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Department of Histology and Embryology Faculty of Medicine,3 Akdeniz University, Antalya 07070, Turkey
Department of Obstetrics and Gynecology,4 Yale University, School of Medicine, New Haven, Connecticut 06520-8063
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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apoptosis, developmental biology, early development, placenta
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, and TNF receptor, and cytoplasmic/nuclear proteins, such as Bcl-2, Bak, Bax, and caspases [1–5]. Apoptosis takes part in both embryonic tissue morphogenesis and adult tissue regulation by balancing homeostasis. The most evident morphological signs of apoptosis are cellular shrinkage, membrane blebbing, nuclear condensation, and fragmentation, which also are the final steps of consequential signaling cascades [6]. Yui et al. [7] suggested that a physiological role of TNF and interferon-
expression in the placental villi may induce apoptotic death of cytotrophoblasts. Vasculogenesis is the process of new blood vessel formation from undifferentiated mesenchymal cells, whereas angiogenesis is the process of new blood vessel formation from preexisting vessels. Both processes occur in direct response to tissue demands and are crucial not only for the pregnancy-associated changes in the reproductive tract but also for embryonic and extraembryonic tissue (i.e., placenta) development. Endometrium, decidua, and placenta are rich sources of angiogenic growth factors [8, 9]. In a recent study, Demir et al. [10] showed that the origin and stages of placental angiogenesis differ from those of the postnatal angiogenesis in terms of cell types and angiogenic factors.
Placental vasculogenesis starts just after the invasion of allantoic mesoderm. Thus, the first structure of tertiary villi emerge around 21 days postconception (dpc) [11–13]. Vasculogenesis consists of three major steps: differentiation of undifferentiated mesenchymal cells to hemangioblasts and then to angioblasts, assembly of primordial vessels setting up a primitive vascular network, and transition from vasculogenesis to angiogenesis. Two forms of angiogenesis have been described: sprouting angiogenesis, and nonsprouting angiogenesis. During the vasculogenesis, first roundish or cord-like masses of hemangioblastic cells can be distinguished, followed by a clearly defined lumen around 28 dpc [9]. However, Demir et al. [12] showed that the first signs of primitive lumen formation start around 23 dpc.
As described above, placental vasculogenesis includes several steps, and collection of very early (22–48 dpc) placental samples to analyze these steps is quite difficult. Moreover, placental angiogenesis is different from conventional angiogenesis as described above, suggesting the existence of additional steps in the process [10]. In the present study, we hypothesized that during placental vasculogenesis and angiogenesis, differentiating endothelial and mesenchymal cells undergo apoptosis during the lumen formation, angiogenic cell cord connection, and vascular branching steps. To our knowledge, no studies have described any apoptotic signs through placental vasculogenesis and/or angiogenesis. Therefore, our aim was to analyze molecular and morphological markers of apoptosis using immunohistochemistry, TUNEL, histochemistry, and electron microscopy during very early stages of placental vasculogenesis.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Human placental tissues from the first trimester (n = 15 total from 22, 24, 26, 27, 29, 32, 36, 40, 42, 48, and 52 dpc) were collected from clinically normal pregnancies, which were terminated voluntarily by dilation and curettage. The distribution of the number of samples according to dpc was two samples each from 32, 36, 42, and 48 dpc and one sample each from 22, 24, 26, 27, 29, 40, and 52 dpc. The samples were classified as Pregnancy Weeks 3–4 (22–29 dpc), Pregnancy Weeks 5–6 (32–40 dpc), and Pregnancy Weeks 7–8 (42–52 dpc). Informed consent was obtained from each patient before obtaining the placenta. Consent forms and protocols were approved by the Ethical Committee of Akdeniz University. Mean age of the patients was 32.5 yr (range, 28–35 yr). The tissues were placed in PBS (pH 7.4) and transported to the laboratory for separation of placental villous from other tissues, such as decidua or amnion. Tissues were fixed in Bouin fixative and embedded into paraffin for immunohistochemistry or were fixed by immersion in 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) for ultrastructural analysis.
