European Journal of Cardio-thoracic Surgery 28 (2005) 677—684 www.elsevier.com/locate/ejcts
Amniotic mesenchymal cells autotransplanted in a porcine model of cardiac ischemia do not differentiate to cardiogenic phenotypes Saverio Sartore a,*, Maddalena Lenzi a, Annalisa Angelini b, Angela Chiavegato a, Lisa Gasparotto a, Paolo De Coppi c, Roberto Bianco d, Gino Gerosa d a
Department of Biomedical Sciences, University of Padua, Viale G. Colombo, 3, I-35121 Padua, Italy b Institute of Pathologic Anatomy, University of Padua, Padua, Italy c Department of Pediatrics, University of Padua, Padua, Italy d Department of Cardiological, Thoracic and Vascular Sciences, University of Padua, Padua, Italy
Received 17 June 2005; received in revised form 25 July 2005; accepted 27 July 2005; Available online 26 September 2005
Abstract Objective: Transplantation of stem cells in the acute ischemic myocardium (AMI) may play a role in the recovery of cardiac function. Here, we investigated the ability of amniotic fluid-derived mesenchymal cells (AFC) for phenotypic conversion to vascular cells and cardiomyocytes (CM) when autotransplanted in a porcine model of AMI. Methods: Single AFC preparations were taken from 12 fetuses 3 days before normal delivery. AFC were expanded in vitro and stored separately until animals of the original litter weighed 22—25 kg. A new model of AMI, i.e. 45-min circumflex coronary occlusion followed by wall dissection, was used to assess AFC differentiation potential. CMFDA-labeled AFC were autogenically transplanted in the ischemic area 1 week after AMI induction. Thirty days later, pigs were sacrificed and the phenotypic profile of transplanted AFC was assessed and compared to the corresponding pre-injection pattern. Results: AFC showed in vitro to be of mesenchymal type also expressing markers of ‘embryonic stem’ cells (SSEA4 and Oct-4), as well as endothelial (von Willebrand factor, VE-cadherin) and smooth muscle (SM a-actin, SM22) cells. Thirty days after transplantation, in the survived AFC (5 1%) ‘embryonic stem’ cell markers disappeared and mesenchymal cell markers were down regulated with the exception of smooth muscle and endothelial antigens. No evidence for expression of cardiac troponin I was found.Conclusions: In the conditions used in this study, AFC were able to transdifferentiate to cells of vascular cell lineages but not to CM. Thus, porcine AFC may require further ex vivo re-programming to be suitable for therapeutic use in AMI. # 2005 Elsevier B.V. All rights reserved. Keywords: Cell transplantation; Ischemia; Myocardial infarction; Tissue engineering.
1. Introduction Stem cells, directly transplanted or mobilized from cardiac or extra-cardiac derived compartments, have been proposed as innovative and efficacious strategies to replace damaged myocardium and vessels or to limit remodeling of the post-ischemic heart [1]. It has been hypothesized that cells of the cardiogenic lineages can be produced via a transdifferentiation (a phenotypic conversion from an ontogenically distinct cell type to another), fusion (with a differentiated resident cell), growth factor/cytokine stimulation (released from transplanted or resident stem cells) or, perhaps, a combination of these processes [1]. Newly formed CM, vascular smooth muscle or endothelial cells in the ischemic heart can be obtained by transplantation of embryonic or adult (bone marrow-derived or circulating) stem cells, resulting in functional improvement with partial recovery of contractility or reduced cardiac remodeling [2]. * Corresponding author. Tel.: þ39 049 827 6032; fax: þ39 049 827 6040. E-mail address:
[email protected] (S. Sartore). 1010-7940/$ — see front matter # 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.ejcts.2005.07.019
Embryonic stem cells seem to be potentially a better tool than the adult counterpart [2] in terms of cell plasticity and, hence, differentiation potential. In contrast, the use of these cells is hampered by ethical reasons. Thus, other sources of stem cells not sharing these problems, such as cord blood stem cells [3] or cells of the amniotic fluid [4], could be potentially of interest for clinical applications provided that they are able to give structural and functional benefits to the damaged heart once transplanted. Human amniotic fluid cells are endowed with some interesting properties such as: (1) in vitro expression of ‘stemness’ markers [5] as well as mesenchymal and neuroglial markers [4—6], (2) a tolerant-type histocompatibility antigen profile [7], (3) the capacity to switch to CM-like cells in vitro after stimulation with FGF-2, activin A or coculture with rat neonatal CM [8], and (4) the ability of correcting genetically based enzymatic defects once transplanted in injured brain [9]. Before moving on to functional studies, it is preliminarily important to assess the differentiation potential of AFC in vivo, possibly using a model of AMI of clinical relevance. The porcine model appears to be particularly suitable in this
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respect inasmuch as other transplantation studies with stem and non-stem cells were performed [10—18]. In addition, such a model could allow, in perspective, the use of imaging modalities that are clinically important and can validate the putative structural changes inherent to cell transplantation. The aim of this study was to ascertain the ability of AFC to differentiate to vascular cells and CM once transplanted into the infarcted myocardium of a porcine model. Since the amniotic fluid contains a mixture of different cell populations, we have selected a plastic-adherent cell fraction, potentially comparable to the bone marrow MSC already used in other experiments [15,16]. Fig. 1. Experimental design of the in vivo protocol.
