Download Formation of presumptive embryoid bodies in undifferentiated

 Formation of presumptive embryoid bodies in undifferentiated gonadal cell cultures from the embryonic chick due to the presence of primordial germ cells Honors Thesis Presented to the College of Agriculture and Life Sciences, Department of Animal Sciences of Cornell University in Partial Fulfillment of the Requirements for the Research Honors Program by Adrienne M. Chan January 2010 Dr. John E. Parks ABSTRACT Primordial germ cells (PGCs) are pluripotent stem cells and exist as embryonic precursors to germ cells in vivo. PGCs have been shown to differentiate into embryonic germ (EG) cells which can aggregate to form embryoid bodies (EBs) in vitro. EGs within EBs are capable of differentiating into a variety of cell types; thus EBs can be useful for generating large numbers of stem cells for regenerative medicine. The objectives of this study were to harvest and culture primordial germ cells from indifferent gonads of the embryonic domestic chick and assess factors affecting survival, proliferation, and development of gonadal cells (somatic and germ) in vitro. PGCs were identified based on size and staining with pluripotent stem cell markers. Formation of cell aggregates, presumptive EB precursors, was observed throughout all gonadal cultures in the presence of PGCs. Gonadal cultures treated with busulfan to eliminate PGCs had only limited aggregate formation. Cultures supplemented with fibroblast growth factor showed increased aggregate formation. Co‐culture with embryonic testes or ovaries seemed to influence somatic cells surrounding aggregates. PGC‐derived EGs were identified immunocytochemically within aggregates using pluripotent stem cell and germ cell markers (anti‐SSEA1, anti‐SSEA4, and anti‐EMA1). In vitro culture conditions for dissociated embryonic gonad cells in this study may provide a basis for expanding embryonic germ cells and directing their development for germ stem cell development, in vitro gamete production, and other stem cell applications. ACKNOWLEDGEMENTS
First and foremost I offer my sincerest gratitude to my supervisor, Dr. John Parks, who has supported me throughout my thesis with his patience and knowledge whilst allowing me the room to develop my own project. I attribute the achievement of Honors and Distinction in Research to his encouragement and effort. This thesis could not have been written without his exceptional teaching and mentorship. One simply could not wish for a more supportive supervisor. In the laboratory I have been aided by Kim Grant, Mike Simkin, and James Giles. TABLE OF CONTENTS Introduction, 1‐2 Review of Literature, 3‐10 Materials and Methods, 11‐21 Results, 22‐39 Discussion, 40‐42 Literature Cited, 43‐44 INTRODUCTION Primordial germ cells (PGCs) are the embryonic precursors to germ cells (oogonia in the ovaries or spermatagonia in the testes). Because PGCs are pluripotent stem cells, their study, isolation and transfer have many important implications for a variety of biotechnological and biomedical fields. Pluripotency of PGCs means that they have the capacity to develop into all cell types except for extraembryonic tissue. In that respect, PGCs are similar in potential to embryonic stem cells (ES cells) which are pluripotent stem cells derived from the inner cell mass of the blastocyst, an early‐stage embryo. Both isolated ES cells and PGCs have been shown to differentiate into embryonic germ (EG) cells which can aggregate to form embryoid bodies (EBs) in vitro. EBs have been described as three‐
dimensional cystic bodies that comprise the three germ lineages, making them a useful model system to study early embryonic development (1). Immunohistochemical analysis using antibodies against actin, alpha‐1‐fectoprotein, and S100 Dako which are specific for tissues from all three germ layers ‐skeletal muscle, endoderm, and ectoderm respectively‐ show that EG cells within EBs are capable of differentiating into a variety of cell types in vitro (2,3). Thus, EBs can be a useful system for generating large numbers of cells for regenerative medicine. In particular, ES cell‐derived EGs supported within and by EBs have been shown to form functional germ cells in vitro (4,5,6). This study focuses on the differentiation patterns of PGCs isolated from chick embryos when exposed to different culture conditions in vitro. The objective of the study was to determine the fate of undifferentiated PGCs when cultured in medium conditioned by co‐culture with male or female gonads of different developmental ages. Specifically, we asked whether PGCs can be directed to differentiate in vitro into male versus female germ cells (oogonia or spermatogonia). Cell aggregates, which may be the early stages of EB formation, were collected from an initial culture of dissociated day 6 embryonic gonads. Aggregates of PGCs were co‐cultured with intact male or female gonads (testes or ovaries) of 1
different embryonic ages and monitored for differentiation patterns as a function of their co‐culture environment. PGC‐derived EGs within the presumptive primitive EBs were stained with pluripotent stem cell and germ cell markers (anti‐SSEA1, anti‐SSEA4, anti‐EMA1, and anti‐vasa homolog) to confirm their presence in the presumptive EBs. 2
REVIEW OF LITERATURE As one of the two known embryonic sources of pluripotent stem cells, PGCs are a popular subject of stem cell research (the other being the inner cell mass of preimplantation embryos from which ES cells are derived). Pluripotent EG cells derived from PGCs form gametes which impart totipotency (the ability to divide and differentiate into all cell types including extraembryonic tissue) at syngamy and perpetuate the germ line and thus the species. Origin of PGCs Embryonic precursor cells for PGCs originate in the epiblast proximal to extraembryonic ectoderm. The extraembryonic tissues, visceral endoderm (VE) and extraembryonic ectoderm (ExE) are known to be important for the induction of PGCs in mice. The VE regulates the formation of a normal number of PGC precursors, while the ExE regulates their appropriate localization in the epiblast during early development (7). The precursor cells are induced to become PGCs by initiation of a signal transduction cascade by bone morphogenetic proteins (BMP). BMPs are members of a type B growth factor family and function as homodimers or heterodimers to signal through heteromeric receptor complexes and downstream transforming growth factors (8). Specification of cells to become PGCs is associated with fragilis expression in epiblast tissue; after precursor cells become competent to form PGCs, they differentiate into 40‐50 PGCs (9). Migration and proliferation of PGCs PGCs are first identified at the base of the amniotic fold and allantoic bud where they were first regionalized by the ExE. Expression of Blimp 1, a transcriptional repressor that represses genes required for somatic cell differentiation, is associated with the proliferation and migration of PGCs into the 3
mesoderm of the primitive streak. Via morphogenetic movements during gastrulation, active migration by amoeboid‐like movement, and directed movement by cell adhesion and chemo‐attraction by the genital ridge, PGCs migrate from the mesoderm through the endoderm of the yolk sac and hindgut, through the dorsal mesentery, and into the genital ridges. Avian PGC migration also includes transport within the circulation. In the domestic chick, PGCs circulate in the bloodstream during embryonic stage 18 (day 3 of incubation according to Hamburger‐Hamilton stages (10) and migrate into the genital ridge by stage 28 (day 6 of incubation) where they continue to proliferate to several hundred to a few thousand PGCs. Proliferation of PGCs is mediated by elevation in cAMP, pituitary adenyl cyclase activating proteins, and retinoic acid (RA). Proliferation of PGCs ceases shortly after migration to the genital ridges, and PGCs differentiate to gonocytes in the testis or primary oocytes in the ovary (11). Appearance and identification of PGCs PGCs are identified by their relatively large size (18‐20 µm diameter compared to 8‐12 µm diameter of typical somatic cells). The first marker used to identify PGCs in vivo and in vitro was tissue‐
