Stem Cells
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

First published online June 7, 2005
Stem Cells Vol. 23 No. 7 August 2005, pp. 965 -974
doi:10.1634/stemcells.2004-0174; www.StemCells.com
© 2005 AlphaMed Press

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
2004-0174v1
23/7/965    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Reprints/Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bryja, V.
Right arrow Articles by Hampl, A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bryja, V.
Right arrow Articles by Hampl, A.

Abnormal Development of Mouse Embryoid Bodies Lacking p27Kip1 Cell Cycle Regulator

Vítezslav Bryjaa,b, Lukás Cajáneka, Jirí Pacherníka,c, Anita C. Halld, Viktor Horváthe, Petr Dvoráka,b,c, Ales Hampla,b,c

a Center for Cell Therapy and Tissue Repair, Charles University, Prague, Czech Republic;
b Department of Molecular Embryology, Institute of Experimental Medicine, Academy of Sciences of the Czech Republic, Prague, Czech Republic;
c Laboratory of Molecular Embryology, Mendel University Brno, Brno, Czech Republic;
d Laboratory of Molecular Neurobiology, Department of Medical Biochemistry and Biophysics, Karolinska Institutet, Stockholm, Sweden;
e Laboratory of Cytokinetics, Institute of Biophysics, Academy of Sciences of the Czech Republic, Brno, Czech Republic

Key Words. Mouse embryonic stem cells • p27 • Embryoid bodies • Lewis-X • Apoptosis

Correspondence: Ales Hampl, D.V.M., Ph.D., Laboratory of Molecular Embryology, Mendel University Brno, Zemedelská1, 61300 Brno, Czech Republic. Telephone: 420-5-45133297; Fax: 420-5-45133357; e-mail: hampl{at}mendelu.cz


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Conclusion
 References
 
Cultures of three-dimensional aggregates of embryonic stem cells (ESCs) called embryoid bodies (EBs) provide a valuable system for analyzing molecular mechanisms that regulate differentiation of this unique cell type. Cyclin-dependent kinase inhibitor p27Kip1 (p27) becomes elevated during the differentiation of mouse ESCs (mESCs). In this study, various aspects of differentiation of EBs produced from normal and p27-deficient mESCs were analyzed to address the biological significance of this elevation. It was found that EBs lacking p27 grew significantly bigger, but this was not accompanied by detect-able abnormalities in the activities of cyclin-dependent kinases (CDKs). In most EB cells, downregulation of activating cyclins rather than upregulation of inhibiting p27 is probably responsible for lowering the activity of their CDKs. Abnormalities in the development of specific cell lineages were also observed in p27-deficient EBs. These included elimination of cells positive for cytokeratin endo-A (TROMA-I) and increased proliferation and formation of cavities originating from cells positive for Lewis-X. Our data also suggest that although two different pools of Lewis-X–expressing cells, cluster forming (ESC-like) and cavity forming (neural progenitors), normally exist in EBs, the absence of p27 leads to the enhancement of only the neural pool. No failure was found when the neurogenic capacity of p27-deficient mESCs was tested using various protein markers. Together, our data point to a dual role of p27 in mESCs, with one role being in the regulation of proliferation and the other role in establishing some other aspects of a differentiated phenotype.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Conclusion
 References
 
Cell cycle regulatory protein p27Kip1 (from this point referred to as p27) belongs to a Cip/Kip family of cyclin-dependent kinase (CDK) inhibitors. Because its best-studied function is inhibiting the activity of cyclin-CDK complexes that drive the progression of the cell cycle, p27 is mostly understood as a negative regulator of cell proliferation (for review, see [1]). The phenotype of p27-null animals, which are significantly larger due to a higher number of cells, supports such a view [2–4]. However, detailed analyses of p27–/– animals have also revealed defects that could not be explained by simple alterations in cell proliferation. Specifically, it was shown that processes such as cellular differentiation and apoptosis were severely affected in p27-deficient mice [5–10].

Embryonic stem cells (ESCs) are unique in terms of molecules and mechanisms used to regulate their undifferentiated growth [11–13]. In regard to this fact, we have recently found that rather than for regulation of proliferation of pluripotent mouse ESCs (mESCs), p27 is essential for the in vitro differentiation of such cells to proceed normally. Specifically, when p27-deficient mESCs grown in monolayer are induced to differentiate into extraembryonal endoderm by withdrawal of leukemia inhibitory factor (LIF) combined with retinoic acid treatment, most of these mESCs die by apoptosis before finishing their differentiation program [14]. Although this phenomenon is well-pronounced in vitro, its relevance to the development of the early embryo is limited due to the used differentiation system. It is well accepted that early developmental processes can be mimicked in vitro by culturing multicellular aggregates of mESCs in the absence of LIF and feeder cells [15]. Under such conditions, mESC aggregates give rise to simple embryoid bodies (EBs) containing an outer layer of endodermal cells and a solid core of undifferentiated ectodermal cells. The inner cells of simple EBs subsequently undergo the wave of programmed cell death to form cystic EBs, a process called cavitation [16]. Because cystic EBs are cultured in vitro, they provide a unique tool for the analysis of the impact of individual genes and proteins on proliferation, differentiation, and apoptosis in early development (for review, see [17]).

In this study, the phenotypes of EBs produced from normal and p27-deficient mESCs were analyzed in detail to improve our understanding of the significance of p27 for early embryogenesis. The data here show that in the absence of p27, the EBs reach a bigger size than their normal counterparts, most likely due to enhanced cavitation affecting primarily Lewis-X–positive cells.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Conclusion
 References
 
Cell Culture
Both normal and p27-deficient mESCs [14] were propagated in an undifferentiated state by culturing on gelatinized tissue culture plastic in Dulbecco’s modified Eagle’s medium containing 20% fetal calf serum, 100 mM nucleosides, 0.05 mM ß-mercaptoethanol, 100 iu/ml penicillin, and 0.1 mg/ml streptomycin (referred to as complete DMEM), supplemented with 1,000 U/ml LIF. To accomplish differentiation in EBs, mESCs were trypsinized, depleted of LIF, and plated onto bacteriological dishes that promote formation of floating cell aggregates (5 x 104 cells/ml). EBs were transferred daily to fresh medium in a new bacteriological dish. Neural differentiation in EBs was accomplished using a modified protocol of Bain et al. [18] that included the following: (a) 4-day culture under nonadherent conditions in complete DMEM without LIF; (b) 4-day culture under nonadherent conditions in complete DMEM supplemented with 1 µM retinoic acid; and (c) further culture on gelatinized tissue culture dishes in DMEM/F12 (1:1) medium supplemented with insulin, transferrin, and selenium (ITS supplement; Invitrogen Life Technologies, Carlsbad, CA, http://www.invitrogen.com) and antibiotics as above (referred to as ITS medium). ITS medium was then changed at 3-day intervals.