2. Material and methods 2.1. Cesarean section and collection of amniotic fluid A pregnant Large White sow underwent a cesarean section three days before term. All procedures were approved by the Italian Ministry of Public Health and performed in accordance with the ‘Guide for the Care and Use of Laboratory Animals’ prepared by the Institute of Laboratory Animal Resources, National Research Council (published by the National Academy Press, revised 1966). The pig used in this study originated from a colony in which MHC class II antigen, as determined by indirect immunofluorescence with a specific anti-porcine antibody (Serotec, Oxford, UK), is expressed in 100% of bone marrow-derived MSC. The pig was pre-medicated with azaperone, midazolane and ketamine hydrochloride. Anesthesia was induced with a ventilation mask using 4.0% isofluorane. The animal was intubated with a cuffed endotracheal tube and ventilated with 100% oxygen. Anesthesia was maintained with 1.5% isofluorane. Electrocardiogram and respiration were monitored by a multiple channel recorder. By abdominal hysterectomy, 15 fetuses and the respective amniotic fluids (about 50 ml) were rapidly collected. All piglets were reanimated and kept in a controlled environment for a few hours and then transferred to a foster-mother in the animal facility where they were kept for 3 months.
angiography, a guide wire was positioned in it and a balloon (Opensail, 2.5 mm of diameter) was advanced in the lumen. Occlusion of this vessel branch was achieved by increasing the intraluminal pressure to 8 atm and confirmed by angiography. After 45 min of occlusion, the pressure was further increased to 20 atm to induce wall dissection and subsequent thrombosis. Electrocardiographic monitoring showed ST-segment change and occlusion of the vessel was confirmed by angiography. Episodes of ventricular tachycardia and fibrillation were terminated with lidocaine and external defibrillation. Finally, a 6-Fr pigtail catheter was retrogradely advanced into the left ventricle and a cardiac ventriculography was obtained to confirm the segmental dysfunction of the left ventricular wall caused by coronary artery occlusion. After intervention, pigs received analgesics (Buprenex, 0.03 mg/kg) and cortisone (1 mg/kg) before returning to the animal facility. One of the operated animals was sacrificed 2 h after injection to identify the AFC distribution with respect to the ischemic region and to evaluate the surviving level of CMFDA-labeled AFC 30 days after transplantation (see Fig. 1). Two animals were injected with aMEM solution only. Two pigs were sacrificed 7 and 37 days, respectively, after induction of AMI to study the time-course of infarction in absence of AFC transplant. Two AFC preparations were also delivered in an autogenic manner, to the heart of intact animals.
