to 5c). Similar dose-dependent SMC commitment was observed when CD34+ cells from another patient were used (Figure II in the online-only Data Supplement). These findings suggest that transplanted CD34+ cells may have dose-dependent potency of differentiation into SMCs.
Transplanted hCD34+ Cells Dose-Dependently Differentiate Into ECs
We confirmed the specificity of UEA-1 staining for human ECs using double immunohistochemistry with UEA-1 and HNA (Figure 6a to 6e). Differentiated human ECs derived from the transplanted CD34+ cells were observed in the vasculatures within the peri-infarct area by UEA-1 staining. Identified UEA-1–positive cells were greater in higher-dose groups than lower-dose groups (high group, 3373±363; mid group, 980±211; low group, 226±35; PBS group, 0±0/mm2; P<0.05 for high versus mid and mid versus low groups). In contrast, differentiated human ECs were not identified in the PBS group (Figure 6f, 6g). Similar dose-dependent endothelial commitment was observed when CD34+ cells from another patient were used (Figure III in the online-only Data Supplement).
Thus, locally transplanted CD34+ cells were incorporated not only into ECs but also into mature SMCs, resulting in contribution to vasculogenesis in ischemic myocardium.
FISH Analysis of Transplanted CD34+ Cell–Derived Cardiomyocytes
To determine whether cardiac repair occurred through cell fusion in MI tissue, we performed FISH with human Y chromosomes and rat genome probe. The specificity of the probes was tested in tissues of normal rat heart and rat heart immediately after human cell transplantation. We confirmed that these 2 probes did not cross-react with cells of the other species (data not shown). The FISH analysis revealed the existence of cardiomyocytes in which human Y chromosome was paired with rat genome (cell fusion) as well as those without genome (no fusion) (Figure 7a to 7d). These findings indicate that both cell fusion and multilineage differentiation without fusion may be involved in transformation of transplanted CD34+ cells into cardiomyocytes.
Dose-Dependent Gene Expression of Human-Specific Cardiac, Smooth Muscle, and Endothelial Lineage Cell Markers in Rat Ischemic Myocardium After CD34+ Cell Transplantation
To further ensure the immunohistochemical results with regard to cardiomyogenesis and vasculogenesis by the molecular approach, we performed RT-PCR with rat ischemic myocardium by using human-specific primer BNP, cTn-I, MHC-α, MHC-β, and Nkx 2.5 as human cardiomyocyte lineage markers, human-specific primer sm22α and SMA as human SMC markers, and human-specific primer CD31 and KDR as human EC markers.
The RT-PCR analysis revealed dose-dependent expression of human-specific cardiomyogenic, arteriogenic, and vasculogenic genes in rat ischemic myocardium after human CD34+ cell transplantation with the use of the NIH Image program (version 1.62) (Figure 8a and 8b). Notably, gene expression of all markers except hSMA was not detected in freshly isolated CD34+ cells (Figure 1c) but was dose-dependently augmented in ischemic myocardium after cell transplantation.
Abrupt occlusion of coronary arteries causes MI, which leads to massive cardiomyocyte loss and consequently deterioration of cardiac function because cardiomyocytes have severely limited capacity to be divided and thus replace the damaged tissue. Progressive heart failure is a major cause of death or frequent hospitalization in patients after MI. Although MI is classified as vascular (coronary artery) disease, therapeutic strategies should be focused on regenerating not only blood vessels but also cardiac muscle to improve the poor prognosis of the disease.