nonspecific alkaline phosphatase. More recently, cell‐surface markers such as SSEA1 and 4, 4C9, EMA1, and ckit were used to identify mouse PGCs as well as intracellular proteins including mouse vasa homolog, germ cell nuclear antigen, and Oct4. Immunoaffinity purification, density gradient centrifugation (Percoll), magnetic cell sorting, and fluorescent‐activated cell sorting have been used to partially purify isolated PGCs (12). Derivation of pluripotent EG cells from PGCs PGCs in the chick embryo can be isolated at stage 16‐17 from the bloodstream or stage 28‐29 from the genital ridge. Park and Han (2) cultured isolated gonadal PGCs (gPGCs) in EG cell culture medium which consisted of DMEM medium (Gibco) supplemented with fetal bovine serum, chicken 4
serum, sodium pyruvate, L‐glutamine, 2‐mercaptoethanol, streptomycin, penicillin, human stem cell factor, leukemia inhibitory factor, bovine basic fibroblast growth factor, interleukin‐11, and insulin‐like growth factor‐I. Under the culture conditions of the study, EG cell colonies were derived. EG cells where identified by PAS staining, anti‐SSEA‐1 antibody screening, proliferation assays using thymidine analog bromodeoxyuridine (BrdU) which incorporates into S‐phase nuclei, and alkaline phosphatase activity. Expression of the SSEA‐1 epitope, a criterion for distinguishing pluripotent stem cells, was observed in chicken EG cell colonies. EG cells stained weakly for alkaline phosphatase activitiy and PAS reaction which stains for glycogen in cytoplasm, though high levels of AP activity associated with ES and EG cultures are observed in other studies including those using human PGCs (13). Continuous proliferation of EG cells after several passages on mitotically active chicken embryonic fibroblasts (CEF) was confirmed using BrdU. The study defined and established an effective EG cell culture medium with important growth factors for the survival and proliferation of gPGCs including SCF, LIF, bFGF, IL‐11, and IGF‐I. Without IL‐11 and IGF‐I, colonization of chicken EG cells was not observed (2). Formation and differentiation of EBs from PGCs EBs are induced to form from PGC and ES cell cultures using complete growth medium lacking LIF and embryonic fibroblasts. These conditions induce differentiation, permit suspension (low‐
attachment) cultures, and cause cells to form aggregates (EG cell colonies). EG cell colonies are described as uniformly round, multi‐layered, and well‐delineated on the order of 50‐150 µm in diameter (2). These cell aggregates derived from PGC or ES cell cultures can be dissociated using standard trypsin dissociation procedures to establish single‐cell suspensions of ES cells (<20 µm in size). The single‐cell suspensions can be used to produce hanging drop cultures which can then be transferred and resuspended in 1% agar treated low adhesive plates with medium lacking LIF (2). Within 4 days of culture with 24h to 48h medium changes, formation of EBs are observed. EBs from cultured chicken 5
PGCs have been characterized with PAS, AKP, c‐kit, and SSEA‐1 staining in the lab of Wu and Zhang (9). Positive staining of PAS, AKP, c‐kit, and SSEA‐1 in PGC‐derived EBs indicated the pluripotent cells ability to undergo self‐renewal and express specific markers of stem cells. EBs plated into 96‐well plates were allowed to attach and differentiate. Positive immunocytochemical staining of differentated EBs for expression of neuron specific enolase and keratin indicated that EBs could effectively differentiate into different germ lines including epithelial, neuronal, and skeletal myoblast cells. Park and Han show that EBs derived from chick EG cells strongly stained for muscle‐specific actin, endoderm‐specific alpha‐1‐
fetoprotein, and ectoderm‐specific S100. Furthermore, rhythmic beating cells at beating rates of 20 to 100 times per min were observed in some EBs (3). EB morphology and diffusive transport of inductive biochemicals for support of stem cell (PGC‐ or ES cell‐
derived EG cell) differentiation Sachlos and Auguste (1) provided an excellent description of EB morphology and EB shell characterization through SEM images, immunofluorescence staining, and diffusion profiles. EBs were monitored for structural changes and differentiation of hES cells into all 3 germ layers. They report that day 1 EBs undergo remodeling (day 0 was designated as the day that undifferentiated hES cells were transferred to a culture well lacking LIF and bFGF), day 3 EBs have a synthesized fibrous extracellular matrix coating (ECM), day 7 EBs have a smoother surface topography because spaces between cells are filled with ECM, and day 14 EBs have indistinguishable cell boundaries due to increased ECM deposition. A day 7 EB has a dense shell of 20 µm thickness consisting of an ECM comprised of collagen type I, a squamous cellular layer with tight cell‐cell adhesions positive for E‐cadherin, and collagen type IV lining indicative of a basement membrane. Sachlos and Auguste warn of the decreased ability of diffusive transport due to the characteristics of the thick shell membrane. They found that permeabilizing the shell membrane with collagenase treatment or inhibiting the shell membrane formation with noggin, a 6
BMP inhibitor, restores diffusive transport ability. Despite the near non‐permeable shell membrane reported by Sachlos and Auguste, others have relied on EB diffusive transport of biochemicals to support and induce stem cell differentation. Formation and differentiation of EBs has been a strategy for stem cell differentiation. EBs sustain blood development, and by diffusive transport of inductive biochemicals, EBs may support stem cell differentiation. Germ cells have been derived from ES cells within EBs by imposing specific culture conditions. Oocyte‐like cells were derived from mouse ES cells after co‐culturing ES cell‐derived EBs with ovarian granulosa cells. Qing and Shi (14) demonstrated that the induced oocyte‐like cells from mouse ES cells expressed oocyte‐specific genes Figα, GDF‐9, and ZP1‐1 but not testis‐specific genes acrosin, Sex‐determining Region Y (SRY), haprin (a gene that encodes a haploid germ cell‐specific RING finger protein and is transcribed exclusively in testicular germ cells after meiotic division), and RET finger protein (Rfp) which was reported to be expressed in different stages of spermatogenesis (15) by RT‐PCR analysis. In their study, EBs were first formed from ES cells and were stimulated to cause PGC proliferation by adding retinoic acid (RA) to culture medium. EBs were then co‐cultured with granulosa cells or cultured in granulosa cell‐conditioned medium. After 4d culture, germ cell colonies within the EBs were identified by immunocytochemistry assays for expression of mouse vasa homolog (Mvh), a post‐migration PGC marker and SCP3, a meiosis‐specific marker. Germ cell colonies were then checked for expression of oocyte‐specific genes and testis specific genes. Cells within EBs cultured in granulosa conditioned medium did not differentiate into cells that expressed oocyte‐specific markers, whereas cells within EBs co‐cultured with granulosa cells did. The results demonstrated that granulosa cells were effective in inducing the differentiation of ES cell‐derived PGCs within EBs into oocyte‐like cells through direct cell‐to‐cell contacts (14). Hubner and Fuhrmann demonstrated that oogonia derived from mouse ES cells can enter meiosis, recruit adjacent cells to form follicle‐like structures, and later develop into blastocysts with fertilization. The study provides evidence of increased estradiol production of derived 7
follicle‐like structures, micrographs showing the derived follicle‐like structures’ similarity to early primary and secondary follicles, and increased expression of GDF‐9, aromatase, Cyp17, and StAR. Expression of ZP1‐3, Figα, Oct4, and SCP3, characteristic of oocytes and germ cells undergoing meiosis, was detected. Blastocyst‐like structures derived from ES cells were morphologically similar to and expressed molecular markers similar to mouse preimplantation embryos (16). Male gametes can be derived from ES cells after EB formation as well. Geijsen and Horoschak demonstrated that EBs support maturation of PGCs into haploid male gametes. The derived germ cells during EB development showed expression of germ‐cell‐specific genes in the male gonad: Piwil2, Rnf17, Rnh2, Tdrd1, and Tex14. The male gametes were able to restore the somatic diploid chromosome complement when injected into oocytes, and blastocysts developed (6). EBs can support stem cell differentiation into a variety of cell types based on culture conditions. Wang and Jiao demonstrated that chick EGs can be directed to differentiate into neural cells using RA induction in vitro. EBs grown in suspension and subjected to RA induction form neurosphere colonies which are composed of neural progenitor cells. Neurons, astrocytes, and oligodendrocytes were evident on day 9 after RA induction in EB cultures (17). PGCs in transgenic research and endangered species conservation Experiments to generate germline chimeras by the transfer of PGCs have been successful within and between avian species, pigs, mice, and zebrafish. This technology has potential use in the preservation of threatened or endangered species given that adult stem cells (perhaps spermatogonial) can be isolated and induced to become PGCs in culture. Those stem cell‐derived PGCs containing the genome of the endangered species can then be transferred into a more common recipient host species that is at the embryonic stage of PGC migration. Blood samples, which contain circulating PGCs from stages 16‐17 (56h development) in the chick embryo according to Hamburger and Hamilton, can be 8