2.2. Induction of AMI 2.3. Preparation of AFC Cesarean-born animals enrolled in this experiment, weighing about 22—25 kg, were subjected to AMI. Premedication and anesthesia were given as described above. Antibiotics (Cefazolin, 35 mg/kg) were given the day before surgery and continued up to 5 days post-surgery. AMI was obtained by a percutaneous procedure involving an intraluminal thrombotic occlusion of the 3rd lateral branch of circumflex coronary artery. In this procedure, the left femoral artery was isolated and cannulated with an introduction sheath (percutaneously accessed by needle puncture, in which a flexible guide wire was advanced, then the needle was withdrawn and the introducing sheath placed over the guide wire). Through the sheath, a JR 3.5 6F guiding catheter was used to cannulate the left main stem under fluoroscopic guidance. Once the circumflex branch of choice became visible by
Samples of amniotic fluids taken from the different fetuses were diluted with PBS pH 7.2 (1:2, v/v) and then spun down at 1000 rpm in Eppendorf 5804R Centrifuge. Pellets were resuspended in the Amniotic Culture Medium (ACM; 63% aMEM (Life Technology, Gaithersburg, MD), 20% of Chang Medium (Chang B plus Chang C; Irvine Scientific, Santa Ana, CA), 15% of fetal bovine serum (FBS) containing streptomycin, penicillin and L-glutamine. Cell seeding was performed on Falcon petri dishes (Becton Dickinson, Los Angeles, Ca) at a density of 4.5102 cells/cm2. After 3 days, non-adherent cells and debris were discharged and the adherent cells cultivated until preconfluency. Adherent cells were detached from the plastic substrate using 0.05% trypsin and 0.02% sodium-EDTA (Life Tech). Cells grown in an incubator at 378C with 95% air and 5% carbon dioxide were
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passaged until approximately 10—12 million of AFC for each animal were collected (5—6 passages).
tissue blocks of animals sacrificed 2 h vs. 30 days from AFC transplantation.
2.4. In vitro phenotypic profile of AFC
2.6. Immunocytochemical analysis of AMI lesion
Stability of antigenic marker expression in ACM was assayed by culturing AFC at each passage onto 1% gelatincoated glass coverslips for 5 days. Propensity of AFC to vascular cell phenotypic conversion was evaluated by seeding cells (3103 cells/cm2) in Endothelial Growth Medium (Promocell, Heidelberg, Germany), containing 20% FBS (for endothelial cells), or in Smooth Muscle Cell Medium, containing DMEM and 10% FBS (for smooth muscle cells) on gelatin-coated Falcon petri dishes. Cells were fixed after 7 days of culture. Coverslips with AFC grown in ACM or in the two differentiating media were fixed in 1.5% p-formaldehyde in PBS and tested in single or double immunofluorescence using anti-PECAM/CD31 (Chemicon, Temecula, Ca), anti-VEcadherin (Santa Cruz Biotech, Santa Cruz, Ca), anti-Oct4 (Chemicon), anti-SM a-actin (Sigma, St Louis, Mo) and SM-E7 anti-SM myosin [19] and IgG anti-mouse or anti-rabbit rhodaminated or fluoresceinated secondary antibodies (Chemicon). Distribution of antigens was studied using a Zeiss Axioplan epifluorescence microscope (Zeiss, Oberkochen, Germany), and images were acquired using a Leica DC300F digital videocamera (Leica, Wetzlar, Germany).
Cardiac cryosections from AMI hearts were fixed in 1.5% pformaldehyde in PBS pH 7.2 and then incubated with the primary antibody for 30 min at 37 8C. After two rinses in PBS, cells were incubated with peroxidase-labeled secondary antibody (Dako, Glostrup, Denmark) to the monoclonal primary antibody according to Chiavegato et al. [19]. The primary antibodies were the followings: anti-cardiac troponin I [20]; anti-EIIIA fibronectin (a generous gift of Dr Luciano Zardi, Genoa, Italy); anti-desmin (Chemicon); anti-procollagen I (Iowa Hybridoma Bank; Iowa City); anti-SM a-actin (Sigma); anti-von Willebrand factor (Dako), anti-CD45 (Serotec); 1B8 anti-SM22 [19]. Optical images were acquired by a Leica DMR microscope connected to a Leica DC300 videocamera (Leica, Wetzlar, Germany).