Compelling evidence suggests that transplantation of bone marrow–derived CD34+ cells or cultured EPC-enriched population contributes to preservation of LV function after MI through enhancing ischemic neovascularization.10–12 The mechanism of this therapeutic effect was previously considered to be incorporation, differentiation, and proliferation of EPCs for new blood vessel formation.9,11,26 Recently, Badorff et al27 reported in vitro transdifferentiation of EPCs into functional cardiomyocytes. Yeh et al16 also demonstrated in vivo differentiation of CD34+ cells into cardiomyocytes and SMCs in a mouse model of acute MI. Regeneration of SMCs as well as ECs may result in mature vasculogenesis, which is more potent for blood flow recovery in ischemic myocardium compared with capillary formation by EC-only regeneration. These findings lead to a novel concept that CD34+ cell transplantation may contribute to cardiomyogenesis and vasculogenesis, which may be an ideal strategy to treat MI. On the other hand, Balsam et al18 and Murry et al19 reported that mouse bone marrow HSCs isolated as Lin−c-Kit+ cells or c-Kit+Thy1.1loLin-Sca-1+ cells do not transdifferentiate into cardiomyocytes in infarcted myocardium. Several points should be considered carefully with regard to this discrepancy, such as the difference in species (human versus murine), cell populations (CD34+ cells versus Lin−c-Kit+ cells or c-Kit+Thy1.1loLin-Sca-1+), cell doses to transplant, or cell delivery methodologies. Given the controversy, the question is whether improvement of myocardial function after EPC transplantation was due to myocardial preservation through the signal from enhanced vasculogenesis as well as due to regenerative cardiomyogenesis by transplanted cells. To solve this issue, we tried to confirm the lineage potency and the tissue plasticity of CD34+ cells by transplanting the cells into immunodeficient rats with MI in a dose-ranging fashion. From a practical point of view, a significant contribution of cardiomyogenesis and vasculogenesis to LV functional recovery after MI may not be expected if such translineage differentiation is a rare event after CD34+ cell transplantation. To detect the translineage differentiation of human CD34+ cells into rat myocardium, we performed not only immunohistochemistry but also RT-PCR for human-specific markers of cardiomyocytes, SMCs, and ECs. These sensitive assessments revealed dose-dependent augmentation of cardiomyogenesis and vasculogenesis of human CD34+ cells in rat infarcted myocardium. The translineage potential was accompanied with dose-dependent enhancement of capillary density, inhibition of LV fibrosis, and preservation of LV function. These findings suggest that transplantation of a higher dose of CD34+ cells may be more potent for therapeutic application to the damaged myocardium than a lower dose. Another interesting finding in this study is that these favorable effects of CD34+ cells were not significantly observed in the low-dose group (1×103 cells/kg). To our knowledge, information about a noneffective dose of CD34+ cells has never been provided. Taken together, the present results strongly suggest the therapeutic importance of the cell dosage in the actual clinical application.
It is unclear what mechanism of CD34+ cells is involved in multilineage commitment and significant incorporation for functional organogenesis. One mechanism for multilineage commitment is the translineage differentiation of already committed EPCs or HSCs among CD34+ cells through transdifferentiation or cell fusion. Recently, Zhang et al17 demonstrated that 70% of newly formed cardiomyocytes derived from CD34+ cells were developed through a cell fusion mechanism between human and mouse cells, whereas CD34+-derived ECs are mainly not developed by cell fusion. The FISH analyses in this study revealed that the mechanism of cardiomyogenic plasticity of CD34+ cells involves both cell fusion and the multilineage differentiation without fusion, although the functional contribution of both mechanisms to myocardiogenesis remains uncertain.
The other possible mechanism of multilineage differentiation is due to the original multipotency of the CD34+ cell population. Recently, peripheral blood CD34+ cells were proved to contain a cell fraction expressing not only hematopoietic and endothelial but also cardiac, muscle, liver, and neural lineage markers after mobilization following G-CSF administration or myocardial ischemia,28,29 whereas this issue is not determined if multilineage cells are derived from pluripotent stem cell or various lineage progenitor cell mixtures in CD34+ cells. When the magnificent incorporation of CD34+ cell–derived cardiomyocytes and SMCs is taken into account, the mechanism is likely due to programmed lineage commitment in the myocardial ischemia environment, although we have not defined each responsible cell fraction for lineage diversification in CD34+ cells.
The cooperative signal from vasculogenesis to cardiomyogenesis must also be considered further in the regenerative process through multilineage commitments by CD34+ cells. Cardiomyogenesis and vasculogenesis are closely regulated in terms of microenvironmental interaction during the developmental stage. Recently, Shen et al30 proved the significance of vascular signals for postnatal neural stem cell biology, as formerly indicated in the case of liver and pancreas development in embryo.31,32 Microenvironmental interaction between myocardial and vascular lineage cells involves not only paracrine regulatory factors but also direct cellular communications in developing CD34+ cells. We speculate that an enhanced vasculogenesis signal may exert cellular commitment and development of CD34+ cells into myocardial cells as a cooperative organogenesis mechanism.
In conclusion, after transplantation of bone marrow–derived CD34+ cells, the collaborative multilineage differentiation potential of CD34+ cells not only into ECs but also into cardiomyocytes and SMCs was enhanced by cell dose escalation and was conducive to heart regeneration in terms of functional and histological recovery through vasculogenesis and cardiomyogenesis.
This work was supported by Health and Labor Sciences research grants (H14-trans-001, H17-trans-002) from the Japanese Ministry of Health, Labor, and Welfare. We thank Yumiko Masukawa and Tomoko Itoh for secretarial assistance. Human-specific antibody against brain natriuretic peptide (hBNP) was a generous gift from Dr Hiroshi Itoh of Kyoto University, Kyoto, Japan.
The online-only Data Supplement can be found at http://circ.ahajournals.org/cgi/content/full/113/10/1311/DC1.
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