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Embryo |
Division of Reproductive Sciences,3 Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, Oregon 97006
Department of Obstetrics/Gynecology and Physiology and Pharmacology,4 Oregon Health & Science University, Portland, Oregon 97201
Department of Physiology,5 Keio University School of Medicine, Shinjuku-ku, Tokyo 160-8582, Japan
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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-fetal protein, cardiac troponin I, and vimentin) representative of all three embryonic germ layers. Greater than 70% of the expanded cell populations expressed antigenic markers (nestin and musashi1) for NPCs. After removal of FGF-2, approximately 70% of the NPCs differentiated into neuronal phenotypes expressing either microtubule-associated protein-2C (MAP2C) or neuronal nuclear antigen (NeuN), and approximately 28% differentiated into glial cell types expressing glial fibrillary acidic protein. Small populations of MAP2C/NeuN-positive cells also expressed tyrosine hydroxylase (
4%) or choline acetyltransferase (13%). These results suggest that monkey ES cells spontaneously differentiate into cells of all three germ layers, can be induced and maintained as NPCs, and can be further differentiated into committed neural lineages, including putative neurons and glial cells.
differentiation, embryonic stem cells, glial cells, monkey, neurons
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Recently, ES cell lines were established from primate embryos, including rhesus [5], marmoset [6], and cynomolgus monkeys [7] as well as humans [8, 9]. The derivation of human ES (hES) cells, together with recent developments in hES cell culture, such as embryoid body (EB) formation [10] and differentiation in vitro, demonstrate that hES cells, like their mouse counterparts, can differentiate into cells of multiple lineages in vitro, including neurons and glial cells [11–14]. Whereas the potential application of hES cells for transplantation in patients with neurodegenerative diseases or spinal injuries is clearly recognized, clinical application of ES cell-based therapy should be preceded by a thorough understanding of the growth and developmental fate of ES cells and their derivatives in vivo. Questions regarding the potential of transplanted primate ES cells to become neoplastic, their functional capacity, and the potential for immune rejection must be answered first in a nonhuman primate model [15]. In this regard, rhesus macaques resemble humans closely in anatomy, physiology, and genetic makeup, and monkey ES cell lines are available [16].
The directed differentiation of ES cells toward neural lineages has been studied in rodents, and several differentiation paradigms have emerged. For instance, ES cell aggregates cultured in medium containing retinoic acid undergo neuronal differentiation [17], and serum-free sequential culture with fibroblast growth factor (FGF)-2 has been used to isolate and enrich nestin-immunoreactive neural progenitors and neurons [18]. Neuronal differentiation of monkey ES cells, including tyrosine hydroxylase (TH)-positive phenotypes, has also been reported by coculturing monkey ES cells with bone marrow stromal cells [19]. Recently, serum-free culture conditions have been adapted to the production of a highly enriched neural population of hES cells [12, 14].
In the present study, we investigated the differentiation potential of monkey ES cells into EBs, neural progenitor cells (NPCs), and putative neural cells. We demonstrate that 1) monkey ES cells spontaneously give rise to cells of all three germ layers and neural lineages within EBs; 2) enriched populations of proliferating neural progenitors can be obtained via a combination of mechanical isolation, serum-free culture, and cell proliferation in the presence of FGF-2; 3) cryopreservation of monkey ES cell-derived NPCs is possible, with recovery of a high yield of viable NPCs; and 4) fresh or frozen ES cell-derived NPCs can be induced to differentiate into putative neurons and glial cells in vitro.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Monkey ES Cell Culture and EB Formation (Step 1)
The procedures for monkey ES cell culture were similar to those described by Thomson et al. [5]. Briefly, monkey ES cells (cell line 366.4 [16]) were grown on mitomycin C-treated (5 µg/ml, 37°C for 30 min; Sigma, St. Louis, MO) mouse embryonic fibroblast (MEF) feeder layers in gelatin-coated tissue culture dishes (Nalge Nunc International Co., Naperville, IL). The ES cell culture medium consisted of 80% Dulbecco modified Eagle medium (DMEM; with L-glutamine and glucose and without sodium pyruvate; Invitrogen, Grand Island, NY) supplemented with 20% fetal bovine serum (FBS; Hyclone, Logan, UT), 0.1 mM ß-mercaptoethanol (Sigma), 1% nonessential amino acids (Invitrogen), and 2 mM glutamine (Invitrogen). The medium was changed daily, and ES cell colonies were split every 5–7 days by incubation in collagenase IV (1 mg/ml, 37°C for 10–20 min; Invitrogen) and replating collected cells after centrifugation onto dishes with new MEF feeder cells.