Immunohistochemistry
Serial sections were collected on poly-L-lysine-coated slides (Sigma, St. Louis, MO), dewaxed, dehydrated, and placed in citrate buffer. Immunohistochemical detection procedures have been described previously [11]. To unmask antigens, an antigen-retrieval procedure was performed by treating the samples twice in a microwave oven at 750 W for 5 min each time. After cooling for 20 min at room temperature, the sections were washed in PBS and then kept in 3% H2O2 for 15 min to remove endogenous peroxidase activity, followed by three washes with PBS. After blocking with 5% normal horse serum to reduce nonspecific binding, sections were incubated with mouse anti-CD31 (Labvision, Fremont, CA) and mouse anti-CD34 (Dako A/S, Glostrup, Denmark) primary antibodies at 4°C overnight. Thereafter, sequential 30-min incubations were performed with biotinylated secondary antibody (Vector Laboratories, Burlingame, CA) and peroxidase-labeled streptavidin. The resulting signal was developed with diaminobenzidine (Vector Laboratories) and mounted with glycerol-gelatin (Sigma). Negative-control staining was performed by replacing the primary antibody with the appropriate nonimmune immunoglobulin G isotype. Photomicrographs were taken with an Axioplan microscope (Zeiss, Oberkochen, Germany).
TUNEL-Immunohistochemistry Double-Labeling
Apoptosis in placental tissues was detected by enzymatic labeling of DNA strand breaks using TUNEL, which was conducted as described previously [14]. Serial paraffin sections (thickness, 5 µm) from the placental tissues were cut and taken onto the slides covered with poly-L-lysine, and after drying, the slides were left in the incubator at 45°C overnight and at 60°C for 1 h. After deparaffinization and dehydration, slides were washed twice in PBS for 5 min. Following the incubation of slides with the permeabilization solution (0.1% Triton X-100 in 0.1% sodium citrate) for 8 min at 4°C and washing twice with PBS for 5 min, the labeling reaction was performed using 50 µl of TUNEL reagent for each sample except for the negative control, in which reagent without enzyme was added and incubated for 1 h at 37°C. Following PBS washing, slides were incubated with converter reagent for 30 min at 37°C. After washing, color development for localization of cells containing labeled DNA strand breaks was performed by incubating the slides with Fast Red substrate solution for 10 min. The TUNEL labeling was conducted using a Cell Death Detection kit (Roche, Mannheim, Germany) and performed according to the manufacturer's instructions. Then, the same slides were processed for CD31 immunohistochemical staining through the same protocol described above.
Hematoxylin-Eosin Staining
Sections (thickness, 5 µm) were cut from paraffin blocks and collected on poly-L-lysine-coated slides (Sigma). Sections were dewaxed in xylene, dehydrated in descending alcohol series, and stained by a routine hematoxylin-eosin staining technique as described previously [15].
Electron Microscopy
Samples of human placental tissues were fixed by immersion in 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) at room temperature for 4 h and postfixed in 1% phosphate-buffered osmium tetroxide for 2 h. The specimens were dehydrated in ascending concentrations of ethanol and embedded in araldite-epoxy resin. Semithin sections (thickness, 0.5 µm) were stained with toluidine blue. Ultrathin sections (thickness, 70 nm) were double-stained with uranyl acetate and lead citrate [16, 17].
Quantitative Analysis of Apoptotic Index
Quantification of apoptotic cells was accomplished by counting the cells involving apoptotic bodies in angiogenic cell cords, main vascular patterns, vasculatures with advanced lumen, and stromal cells among the branching vascular areas sighted in the microscopic field. Then, the apoptotic cell number was divided to total cells in the related area for determination of the apoptotic percentage. Two investigators who were blinded to the slides analyzed two sections from each sample and five areas for each of the vasculogenic stages in each section. The number of cells varied depending on the areas. However, all cells in the areas were counted, and the ratios of apoptotic cells to total cells were calculated.
Statistical Analysis
Quantitative analysis of apoptotic cell number was normally distributed as assessed by the Kolmogorov-Smirnov test. Both ANOVA and post-hoc Tukey test for pairwise comparisons were used in statistical analysis. A level of P < 0.05 was considered to be significant. Statistical calculations were performed using Sigmastat for Windows, version 2.0 (Jandel Scientific Corporation, San Rafael, CA).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Using semithin sections and CD31 immunohistochemistry, vasculogenic and angiogenic areas were determined and classified as described previously [10, 12]. Development and recruitment of hemangioblastic progenitor cells, formation of hemangioblastic cell cords, differentiation of primitive endothelium, formation of primitive lumen, endothelial tube elongation, and angiogenic branching were determined [10, 12, 18, 19]. In mesenchymal villi, islands of hemangioblastic progenitor cells containing two to four cells were determined morphologically and by using CD31 immunoreactivity from 22 to 48 dpc (Fig. 1, a and b). We observed cells elongating from one island to others (Fig. 1, c and d). In addition, main vascular patterns including angiogenic cords with or without primitive lumen, angiogenic branching, and connecting tubes were observed in the semithin and CD31-stained placental sections of villi (Fig. 1, e and f). The hemangioblastic and hematopoietic aspects of the cells in the villous core were further confirmed according to the intensity of immunoreactivity with anti-CD34 antibody (Fig. 1, g and h).