Western Blotting
For Western blot analysis, cell samples were prepared as follows: mESCs and/or EBs were washed twice with phosphate-buffered saline (PBS) (pH 7.2) and lysed in 100 mM Tris/HCl (pH 6.8), 20% glycerol, and 1% SDS. Protein concentrations were determined using DC Protein Assay Kit (Bio-Rad, Hercules, CA, http://www.bio-rad.com). Equal amounts of total protein were subjected to 10% SDS-PAGE, electrotransferred onto Hybond-P membrane, immunodetected using appropriate primary and secondary antibodies, and visualized by ECL+Plus reagent (Amersham, Aylesbury, U.K., http://www1.amershambiosciences.com) according to manufacturer’s instructions. When required, membranes were stripped in 62.5 mM Tris/HCl (pH 6.8), 2% SDS, and 100 mM ß-mercaptoethanol, washed, and reblotted with another antibody. The antibodies used were as follows: mouse monoclonal antibody to cyclin D1 (sc-450), rabbit polyclonal antibodies to Oct-4 (sc-9081), cyclin E (sc-481), PARP (sc-7150), and CDK4 (sc-260), and goat polyclonal antibodies to CDK2 (sc-163-G), GAP-43 (sc-7457), and GFAP (sc-6170) (purchased from Santa Cruz Biotechnology, Santa Cruz, CA, http://www.scbt.com); mouse monoclonal antibody to p27 (K25020) (purchased from Transduction Laboratories, Lexington, KY, http://www.bdbiosciences.com); mouse monoclonal antibodies to cyclin A (Ab-1,E23) and cyclin D2(Ab-4,DCS-3.1+DCS-5.2)(purchased from Neomarkers, Fremont, CA, http://www.neomarkers.com); mouse monoclonal antibody to N-CAM (C0678) (purchased from Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com), mouse monoclonal antibodies to E-cadherin (C20820 [GenBank] ) and N-cadherin (C70320 [GenBank] ) (purchased from Becton, Dickinson and Company, San Jose, CA, http://www.bd.com); mouse monoclonal antibody to cyclin D3 (DCS-22) (generously provided by Dr. Lukas, Danish Cancer Society, Copenhagen, Denmark); and cytokeratin endo-A (TROMA-I) antibody (specified in Morphological, Immunohistochemical, and Immunocytochemical Analyses section). After immunodetection, each membrane was stained by amidoblack to confirm equal protein loading.

Immunoprecipitation and Kinase Assays
For these assays, mESCs and/or EBs were extracted for 30 minutes in ice-cold lysis buffer (50 mM Tris/HCl [pH 7.4], 150 mM sodium chloride, 0.5% Nonidet P-40, 1 mM ethylenediaminetetraacetic acid, 0.1 mM dithiothreitol, 50 mM sodium fluoride, 8 mM ß-glycerophosphate, 100 mM phenylmethylsulfonyl fluoride,1 µg/ml leupeptin, 1 µg/ml aprotinin, 10 µg/ml soybean trypsin inhibitor, and 10 µg/ml tosylphenylalanine chloromethane). Extracts were cleared by centrifugation at 15,000g for 5 minutes at 4°C and stored at –80°C until use. After thawing, protein concentrations were determined using a DC Protein Assay Kit (Bio-Rad). Extracts were subjected to initial absorption with protein G agarose beads and then incubated with appropriate antibody for 1 hour in an ice bath. Immunoprecipitates were collected on protein G agarose beads by overnight rotation, washed four times with lysis buffer, resuspended in 2 x Laemmli sample buffer, and subjected to SDS-PAGE followed by Western blot analysis. For kinase assays, immunoprecipitates were prepared as above, except that the last two washes were done using kinase assay buffer (50 mM HEPES [pH 7.5], 10 mM MgCl2, 10 mM MnCl2, 8 mM ß-glycerophosphate, and 1 mM dithiothreitol). For CDK2, kinase reactions were carried out for 30 minutes at 37°C in a total volume of 25 µl in kinase assay buffer supplemented with 100 µg/ ml histone H1 (type III-S) and 40 µCi/ml [32P]ATP. For CDK4, kinase reactions were carried out for 30 minutes at 30°C in a total volume of 25 µl in kinase assay buffer supplemented with 160 µg/ ml GST-pRb (type III-S) and 40 µCi/ml [32P]ATP. Reactions were terminated by addition of 2 x Laemmli sample buffer, and each reaction mix was subjected to SDS-PAGE and autoradiography. When required, the intensities of signals were assessed by densitometry using Intelligent Quantifier software (BioImage, Ann Arbor, MI, http://www.bioimage.com).

Morphological, Immunohistochemical, and Immunocytochemical Analyses
In culture, developing EBs were examined on Olympus SZH10 microscope and photographed using an Olympus Camedia 3030 digital camera (Olympus C&S, Ltd., Prague, http://www2.olympus.cz). For size analysis, a minimum of 200 EBs on 10 independent pictures were measured using Image-Pro Plus software (Media Cybernetics, Silver Spring, MD, http://www.mediacy.com). For indirect immunofluorescence, EBs were fixed overnight in ice-cold buffered 4% paraformaldehyde (pH 7.4), slowly dehydrated in ethanol, and embedded in polyester wax (Agar Scientific, Essex, U.K., http://www.agarscientific.com). Ten-micrometer-thick sections were made and placed onto egg white–coated slides, dewaxed with ethanol, and rehydrated. The sections were quenched for 1 hour at room temperature (RT) in 1% bovine serum albumin (BSA) in PBS (pH 7.4), incubated with appropriate primary (overnight at 4°C) and fluorescein isothiocyanate (FITC)–conjugated secondary (1 hour at RT) antibodies, and mounted to Mowiol (Hoechst, Frankfurt, Germany) containing 1,4-diazobicyclo-[2.2.2]-octane to prevent fading. Negative controls in which a primary antibody was omitted were always included. The reactions were examined using an Olympus Fluoview confocal laser-scanning microscope system.