obtained from fertilized eggs via the forging of microsurgical techniques. Immunoaffinity purification, density gradient centrifugation (Percoll), magnetic cell sorting, and fluorescent‐activated cell sorting can be used to partially purify isolated PGCs from blood samples. Purified PGCs can then be transferred into the bloodstream of a host species at the same developmental stage, so that transferred PGCs can migrate to the genital ridges and be perpetuated by the host. Busulfan emulsions and blood extractions from the host are methods to diminish the host’s endogenous EGs. Prior to the PGC injection into the host species, the host may be subjected to a busulfan emulsion to destroy the host’s endogenous PGCs, or a minimal amount of blood containing a proportion of endogenous circulating PGCs may be drawn from the host and discarded. At stage 16‐17 in the chick embryo, only 40‐50 PGCs are circulating within the bloodstream; thus, drawing a few µls of blood and injecting several thousand purified PGCs from the species of interest into the host’s circulation will reduce the host’s endogenous germ cell propagation. Successful PGC transplants can lead to the salvation and production of fertile adult germ cells of donor species by a more common and affordable host species (18). Transfer of PGCs in avian species For avian species, PGCs derived from female chick embryos have been shown to preferentially migrate to the left genital ridge when transferred to either male or female recipient embryos (19). This observation suggests that female PGCs have some cell autonomous factor that directs migration to the left genital ridge in male and female recipients since the left gonad is the only functional gonad in the genetic female. The suggested advantage for this preferential migration to the left gonad is to enable a more efficient assembly of gametes in the female. Though female PGCs preferentially migrate to the left genital ridge in transplant experiments, female PGCs are not hardwired to become female gametes. PGCs derived from female embryos have the capacity to become male germ cells after they are transplanted into the testes of chickens. PGCs isolated from 2.5 day embryos (stage15) through blood 9
collection via micromanipulation were sexed by PCR. Germline chimeric chickens were generated by microinjecting 50‐200 pooled female donor PGCs into the bloodstream of male recipient embryos. Recipient male embryos were then incubated until hatching. W‐bearing sperm in semen of sexually mature male chimeric chickens were detected by PCR. The W chromosome is the female determining chromosome in avian species. W‐bearing spermatagonia, spermatocytes, and round spermatids were observed in the seminiferous tubules of male chimeras at 31%, 33%, and 28% of total like cells observed. However, W‐bearing elongating spermatids and spermatozoa were observed at a much lower frequency (8% and 0.2%) suggesting that female PGCs in male gonads were capable of completing the first and second meiotic divisions in the male environment, but were less capable of undergoing complete spermiogenesis (20). The implications of Tagami’s experiment offer a piece to the puzzle for understanding the differentiation processes of female PGCs. Female PGCs were affected in their differentiation process when exposed to the male embryonic environment in vivo. 10
MATERIALS AND METHODS I. Isolation and Dissociation of Chick Embryonic Gonads Egg source Fertilized eggs from White Leghorn hens were obtained from the Cornell University Poultry Biology Laboratory. Fertile eggs were acquired on day 6, 9 or 12 of incubation (stage 28‐29, 35‐36, and 38 embryos) according to the developmental stages described by Hamburger and Hamilton (10). Gonads were excised aseptically and PGCs were recovered on the same day. Recovery of embryonic gonads Eggs were positioned with their long axis vertical and large end up upon delivery so that embryos rotated to the top of the egg. Eggs were wiped thoroughly with 70% ethanol then transferred to a sterile laminar flow hood. Eggs were windowed by cutting a 20 mm diameter opening at the top of the egg using sterile scissors. The inner shell membrane was removed and the intact embryo was separated from its vasculature and extraembryonic membranes (allantois, chorion, amnion and yolk sac) using sterile Adson tissue forceps. Detached embryos were gently removed from windowed eggs using a sterile Fisher scoopula. Each embryo was pinned in a supine position on a Petri dish containing stained solid paraffin. If needed, embryos were gently swirled in sterile Dulbecco’s Phosphate Buffered Saline without calcium chloride and magnesium chloride (PBS, Sigma, D5652) to remove yolk that may have been carried over from the eggs. Pins were placed near the caudal and rostral ends of the embryo to secure the embryo in place. Using a 27 ½ or 30 gauge (25 mm) beveled needle, the abdominal wall and gastrointestinal tract were removed to reveal the underlying mesonephros and gonads. The mesonephroi were clearly identified in 6 day embryos as two large, elongated structures stippled red. Gonads were attached on the ventromedial surface of each mesonephros. Using the beveled needle, 11
connective tissue was gently separated both from between the gonad and the mesonephros and the area between the two gonads (Figure 1). Once near total detachment from connective tissue, the gonads were removed using jewelers forceps and placed in a sterile 1 ml microfuge tube containing PBS. The same procedure was implemented for dissection of embryonic gonads from day 9 and 12 embryos. Male and female gonads were easily distinguished from each other at this stage because males had two gonads of approximately equal size whereas females had a distinctly enlarged left gonad and a small, regressing right gonad. Male and female gonads, excluding the regressing right ovary in females, were collected separately and placed in separate wells of a polystyrene, flat bottom 24‐well culture plate with 16mm well diameter (Corning, 25830) with 0.5 ml StemPro medium lacking LIF (Gibco, 10565‐018). Two to four gonads of either male or female gonads were placed in each well. Gonadal dissociation Dissected gonads were transferred with a 1 ml micropipettor and sterile tip into a sterile 70 mm watch‐glass containing 1 ml 0.125% collagenase (C9407, Sigma‐Aldrich) which was then placed in a 37⁰C incubator (5% CO2 in air) for 5‐10 min with intermittent gentle pipetting using a sterile gel‐loading tip to help dissociate the tissue. Dissociation of tissues was monitored in a sterile culture hood with a dissecting microscope. When dissociation was near completion, the collagenase solution was diluted with PBS containing 2% bovine serum albumin (BSA, A9647100G, Sigma Aldrich) for subsequent immunohistochemical staining. Trypsin (0.25%, T0646, Sigma‐Aldrich) was also used to dissociate gonadal tissue but generally compromised the cells. Fluorescence microscopy of immunostained cells subjected to trypsin treatment showed fluorescent fragments but few intact cells. Apparently dilution of the trypsin solution with PBS but without inactivation, i.e. soybean trypsin inhibitor or DMEM/F12 with GlutaMax medium, did not stop enzymatic activity and cells were lysed. Thus, collagenase treatment was preferred because most cells were not damaged. 12
B
A
Figure 1: A) Day 6 embryonic gonads (red arrows) attached on the ventromedial surface of each mesonephros (yellow arrows). B) Excised day 6 embryonic gonad. Scale bar is 0.10 mm. 13