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Molecular Evidence for Apoptosis During Vasculogenesis
To detect the apoptotic cells in vasculogenic areas at the molecular level, TUNEL-CD31 double-staining was used. No TUNEL-positive cell was observed in hemangioblastic progenitor cells observed alone. Moreover, cell islands, formed by recruitment of hemangioblastic progenitor cells, also were negative for TUNEL but positive for CD31 (Fig. 2a). When the later stages of vasculogenesis were analyzed with TUNEL at the stage of primitive vascular lumen formation, some cells located at the center of the primitive lumen were TUNEL-positive (Fig. 2, b–d). Interestingly, some of the stromal cells located between vasculogenic areas during the endothelial tube elongation and angiogenic branching also were TUNEL-positive (Fig. 3, a–c). However, in reverse correlation to vessel maturation, significantly fewer apoptotic nuclei were observed (Fig. 3d).
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To confirm these findings further, morphological indicators of apoptosis, such as cell shrinkage, nuclear segmentation, and nuclear bodies, were analyzed in slides stained with hematoxylin-eosin. The morphological findings and localization for apoptotic cells were similar to the results obtained from TUNEL-CD31 double-labeling (Fig. 4).
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Ultrastructural Evidence for Apoptosis During Vasculogenesis
Electron-microscopic findings supported the light-microscopic data presented above. During the early stages of placental vasculogenesis, groups of or single hemangiogenic cells derived from undifferentiated mesenchymal cells were noticed in the core of mesenchymal villi. The hemangiogenic cells attached to each other and formed cell islands and hemangioblastic cell cords close to the cytotrophoblastic layer (Fig. 5a). Moreover, the primitive lumen of the developing vessel structure emerging from the dilating intercellular space of hemangioblastic cells also was observed (Fig. 5b). Interestingly, during the lumen formation, noticeable signs of apoptosis, such as chromosome condensation, nuclear shrinkage, and apoptotic bodies, were detected (Fig. 5c). Also, some cell debris that likely belonged to the very late stage of apoptosis was observed around the main vascular pattern, with hemopoietic cell series at different stages of maturation (Fig. 5d).
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Quantitative Analysis of Apoptotic Cell Index
Two villous types, mesenchymal villous and immature-intermediate villous, were analyzed for apoptotic cell ratio. The ratio was calculated in four different areas of villi classified as follows: the angiogenic cell cords, the main vascular pattern (at the stage of lumen formation), in the vasculature with advanced lumen, and the stromal cells among the branching vasculogenic areas. All four areas were analyzed in the immature-intermediate villi, whereas only angiogenic cell cords and main vascular pattern areas were analyzed in the mesenchymal villi. The highest ratio for apoptotic cells was found in the villous core among the vascular branching areas and then in the primitive capillary lumen (Fig. 6). These values are significantly different from those of angiogenic cell cords and vasculatures with advanced lumens in the immature-intermediate villi (P < 0.05) (Fig. 6). Moreover, a significantly higher apoptotic index was found in the main vascular patterns of the immature-intermediate villi than those of mesenchymal villi (P < 0.05). However, no significant difference for the apoptotic index was found between the angiogenic cell cords of mesenchymal and immature-intermediate villi (Fig. 6). Furthermore, no significant difference was detected in total apoptotic cell index when compared according to the day of pregnancy. When the index was analyzed according to the vasculogenic stages across time, the apoptotic index showed no significant increase in angiogenic cell cord, main vascular pattern, and vasculature with advanced lumen stages as pregnancy advanced (Fig. 7). However, the index was increased significantly in vascular branching areas starting from Pregnancy Week 6 toward Pregnancy Weeks 7–9 (Fig. 7).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Demir et al. [10] showed that angiogenic cells and cell cords are established as "main vascular pattern-like structures," and these structures form the main route for vessel segments and new blood vessel tubes to attach. Furthermore, the growth occurs by cell budding and outgrowths in the longitudinal axis of the villous growth. Numerous microvessels and vasculogenic areas interconnect by various bridge-like connector cells and/or microvessel connecting tubes and participate in generating the early placental vascular network [10, 25].
