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First published online March 22, 2007
Stem Cells Vol. 25 No. 7 July 2007, pp. 1690 -1696
doi:10.1634/stemcells.2006-0607; www.StemCells.com
© 2007 AlphaMed Press

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TECHNOLOGY DEVELOPMENT

Facilitated Expansion of Human Embryonic Stem Cells by Single-Cell Enzymatic Dissociation

Catharina Ellerströma,b, Raimund Strehla, Karin Noakssona, Johan Hyllnera, Henrik Sembb

aCellartis AB, Göteborg, Sweden;
bStem Cell Center, Lund University, Lund, Sweden

Key Words. Human embryonic stem cell • Human feeders • Single cells • Enzymatic dissociation • Expansion

Correspondence: Henrik Semb, Ph.D., Stem Cell Center, Lund University, BMC, B10, SE-221 84 Lund, Sweden. Telephone: 46-46-222-31-59; Fax: 46-46-222-36-00; e-mail: henrik.semb{at}med.lu.se

Received September 28, 2006; accepted for publication March 13, 2007.
First published online in STEM CELLS EXPRESS   March 22, 2007.


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Traditionally, human embryonic stem cells (hESCs) are propagated by mechanical dissection or enzymatic dissociation into clusters of cells. To facilitate up-scaling and the use of hESC in various experimental manipulations, such as fluorescence-activated cell sorting, electroporation, and clonal selection, it is important to develop new, stable culture systems based on single-cell enzymatic propagation. Here, we show that hESCs, which were derived and passaged by mechanical dissection, can be rapidly adjusted to propagation by enzymatic dissociation to single cells. As an indication of the stability of this culture system, we demonstrate that hESCs can be maintained in an undifferentiated, pluripotent, and genetically normal state for up to 40 enzymatic passages. We also demonstrate that a recombinant trypsin preparation increases clonal survival compared with porcine trypsin. Finally, we show that human foreskin fibroblast feeders are superior to the commonly used mouse embryonic fibroblast feeders in terms of their ability to prevent spontaneous differentiation after single-cell passaging. Importantly, the culture system is widely applicable and should therefore be of general use to facilitate reliable large-scale cultivation of hESCs, as well as their use in various experimental manipulations.

Disclosure of potential conflicts of interest is found at the end of this article.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Pluripotent human embryonic stem cells (hESCs) are derived from the inner cell mass of blastocysts and have the theoretical capacity for indefinite, undifferentiated proliferation in vitro. hESCs have the unique potential to give rise to all cell types of the body. To maintain hESCs in an undifferentiated state in vitro, the cells have traditionally been maintained on mouse embryonic fibroblast (MEF) feeders and have been passaged by manual microdissection and transfer of individual colonies [1–4]. In addition, feeder-free culture conditions for hESCs, using MEF-conditioned or growth factor-supplemented defined media, have been used [5–11]. Whereas these methods can be used for experiments that do not require large-scale production of undifferentiated hESCs, gene targeting, drug discovery, in vitro toxicology [12, 13], and future potential clinical applications will require new methods for the stable large-scale expansion of hESCs, including enzymatic passaging. Indeed, enzymatic expansion of hESCs has been reported by several groups [5–8, 14, 15]. However, technical disadvantages have become apparent, such as the necessity to maintain the hESCs in clusters within a narrow size range during passage [6–8], the necessity to adapt the cells to enzymes directly after the establishment of a new hESC line [14], a lengthy adaptation phase [15], and relatively low passage ratios [7, 14, 16]. Furthermore, concerns have been raised that these culture methods may result in genetic and epigenetic changes [14, 15, 17, 18]. For example, recent studies suggest that enzymatic dissociation of hESCs may lead to abnormal karyotypes [9, 14, 15, 17, 19]. Consistent with these observations, the potential transformation of hESCs toward a culture-adapted or in vitro-adapted cell line, which may start to resemble embryocarcinoma cells with changes in its differentiation potential, has also been reported [20].

It is known that hESCs depend on cell-cell interactions as well as para/autocrine signals and that survival of hESCs at clonal density is very low [21]. Therefore, the cellular microenvironment composed of the substrate that the cells grow on and the culture medium will play a major role in the survival of single cells. Thus, we considered it important to develop a highly supportive culture environment, which would allow robust large-scale expansion of undifferentiated hESCs without compromising the pluripotency or genetic stability over extended culture periods. In addition, the method should be widely applicable to already established hESC lines and only demand a short period of transfer adjustment from the traditional propagation method to the new culture system. We considered it to be an additional advantage if the new system could support single-cell dissociation of hESCs, because that would facilitate hESCs expansion, cell sorting, and defined seeding for multiwell plate assays and enable automatization of culture procedures and clonal expansion (e.g., in clonal assays and electroporation followed by clonal selection).

Here, we describe a stable single-cell enzymatic dissociation (SCED) culture system, which supports long-term maintenance and efficient expansion of undifferentiated, pluripotent hESCs. We evaluated the method using several hESC lines and demonstrated that the method is robust and generally applicable to all cell lines tested.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Culture of Human Foreskin Fibroblast Feeders and Preparation of High-Density Feeder Layers
Commercially available human foreskin fibroblast feeders (HFFs) were obtained from the American Type Culture Collection (CRL-2429; Manassas, VA, http://www.atcc.org) and were cultured in Iscove's modified Dulbecco's medium (Invitrogen, Paisley, U.K., http://www.invitrogen.com), supplemented with 10% fetal bovine serum (FBS) (Invitrogen) and 1% penicillin-streptomycin. The cells were passaged using 1x trypsin-EDTA (Invitrogen). Confluent monolayers of HFFs were treated with mitomycin-C (Sigma-Aldrich, St. Louis, http://www.sigmaaldrich.com) and plated on 0.1% gelatin (Sigma-Aldrich)-coated in vitro fertilization (IVF) dishes at a density of 70,000 HFFs per cm2 in VitroHES medium (Vitrolife AB, Kungsbacka, Sweden, http://www.vitrolife.com) supplemented with 4 ng/ml human recombinant basic fibroblast growth factor (hrbFGF) (Invitrogen). We used HFFs from passages 5–16 because that range has been reported to support undifferentiated growth of hESCs [22, 23].