PGC and somatic cell counts Two 6d gonads were dissociated and suspended in 0.1 ml PBS in a microfuge tube. Using a 10 µl pipettor, 10 µl of the gonadal cell suspension was placed on a standard hemocytometer. Suspected PGCs and somatic cells were counted based on qualitative size differences (PGCs should be twice as large as somatic cells). Cells were counted in the 4 large corner squares (each 1/25 mm2) of the hemocytometer. For cells that overlapped a ruling, cells were counted if it overlapped the top or right ruling, and were not counted if they overlapped the bottom or left ruling. An average of 3 different cell counts was computed for each pair of 6 day gonads, and computations were adapted from an introductory biology laboratory methods guide by David Caprette, Rice University (21). II. Identification of PGCs Immunohistochemical staining After tissue dissociation, cells were aspirated using a 100 µl micropipette with a gel‐loading tip. Cells were distributed to 10 0.5 ml microfuge tubes with 0.5 ml PBS and centrifuged at 200 x g for 5 min. The supernatant was carefully removed without disturbing the cell pellet and discarded, leaving 0.1 ml with cell pellet at the bottom of each tube. The cell pellet was resuspended in the remaining 0.1 ml by gentle pipetting using a 20 µl micropipette. For immunohistochemical staining, 1:2, 1:10, and 1:50 dilutions (v/v) of SSEA‐1 (MC480, Developmental Studies Hybridoma Bank (DSHB), University of Iowa, Dept. of Biology, 27 µg/ml Ig), SSEA‐4 (MC81370, DSHB, 49 µg/ml Ig), and EMA‐1 (DSHB, 40 µg/ml Ig) were used as primary antibodies. Use of these monoclonal antibodies for identifying avian primordial germ cells was validated using domestic Japanese quail (22). Tubes were designated as follows with 50 µl of diluted primary antibody added to each tube: Tube 1 SSEA‐1 at 1:2 dilution (16 µl per 34 µl 2% BSA‐PBS) Tube 2 SSEA‐1 at 1:10 dilution (5 µl per 45 µl 2% BSA‐PBS) 14
Tube 3 SSEA‐1 at 1:50 dilution (1 µl per 49 µl 2% BSA‐PBS) Tube 4 SSEA‐4 at 1:2 dilution (16 µl per 34 µl 2% BSA‐PBS) Tube 5 SSEA‐4 at 1:10 dilution (5 µl per 45 µl 2% BSA‐PBS) Tube 6 SSEA‐4 at 1:50 dilution (1 µl per 49 µl 2% BSA‐PBS) Tube 7 EMA‐1 at 1:2 dilution (16 µl per 34 µl 2% BSA‐PBS) Tube 8 EMA‐1 at 1:10 dilution (5 µl per 45 µl 2% BSA‐PBS) Tube 9 EMA‐1 at 1:50 dilution (1 µl per 49 µl 2% BSA‐PBS) Tube 10 unlabelled cells (control) Cells were incubated with their respective primary antibody at 8°C for 20 min. Cells were washed twice (centrifuged at 200 x g for 5 min) with 2% BSA‐PBS. The supernatant was carefully removed without disturbing the cell pellet and discarded, leaving a residual volume of 0.1 ml after each wash. After the final wash, the cell pellet was resuspended in the residual 0.1 ml by gentle pipetting. A 1:4 dilution (10 µl per 40 µl 2% BSA‐PBS) of Alexa Fluor 488‐labelled goat anti‐chicken IgG antibody (MP11039, Invitrogen/Molecular Probes, 2mg/ml) was used as the secondary antibody. Anti‐
chicken secondary antibody was used based on a broader survey of pluripotent cell markers in which chicken antigen was used where available. We continued to use in our study because chicken PGCs stained specifically in dissociated testis cells with its use, although use of goat‐anti‐mouse secondary antibody is now preferred (22). Each tube received 50 µl diluted secondary antibody. Cells were incubated with the secondary antibody at 8⁰C for 15 min. Cells were washed twice with 2% BSA‐PBS. Fluorescence microscopy Final PBS supernatant was carefully removed, leaving 0.1 ml containing stained cells at the bottom of microfuge tubes. Cells were resuspended by gentle pipetting. Using a 20 µl micropipette, 8 µl of cell suspension was transferred onto microscope slide. A 22 mm2 cover slip (Corning Labware and Equipment) was placed on top of the drop, and the coverslip borders were sealed with clear nail polish 15
to prevent dessication and extend time for microscopic examination. Slides were examined using fluorescence microscopy (Nikon Eclipse, E600, B‐2E/C FITC filter, EX 46115‐495) immediately after immunostaining or the day after. If cells were examined the next day, cells were fixed for 3 min in 4% buffered paraformaldehyde, then washed twice with PBS. Cells were stored in 1 ml PBS at 8⁰C in the dark. Cells were first located using brightfield optics. PGCs were ~2 times larger in diameter than somatic cells; this is both an observed estimate and a documented difference in diameter from somatic cells. After PGCs were identified by size using brightfield optics, fluorescence optics were used to detect SSEA positive cells, stained with Alexa Fluor 488‐conjugated secondary antibody. SSEA positive cells fluoresce bright green (emission optimum 519 nm) against a dark background. III. Cell culture of PGCs using different media Cell culture of PGCs Dissection of gonads for the purpose of cell culture was completed using sterile technique; all instruments were autoclaved prior to use and immersed in 70% ethanol between use during a concomitant period. For a typical experiment, 8 gonads were dissociated with collagenase as described above for 10‐15 min or until intact tissue was no longer visible. After the final wash, the cell pellet in a residual 0.1 ml volume was resuspended by gentle pipetting. Cells were transferred as a 25 µl suspension to a 4‐well culture plate (Nunclon Surface, DK‐4000) containing 0.5 ml stem cell medium prepared from a StemPro hESC SFM kit (A10007‐01, Gibco)and pre‐warmed to 37°C per well. Cells were incubated in an atmosphere of 5% CO2 in air at 37°C. Medium was changed every 24 hr as follows: (1) culture plates were removed from the CO2 incubator and wiped with 70% ethanol before moving to culture hood, (2) attached cells were located in culture well using a dissecting microscope (Nikon,SMZ‐
U), (3) spent medium was removed with a 1000 µl pipettor while avoiding attached cells on the bottom 16
of wells, (4) 0.5 ml fresh medium pre‐warmed to 37°C was added to wells, and (5) culture plates were wiped with 70% ethanol and returned to the CO2 incubator. Effect of LIF and FGF on PGC cultures To each well of a 4‐well plate was added cells from dissociated 6 day gonads and 0.5 ml of complete StemPro culture medium consisting of DMEM/F12 with GlutaMax (1X) and 1.8% BSA with 1) 1X LIF supplement (10X StemPro hESC SFM Growth Supplement added as 0.1 ml/ml of medium) or 2) 1X LIF and FGF‐basic (8 ng/ml). Complete medium was prepared according to the StemPro hESC SFM manual (23). Cultures with each media were monitored daily for proliferation and structural development (cell aggregate formation). Counting of cell aggregates in cultures A transparency of a 10x10 square centimeter graph paper was placed beneath a cell culture plate and viewed under a dissecting microscope. Cell aggregates were counted within 4 random 1x1 mm2 areas per well. Cell aggregates were counted if they were within the 1x1 mm2 area and on the upper and left side of each square. An average was computed for every 4‐well culture (4 1‐mm2 areas x 4 wells for a total of 16 counts). Aggregates were counted on day 1 and 4 of incubation for all PGC cultures. IV. Cell cultures with busulfan treatment for inhibition of cell aggregate formation Cell cultures were treated with busulfan to determine whether the presence of PGCs in the initial culture was required to induce cell aggregate formation. Busulfan is known to first destroy mitotically active cells (i.e. stem cells including PGCs and tumor‐forming cells). A stock solution of 1 mg/100 µl busulfan in DMSO was prepared. Working solutions were prepared by adding 0, 40, 60, or 80 17
µl of stock busulfan/DMSO to 1 ml StemPro medium supplemented with LIF and FGF in microfuge tubes. These dilutions were adopted from an in vitro study on pharmacodynamics of busulfan for leukemia treatments (24). Cells were incubated in the 4‐well format described previously with control or busulfan treated medium for 2h at culture conditions, then washed 3 times using StemPro medium supplemented with LIF and FGF. Cells were then incubated overnight and examined for differences in proliferation and structural formation. V. Directed differentiation of PGCs within cell aggregates co‐cultured with ovaries or testes Day 9 and day 12 gonads were used as separate co‐culture systems to assess whether a stage of development directs the differentiation of PGCs within the cell aggregate. Day 8‐9 for the embryonic chick is the stage in which gonadal differentiation is observable; the right ovary in females is slightly smaller than the left, and the testes in males are equal in size. Day 12 for the embryonic chick is a later stage in which gonadal differentiation is complete; the left ovary is several‐fold larger than the regressing right gonad. After 5 days of culture, large cell aggregates (100‐200 µm diameter) which formed over attached, non‐confluent somatic cells in LIF + FGF culture conditions were selectively picked up using a 100 µl pipet tip. Four to five aggregates were placed in a 0.4 µm pore size, PET track‐