For EB formation, entire ES cell colonies were loosely detached from MEF feeder cells by exposure to collagenase IV (1 mg/ml, 37°C for 10–20 min) and carefully aspirated into a micropipette, rinsed in ES medium three times, and cultured in hanging drops of ES medium (40 µl/drop) on the inner surface of 60-mm culture dish lids. The culture medium was changed daily.
Selection of NPCs (Step 2)
The EBs maintained for a total of 4 days (2 days in hanging drops and 2 days in suspension) were plated onto gelatin-coated culture dishes in ES medium. After 24 h of culture to allow cell attachment and surface spreading, ES medium was replaced with serum-free ITSFn medium containing DMEM/F12 (1:1) supplemented with insulin (10 µg/ml), sodium selenite (6.7 pg/ml), transferrin (5.5 µg/ml), and fibronectin (5 µg/ml; Invitrogen). The resulting EB outgrowths were maintained in serum-free ITSFn medium for 7 days, with medium replenishment every 2 days. For immunocytochemistry (ICC), NPCs were plated onto polyornithine- and laminin-coated glass coverslips (polyornithine, 15 mg/ml; laminin, 1 mg/ml; Sigma).
Expansion of NPCs (Step 3)
The EB outgrowths were incubated with 1–2 ml of collagenase IV (1 mg/ml, 5–10 min) followed by the addition of DMEM/F12 (1:1) plus 10% FBS. Dispersed cell clumps were collected by centrifugation (1000 x g, 10 min) and replated onto polyornithine- and laminin-coated six-well plates or glass coverslips in N2 medium (DMEM/F12 [1:1] supplemented with laminin [1 µg/ml; Invitrogen], FGF-2 [10 ng/ml; R&D Systems, Minneapolis, MN], and N2 supplement [1%; Invitrogen]). The medium was changed every 2 days, whereas FGF-2 was added daily. Cells plated onto coverslips were fixed in 2% paraformaldehyde for 10 min and held in 30% ethylene glycol and 20% glycerol in 0.05 M sodium phosphate buffer at -20°C for ICC analysis.
Differentiation of NPCs into Neural Lineages (Step 4)
The NPCs, either attached on substrates or developing as suspended spherical structures, were cultured in N3 medium that was changed every 2 days for 7–12 days until cells were fixed for ICC assays. The N3 medium was the same as the N2 medium, but it did not contain FGF-2.
Low-Temperature Storage of NPCs
The NPCs maintained in N2 medium were dissociated into small cell clumps by exposure to 0.05% trypsin and 0.04% EDTA in PBS and were neutralized in DMEM/F12 (1:1) containing 10% FBS. Cells were collected by centrifugation and transferred into a 1.2-ml cryovial (Nalge Nunc International) containing 1 ml of freezing medium (90% serum and 10% dimethyl sulfoxide [DMSO; Sigma]). Vials were slowly cooled (1°C/min) to -80°C and stored in liquid nitrogen. Thawing was in a 37°C water bath, and the freezing medium was diluted gradually with 10 ml of culture medium. Cells were plated as described above or incubated in culture medium containing 0.1% trypan blue solution (Sigma) for evaluation of viability.
Spherical structures in expanded NPC cultures (Step 3) were vitrified by incubation for 3 min in 10% glycerol in Hepes-buffered Tyrode albumin lactate pyruvate medium containing 20% fetal bovine serum (TH20), 3 min in 10% glycerol and 20% ethylene glycol in TH20, and then briefly (30–60 sec) in 25% glycerol plus 25% ethylene glycol in TH20. Spherical structures were drawn into a serological pipette and allowed to settle toward the tip before individual drops were slowly released directly into liquid nitrogen. These solid drops were collected with precooled forceps and sealed in liquid nitrogen-filled cryovials for storage. Warming and recovery of spherical structures was performed by pouring the vitrified drops and liquid nitrogen into a container. The solid drops were quickly moved into a solution of 0.5 M sucrose/TH20 for 3 min at room temperature and then transferred to 0.25 M sucrose and 0.125 M sucrose for 3 min each at room temperature before being rinsed and cultured.