For immunocytochemistry on immobilized EB cells, EBs were disintegrated by trypsinization, and resulting cells were allowed to adhere for 5 hours onto gelatinized microscopic coverslips. Then the cells were fixed for 30 minutes in ice-cold buffered 2% paraformaldehyde (pH 7.4), rinsed with PBS, and blocked for 1 hour at RT in PBS containing 1% BSA, 0.05% Tween, and 0.01% sodium azide. After incubation with appropriate primary antibody (overnight at 4°C) and FITC-conjugated secondary antibody (1 hour at RT), cells were extensively washed with 0.05% Tween in PBS and mounted and analyzed as above for sectioned EBs.

For immunohistochemical analysis of embryonic brains, E10.5 embryos of CD1 strain of mouse were fixed in 4% paraformaldehyde (4 hours, 4°C), rinsed with PBS, and cryoprotected in 20% sucrose in PBS overnight. The embryos were then embedded in OCT mounting medium, and 14-µm coronal sections were cut on a cryostat. The sections were rinsed in PBS, blocked in 5% goat serum in PBS, and incubated overnight with appropriate primary antibody in PBS containing 3% BSA and 0.05% Triton. Sections were then washed in PBS, incubated with the secondary antibody, and, after Hoechst counterstaining, mounted in PBS and glycerol (1:9).

Mouse monoclonal antibody to SSEA-1 (TEC-01) was generously provided by Dr. Draber (Institute of Molecular Genetics, Academy of Sciences of the Czech Republic, Prague, Czech Republic). The hybridoma TROMA-I developed by Drs. Brulet and Kemler, the hybridoma FORSE-1 developed by Dr. Patterson, and mouse monoclonal antibody against {alpha}-nestin (Rat-401) developed by Dr. Hockfield were obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the National Institute of Child Health and Human Development and maintained by the University of Iowa, Department of Biological Sciences (Iowa City, IA, http://www.uiowa.edu/~dshbwww).

Flow Cytometry and Cell Sorting
For analysis by flow cytometer, EBs were reduced to single-cell suspension by trypsinization, fixed for 1 hour in ice-cold buffered 2% paraformaldehyde (pH 7.4), rinsed with PBS, and blocked for 1 hour at RT to avoid nonspecific interactions in PBS containing 1% BSA, 0.05% Tween, and 0.01% sodium azide. After incubation with appropriate primary antibody (overnight at 4°C) and FITC-conjugated secondary antibody (1 hour at RT), cells were washed four times with 0.5% BSA in PBS, resuspended in PBS, and subjected to analysis by flow cytometer equipped with cell-sorting and cell-concentrating units (argon ion laser, 488 nm for excitation, FACSCalibur; Becton, Dickinson and Company). Negative controls represent samples in which a primary antibody was omitted. Cell debris was excluded by appropriately increasing the forward scatter. Cells from gate G2 = R2 (FL-1 [Height] positive) and G3 = NOT R2 (FL-1 [Height] negative), respectively, on side scatter versus FL-1 (Height) plot were continually sorted out with maximum sorting ratio, 250 events per second, to cell culture inserts with PET track-etched membrane (12-well format, 1.0- µm pore size; Falcon, Becton, Dickinson and Company).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Conclusion
 References
 
p27-Deficient EBs Grow Bigger Than Their Normal Counterparts
Because p27 is known primarily for its proliferation-regulating properties, it was first necessary to determine whether the absence of p27 results in the alteration of growth of EBs. EBs were allowed to form from both normal and p27–/– mESCs, and they were digitally photographed at day 10 of their development. The average diameter of EBs in each wild-type and p27–/– group was then determined using image-analysis software while assuming the EBs to be of round shape. Because data of both normal and p27-deficient cells showed log-normal distribution (Kolmogorov-Smirnov test; p = .02 for p27+/+ data and p = 1.5 x 10–8 for p27–/– data), they were ln-transformed (Kolmogorov-Smirnov test of ln-transformed data; p = .49 for p27+/+ data and p = .07 for p27–/– data) and compared by Student’s t-test. As demonstrated in Figure 1Go, p27-deficient EBs grew significantly bigger than normal EBs (two-tailed Student’s t-test of ln-transformed data sets; p = 8.8 x 10–16).



View larger version (36K):
[in this window]
[in a new window]
  		Figure 1. Effect of p27 on size of EBs. Normal and p27-deficient EBs were cultured up to day 10, photographed, and analyzed by image analysis software. Typical EBs are shown (A), and statistical analysis (B) of diameter of EBs (box-and-whisker plot; median, 25% to 75% interval, minimal and maximal values) is presented. Abbreviations: EB, embryoid body; K-S, Kolmogorov-Smirnov test of normal distribution of data.

 
The Activities of CDKs Are Not Deregulated in p27–/– EBs
Increased activities of CDK2 and CDK4 resulting in hyperplasia of various tissues or organs are typical for mice that are deficient in p27 [2, 4, 10, 14]. Therefore, it was important to determine whether the bigger size of p27–/– EBs observed here was correlated with upregulated CDK activities. Undifferentiated mESCs and EBs at days 5, 10, and 20 of their development, both wild-type and p27-deficient, were analyzed to address this question. The following parameters were determined: (a) levels of CDK2, CDK4, cyclin A, cyclin E, all three D-type cyclins, and p27; (b) kinase activities associated with CDK2, CDK4, cyclin A, and cyclin E; and (c) the amounts of p27 associated with the regulators listed under (b). During the development of wild-type EBs, the amount of p27 physically bound to CDK2, CDK4, cyclin A, and cyclin E, respectively, dramatically increases, corresponding to the elevation of the total amount of p27 (Fig. 2Go). Except for a generally lower amount of cyclin D3 in p27-deficient mESCs and EBs that can be well explained by the molecular mechanism described by us previously [10, 14], the behavior of no other cyclin was significantly affected by the absence of p27 (Fig. 2Go). Total amounts of all cyclin A, cyclin E, cyclin D1, and cyclin D3 were downregulated with the progressing development of EBs (Fig. 2Go). On the other hand, cyclin D2 followed the opposite pattern, being undetectable in mESCs and increased in EBs (Fig. 2Go). Downregulation of the kinase activities of CDK2, cyclin A, cyclin E, and CDK4 took place in developing EBs irrespectively of their p27 genotype (Fig. 2Go). These data document that the abnormal growth of EBs produced from p27-deficient mESCs cannot be due to hyperactivation of major CDKs. Furthermore, the results suggest that downregulated cyclins rather than elevated p27 are responsible for the decline of CDK activities that accompanies EB development.