etched membrane cell‐culture insert (Falcon, #353095) with pre‐warmed StemPro medium without LIF above embryonic gonads from day 9 or day 12 embryos with StemPro medium without LIF in a 24‐well format. Dissected embryonic gonads from day 9 and day 12 embryos were placed in a polystyrene, flat bottom 24‐well culture plate with 16mm well diameter (Corning, #25830) on the day that cell aggregates were placed in the cell culture insert. Two to four male or female gonads were placed in each well. Cell aggregates in co‐culture systems were examined daily for developmental changes in morphology, and a single cell aggregate was picked using a 100 µl tip for partial dissociation and 18
immunostaining to identify EG cells (originally PGCs). Cells were cultured for 3 weeks with weekly medium changes. VI. Staining of cell aggregates and microscopy Staining of cell aggregates Cell aggregates from different days of incubation and culture conditions (busulfan treated or gonadal co‐cultures) were stained with dilutions of 1:100 rabbit polyclonal antibody to DDX4/Mvh (Abcam, ab13840) and 1:10 SSEA1 separately. Stained aggregates were incubated overnight, then washed twice in PBS with 2% BSA. A 1:5 dilution of Alexa Fluor 488‐labelled goat anti‐chicken IgG antibodies was used as the secondary antibody. Aggregates were incubated for 15‐20 min with secondary antibody, then washed twice with 2% BSA. In some instances, cells were fixed with 4% paraformaldehyde after staining steps before observation with a fluorescence microscope. Fixed cells were also stained with a 1:100 dilution of propidium iodide (PI) (Invitrogen, L7011, 2.4 mM solution in water). Fluorescence microscopy Stained cell aggregates were collected in a watch‐glass with 1.0 ml PBS. Individual aggregates were picked up using a 20 µl micropipette and placed with 10 µl of PBS onto a microscope slide. A 22 mm2 cover slip was placed on top of the cell aggregate using Vaseline/paraffin “pedestals” on each corner to prevent air bubbles and protect the aggregates. Aggregates were first located using brightfield optics, then with the green filter of the fluorescence microscope to detect Alexa Fluor 488 conjugated to the secondary antibody, and then with the red filter to detect propidium iodide staining. Stained EGs (differentiated PGCs) fluoresce bright green (emission 519 nm) against a dark background, and PI‐
stained nuclei of all cells fluoresce bright red. 19
VII. Click‐iT EdU Cell Proliferation Assay The Click‐iT EdU proliferation assay (Invitrogen, MP 10083) was performed according to the protocol provided by Invitrogen. “EdU (5‐ethynyl‐2’‐deoxyuridine) is a nucleoside analog of thymidine and is incorporated into DNA during active DNA synthesis. Detection is based on a click reaction, a copper catalyzed covalent reaction between an azide and an alkyne. In this application, the EdU contains the alkyne and the Alexa Fluor® dye contains the azide. The small size of the dye azide allows for efficient detection of the incorporated EdU.” (25) Labeling Cells with EdU Gonadal cells from 6 day embryos were plated on coverslips in the 4‐well format described in StemPro medium supplemented with LIF and FGF for 2 days. On the second day of incubation after numerous cell aggregates formed, spent media was replaced with an equal volume of prewarmed 4X EdU solution in complete medium (concentration of 40 µM) to the volume of media containing cells to obtain a 2X EdU solution (final concentration of 20 µM). Cells were incubated in 2X EdU solution under culture conditions for 1 day. Cell Fixation and Permeablilization After incubation, media containing EdU was removed and 0.7 ml 4% buffered paraformaldehyde in PBS was added to each well containing coverslips. After 15 min incubation at room temperature, fixative was removed and cells were washed twice in each well with 0.7 ml 3% BSA in PBS (3 min for each wash). Wash solution was removed and 0.7 ml 0.5% Triton X‐100 in PBS was added to each well. Cells were incubated in Triton permeablization buffer for 20 min at room temperature. The permeabilization buffer was removed and cells in each well were washed twice with 0.7 ml 3% BSA in PBS. 20
EdU Detection For 4 coverslips, 2 ml Click‐iT reaction cocktail was prepared from 1.8 ml 1X Click‐iT reaction buffer, 80 µl CuSO4, 10 µl Alexa Fluor azide, and 200 µl reaction buffer additive (reagants provided in Click‐iT kit) and protected from light. Wash solution was removed and 0.5 ml Click‐iT reaction cocktail was added to each well containing a coverslip within 15 min of cocktail preparation. The plate was rocked briefly to insure that the reaction cocktail was distributed evenly over the coverslip. Cells were incubated for 30 min at room temperature, protected from light. After incubation, the reaction cocktail was removed and each well was washed once with 0.7 ml 3% BSA in PBS and again in 0.7 ml PBS. DNA Staining Hoeschst 33342 solution provided in Click‐iT kit was diluted 1:2000 in PBS to obtain a 1X Hoechst 33342 solution (5 µg/ml final concentration). Wash solution was removed and 0.7 ml 1X Hoeschst 33342 solution was added to each well. Cells were incubated for 30 min at room temperature, protected from light. Hoeschst 33342 solution was removed and cells were washed twice with 0.7 ml/well PBS. Imaging and Analysis Using jeweler’s forceps, coverslips were removed from wells and placed right‐side down on a microscope slide with an 8 µl drop of PBS so that cells were between the coverslip and slide. Slides were examined using fluorescence microscopy to detect stained cells. The approximate fluorescence excitation/emission maxima for Alexa Fluor 594 for EdU detection is 590 nm/615 nm (fluoresce bright red), and for Hoechst 33342 stain bound to DNA is 350 nm/461 nm (fluoresce bright blue). 21
RESULTS I. Identification and Characterization of Isolated PGCs PGC and somatic cell counts Concentration of gonadal cells in cell suspensions prepared from two 6d chick gonads was determined using a hemacytometer and phase contrast microscope. In a 0.1 ml suspension of 6 day gonadal cells, cell concentration was 66,875,000 somatic cells/ml (n=3). Counting of PGCs was difficult due to flow concentration and aggregation of cells. Based on the literature, PGCs are present as 7,400 ± 1,300 (0.74± 0.13%) per 1 x 106 gonadal cells at this stage of development (26). Fluorescence microscopy PGCs stained with 1:2 and 1:10 dilutions of primary antibody against SSEA‐1, 1:2 and 1:10 dilutions of SSEA‐4, and a 1:2 dilution EMA‐1 antibody fluoresced brightly and selectively. Other experimental concentrations of primary antibodies showed either very faint fluorescence or no fluorescence. Poling and Parks (22) demonstrated that SSEA‐1 was expressed at higher levels than SSEA‐
4, EMA‐1, and CD‐9 antigens in cells of the female embryonic gonad and testicular cells from Japanese quail (Cotumix japonica). Observations in the present study are consistent with these earlier reports. Stained cells were approximately twice as large in diameter as smaller, presumed somatic cells which showed no fluorescence after staining procedure. Figure 2 shows single stained fluorescent cells with non‐fluorescent cells in the same field. The fluorescence indicates the cell’s pluripotency and presumed PGC identity. Larger cells of approximately 20 µm were consistently positive (fluorescent) for pluripotent cell markers except when located within an undissociated cell cluster. Cells incubated with only secondary antibody and no primary antibody (control) did not show fluorescence (not shown in figure). 22
A
B
C
D
E
F
G
H
I
J
K
L
Figure 2: Brightfield and fluorescent images of titrated primary antibodies. A 1:2 dilution SSEA‐1 brightfield, B 1:2 dilution SSEA‐1 fluorescent, C 1:10 dilution SSEA‐1 brightfield, D 1:10 dilution SSEA‐1 fluorescence, E 1:2 dilution SSEA‐4 brightfield, F 1:2 dilution SSEA‐4 fluorescence, G 1:10 dilution SSEA‐4 brightfield, H 1:10 dilution SSEA‐4 fluorescence, I 1:2 dilution EMA‐1 brightfield, J 1:2 dilution EMA‐1 fluorescence, K 1:10 dilution EMA‐1 brightfield, L 1:10 dilution EMA‐1 fluorescence. Scale bars are 20 µm. 23