Spherical structures were also frozen using equilibrium, controlled-rate cooling after incubation in 1.5 M ethylene glycol and 0.1 M sucrose for 20–30 min. Cryovials were cooled to -7°C at -2°C/min, seeded and further cooled to -40°C at -0.3°C/min, and then cooled again to -100°C at -4°C/min before storage in liquid nitrogen. Vials were thawed in a 37°C water bath, and the spherical structures were washed through progressively lower concentrations of cryoprotectant media (1.5, 1.0, 0.5, and 0 M ethylene glycol) before viability analysis.
ICC Staining
The procedures for ICC staining were similar to those described previously [20]. Briefly, 2% paraformaldehyde-fixed ES cells, EBs, or their derivatives cultured on polyornithine- and laminin-coated glass coverslips were rinsed twice with PBS (5 min each rinse) and once with 0.1% Triton-X 100 in PBS (5 min at room temperature). After treatment with 10% normal serum for 20 min at room temperature, cells were incubated with primary antibodies for 24–48 h at 4°C, washed twice with PBS (5 min at room temperature for each wash), and incubated either with biotinylated (for embryonic antigens) or fluorophore-conjugated (for other protein markers) second antibodies for 1 h (in the dark for immunofluorescence). Biotinylated second antibodies were linked to the biotin/avidin system (Vectastain; Vector Laboratories, Burlingame, CA) before signal amplification with 3',3'-diaminobenzidine (DAB; Vector Laboratories) according to the manufacturer's protocols. After three washes in PBS (5 min each wash), cells were counterstained with 4',6-diamindino-2-phenylindole dihydrochloride (DAPI; 300 nM; Molecular Probes, Inc., Eugene, OR), and the coverslips were mounted onto glass slides with a glycerol-based mounting solution containing 2.5% polyvinyl alcohol and 1,4-diazabicyclo[2,2,2] octane (Sigma). Mounted slides were placed under a hood at room temperature (for biotin/avidin/DAB) before image capture with light microscopy or in a light-proof box overnight at 4°C with storage at -20°C for immunofluorescence or confocal microscopy.
The primary antibodies used were against SSEA-1, SSEA-3, SSEA-4, TRA-1-60, and TRA-1-81 (mouse monoclonal, kindly provided by Dr. Peter W. Andrews, University of Sheffield, Sheffield, U.K.), Oct-4 (Santa Cruz Biotechnology, Santa Cruz, CA), -fetoprotein (mouse monoclonal, 1:500; Sigma), cardiac troponin I (mouse monoclonal, 1:50; Santa Cruz Biotechnology), vimentin (mouse monoclonal, 1:5; Hybrid Bank C1B7, Developmental Studies Hybridoma Bank, Iowa City, IA), human nestin (rabbit polyclonal, 1:2000; Nakama et al., unpublished results), musashi1 (rat monoclonal, 1:1000), neuronal nuclear antigen (NeuN; mouse monoclonal, 1:50; Chemicon, Temecula, CA), microtubular associated protein-2C (MAP-2C; mouse monoclonal, 1:500; Chemicon), glial fibrillary acidic protein (GFAP; mouse monoclonal, 1:100; Chemicon), choline acetyltransferase (ChAT; sheep polyclonal, 1:100; Chemicon), and TH (sheep polyclonal, 1:100; Chemicon). Secondary antibodies conjugated with fluorescein isothiocyanate (green), Cy5 (red), Texas Red (red), and AMCA (aminomethylcoumarin ultraviolet-purple) were purchased from Jackson Laboratories (West Grove, PA), and those conjugated with Alexa Fluor 488 (green) were from Molecular Probes. Images were obtained from a Leica confocal system (Leica, Heidelberg, Germany) at an original magnification of 20x or 40x with individual filter sets for each channel. Fluorescent cells, visualized using confocal microscopy, were quantified in at least five nonoverlapping fields in each sample. The total number of cells in each field was determined by counting DAPI-positive cell nuclei. Negative controls for each fluorophore-conjugated secondary antibody, carried out without the addition of the primary antibody, detected no nonspecific binding of the secondary antibodies. The expression of alkaline phosphatase in monkey ES cells was detected following fixation of cells with 100% ethanol using a Vector blue kit (Vector Laboratories) according to the manufacturer's instructions.