View larger version (49K):
[in this window]
[in a new window]
  		Figure 2. Analysis of cyclin-CDK complexes in normal and p27-deficient EBs. EBs were differentiated from normal and p27-deficient mES cells (D0) and lysed at day 5 (D5), day 10 (D10), and day 20 (D20) of differentiation. Cell lysates were used to immunoprecipitate CDK2, cyclin A, cyclin E, and CDK4. The kinase activities toward histone H1 or GST-Rb were determined by autoradiography. The amount of p27 in immunoprecipitates was determined by Western blotting. The total amount of CDK2, cyclin A, cyclin E, CDK4, D-type cyclins, and p27 as determined by Western blotting is also shown. Densitometry was used to assess the intensity of autoradiographic signal. The average density of p27+/+ (D0) sample was defined as 1.0, and from this value all other values were calculated. Data represent the means, with standard deviations indicated by error bars. Western blots and autoradiograms are representative of at least three independent replicates. Abbreviations: EB, embryoid body; IP, immunoprecipitate; WB, Western blotting.

 
The Absence of p27 Prevents Normal Expression of Cytokeratin Endo-A (TROMA-I) in Developing EBs
We have previously shown that p27-deficient mESCs suffer from apoptosis when they are induced to differentiate into TROMA-I–positive extraembryonal endoderm in monolayer culture [19]. When wild-type mESCs were allowed to differentiate into EBs, strong positivity for TROMA-I was detected by Western blot as early as at day 5 of differentiation and stayed unchanged until the end of culture at day 20 (Fig. 3AGo). In contrast, only very low expression of TROMA-I was reached in EBs produced from p27-deficient mESCs, although mESCs of both genotypes progressed in their differentiation, as documented by downregulation of the stem cell marker Oct-4 (Fig. 3AGo). We next proceeded to determine whether a low proportion of TROMA-I–positive EBs, or decreased numbers of TROMA-I–positive cells in EBs, underlay the lower levels of cytokeratin endo-A determined above. Immunohistochemical visualization of cytokeratin endo-A on sectioned EBs produced by 10-day differentiation of normal and p27–/– mESCs was used to address this question. More than 90% of EBs of wild-type genotype contained TROMA-I–positive cells, with these cells being abundantly distributed in the superficial layer of the EBs (Figs. 3B, 3CGo). In contrast, only approximately 10% of p27–/– EBs contained TROMA-I–positive cells, which were invariably restricted to only dispersed individual cells (Fig. 3CGo).



View larger version (29K):
[in this window]
[in a new window]
  		Figure 3. Analysis of expression of TROMA-I in normal and p27-deficient EBs. (A): EBs were differentiated from normal and p27-deficient mouse embryonic stem cells (D0) and harvested at day 5 (D5), day 10 (D10), and day 20 (D20) of differentiation. The expression of Oct-4 and TROMA-I was analyzed by Western blotting. (B): EBs (day 10) were fixed, sectioned, and immunohistochemically stained using antibody against TROMA-I (green) and by PI (red). Typical pattern of TROMA-I staining for p27+/+ and p27–/– EBs is shown. (C): EBs containing at least some TROMA-I–positive cells were considered as TROMA-I–positive, and their proportion within total number of EBs was calculated. n indicates number of analyzed EBs. Abbreviations: EB, embryoid body; PI, propidium iodide; TROMA-I, cytokeratin endo-A.

 
Lewis-X–Positive Cells Form Cavities in p27-Deficient EBs
The abnormality in development of TROMA-I–positive cells itself could not explain the overgrowth of EBs lacking p27. Therefore, the question remained whether the bigger size of EBs may mirror an abnormal expansion of defined cell types. We have collected a large selection of antibodies against lineage-specific markers and applied them on the sectioned EBs as above to address this question. Interestingly, of all tested markers, only the early neural antigen identified by the antibody FORSE-1 [19] revealed a significant difference between normal and p27-deficient EBs. Specifically, while in normal EBs FORSE-1 positivity was limited to clusters of 10 to 50 cells, under p27–/– conditions, the FORSE-1 positivity was redistributed to both surround and fill the cavities in most EBs (Fig. 4AGo). Such redistribution was invariably accompanied by a loss of FORSE-1–positive cell clusters in p27–/– EBs. A similar phenotype was observed in all three tested p27-deficient mESC lines.



View larger version (54K):
[in this window]
[in a new window]
  		Figure 4. Expression of FORSE-1 and TEC-01 in normal and p27-deficient EBs. EBs were differentiated from normal and p27-deficient mouse embryonic stem cells, and at day 10 they were fixed, sectioned, and immunohistochemically stained using antibody against FORSE-1 and TEC-01, respectively (green), and PI (red). (A): Typical patterns of FORSE-1 staining are shown. Depending on distribution of FORSE-1–positive cells, two phenotypes of FORSE-1–positive EBs were distinguished: cluster (typical for p27+/+ EBs; EB contains a compact aggregate of 5 to 100 FORSE-1–positive cells without a cell-free space inside the group of positive cells) and cavity (typical for p27–/– EBs; area of FORSE-1–positive cells is characterized by the presence of cell-free space and expression of FORSE-1 is often polarized, having its maximum in lumen of the cavity). Proportion of individual phenotypes of FORSE-1–positive EBs is presented as percentages of EBs typical by clusters and cavities, respectively. n indicates number of analyzed EBs for each genotype. Data represent the means, with standard deviations indicated by error bars. (B): Serial sections of EBs were stained either for FORSE-1 or TEC-1 (green) and compared. Cell nuclei were stained with PI (red). Representative sections are shown. Abbreviations: EB, embryoid body; PI, propidium iodide.