2.5. Labeling and injection of AFC To identify the transplanted cells in the recipient myocardium, AFC were labeled with CMFDA (chloromethylfluorescein diacetate) following the instructions furnished by the manufacturer (Molecular Probes, Eugene, Or). For transplantation, cells taken from cultures at confluent conditions were dissociated from the petri dishes using 0.025% trypsin-EDTA and spun at 1050 rpm for 10 min to remove the enzyme. The pellet was rinsed twice in PBS and cells suspended in alpha MEM medium at a concentration of 7.5106 cells/ml. Trypan blue staining of CMFDA-labeled cells showed that >90% of AFC were viable. One week after AMI induction (see Fig. 1), pigs destined to receive AFC transplants were examined by echocardiography and ventriculography to confirm and localize the infarcted region. Subsequently, with the animals under general anesthesia, the infarcted area was exposed through a left thoracotomy. The various cell suspensions were autogenic injected in the border zone of ischemic region (1 ml, 7.5106 cells, distributed in 3—4 sites) of the corresponding animals using an insulin syringe and a 27-gauge needle. Culture medium (aMEM) was used as control. Area of cell transplantation or culture medium injection was marked with a Prolene suture for subsequent identification at the time of sacrifice. Then, the chest was closed leaving the pericardial wound open and the air evacuated from the cardiac cavity. After intervention, pigs received analgesics (Buprenex, 0.03 mg/kg) and cortisone (1 mg/kg) before returning to the animal facility. AMI pigs with transplanted AFC were euthanized after 30 days from cell injection. Score of survived CMFDA-stained AFC in AMI hearts was performed by counting positive cells in four randomly selected high-power fields taken from 10 cryosections per
2.7. Immunocytochemical analysis of AFC before and after transplantation AFC to be used in cell transplantation were enzymatically detached from petri dishes and studied for their immunophenotypic properties by cytospin preparations obtained with a Shandon Cytospin 4 centrifuge. Cytospin-obtained cells were fixed in 1.5% p-formaldehyde in PBS pH 7.2 and then incubated with the primary antibody (antigens of the cardiogenic and non-cardiogenic cell lineages as well embryonic and hematopoietic antigens; see Table 1) for 30 min at 37 8C. Cells were incubated with rhodaminated secondary antibody (Chemicon) to the monoclonal or polyclonal primary antibodies. CMFDA-labeled cells in the transplants were detected by FITC-conjugated rabbit Table 1 Immunophenotypic characterization of AFC as determined in vitro (tissue cultures and cytospins) and in vivo (CMFDA-positive transplanted cells in the ischemic myocardium) Antigens
Cell cultures
Cytospins
CMFDAþ-transplanted cells
CD34 CD45 CD117/c-kit Sca-1 SSEA4 Oct4 Vimentin CD105/Endoglin Thy-1/CD90 Desmin CardiacTnI SM a-actin SM22 SM myosin CD31/PECAM von Willebrand factor VE-cadherin
— — — — þ þ þþþ — þþþ n.d. — þ þþþ n.d. þ þþþ þ
— — — — þ þ þþþ þþ þþþ — — þþ þþ — þ þþ þþ
— — — — — — — þ þ n.d. — þ — — þ þ þ
n.d., Not determined; —, no cell labelled; þ, up to 30% labelled cells; þþ, from 30 to 60% of cells are labelled; þþþ, 60—90% of cells are labeled. Note that the expression of antigenic markers in AFC reported in the Cell Cultures column was evaluated after 15 days in cultures, keeping AFC on confluent conditions and in the presence of ACM.
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anti-CMFDA antibody (Molecular Probes) in double labeling experiments [19]. The primary antibodies used in the immunofluorescence experiments were the following: anti-CD34 (Becton Dickinson); anti-CD45 (Serotec); anti-ckit/CD117 (Dako); anti-Oct4 (Chemicon); anti-SSEA4 (Chemicon); anti-Sca-1 (Cedarlane, Hornby, Ont., Canada); antiThy-1/CD90 (Cymbus, Chandlers Ford, UK); 1B8 anti-SM22 [19]; anti-SM a-actin (Sigma); SM-E7 anti-SM myosin [19]; anti-PECAM/CD31 (Chemicon); anti-VE-cadherin (Santa Cruz); anti-von Willebrand factor (Dako); anti-vimentin (Dako); anti-desmin (Chemicon); anti-endoglin/CD105 (Cymbus). Immunofluorescence observations were carried out using a Zeiss Axioplan epifluorescence microscope and results obtained re-evaluated by Leica SPS2SL confocal microscope. Semiquantitative evaluation of antigenic expression profile in cytospins, cell cultures and transplants of AFC (Table 1) was carried out by manually counting positive cells with respect to Hoechst-stained nuclei by two independent operators. Cells in four randomly selected high-power fields, five cytospin preparations or cryosections, were averaged. There was a total of at least 100 cells in each field.