Statistical Analysis
Results are expressed as the mean ± SEM. Differences were evaluated by Student t-tests. Significance was set at P < 0.05.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The ES cells maintained on feeder layers of MEF formed colonies with individual cells containing large nuclei, prominent nucleoli, and distinctive cell boundaries (Fig. 2A). The cell line R366.4 has a 42,XY karyotype [16] and was at passage numbers 31–36 when placed in culture, where it has been maintained for more than 8 mo in vitro. Maintenance of ES cells in an undifferentiated state was monitored by visual observations of their morphology and the expression of primate ES cell-specific antigens TRA-1-60, TRA-1-81, and SSEA-4 (Fig. 2, B–D). In contrast to mouse ES cells, monkey ES cells did not express SSEA-1 (Fig. 2E). Monkey ES cells also expressed alkaline phosphatase (Fig. 2F) and the POU domain gene product, Oct-4 (Fig. 2G).
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EB Formation
The initial step in spontaneous ES cell differentiation was colony removal from the feeder layer with transference of isolated ES cell clumps to hanging drops of ES medium containing serum. Efforts to completely disassociate ES cells into single cells (n = 50, two replicates) before culture in hanging drops were unsuccessful, because the dissociated cells failed to aggregate and degenerated. After 2–3 days of culture in hanging drops, 76% ± 6% (271/357, three replicates) of the ES cell clumps had aggregated into early or simple EBs (Fig. 3A) as defined by their uniform appearance and lack of a central cavity. On ICC characterization of early EBs (5 days of culture in hanging drops, n = 5), only one EB expressed the ectodermal marker, vimentin, and none expressed cardiac troponin 1 as a mesodermal marker or -fetoprotein, an endodermal marker. In an effort to allow more extensive differentiation, simple EBs were maintained in culture for up to 13 days, resulting in many of them (42–76%) progressing to a more complex structure containing a dark central core and a visible cavity (complex/cystic EBs) (Fig. 3B). In contrast to early EBs, all complex/cystic EBs (total of 15 days of culture in hanging drops, n = 7) expressed all three of the germ layer markers (Fig. 4, A–C). The NPC markers, nestin (Fig. 4D) and musashi1 (Fig. 4E), and a neuronal cell marker, neuron-specific enolase (Fig. 4F), were also detected in all cystic/complex EBs.
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Neural Progenitor Cells
The next step in differentiation was serum removal and culture in the presence of ITSFn medium. On culture of early EBs in ITSFn medium, a distinctive pattern of cells with varied morphologies emerged that was indicative of differentiation: tightly packed epithelial-like and fibroblast-like cells at the periphery of the outgrowth and small elongated cells in the center. By Days 5–7, tubular structures of tightly packed columnar cells appeared in these EB outgrowths (Fig. 5A), which proliferated on addition of FGF-2 to the culture medium. Additional culture in N2 medium resulted in the formation of suspended spherical structures (Fig. 5B) and adherent NPCs (Fig. 5C). The adherent cell population could be cultured and passaged on reaching approximately 70% confluency. The spherical structures responded to FGF-2 stimulation with an increase in size; quartered or halved sections grew back to their original size. The ICC characterization of the expanded cell populations from the entire EB outgrowth indicated that 66% ± 6% (1070/1622, two replicates) and 55% ± 4% (939/1710, two replicates, P > 0.05) of the cells were positive for the NPC markers, nestin and musashi1, respectively (Fig. 6, A and B). In an effort to enrich NPCs, mechanical isolation of tubular structures from EB outgrowths, as seen in Figure 5A, was undertaken with subsequent culture in N2 medium containing FGF-2. Expansion of the cell population in the spherical structures that formed was achieved by dissecting the spheres (diameter, >0.7 mm) into halves or quarters under a dissecting microscope using a scalpel blade. The expanded cell populations from mechanically harvested tubular structures showed increased percentages of cells positive for nestin (78% ± 4%, 587/749, three replicates) and musashi1 (71% ± 8%, 474/667, three replicates), respectively.