 
The antigen detected by FORSE-1 antibody involves the oligosaccharide epitope called Lewis-X [20]. However, as the structures that carry the Lewis-X epitope may modify antibody binding, we wanted to confirm the authenticity of the epitope by an independent reagent. We used the monoclonal antibody TEC-01 developed by Draber and Pokorna [21] that is specific for the Lewis-X epitope (identical to the marker of pluripotent cells often referred to as SSEA-1 [22]). The specificity of FORSE-1 and TEC-01 antibodies was investigated on serial sections made from EBs produced by 10-day differentiation of normal and p27–/– mESCs. As demonstrated in Figure 4BGo, identical structures, both cells and extracellular matrix, were recognized by FORSE-1 and TEC-01 antibodies. Therefore, in the below text we refer to these structures as Lewis-X–positive.

Lewis-X–Positive Populations in Normal and p27-Deficient EBs Are Different from Each Other
To answer the question of whether increased proliferation within the Lewis-X–positive cell population was involved in establishing the observed phenotype, EBs at days 4, 5, 6, and 10 of their development were trypsinized and the proportion of Lewis-X–positive cells was determined by immunocytochemical staining of coverslip-immobilized cells. As demonstrated in Figure 5AGo, the number of Lewis-X–positive cells gradually decreased in both normal and p27-deficient EBs down to 3.5% at day 10 of differentiation. Interestingly, although the general trend to decrease was common to EBs of both genotypes, a slightly higher proportion of Lewis-X–positive cells was observable in p27-deficient EBs between days 4 and 6 of their development. This tendency was statistically significant at day 6 and suggests that higher proliferation of Lewis-X–positive cells may contribute to the formation of the cavities in p27-deficient EBs.



View larger version (29K):
[in this window]
[in a new window]
  		Figure 5. Properties of Lewis-X–positive cells in normal and p27-deficient EBs. (A): Normal and p27-deficient EBs were trypsinized at days 4, 5, 6, and 10 (D4, D5, D6, D10), adhered on microscopic cov-erslips, and stained with antibody against Lewis-X. A total of 1,000 cells at two independent regions of the coverslip was counted, and the proportion of Lewis-X–positive cells was determined. The graph represents means and standard deviations obtained from two independent experiments. Statistically significant difference between normal and p27-deficient cells is indicated by an asterisk (Student’s t-test, p < .05). (B): Normal and p27-deficient EBs were cultured until day 10, trypsinized, and stained with antibody against Lewis-X. Cell suspension was subjected to flow cytometric analysis, and Lewis-X–positive cells (region R2 in B) were sorted out. (C): The amounts of p27 in Lewis-X–positive (inside R2 region in B) and Lewis X–negative cells (outside R2 region in B) were analyzed by Western blotting. (D): The levels of p27, Oct-3/4, N-cadherin, E-cadherin, and PARP were determined in sorted Lewis-X–positive cells (region R2 in B) by Western blotting. This experiment was repeated twice, and, except for N-cadherin, the results of only one experiment are presented. Abbreviations: EB, embryoid body; PARP, poly (ADP-ribose) polymerase.

 
Still, the observed changes in the Lewis-X–positive population also suggest that qualitative differences may exist between the cluster- and cavity-forming Lewis-X–positive cells. To address this issue, Lewis-X–positive cells were sorted out using flow cytometry from 10-day-old EBs (Fig. 5BGo) and then were analyzed for the expression of several markers by Western blotting. When compared with Lewis-X–negative cells, in normal EBs Lewis-X–positive cells contain much higher amounts of p27 (Fig. 5CGo) that may make Lewis-X–positive cells more vulnerable to a loss of p27. Within normal EBs, Lewis-X–positive cells, but not Lewis-X–negative cells, express Oct-3/4 (not shown). In the absence of p27, the level of Oct-3/4 is lowered (Fig. 4DGo). Also, the levels of lineage-specific cadherins are slightly changed, with neural N-cadherin being upregulated and epithelial E-cadherin being downregulated (Fig. 4DGo). The level of nuclear protein poly(ADP-ribose) polymerase (PARP) that was used as a loading control has remained unchanged in p27-deficient Lewis-X–positive cells. Taken together, these data strongly suggest that the Lewis-X–positive populations in normal and p27-deficient EBs are not identical. The cluster-forming population (most abundant in normal EBs) that has a high level of Oct-3/4 (approximately one fourth of the level typical for undifferentiated mESCs, not shown) most likely represents remnants of undifferentiated ESCs, whereas the cavity-forming population (most abundant in p27-deficient EBs) seems to have a neural phenotype and thus may represent neural progenitor/ stem cells.

Nestin Is Expressed in the Cavities of p27-Deficient EBs
Lewis-X was previously found to be associated with certain regions of the developing nervous system [23, 24], and it was also identified as a marker of adult neural stem cells [25]. To further address whether Lewis-X–positive cavity-forming cells could represent neural progenitors, the expression of nestin was determined in the set of p27-deficient 10-day-old EBs. Nestin-positive cells were clearly found in the cavities of p27-deficient EBs (Fig. 6AGo), again suggesting Lewis-X–positive cells that are associated with cavities to be neural progenitors/stem cells.



View larger version (44K):
[in this window]
[in a new window]
  		Figure 6. Neural differentiation of normal and p27-deficient EBs. (A): EBs that were obtained by 10-day-long differentiation of p27-deficient mESCs were fixed, sectioned, and immunohistochemi-cally stained by antibody against nestin (green) and PI (red). (B, C): Mouse E10.5 embryos were fixed and coronal cryosections of embryonal brain were stained with Lewis-X–specific and nestin-specific antibody, respectively (red). Cell nuclei were counterstained with Hoechst (blue). The pattern of Lewis-X (B) and nestin (C) expression in ventral cerebral cortex is shown. Nestin-positive processes coming from ventricular area are indicated by arrows. (D): Normal and p27-deficient mESCs were differentiated according to protocol (for details, see Materials and Methods). Normal and p27-deficient mESCs (D0) and EBs were harvested at day 8 (D8), day 15 (D15), and day 25 (D25) of differentiation, and the expression of neural markers N-CAM, GAP-43, and GFAP was analyzed by Western blotting (E). Abbreviations: EB, embryoid body; mESC, mouse embryonic stem cell; PARP, poly (ADP-ribose) polymerase; PI, propidium iodide.