II. Cell culture of PGCs using different media Dissociated cells of 6 day embryonic gonads originally put into culture consisted of numerous 8‐
10 µm diameter somatic cells and few 20 µm diameter PGCs based on measurements from Nikon DS Camera Head DS‐Fi1 and DS Camera Control Unit DS‐L2 (Figure 3). These measurements were consistent with literature on PGCs. Effect of media on gonadal cell cultures Qualitative observations of cultures supplemented with only LIF were from 20 cultures in the 4‐
well format described (e.g. slightly less than 80 wells accounting for culture contamination). During a 4 day culture, most aggregates were on the order of 80 ‐100 µm diameter; only a very few cell aggregates were greater than 100 µm diameter (Figure 4). Clumps of cell aggregates ranging from 100‐200 µm in length were observed after 3 days of incubation. Cell aggregates were uniformly round, multi‐layered, and well‐delineated. In a 1x1 mm2 area of the culture plate, a mean of 3.4 ±1.9 aggregates formed after 1 day of incubation and increased steadily to a mean of 6.6 ±1.8 aggregates after 4 days of incubation. These data were obtained from a single 4‐well culture (16 counts total as described in M&M) that demonstrated typical characteristics of 4‐well cultures using media supplemented with only LIF. Cultures supplemented with only LIF had a substantially lower rate of proliferation of somatic cells and formation of cell aggregates compared with cultures supplemented with LIF and FGF. 24
Figure 3: Low magnification of dissociated cells from day 6 embryonic gonads initially put into culture for all experiments, consisting of somatic cells and a few PGCs (see Figure 2E). Green bars (Legends 1‐4) are measurements of ~12 µm from the Nikon DS Camera Control Unit. While measurements of somatic cells and PGCs were generally consistent with sizes in literature (8‐10 µm diameter somatic cells and 20 µm diameter PGCs), identification of individual cells based on size was difficult in unstained preparations due to incomplete dissociation of all cells. Scale bar (5) is 100 µm. 25
A
B
C
D
Figure 4: Dissociated cells of 6 day embryonic gonads originally put into culture using StemPro medium with LIF and no FGF after: A) 1 day, B) 2 days, C) 3 days, and D) 4 days in culture. Cell aggregates were attached to the bottom of the culture wells. Scale bars are 100 µm. 26
Qualitative observations of cultures supplemented with LIF and FGF were from 10 cultures in the 4‐well format described (e.g. slightly less than 40 wells accounting for culture contamination). Cultures supplemented with LIF and FGF formed well‐defined, uniformly round cell aggregates after 1 day of incubation (Figure 5). By 2 days of incubation, some cell aggregates in FGF culture medium reached over 200 µm in diameter; aggregates in medium lacking FGF only reached around 170 µm in diameter after 4 days of incubation. Clumps of cell aggregates ranging from 100‐200 µm in length were observed after 1 day of incubation; cell aggregates formed even larger clumps ranging from 200‐500 µm in length after 5 days of incubation in medium supplemented with LIF and FGF. Based on subjective assessment, FGF supplementation increased the rate of cell aggregation formation several‐fold in comparison to cultures without FGF. In a 1x1 mm2 area of the culture plate, a mean of 19.4±7.2 aggregates formed after 1 day of incubation. This data was obtained from a single 4‐well culture (16 counts total as described in M&M) that demonstrated typical characteristics of 4‐well cultures using media supplemented with LIF and FGF. A mean number of aggregates after 4 days of incubation was not recorded because consistent counting was not achievable using the same methods due to clumping and increased sizes of aggregates. However, there was an observable increase in cell aggregate formation throughout 4 days of incubation. This observation was supported by the Click‐iT EdU proliferation assay and Hoechst 33342 DNA staining (Figure 6). Aggregates were counterstained with Hoechst 33342, nuclear stain, to observe total cells. A high percentage of EdU‐Alexa Fluor 594 stained cells fluoresced bright red within cell aggregates on day 3 of incubation. 27
A
B
C
D
Figure 5: Dissociated cells of 6 day embryonic gonads initially put into culture using StemPro medium supplemented with LIF and FGF after: A) 1 day, B,C) 2 days, D) 3 days, and E) 4 days in culture. By day 2, some cell aggregates are over 200 µm in diameter (B). Green bars are E
F
measurements of aggregate width made with the Nikon DS Camera Control Unit. Cell aggregates were attached to the bottom of the well. Scale bars are 100 µm. 28
A
B
C
D
E
F
Figure 6: Click‐iT EdU proliferation assay 3rd day of incubation. A,B) EdU‐Alexa Fluor 594 stained cells fluoresce bright red. C,D) Hoechst 33342 stain bound to DNA fluoresces bright blue. E,F) Merged images of EdU‐Alexa Fluor 594 and Hoechst 33342 stained cells (A,C and B,D). Scale bars are 100 µm. 29
III. Effect of busulfan treatment on cultured 6d chick gonadal cells Cell cultures conditioned with LIF and FGF supplements treated with 0, 40, 60, and 80 µg/ml busulfan in DMSO were examined for differences in proliferation and aggregate formation (Figure 7). Difference in aggregate formation was striking. While a single 1 mm2 area of a culture plate with a uniform distribution of cells and cell aggregates contained approximately 20 cell aggregates, only 3 cell aggregates were seen in 40 µg/ml, 2 cell aggregates were seen in 60 µg/ml, and 2 cell aggregate were seen in 80 µg/ml busulfan treated cultures after 1 day incubation throughout the entire well. Though there was an observed decrease in somatic cell number, somatic cells 10‐12 µm diameter were still present after 1 day incubation, thus it was assumed that the two hour busulfan treatment did not disturb somatic cell survival. The busulfan treatment was presumed to have affected stem cell proliferation by destroying PGCs in the initial culture. Few cell aggregates formed after 1 day culture which could be explained by the presumed absence of PGCs from the busulfan treatment. 30
A
B
D
C
Figure 7: Busulfan treated cell cultures. A) 0 µg/ml (control), B) 40 µg/ml, C) 60 µg/ml, and D) 80 µg/ml busulfan in DMSO after 1 day of incubation. Cell aggregates formed in 0 µg/ml busulfan treated culture (control) after 1 day of incubation. Only 3 cell aggregates were seen in 40 µg/ml, 2 cell aggregates were seen in 60 µg/ml, and 2 cell aggregates were seen in 80 µg/ml busulfan treated cultures (4 well culture plate) after 1 day of incubation. Scale bars are 100 µm. 31
IV. SSEA‐1 staining of cell aggregates formed in cultures conditioned with LIF and FGF Cell aggregates were stained with SSEA1 overnight. There were approximately 3‐4 SSEA1‐
positive fluorescing cells within a cell aggregate at a given field of focus (Figure 8). This would suggest that the cell aggregate contained pluripotent stem cells. These SSEA1‐positive cells were located throughout the 3‐dimensional cell aggregate. Though cell aggregates fluoresced diffusely after immunocytochemical staining, bright SSEA1‐positive fluorescing cells were still detectable. A clump of 3 cell aggregates stained without primary antibody (SSEA‐1) and only Alexa Fluor 488 conjugated to the secondary goat anti‐chicken antibody (Figure 8G) showed all cells fluorescing diffusely. Unstained cell aggregates revealed no auto‐fluorescence (Figure 8H) which indicated that the secondary antibody was binding non‐specifically to cells. V. SSEA‐1 staining of cell aggregates culture with busulfan The few cell aggregates that formed in the busulfan treated culture were stained with SSEA1 to detect presence of any pluripotent EG cells. Fluorescence microscopy revealed very few fluorescing cells within the aggregates (Figure 9) compared to aggregates in cultures not treated with busulfan (Figure 8). This is a qualitative assessment; fluorescing cells were not counted due to limitations of using fluorescent optics on the large 3‐dimensional aggregates. Presumed SSEA1‐positive EG cells were considered to be PGCs that survived the busulfan treatment. 32
A
B
C
D