the original litter were randomly selected as candidates of autogenic cell transplantation with their own CMFDA-labeled AFC preparations after induction of AMI. 3.2. Characterization of AMI model The transient blood flow obstruction followed by wall dissection applied to induce the AMI model in pig gave rise to a thrombus formation (Fig. 2B—D) and a reproducible ischemic lesion which involved the lateral region of the left ventricle showing a marked wall thinning out (Fig. 2E and F). Cross-sections of the ventricular wall at day 7 postischemic time revealed that the infarction is predominantly transmural (not shown). Histological staining with hematoxylin—eosin showed that the ischemic region is characterized by the absence of CM (no immunoreactivity for cardiac troponin I and desmin), presence of granulation tissue (i.e. myofibroblasts identified by immunoreactivity for SM a-actin and SM22; neoangiogenesis identified by immunoreactivity for von Willebrand factor; fibrous tissue identified by immunoreactivity for fibronectin EIIIA isoform
3. Results 3.1. Animal data Three out of the 15 animals obtained by cesarean section died within a few days after birth (Fig. 1). The 12 surviving animals were operated via the percutaneous procedure which, in our hands, was not life-threatening and allowed for a clear cut recognition of the ischemic area and a safe positioning of AFC injections in the border zone. Six pigs of
Fig. 2. Results of percutaneous catheterism on circumflex coronary artery at day 37 after intervention. (A) Control vessel; (B) obliterated vessel (arrowhead); hematoxylin—eosin (C) and Azan Mallory (D) staining of the vessel showed in B, in which a thrombus completely occludes the lumen (arrowheads). (E) Lateral view of the heart showing the ischemic region (rectangle). (F) Cross-section of the heart showing localization of the ischemic region and thinning of the cardiac wall (arrowhead).
Fig. 3. Immunocytochemical patterns of cardiac cryosections from day 7 postAMI stained with antibodies to cardiac troponin I (cTnI; A), desmin (B), SM aactin (C), SM22 (D), von Willebrand factor (vWf; E), fibrionectin isoform EIIIA (F), procollagen I (G) and CD45 (H). Asterisks indicate the ischemic region. Note that CM have almost disappeared and a granulation tissue is now visible. A few inflammatory cells are detectable in H. Bar, 150 mm.
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and procolloagen I) and a few inflammatory, mainly CD45positive, cells (see Fig. 3). Histological examination of cryosections after 37 days of ischemia showed the presence of some vessels and myofibroblasts embedded into an organizing fibrous tissue (not shown). 3.3. Immunophenotyping of AFC before transplantation Fifteen days after the initial plating, two major morphologically distinguishable AFC populations appeared in vitro: one is of spindle-shape and the other is a polygonaltype (Fig. 4B). The polygonal-type cells, often grouped in clusters, are predominantly expressed in confluent cultures (Fig. 4C). There was no difference in the relative proportion of the two populations after passaging, but some AFC preparations from different animals presented almost exclusively spindle-shaped cells. Immunophenotyping of cultured, confluent AFC in ACM did not show any correlation between antigenic marker distribution (at least for the markers shown in Table 1) and morphological features. Interestingly, some markers (e.g. CD105/endoglin) disappeared when AFC were cultured on a different substrate (non-tissue culture Petri dishes vs. glass coverslips; not shown). When the AFC were grown in differentiating media, e.g. Endothelial Growth Medium or Smooth Muscle Cell Medium, both in sub- and confluent conditions, the expression of vascular cell markers is enhanced and occurs earlier than in
Fig. 4. Phase contrast aspects of in vitro grown AFC as seen at 7 (A), 15 (B) and 20 (C) days of culture in Amniotic Culture Medium. In panel A, black arrowhead in A shows a cell with a neuron-like morphology whereas white arrowhead indicates a spindle-shaped cell. Arrows in B and C indicate clusters of polygonal-shaped cells particularly abundant in confluent cultures (C). Phase contrast (D and G) and immunofluorescence staining (E and F;H and I) of 7day cultured porcine AFC in Endothelial Growth Medium (D—F) or Smooth Muscle Cell Medium (G—I). Note the different morphology of AFC when grown in the two different media. AFC grown in Endothelial Growth Medium where all Oct4 and PECAM positive (E) whereas VE-cadherin (F) is heterogeneously expressed (red arrowhead in F indicates a negative cell for this marker). In Smooth Muscle Cell Medium, a minority of AFC shows to be positive for SM aactin (blue arrowhead in H indicates a negative cells for this marker) and all are Oct4þ (H) SM myosin is, however, undetectable (I). Bars, 90 mm (A—C), 60 mm (D—G) and 70 mm (E and F, H and I). Hst, Hoechst staining. (For interpretation of the reference to color in this legend, the reader is referred to the web version of this article.)