 
To identify cells that could represent in vivo analogs of nestin/Lewis-X–positive cells in the cavities of p27-deficient EBs, serial sections were made from the brain of E10.5 mouse embryos and were analyzed for the expression of Lewis-X and nestin. Lewis-X expression was localized to the surface of ventricles in certain parts of the embryonic brain (cerebral cortex, caudal part of dorsal telencephalon, ventral lining of fourth ventricle, and third ventricle lining). Although the expression of nestin was more widespread throughout the brain and the regions of Lewis-X and nestin expression were not completely overlapped, we clearly observed the regions where double-positive cells were residing. As demonstrated on coronal sections of ventral cerebral cortex (Fig. 6BGo), Lewis-X staining was highly polarized toward the ventricle and resembled the staining of the EB cavities (Fig. 4Go). In the corresponding region of the brain, we detected nestin-positive processes coming from the ventricular zone (Fig. 6CGo). These processes most likely belong to the radial glial cells that have their nuclei in the Lewis-X–positive zone [26] and have been shown previously to be positive for nestin [27]. Although similar processes positive for nestin are not developed in the EBs, presumably because the precise spatial organization of the neural tube is lost in the EBs, we hypothesize that the Lewis-X–positive cavities in EBs may represent in vitro analogs of embryonic ventricles.

Neural Differentiation of p27-Deficient ESCs Is Not Affected
This led us to hypothesize that alterations occurring to Lewis-X–positive cells in p27-deficient EBs may be reflected by some abnormalities in their neurogenic potential. To address this issue, mESCs of both genotypes were allowed to differentiate under the conditions that promote neural differentiation (Fig. 6DGo). Resulting cells were harvested at days 8, 15, and 25 of differentiation and were Western blotted for the expression of the following neural markers: N-CAM (neuron/glia-specific), GAP-43 (neuron/ glia-specific), and GFAP (astrocyte-specific). As demonstrated in Figure 6EGo, p27-deficient mESCs showed no apparent abnormality in the expression of any of these neural markers compared with their normal counterparts. In other words, despite the molecular characteristics and organization of Lewis-X–positive cells in EBs produced from p27-deficient mESCs, neurogenic potential of p27-deficient mESCs does not seem to be affected.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Conclusion
 References
 
We have shown previously using p27-deficient mESCs that p27 protects differentiating mESCs from apoptosis when they are driven into an endodermal lineage in monolayer culture [14]. In this study, the same normal and p27-deficient mESCs were allowed to form three-dimensional EBs to further investigate the role of p27 in early embryogenesis. Using this differentiation system, we revealed at least three phenomena linked to the absence of p27: (a) EBs were larger, (b) most EBs failed to express the endodermal marker cytokeratin endo-A, and (c) EBs contained an expanded population of Lewis-X–positive cells that often gave rise to cavities.

It is of note that high expression of TROMA-I is typical for the cells of the placenta [28], the development of which is strictly dependent on the cooperative activity of p27 and p57 [29]. In contrast to the cells of the placenta, TROMA-I–positive cells derived from mESCs do not express p57. This fact well explains the crucial role of p27 observed in this study and in our study published before [14] and justifies a major importance of p27 for the differentiation and/or survival of mESC-derived TROMA-I–positive cells.

Increased proliferation of p27-deficient cells is driven by deregulated activities of CDK2 and CDK4 [2, 4, 10]. Surprisingly, we were not able to detect any abnormality in downregulation of CDKs in p27-deficient EBs compared with their normal counterparts. Thus, at least most cells in EBs use a p27-independent mechanism to inhibit their CDKs. Lowering the amounts of activating cyclins is obviously the most likely candidate. However, it is important to realize that our data on cell cycle regulators cannot exclude the existence of a small population of cells that use p27 for regulation of their CDK activities. It is well established that cells of different lineages may differ in their requirements for p27. Recently, proliferation of specific stem and/or progenitor cells of living mice was shown to be enhanced in the absence of p27. These cells include, for example, progenitors originating from bone marrow stem cells [30], transit-amplifying cells during adult neurogenesis [31], and progenitors of oligodendrocytes [32, 33]. It is typical for advanced EBs to contain cells of various developmental lineages, some of which are similar or even identical to the progenitors listed above (reviewed in [17]). Therefore, we propose that it is the enhancement of a pool of specific progenitor cells that underlies the overgrowth of p27-deficient EBs. Due to the minute numbers and temporally restricted occurrence of such cells, they do not significantly alter CDK activities assayed in whole EBs.

To define the progenitor population that is responsible for increased size of p27–/– EBs, we used an immunohistochemical approach to search for cell lineages expanded in p27-deficient EBs. Such a strategy leads to the identification of a cell population positive for Lewis-X that has altered distribution and transiently increased abundance in p27-deficient EBs. Lewis-X is a developmentally regulated carbohydrate structure present on proteins and lipids of cell membranes and extracellular matrix [22]. Whereas in normal EBs, Lewis-X–positive cells are restricted mainly to small compact islets ([34], this study), in p27-deficient EBs, Lewis-X epitopes are present primarily on a population of cells surrounding the cavities and also on the cellular debris and extra-cellular matrix, which fill these cavities. It is of note that cavities not containing cells and/or material positive for Lewis-X epitope are normally present in EBs irrespective of their p27 genotype.