Figure 8: Fluorescent and brightfield images of day 3 cell aggregates stained with primary antibody SSEA1 and Alexa Fluor 488 conjugated to the secondary goat anti‐chicken antibody and/or propidium iodide. A,B) Brightfield and fluorescence images of cell aggregate stained with SSEA1 in two different focal planes (aggregates are 3 dimensional). All cells fluoresce diffusely presumably due to non‐specific staining, but more intense fluorescing cells (arrows) were considered to be SSEA1‐positive pluripotent EG cells (differentiated PGCs). C) Merged brightfield and fluorescent images of cell aggregate stained with SSEA1. D) Fluorescent images of cell aggregates stained with SSEA1 and propidium iodide. All cells that stained with propidium iodide fluoresced red. Reducing the brightness of the fluorescent image of SSEA1‐stained aggregates using the filter for Alexa Fluor 488 revealed a smaller population of more intense green fluorescing cells (presumptive EG cells) that remained visible when merged with the fluorescent image of propidium iodide stained cells. Aggregates on the right are different from the aggregate on the left and exhibited different fluorescence intensity when viewed for Alexa Fluor 488 and different shading when merged with the image of PI‐staining. 33
E
F
G
H
Figure 8 continued: E,F) Merged brightfield and fluorescent images of a cell aggregate stained with SSEA1 in two different focal planes. G) Clump of cell aggregates stained with only Alexa Fluor 488 conjugated to the secondary goat anti‐chicken antibody and no primary antibody demonstrating fluorescence. Fluorescence indicated secondary antibody bound non‐specifically with aggregates, but notably brighter fluorescing individual cells in A‐F (indicated by arrows) are presumed to be stained positive for SSEA1. H) Control: unstained clump of cell aggregates with no auto‐fluorescence. Scale bars are 100 µm. Magnification is the same for all images. 34
A
B
C
D
Figure 9: Fluorescent and brightfield images of 4 separate cell aggregates from A) 40 µg/ml, B,C) 60 µg/ml, D) 80 µg/ml busulfan treated cultures stained with primary antibody SSEA1 and Alexa Fluor 488 conjugated to the secondary goat anti‐chicken antibody. Only 1‐2 aggregates formed in these busulfan treated cultures which was significantly different from the 20 aggregates formed in a culture without busulfan treatment (see figure 7A); however SSEA1 staining of these few aggregates revealed the presence of EG cells that apparently survived busulfan treatment. A,B are merged fluorescent and brightfield images. Aggregates exhibited different fluorescence intensity when viewed for Alexa Fluor 488, so brightness was adjusted to capture image. C,D are fluorescent images of separate aggregates. Bright, intense fluorescing cells were observed (arrows) and assumed to be SSEA1‐positive pluripotent EG cells. Scale bars are 100 µm. Magnification is the same for all images. 35
VI. Effects of embryonic testis and ovary co‐culture on differentiation of embryonic gonad cell aggregates. Cell aggregates from primary cultures of dissociated 6 day gonads were placed in a cell‐culture insert with Stem‐Pro medium supplemented with FGF within a culture well containing the same medium and embryonic gonads from day 9 or day 12 embryos. LIF was omitted from the medium so that differentiation would not be inhibited. After five days of culture without changing medium, cell aggregates within the cell‐culture insert underwent morphological changes in structure depending on the sex of the gonads with which they were co‐cultured. Cell aggregates co‐cultured with 9 or 12 day ovaries Aggregates placed above intact 9 or 12 day ovaries did not appear to change morphologically after 5 days of incubation (Figure 10). A positive propidium iodide stain of cells on the cell aggregate surface revealed the aggregate as a ball of cells after 5 days of co‐culture which was not so different from the aggregates when they were first put into co‐culture. This was observed from three 9 day ovary co‐culture systems and eight 12 day co‐culture systems. Cell aggregates co‐cultured with 9 or 12 day testes Cell aggregates co‐cultured above 9 or 12 day intact testes developed a definitive margin, perhaps a basement membrane, after 5 days of incubation (Figure 11). These observations were from 6 co‐culture systems. In two cultures with 12 day embryonic testis, cells proliferated to fill the space between adjacent aggregates suggestive of sex cord development. This sex cord‐like development was thought to have originated from somatic cells of the dissociated 6 day gonads which made up the original cultures for aggregate formation. This may suggest that the 12 day testes directed the differentiation of somatic cells from the indifferent 6 day gonad into Sertoli cells of the male gonad. 36
Sertoli and germ cell specific markers will be needed to identify these cells which suggest sex cord development. The similar observations between the cell aggregates co‐cultured with 9 day testes and cell aggregates co‐cultured with 12 day testes suggest that there was no critical stage in gonadal development that directed the developmental and differentiation patterns of cell aggregates consisting of EG cells. 37
A
B
Figure 10: Aggregates after co‐culturing with A) 9 day and B) 12 day ovaries for 5 days. C) Fluorescent image of cells stained with propidium iodide on surface of aggregate co‐cultured with 12 day ovaries for 5 days. Scale bars are 100 µm. C
38
A
B
C
D
Figure 11: A) Cell aggregates co‐cultured with intact 9‐day testes developed definitive margins, or basement membranes after 5 days incubation. B) Cell aggregates co‐cultured with 9d testes after 2 days of incubation. Black arrow points to intact gonad below cell insert. C,D) Cell aggregates co‐cultured with 12d testes after 5 days of incubation. Red arrows point to somatic cells that proliferated between the aggregates during the 5 day culture period which suggest sex cord development. Scale bars are 100 µm. 39
DISCUSSION The ability to recover, preserve, culture, and direct development of germ cells from avian species has important implications for domestic poultry production, aviculture, and avian conservation. This study was initiated as part of an effort to develop avian germ cell transplantation procedures as a tool for conserving avian germ plasm from species of special concern. The objectives were to harvest and culture PGCs from domestic chick embryonic gonads at the indifferent gonad stage (stage 28‐29, 6 days of incubation) as a model avian system and assess factors affecting survival, proliferation, and development of gonadal cells (somatic and germ cells) in vitro. Initially, indifferent gonads were recovered from 6 day embryos and dissociated with collagenase. Gonadal cells were then cultured in stem cell medium designed to maintain the pluripotency of PGCs. Culture conditions resulted in gonadal cells spontaneously forming large, round aggregates (80‐100 µm diameter) overlaying a more diffuse layer of somatic cells which adhered to the bottom of the culture plate. These aggregates increased in size (100‐200 µm diameter) and number (two‐fold) over 4 days of culture. Including FGF in the culture medium appeared to increase the number and formation rate of cell aggregates; size and count could not be established after 4 days of incubation due to massive clumping of aggregates. However, the proliferation assay confirmed observations of extensive proliferation of cell aggregates in StemPro medium supplemented with LIF and FGF by day 3 of culture. Cell proliferation should be assessed at earlier and longer periods of incubation for direct measurement of cells synthesizing DNA under the various culture conditions used in this study. Also, cell proliferation of PGCs versus somatic cell should be differentiated. Presumptive PGCs, present as a small percentage of total cells within cell aggregates, were identified using pluripotent stem cell marker SSEA1. Addition of busulfan, a stem cell poison, almost completely eliminated cell aggregate formation. The few aggregates that formed contained very few PGCs based on presence of stem cell marker SSEA1, suggesting a requirement of PGCs in aggregate 40