Fig. 5. Immunofluorescence patterns of cytospin-plated porcine AFC stained with antibodies to vimentin (A), cardiac troponin I (cTnI; B), SM a-actin (C), SM22 (D), von Willebrand factor (vWf; E), and VE-cadherin (F). Note that no cells are reactive for cTnI. Arrowheads indicate cells with double-nuclei content. Nuclei are in blue. Bars, 50 mm (A, C—H) and 80 mm (B).
ACM. Fig. 4E and F shows that all AFC are now labeled with anti-PECAM and most of them are also positive with anti-VEcadherin. Similarly, the majority (about 60%) of AFC grown in the presence of Smooth Muscle Cell Medium are now SM aactinþ but a complete differentiation is not achieved inasmuch as these cells are SM myosin (Fig. 4H and I). All AFC in either cell cultures were Oct4þ (Fig. 4E and H). In cytospin preparations, all cells (obtained from AFC grown in ACM, just before transplantation) appeared to be of mesenchymal type inasmuch as they are vimentin-positive (Fig. 5; Table 1). Part of them was also reactive for the mesenchymal cell markers CD105/endoglin and Thy-1/CD90 (Table 1). They did not express hematopoietic cell markers, such as CD34 and c-kit, but they were variably positive for markers of smooth muscle (SM a-actin and SM22), and endothelial (von Willebrand factor, VE-cadherin, PECAM) cell lineages. Interestingly, minority of cells were also stained for the ‘embryonic markers’ SSEA4 and Oct4 (Table 1). Noticeably, about one-third of AFC collected and examined via cytospins (Fig. 5A and D) or glass coverslips (not shown) displayed a double-nuclei content. This ratio did not change with CMFDA labeling or with the duration of cell culturing. 3.4. Immunophenotyping of AFC after transplantation Before transplanting CMFDA-labeled AFC in the intact or AMI hearts we tested in vitro whether the immunofluorescence signal was still detectable with cell proliferation. AFC labeled with CMFDA were grown in pre-confluent conditions and subjected to repeated passages. After three passages, the immunofluorescence staining was still evident. From the fourth up to the eight passage, the presence of CMFDA in the AFC was detectable only using a FITC-labeled anti-CMFDA antibody (not shown).
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Fig. 6. Cell tracking at 2 h (A) and 30 days (B) of CMFDA-labeled AFC transplanted in ischemic porcine hearts by immunofluorescence. Long rows of transplanted cells are particularly visible in B whereas in A they are dispersed among cardiac tissues. In panel B is shown an example of CMFDAþ cells studied for the expression of differentiation markers and identified by small lettered boxes. Vital CMFDA transplanted AFC are in green and an apoptotic cell in yellow (identified by a yellow arrowhead in panel B). Bars, 110 (A) and 80 mm (B). (For interpretation of the reference to color in this legend, the reader is referred to the web version of this article.)