Lewis-X structure is associated with cells of early embryo; the epitope is first detectable at the late eight-cell stage and is highly expressed in morulae. After implantation, Lewis-X is detectable on cells of the embryonic ectoderm, visceral endoderm, and giant cells of trophoblast. As embryogenesis proceeds, embryonic ectoderm gradually loses Lewis-X antigen, and on day 8 of embryogenesis, the only portion of embryonic ectoderm that is positive for Lewis-X is the neuroectoderm surface. Primordial germ cells are also strongly positive for Lewis-X [35]. High expression of Lewis-X is typical for teratocarcinoma-derived pluripotent cells [22], and Lewis-X was recently identified as a good marker of adult neural stem cells [25]. In this study, to uncover the identity of the cell population that is positive for Lewis-X in EBs, the cells that express Lewis-X were sorted and analyzed for the expression of lineage-specific cadherins and Oct-3/4. Importantly, all three markers were expressed by Lewis-X–positive cells of 10-day-old EBs, suggesting that Lewis-X–positive cells may represent two distinct cell populations that are morphologically manifested in EBs as clusters and cavities. The cluster-forming cells, which express Oct-3/4, are most likely remnants of ESCs that were previously found to be present in EBs until day 15 of their differentiation [36]. In contrast, higher levels of N-cadherin combined with the expression of nestin suggest that the Lewis-X–positive cells that give rise to cavities are neural progenitors. Notably, we have found a similar distribution of Lewis-X and nestin also in vivo in the ventricular lining of embryonal mouse brain.

Obviously, Lewis-X–positive cells show cluster and cavity distribution not only in p27-deficient EBs but also in EBs produced by normal ESCs (Fig. 4AGo). However, whereas in normal EBs the neural progenitors represent only a minute part of the total Lewis-X–positive pool, they are highly enriched in p27-deficient EBs. Although the number of Lewis-X–positive cells increases only slightly under p27-deficient conditions and therefore we cannot completely exclude some unidentified cell fate–determining function of p27, here we suggest that p27 is a negative regulator of proliferation of Lewis-X–positive neural precursors but not of Lewis-X–positive ESCs in the EB. This suggestion seems to be in a good agreement with the previously published findings that p27-deficient mESCs have unaltered proliferation [14], whereas the p27-deficient mice show an increased proliferation of neural progenitors from the subventricular zone and increased formation of neurospheres [31]. Importantly, despite the increased neurogenic capacity of p27-deficient mice, no alterations in their ability to generate neurons and glia were detected [31]. Our experiments using p27-deficient EBs thus support the conclusion that p27 regulates the size of the neural progenitor pool without affecting the development of neurons or glia themselves (Fig. 6EGo).


   CONCLUSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Conclusion
References
 
Together, the experiments included in this study recognize p27 as a cell type–specific regulator of both proliferation and differentiation/survival of mESCs differentiating in culture. In some progenitor cell populations, p27 may serve to prevent uncontrolled growth, as for example of neural progenitors positive for Lewis-X epitope, whereas in other cell populations, such as TROMA-I–positive, p27 is necessary for correct differentiation and/or survival. From a general point of view, our study suggests that the emerging concept of a dual role for p27 also applies to ESCs and their progeny.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Conclusion
 References
 
This research was supported by the Academy of Sciences of the Czech Republic (KJB5039301 and AV 0Z5039906), by the Grant Agency of the Czech Republic (204/01/0905, 524/03/0766, and 524/03/P171), and by the Ministry of Education, Youth, and Sports of the Czech Republic (MSM 432100001 and 1M0021620803). We are very grateful to Dr. Jirí Lukás and Dr. Petr Dráber for providing us with antibodies, to Dr. Josef Bryja and Dr. Ivo Überall for their help with statistical evaluation and image analysis, and to Iveta Nevrivá for excellent technical assistance.


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Conclusion
 References
 

  1. Sherr CJ, Roberts JM. Inhibitors of mammalian G1 cyclin-dependent kinases. Genes Dev 1995;9:1149–1163.[Free Full Text]

  2. Fero ML, Rivkin M, Tasch M et al. A syndrome of multiorgan hyperplasia with features of gigantism, tumorigenesis, and female sterility in p27Kip1-deficient mice. Cell 1996;85:733–744.[CrossRef][Medline]

  3. Kiyokawa H, Kineman RD, Manova-Todorova KO et al. Enhanced growth of mice lacking the cyclin-dependent kinase inhibitor function of p27Kip1. Cell 1996;85:721–732.[CrossRef][Medline]

  4. Nakayama K, Ishida N, Shirane M et al. Mice lacking p27Kip1 display increased body size, multiple organ hyperplasia, retinal dysplasia and pituitary tumors. Cell 1996;85:707–720.[CrossRef][Medline]

  5. Tong W, Kiyokawa H, Soos TJ et al. The absence of p27Kip1, an inhibitor of G1 cyclin-dependent kinases, uncouples differentiation and growth arrest during the granulosa->luteal transition. Cell Growth Differ 1998;9:787–794.[Abstract]

  6. de Koning JP, Soede-Bobok AA, Ward AC et al. STAT3-mediated differentiation and survival and of myeloid cells in response to granulocyte colony-stimulating factor: role for the cyclin-dependent kinase inhibitor p27(Kip1). Oncogene 2000;19:3290–3298.[CrossRef][Medline]

  7. Muraoka RS, Lenferink AEG, Simpson J et al. Cyclin-dependent kinase inhibitor p27Kip1 is required for mouse mammary gland morphogenesis and function. J Cell Biol 2001;153:917–931.[Abstract/Free Full Text]

  8. Lin J, Della-Fera A, Li C et al. P27 knockout mice: reduced myostatin in muscle and altered adipogenesis. Biochem Biophys Res Commun 2003;300:938–942.[CrossRef][Medline]

  9. McAllister SS, Becker-Hapak M, Pintucci G et al. Novel p27kip1 C-terminal scatter domain mediates Rac-dependent cell migration independent of cell cycle arrest functions. Mol Cell Biol 2003;23:216–228.[Abstract/Free Full Text]

  10. Bryja V, Pachernik J, Faldíková L et al. A role of p27Kip1 in maintaining the levels of D-type cyclins in vivo. Biochim Biophys Acta 2004;1691:105–116.[Medline]

  11. Savatier P, Lapillonne H, van Grunsven LA et al. Withdrawal of differentiation inhibitory activity/leukemia inhibitory factor up-regulates D-type cyclins and cyclin-dependent kinase inhibitors in mouse embryonic stem cells. Oncogene 1995;12:309–322.