formation, and possibly the preliminary stages of EBs. In standard protocols, EBs were formed from purified ES cells or PGCs. If the cell aggregates described in the present study were preliminary stages of EBs, then purified populations of ES or PGCs might not be required for EB formation because this procedure did not involve any purification of embryonic gonadal cells. Cell aggregates produced in this study shared many characteristics of EBs in the literature reviewed above: size (80‐200 µm in diameter), 3‐dimensional nature, thick shell, and rapid growth (within 4 days of incubation from a suspension of cells). Though SEM imaging of cell aggregates was not included in the study to determine whether a thick ECM was synthesized around the aggregate, difficulties in staining aggregates is suggestive that an impermeant layer exists around the aggregate. Only after aggregates were treated with collagenase did EG cells within aggregates stain successfully for SSEA‐1. In addition, the protocol for EdU staining called for a permeablization step using 0.5% Triton X‐100, suggesting an impermeant outer layer. One principal difference between aggregate formation in the study and EB formation in the literature is that EB formation requires the removal of LIF in culture medium. Aggregates were formed with LIF included in the medium, which might suggest that aggregates were only precursors to EBs. LIF was removed from culture medium when cell aggregates were co‐cultured with embryonic gonads from 9 or 12 day embryos. When co‐cultured with embryonic testes, aggregates appeared to grow and differentiate somewhat, with qualitatively more well‐defined margins. In addition, difficulties in staining the aggregates that were co‐cultured with embryonic testes with germ cell markers SSEA1 and SSEA4, supported our qualitative characterization of a definitive margin or basement membrane. Sachlos and Auguste described day 7 EBs to have a dense 20 µm thick shell that needed to be perforated using collagenase to facilitate diffusive transport of biochemicals (1). It may be possible that staining of cell aggregates after they were co‐cultured with embryonic gonads for more than 5 days was unsuccessful in this study because the aggregates developed a thick membrane, impermeable to standard staining methods. Dissociation of these cell aggregates and staining for EG cells would help to 41
assess the effects of this study’s co‐culture system on PGC differentiation. In two cultures with embryonic testes, cells proliferated to fill the space between adjacent cell aggregates suggestive of sex cord development. Further characterization of cell aggregates that have been co‐cultured with embryonic gonads using specific cell markers (Sertoli or germ cell), and histological examination of fixed, sectioned aggregates are needed to determine the nature of these structures. Aggregates co‐cultured with embryonic ovaries were more difficult to characterize; based on PI‐positive cells on the aggregate surface, they appeared as a ball of germ and somatic cells surrounded by somatic cells after 5 days of co‐
culture. This appearance has some similarity to early follicle formation. The exact nature of gonadal cell aggregates remains to be determined. These structures have some features consistent with embryoid bodies (EB) and may be influenced or directed to differentiate like EBs depending on the conditioned environment. If the cell aggregates formed in this study contain and support a number of EG cells, then culturing them with embryonic gonads may be a method to direct PGC differentiation. Factors secreted by embryonic gonads may be selectively isolated to determine what is absolutely necessary for the specific sex differentiation of PGCs; eventually, it may be possible to derive sex specific germ cells using only those differentiation factors and a continued line of PGCs. This may be a critical advance for nuclear transfer studies in that a limitless source of PGC‐derived oocytes could be provided. Also, the use of PGC‐derived male gonocytes in testis transplants may be a new source of treatment for infertile males. PGC‐derived male gonocytes may functionally engraft in testes like testis cell transplants which have been shown to functionally engraft in the testes of infertile male recipients (27). It will be important to see whether PGC‐derived oocytes are fertilizable and whether PGC‐derived gonocytes can mature into functional sperm that have the ability to fertilize oocytes. At the very least, the directed differentiation of PGCs in vitro via the culture system of this study may contribute to our knowledge of germ cell development and the factors which commit the germ cell to its respective sex. 42
Literature Cited 1. Sachlos, E., Auguste, D. 2008. Embryoid body morphology influences diffusive transport of inductive biochemicals: A strategy for stem cell differentiation. Biomaterials, 29: 4471‐4480. 2. Park, T.S., and Han, J.Y. 2000. Derivation and characterization of pluripotent embryonic germ cells in chicken. Molecular Reproduction and Development, 56: 475‐482. 3. Wu, Y., Zhang, C. 2008. Formation and differentiation of embryoid bodies from cultured chicken primordial germ cells. Comparative Biochemistry and Physiology Part C: Toxicology and Pharmacology, 148:469. 4. Toyooka, Y., Tsunekawa, N. 2002. Embryonic stem cells can form germ cells in vitro. s.l. : Pro. Natl Acad. Sci. USA, 100: 11457‐11462. 5. Novak, I. 2006. Mouse embryonic stem cells form follicle‐like ovarian structures but do not progress through meiosis. Stem Cells, 24: 1931‐1936. 6. Geijsen, N., Horoschak, M. 2004. Derivation of embryonic germ cells and male gametes from embryonic stem cells. Nature, 427: 148. 7. Chuva de Sousa Lopes, S., and Hayashi, K. 2007. Proximal visceral endoderm and extraembryonic ectoderm regulate the formation of primordial germ cell precursors. BMC Developmental Biology, 7: 140. 8. Xiaoxia Qi, Y., Zhao, G. 2001. Induction of primordial germ cells from murine epiblasts by synergistic action of BMP4 and BMP8B signaling pathways. Developmental Biology, 98: 7858–7862. 9. Lange, UC and Saitou, M. 2003. The Fragilis interferon‐inducible gene family of transmembrane proteins is associated with germ cell specification in mice. BioMedCentral Developmental Biology, 3:1. 10. Hamburger, V., Hamilton, HL. 1951. A series of normal stages in the development of the chick embryo. J. Morph, 88: 49‐92. 11. Parks, J. 2002.Primordial Germ Cell Origin, Migration, and Differentiation; Ontogeny of the Reproductive Sycstem and Sexual Differentiation; Functional Anatomy and Histology of the Testis and Ovary. Animal Science 2250 Lecture 2. Ithaca. 12. Durcova‐Hills, G., McLaren, A. 2006. Isolation and Maintenance of Murine Embryonic Germ Cell Lines. Essentials of Stem Cell Biology (Lanza, R., et.al., ed.). pp.269, Academic Press, Burlington. 13. Shamboltt, M. 1998. Derivation of pluripotent stem cells from cultured human primordial germ cells. Developmental Biology, 95: 13726‐13731. 14. Qing, T., Shi, Y. 2007, Induction of oocyte‐like cells from mouse embryonic stem cells by co‐culture with ovarian granulosa cells. Differentiation, 75:902‐911. 15. Tezel, G., Nagasaka, T. 2002, Differential expression of RET finger protein in testicular germ cell tumors. Pathology International, 52: 623 – 627. 16. Hubner, K., Fuhrmann, G. 2003, Derivation of Oocytes from mouse embryonic stem cells. Science, 300: 1251‐1256. 17. Wang, J. 2009, Directed differentiation of chick embryonic germ cells into neural cells using retinoic acid induction in vitro. Journal of Neuroscience Methods, 177: 168‐176. 18. Jenson, J. 2009. Cornell Animal Science Department Reproduction Seminar Series. Ithaca. 19. Naito, M. 2009, Preferential Migration of Transferred Primordial Germ Cells to Left Germinal Ridge of Recipient Embryos in Chickens. Journal of Poultry Science, 46: 40‐45. 20. Tagami, T. 2007, Differentiation of Female Primordial Germ Cells in the Male Testes of Chicken. Molecular Reproduction and Development, 74: 68‐75. 21. Caprette, D. Laboratory Course Web Pages. [Online] Rice University, January 2007. http://www.ruf.rice.edu/~bioslabs/methods/microscopy/cellcounting.html. 22. Poling, M., Parks, J. 2009. Isolation of adult gonadal stem cells from Cotumix japonica and characterization of embryonic gonadal stem cells in Gallus gallus for xenotransfer. SSR: Poster presentation; Stem cells. Escondido : San Diego Zoo’s Institute for Conservation Research. 43
23. StemPro® hESC SFM Manual.[Online] April 2, 2008. 24. Hassan, Z., Hassan, M. 2001, The pharmacodynamic effect of busulfan in the P39 myeloid cell line in vitro. 8, s.l. : Leukemia. Nature, 15: 1240‐1247. 25. Invitrogen. Click‐iT EdU Imaging Kits Manual. [Online] March 27, 2008. http://probes.invitrogen.com/media/pis/mp10338.pdf. 26. Kim, J.N. 2004, Enriched Gonadal Migration of Donor‐Derived Gonadal Primordial Germ Cells by Immunomagnetic Cell Sorting in Birds. Molecular Reproduction and Development, 68: 81‐87. 27. Brinster, R., Zimmermann, J. 1994, Spermatogenesis following male germ‐cell transplantation. 24, Proc Natl Acad Sci USA, 91:11298‐11302. 44