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Because NPCs represent a convenient starting point for in vitro neural differentiation, a conventional DMSO-based, controlled-rate cooling program was applied to the adherent cells described previously with subsequent storage in liquid nitrogen. On thawing, 75.4% ± 8.7% (four replicates) of the NPCs retained viability as evidenced by trypan blue staining. In two further replicates, a total of 41 free-floating spherical structures were equally divided and stored at low temperature using vitrification or slow-rate controlled freezing. After thawing and culture in N2 media, 20 of 25 vitrified spherical structures (80%) survived and attached. Twelve of 16 conventionally frozen spherical structures (75%) also survived and attached. Thus, these methods of low-temperature storage yield comparable viabilities after thawing. Additionally, on FGF-2 removal, these NPCs stored at low temperature showed the same ability as nonfrozen NPCs to differentiate into neuronal lineages as described below.
Neuronal and Glial Cell Differentiation
Differentiation of NPCs into neuronal and glial cell lineages was initiated by removal of FGF-2 from the NPC culture medium and plating cells onto polyornithine and laminin substrates. Within 3–5 days of FGF-2 removal, neurite-like processes appeared (Fig. 7A). By 10–12 days, extensive fibers emanated from clumps of plated NPCs or from spherical structures (Fig. 7B). Networks of fiber bundles connecting cell clusters or spherical structures were also apparent (Fig. 7C). Approximately 45% ± 5% (801/1780, two replicates) of cells stained positive for markers of mature neurons, MAP-2C or NeuN (Fig. 6, C–E). However, in cell clusters or spherical structures with fiber bundles, approximately 69% ± 3% of cells stained positive for MAP-2C (315/459, five replicate) or NeuN. Interestingly, in other areas on the same coverslip, fewer cells were MAP-2C- or NeuN-positive (data not shown), suggesting that neuronal differentiation induced by this protocol may involve random or local development. However, the population of MAP-2C-positive cells (83% ± 7%, 916/1108, three replicates) or NeuN-positive cells (76% ± 6%, 822/1082, three replicates) increased after selection of NPC-containing tubular structures for culture after FGF-2 removal. At least some of these neuronal cells (NeuN-positive cells) expressed biochemical markers, including TH and ChAT. The number of double-labeled NeuN and TH cells (Fig. 6D) amounted to less than 4% (18/541, three replicates) of the ES cell-derived neuronal population, whereas almost 13% (73/583, three replicates) of NeuN-positive cells were also ChAT-positive (Fig. 5E). Approximately 29% ± 4% (514/1766, three replicates) of cells in the NPC population expressed the astrocyte-specific antigen GFAP (Fig. 6F), indicating that these cells had differentiated into mature glial phenotypes.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The protocol employed was a modification of experimentation conducted in the mouse [18], in which more than 80% of NPCs expressed nestin, a marker of neuroepithelial progenitor cells [21], after selection in serum-free medium and proliferation with FGF-2. Further differentiation and characterization of these cells showed that more than 60% of the population expressed the neuronal marker, MAP-2, after withdrawal of FGF-2. With monkey ES cells, approximately 60% of expanded NPCs expressed nestin (66%) or musashi1 (55%), another neurolineage marker [22], with approximately 45% of the cell population expressing markers of mature neurons after NPC differentiation into neural lineages. This cell population could be enriched greater than 1.5-fold (74–76%) when a mechanical isolation step was introduced, similar to the results of Reubinoff et al. [12] and Zhang et al. [14], who reported that highly enriched (>95%) hES-derived NPCs could be obtained by isolation of tubular structures from hES cell cultures or EB outgrowths. Thus, a combination of serum-free conditions and mechanical isolation is an efficient approach for deriving populations of highly enriched NPCs or putative neurons from primate ES cells. Representatives of the two nervous system lineages, neurons and astrocytes, were present in NPC cultures. The third lineage, oligodendrocytes, was not evaluated here but has been reported at low frequency in hES cell progeny [12]. We made little effort to define nonneural phenotypes, despite the recognition that nestin-positive cells also represent the starting point for derivation of pancreatic islet phenotypes [23, 24].