  12. Stead E, White J, Faast R et al. Pluripotent cell division cycles are driven by ectopic Cdk2, cyclin A/E and E2F activities. Oncogene 2002;21:8320–8333.[CrossRef][Medline]

  13. Faast R, White J, Cartwright P et al. Cdk6-cyclin D3 activity in murine ES cells is resistant to inhibition by p16INK4a. Oncogene 2004;23:491–502.[CrossRef][Medline]

  14. Bryja V, Pacherník J, Soucek K et al. Increased apoptosis in differentiating p27-deficient embryonic stem cells. Cell Mol Life Sci 2004;61:1384–1400.[CrossRef][Medline]

  15. Hogan B, Beddington R, Costantini F et al. Manipulating the Mouse Embryo: A Laboratory Manual. New York: Cold Spring Harbor Laboratory Press, 1994.

  16. Coucouvanis E, Martin GR. Signals for death and survival: a two step mechanism for cavitation in the vertebrate embryo. Cell 1995;83:279–287.[CrossRef][Medline]

  17. Desbaillets I, Ziegler U, Groscurth P et al. Embryoid bodies: an in vitro model of mouse embryogenesis. Exp Physiol 2000;85:645–651.[Abstract]

  18. Bain G, Kitchens D, Yao M et al. Embryonic stem cells express neuronal properties in vitro. Dev Biol 1995;168:342–357.[CrossRef][Medline]

  19. Tole S, Kaprielian Z, Ou SK et al. FORSE-1: a positionally regulated epitope in the developing rat central nervous system. J Neurosci 1995;15:957–969.[Abstract]

  20. Allendoerfer KL, Magnani JL, Patterson PH. FORSE-1, an antibody that labels regionally restricted subpopulations of progenitor cells in the embryonic central nervous system, recognizes the Le(x) carbohydrate on a proteoglycan and two glycolipid antigens. Mol Cell Neurosci 1995;6:381–395.[CrossRef][Medline]

  21. Draber P, Pokorna Z. Differentiation antigens of mouse teratocarcinoma stem cells defined by monoclonal antibodies. Cell Differ 1984;15:109–113.[CrossRef][Medline]

  22. Solter D, Knowles BB. Monoclonal antibody defining a stage-specific mouse embryonic antigen (SSEA-1). Proc Natl Acad Sci U S A 1978;75:5565–5569.[Abstract/Free Full Text]

  23. Ashwell KWS, Mai JK. Developmental expression of the CD15 epitope in the hippocampus of the mouse. Cell Tissue Res 1997;289:17–23.[CrossRef][Medline]

  24. Ashwell KWS, Mai JK. Developmental expression of the CD15-epitope in the brainstem and spinal cord of the mouse. Anat Embryol (Berl) 1997;196:13–25.[CrossRef][Medline]

  25. Capela A, Temple S. LeX/ssea-1 is expressed by adult mouse CNS stem cells, identifying them as nonependymal. Neuron 2002;35:865–875.[CrossRef][Medline]

  26. Noctor SC, Flint AC, Weissman TA et al. Dividing precursor cells of the embryonic cortical ventricular zone have morphological and molecular characteristics of radial glia. J Neurosci 2002;22:3161–3173.[Abstract/Free Full Text]

  27. Lendahl U, Zimmerman LB, McKay RD. CNS stem cells express a new class of intermediate filament protein. Cell 1990;60:585–595.[CrossRef][Medline]

  28. Duprey P, Morello D, Vasseur M et al. Expression of the cytokeratin endo A gene during early mouse embryogenesis. Proc Natl Acad Sci U S A 1985;82:8535–8539.[Abstract/Free Full Text]

  29. Zhang P, Wong C, DePinho RA et al. Cooperation between the Cdk inhibitors p27KIP1 and p57KIP2 in the control of tissue growth and development. Genes Dev 1998;12:3162–3167.[Abstract/Free Full Text]

  30. Cheng T, Rodrigues N, Dombkowski D et al. Stem cell repopulation efficiency but not pool size is governed by p27kip1. Nat Med 2000;6:1235–1240.[CrossRef][Medline]

  31. Doetsch F, Verdugo JM-G, Caille I et al. Lack of the cell cycle inhibitor p27Kip1 results in selective increase of transit-amplifying cells for adult neurogenesis. J Neurosci 2002;22:2255–2264.[Abstract/Free Full Text]

  32. Casaccia-Bonnefil P, Hardy RJ, Teng KK et al. Loss of p27Kip1 function results in increased proliferative capacity of oligodendrocyte progenitors but unaltered timing of differentiation. Development 1999;126:4027–4037.[Abstract]

  33. Casaccia-Bonnefil P, Tikoo R, Kiyokawa H et al. Oligodendrocyte precursor differentiation is perturbed in the absence of the cyclin-dependent kinase inhibitor p27Kip1. Genes Dev 1997;11:2335–2346.[Abstract/Free Full Text]

  34. Esner M, Pachernik J, Hampl A et al. Targeted disruption of fibroblast growth factor receptor-1 blocks maturation of visceral endoderm and cavitation in mouse embryoid bodies. Int J Dev Biol 2002;46:817–825.[Medline]

  35. Fox NW, Damjanov I, Martinez-Hernandez A et al. Immunohistochemical localization of early embryonic antigen (SSEA-1) in postimplantation mouse embryos and fetal and adult tissues. Dev Biol 1981;83:391–398.[CrossRef][Medline]

  36. Jackson M, Krassowska A, Gilbert N et al. Severe global DNA hypomethylation blocks differentiation and induces histone hyperacetylation in embryonic stem cells. Mol Cell Biol 2004;24:8862–8871.[Abstract/Free Full Text]

Received August 3, 2004; accepted for publication March 8, 2005.




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
2004-0174v1
23/7/965    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Reprints/Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bryja, V.
Right arrow Articles by Hampl, A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bryja, V.
Right arrow Articles by Hampl, A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
STEM CELLS THE ONCOLOGIST CME ALPHAMED PRESS JOURNALS
http://www.peprotech.com/