There was a considerable number of CMFDA-labeled cells 2 h after transplantation in the border zone of the ischemic region (Fig. 6A) whereas AFC were not found in cardiac regions remote from the injection sites. A large number of cells, morphologically identified as apoptotic cells, and few CMFDA-labeled cells could be detected when AFC were injected in the intact heart (not shown). As a general rule, AFC at day 30 post-implantation time were mostly found grouped in long rows in close vicinity of newly formed fibrous tissue (Fig. 6B) and very rarely seen apposed to the survived myocardial tissue of the border zone. CMFDA-positive cell aggregates were frequently observed, but without any evidence for organized tissue structures such as blood or lymphatic vessels. About 5 1% of the CMFDA-labeled cells present 2 h after transplantation in AMI hearts were still detectable 30 days later. Results of double immunofluorescence experiments of immunophenotyping of tracked AFC using the panel of antibodies as differentiation markers are shown in Fig. 7 and summarized in Table 1. Noticeably, the ‘embryonic stem’ cell markers SSEA4 and Oct4 completely disappeared and the mesenchymal markers vimentin (Fig. 7A—C), CD105/endoglin and Thy-1/CD90 (Table 1) were lost or decreased. AFC immunoreactive for cardiogenic cell lineage markers were in general less numerous with respect to the corresponding cells examined in cytospins or in tissue culture. In particular, the CMFDA-tracked cells did not develop any immunostaining for the myocardial marker cardiac troponin I (Fig. 7D—F). As to the endothelial (PECAM, von Willebrand factor, VEcadherin) and smooth muscle (SM22 and SM myosin) cell lineage markers, we observed a loss/decrease of reactivity with the corresponding antibodies. The only exception is represented by SM a-actin, which is still detectable (Fig. 7G—I). Altogether, the CMFDA-positive cells also reactive for the vascular cell markers shown in Fig. 7 are less than 2%. AMI pigs injected with aMEM showed a immunophenotypic pattern very similar to the day 30 ischemic heart (not shown). AFC injected in the nonischemic control heart gave an immunophenotypic profile similar to that of AFC-transplanted ischemic hearts (not shown).
Fig. 7. Phenotypic expression of mesenchymal (A—C), myocardial (D—F), smooth muscle (G—O) and endothelial (P—R) cell markers in CMFDA-labeled AFC transplanted in ischemic porcine hearts 30 days after injection. A—C, Vimentin; D—F, cardiac troponin I (cTnI); H and I, SM a-actin; J—L, SM myosin; M—O, SM22; P—R, PECAM. Green arrowheads indicate transplanted AFC exclusively positive for CMFDA whereas yellow arrowheads identify transplanted CMFDA-labeled AFC and also reactive with other antibodies. Nuclei are stained blue by Hoechst staining. Bar, 80 mm. (For interpretation of the reference to color in this legend, the reader is referred to the web version of this article.)
4. Discussion In this study, we investigated the differentiation potential of porcine AFC in vitro and in an in vivo model of AMI with the aim to evaluate the possible use of these immature stem cells for cellular therapy of myocardial ischemia. We selected the plastic-adherent AFC and autotransplanted these cells labeled with the cell tracker CMFDA in a novel model of AMI. Different porcine models of ischemic injury have been previously described to study the structural and functional consequences of CM/stem cell transplantation, namely surgical [13,15,16], microcoil embolization [21], interluminal coil occlusion and Gelfoam gelatine embolization [12], and ameroid constrictor implantation [14,17,18]. This
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protocol is safe, reproducible and gives almost no pericardial adherence formation and contrary to the coronary artery ligature does not form tenacious adherences and does not give rise to uncontrolled arrhythmias [21]. In vitro, the selected porcine AFC grown in ACM and in confluent conditions displayed mostly the immunophenotypic profile of the ‘classical’ bone marrow-derived mesenchymal stem cells (MSC), i.e. reactivity for vimentin, CD105/endoglin and CD90/Thy-1 [2] whereas the hematopoietic antigens CD34, Sca-1 and CD117/c-kit were absent [6]. Interestingly, markers of endothelial (von Willebrand factor, VE-cadherin, CD31/PECAM) and smooth muscle (SM a-actin, SM22) cell lineages are also expressed. None of the differentiation antigens, however, are entirely shared by the whole AFC population, which is also morphologically heterogeneous (Fig. 4 and [22]). The presence of the ‘embryonic stem’ cell associated cell surface marker SSEA4 along with Oct4 transcription factor suggests that porcine AFC closely resemble the human multipotent AFC [5,23]. This is confirmed by the in vitro experiments performed with Endothelial Growth Medium and Smooth Muscle Cell Medium: AFC are capable of giving rise to an endothelialand smooth muscle cell-like phenotypic conversion in the presence of Oct4. We do not say whether: (1) prolonged cultivation of AFC in these media can complete the