Differences in the differentiation of monkey and mouse ES cells noted here may suggest suboptimal conditions in the neuronal induction medium or reflect a higher degree of heterogeneity in monkey ES cell populations. Thus, monkey EBs after expansion may contain fewer NPC progenitor cells. The ICC analyses clearly demonstrated that complex/cystic, but not early, monkey EBs (Day 15 EBs) contained cells expressing antigenetic markers representative of all three germ layers. Precursors of other cell lineages or undifferentiated phenotypes likely compose the majority of the cell population within the early EBs used in NPC generation. Therefore, a lower rate of neuronal differentiation would be expected after expansion and differentiation. This is an active area of study along with the characterization of individual ES cell lines and their ability to differentiate into neural lineages.
Two NPC forms were observed during the expansion stage. One type grew as spherical structures, which may be analogous to what others [25, 26] have referred to as neurospheres, and a second grew as an adherent monolayer. We noted that both forms of NPCs can be proliferated in vitro and that both can give rise to differentiated phenotypes. The importance of these findings is now under investigation. In the context of cell-based therapy, in which large numbers of cells may be required to treat acute conditions over relatively short time frames, low-temperature storage protocols will be invaluable. Of course, ES cells can be cryopreserved, although weeks are required to establish viable cultures after thawing. In the present study, we found that both forms of NPCs could be stored using conventional equilibrium cooling or vitrification with rapid recovery of cultures in high yield. This capability allows the accumulation of cells for study with a significant savings in time and, ultimately, may be important in the context of cell-based therapy, in which large numbers of cells may be required to treat acute conditions over relatively short time frames.
In the present study, 60–80% of the ES cell-derived cells from NPCs were positive for the neuronal markers, MAP-2C or NeuN, whereas only a fraction expressed TH (4%), an anabolic enzyme involved in dopamine production, or ChAT (13%), which is likewise integral to choline formation. Additional methods to enrich a specific neuronal phenotype in the mouse (i.e., TH-positive neurons) have been reported. For example, high yields of dopaminergic neurons have been obtained after the addition of sonic hedgehog and fibroblast growth factor-8 (FGF8) into ascorbic acid-containing cultures of proliferating ES-derived NPCs [27]. Furthermore, Kawasaki et al. [19, 28] demonstrated that both mouse and monkey ES cells could be directed to differentiate into dopaminergic neurons on coculture with a stromal cell (PA-6 cell) feeder layer; indeed, 35% TH-positive cells were recovered from monkey ES cell cultures.
Additional characterization of functional neuronal phenotypes will help to define the extent of monkey ES cell neuronal differentiation following the current protocol, including gene expression profiles using reverse transcription-polymerase chain reaction and a determination of physiological behavior. Specifically for dopaminergic neurons from monkey ES cells, this should include a determination of dopamine concentrations in cells and culture medium after induced release by reverse-phase high-performance liquid chromatography [29] and functional capacity using electrophysiological methods [27, 30]. Before use of ES cell-derived dopaminergic neurons in cell-based therapy for Parkinson disease, highly purified cell preparations will be required.
In summary, our results demonstrate that rhesus monkey ES cells are able to generate proliferating NPCs via EB formation and NPC selection and expansion. Furthermore, ES cell-derived NPCs can be stored frozen in highly purified populations before they are subjected to neural differentiation. Enrichment and isolation of monkey ES cell-derived neural stem/progenitor cells, neurons, or glial cells in vitro [31–33] will facilitate the potential application of stem cell-based therapy in the treatment of neurodegenerative diseases.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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2 Correspondence: Don Wolf, Oregon National Primate Research Center, 505 NW 185th Avenue, Beaverton, OR 97006. FAX: 503-533-2494; wolfd{at}ohsu.edu
Received: 9 October 2002.
First decision: 5 November 2002.
Accepted: 27 November 2002.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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