In the classical steps of placental vasculogenesis, emergence of the first precursors of fetal endothelium in the villous core, the so-called hemangioblastic cell cords, can be demonstrated as early as 15–21 dpc [12, 26, 27]. These cells have no cellular extensions, but a few organelles form string-like aggregates of polygonal cells. Desmosomes or band-like junctions that resemble tight junctions bridge the narrow intercellular spaces between these cells. All these steps are confirmed by our present findings, even though we observed no signs of apoptosis either morphologically or molecularly. By 28 dpc, a formation of long, polygonal lumen with surrounding endothelial cells occurs [26, 27]. During this phase, hematopoietic stem cells become visible in the capillary lumen. These cells are not yet circulating, because no anatomical connection exists via the cord [12, 27, 28]. Our findings showing the presence of apoptotic nuclei and TUNEL-positive cells at the center of the capillary lumen suggest that some of the differentiating cells in the islands undergo apoptosis so that the primitive lumen of the vessel may form. The establishment of intercellular spaces between hemangiogenic cell cords and the connection to and fusion with each other to form the presumptive vessel lumen involve various steps [10, 12, 29]. In the present study, we showed that programmed cell death, apoptosis, takes place in this process as a novel approach for lumen formation during vasculogenesis. All findings obtained from electron microscopy, immunohistochemistry, and TUNEL analysis support this hypothesis.
After the assembly of primordial vessels, the primitive vascular network, vasculogenesis advances to angiogenesis [9]. In normal human pregnancy, capillary growth has a biphasic property. It involves an initial phase of branching angiogenesis, followed by a phase of nonbranching angiogenesis. In other words, villous morphology partly reflects the underlying angiogenic processes [30–36]. The endothelial tube segments formed by vasculogenesis are transformed into primitive capillary networks, which occurs by the balanced interaction of two parallel mechanisms: elongation of preexisting tubes by nonbranching angiogenesis, and ramification of these tubes by lateral sprouting (sprouting angiogenesis) [36]. The TUNEL-positive cells that we have observed on the elongation sites of the primitive tubes and among the primitive tubes that are likely to connect with each other suggest a possible mechanism for apoptosis that would open physical spaces. Similar results from electron microscopy and hematoxylin-eosin staining further enhance this possibility.
Many growth factors, such as basic fibroblast growth factor, vascular endothelial growth factor, placental growth factor, TNF, and their receptors, have been reported to stimulate vasculogenesis and angiogenesis in the early placental development [10, 18, 20, 34, 37–39]. None of these molecules has been proposed for the apoptotic signaling cascade except for TNF. However, we do not know whether the TNF-related apoptotic cascade is involved in these processes. On the other hand, several studies have suggested that increased apoptosis in endothelial cells inhibits angiogenesis [40–45]. The p38 mitogen-activated protein kinase mutant placentas display a lack of vascularization and increased rates of apoptosis, suggesting a defective placental angiogenesis [43]. Moreover, the regulation of endothelial cell apoptosis is considered to be a potential therapeutic target because endothelial cell apoptosis diminishes neovascularization in the adult organism. The induction of endothelial cell apoptosis may limit unwanted neovascularization of tumors. In contrast, the prevention of endothelial cell apoptosis may improve angiogenesis and vasculogenesis in patients with ischemia [44]. However, according to our findings, it should be considered that placental vasculogenesis requires apoptosis to have a normal and vigorous vessel development. Beck et al. [45] have shown that Ang-2 induction may cause endothelial cell proliferation or apoptosis depending on the presence or absence of vascular endothelial growth factor during angiogenesis. Those authors suggest that this apoptotic stimulus of Ang-2 is an inhibitory signal for angiogenesis.
In conclusion, the present study revealed, to our knowledge for the first time, that apoptosis is involved in the very early stages of placental vasculogenesis and angiogenesis. Although the present study divulges a new approach that suggests a role for apoptosis in these processes, it is not known which signaling mechanisms are involved or how the apoptotic cascades are triggered. Further molecular studies should be performed to answer these questions.
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FOOTNOTES |
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2 Correspondence: Umit A. Kayisli, Department of Histology and Embryology Faculty of Medicine, Akdeniz University, Antalya 07070, Turkey. FAX: 90 242 227 4486; uali{at}akdeniz.edu.tr
Received: 7 August 2004.
First decision: 21 September 2004.
Accepted: 5 November 2004.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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and -interferon against primary human placental trophoblasts. Placenta 1994 15:819-835[Medline]This article has been cited by other articles:
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