differentation-maturation process that characterizes the vascular cell lineages, and (2) this event is accompanied by down regulation of Oct4 expression. The results of the in vivo study (obtained with AFC previously grown in ACM) suggest that these cells did not show a particular tendency to differentiate giving rise to cells of the cardiogenic lineages. In fact, after 30 days of transplantation in the ischemic myocardium, ‘embryonic stem’ cell and mesenchymal cell markers were absent or reduced, expression of differentiation markers of endothelial and smooth muscle cell lineages was diminished (Table 1). Interestingly, Oct4 and SSEA4 immunoreactivity was undetectable in all survived, transplanted cells. This may suggest that transplanted AFC not expressing originally these markers are unsuitable for phenotypic conversion to vascular cells. Alternatively, the original Oct4-positive AFC may have down regulated the expression of these markers as a consequence of vascular cell commitment induced by the myocardial microenvironment and, perhaps, they are differentiating cells. To sum up, contrary to the porcine bone marrow-derived MSC [13,14, and our own unpublished results], porcine AFC could not increment vascular structures in the ischemic/postischemic environment. As to AFC transdifferentiation to CM, our data suggest a different scenario with respect to porcine MSC [15,16] or human AFC [8]. In fact, we were unable to detect any antigenic expression of cardiac troponin I (cardiac troponin T or connexin43; not shown) in transplanted porcine AFC, at least after 30 days of transplantation. Though possible, it seems unlikely that the post-ischemic environment can influence the potential stem cell progression to the vascular and/or CM phenotypes. We took into account the suggestions raised by Lee and Makkar [24] about the benefits that could be attained when stem cell transplantation in AMI is carried out between days 7 and 14 post-injury time, i.e. when the inflammatory response is declining and reparative process is at its beginning. Our
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immunocytochemical data (Fig. 3) confirm that at the 7th day post-injury, when the AFC were transplanted, granulation tissue was particularly evident. It is well known that from this, tissue growth factors and cytokines are abundantly released and, hence, possibly sustaining transplanted cell growth/survival. In our model, however, these factors seem unable to induce a further differentiation of AFC. That the post-ischemic microenvironment is unable to influence the phenotypic potential of AFC is also demonstrated by the substantial similarity of the differentiation patterns of these cells in the ischemic vs. non-ischemic heart. Our data are apparently in contrast to those of Zhao et al. [8] who used freshly isolated ‘human amniotic mesenchymal cells’ injected in the ischemic rat heart and found that 1 or 2 months after xenotransplantation, these cells improved their myocardial commitment by expressing two markers of differentiated CM, i.e. atrial natriuretic protein and bmyosin heavy chains. This discrepancy could be explained by the inherent structural characteristics of mesenchymal cells obtained from the amniotic fluid (this study) and the placenta [8]. In fact, placental-derived mesenchymal cells, prepared by enzymatic digestion, express the cardiomyogenic transcription factors GATA-4 and Nkx2.5 [25], which can switch on these cells a peculiar propensity to differentiate along this lineage. We cannot exclude, however, that AFC may be phenotypically unstable and cell passaging to obtain enough cells for transplantation may compromise their capacity for engrafting and differentiate to cardiogenic cell lineages as might be inferred from the different phenotypic patterns observed in cytospins vs. cell cultures. Obviously, the incapability of AFC to be phenotypically converted to CM in our model (at least at the time point used in this study) does not deny the possibility that paracrine effects produced by transplanted AFC are indirectly able to influence the functional recovery of post-ischemic lesion [12]. In conclusion, our results indicate that despite the high expansion capacity of AFC these cells, at least in the swine and in naı¨ve, non-manipulated conditions are inadequate to support regeneration of cardiogenic cells. AFC do not seem to survive long enough in the periphery of the ischemic lesion and to give rise to a target-specific, committed progeny. Further ex vivo genetic and epigenetic manipulations might be needed to implement the surviving and differentiation potential of AFC. Acknowledgements We wish to thank Prof. R. Busetto and Dr A. Ramondo for setting up the AMI model in pig. We are indebted to Dr Giovanni Gaudenzi for staff organization. Dr Paolo De Coppi has been supported by ‘Fondazione Citta ` della Speranza’. This project was funded by a grant from Istituto Superiore di Sanita `, Programma sulle Cellule Staminali, Convenzione n. CS 18 to S. Sartore.
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