hESC Lines Used in the Experiments
The hESC lines SA001, SA002, SA002.5, SA121, and SA167 (Cellartis AB, Göteborg, Sweden, http://www.cellartis.com) were established and characterized as previously described [3, 6, 24–29]. Prior to the experiments, all lines were maintained in IVF dishes on mitomycin-C-inactivated early passage (passage 2–3) MEF (F1 hybrid CD1xC57BL/6; Charles River Laboratories, Sulzfeld, Germany, http://www.criver.com) feeder layers (45,000–60,000 cells per cm2) in VitroHES medium supplemented with 4 ng/ml hrbFGF and mechanically passaged by microdissection every 4–5 days. Throughout this study, cultures were maintained in an incubator at 37°C, 95% humidity and 5% CO2.

Transfer of hESCs to the SCED Culture System
To transfer manually passaged hESCs to the SCED culture system, the cells were washed once in 1x phosphate-buffered saline (PBS) (Invitrogen), after which 0.5 ml of 1x TrypLE Select (containing 1 mM EDTA; TrypLE Select; Invitrogen) or 1x trypsin/EDTA (0.25% trypsin, 1 mM EDTA; Invitrogen) was added to each IVF dish and incubated at 37°C for 3–5 minutes. When the hESC colonies started to round up from the feeder layer, the cell sheet was broken apart to a single-cell suspension by trituration with a pipette. After centrifugation (400g for 5 minutes), the supernatant was discarded, the hESC pellet was resuspended in VitroHES medium, and the single-cell suspension was plated onto IVF dishes containing a high-density HFF layer (70,000 HFFs per cm2). For this initial passage, split ratios of 1:2 to 1:8 were used.

Continued Culture and Expansion of hESCs in the SCED Culture System
hESCs were expanded in the above culture system consisting of high-density HFF cells and VitroHES medium supplemented with 4 ng/ml hrbFGF. After enzymatic dissociation for 4–6 minutes at 37°C in 1x TrypLE (Invitrogen) or 1x trypsin/EDTA (Invitrogen), single cells were seeded onto high-density HFFs (70,000 HFFs per cm2) at routine split ratios between 1:4 and 1:40. Culture medium was replaced with fresh VitroHES + 4 ng/ml hrbFGF every 2–3 days. Depending on the growth speed of the individual hESC line, the cells were passaged every 6–12 days. The hESC line SA001 was used to obtain proof of principle data for the SCED system, whereas SA121, SA002, SA002.5, and SA167 were used to confirm that the observed advantages were not restricted to the hESC line initially tested. SCED-cultured hESC lines were frozen and thawed as previously described [14].

Immunohistochemical and Histochemical Analysis of hESCs
The hESCs were regularly characterized immunohistochemically following every 5–10 SCED passages. hESC cultures were fixed in 4% paraformaldehyde for 15 minutes, permeabilized for 5 minutes in 0.5% Triton X-100 solution (Sigma-Aldrich) and subsequently blocked with 5% FBS in PBS (Invitrogen). The cells were incubated with primary antibody solution overnight at 4°C. The primary antibodies used were specific for Oct-4, TRA-1-60, TRA-1-81, SSEA-1, SSEA-3, and SSEA-4 (Santa Cruz Biotechnology, Santa Cruz, CA, http://www.scbt.com). Incubation with fluorescein isothiocyanate-, tetramethylrhodamine B isothiocyanate-, or Cy3-conjugated secondary antibodies (Jackson Immunoresearch Laboratories, West Grove, PA, http://www.jacksonimmuno.com) was performed for 60 minutes at room temperature. Cell nuclei were counterstained with 4,6-diamidino-2-phenylindole (DAPI) (Sigma-Aldrich). The activity of alkaline phosphatase was determined using an alkaline phosphatase activity detection kit according to the manufacturer's instructions (Sigma-Aldrich). Stainings were evaluated and documented using a Nikon Eclipse TE-2000 U fluorescence microscope (Nikon, Tokyo, http://www.nikon.com).

Reverse Transcription-Polymerase Chain Reaction
Primers for the following genes were used: Oct-4 (POU5F1), NANOG, Cripto (TDGF1), and AFP. The design and evaluation of theses primers, as well as the procedure for RNA extraction and reverse transcription, have previously been reported [25]. Polymerase chain reaction (PCR) conditions were optimized, and standard curves were generated as described [30]. All PCRs were performed with SYBR Green I chemistry in a Rotorgene 3000. For PCR, 1x Jump Start buffer 10x (Sigma-Aldrich), 3 mM MgCl2 (Sigma-Aldrich), 0.3 mM dNTP mig (Sigma-Aldrich), 0.4x SYBR Green (Molecular Probes Inc., Eugene, OR, http://probes.invitrogen.com), 0.4 µM forward primer (MWG Biotech, Ebeersberg, Germany, http://www.mwg-biotech.com), 0.4 µM reverse primer (MWG Biotech), 0.004 U/µl Jump Start taq polymerase (Sigma-Aldrich), and 2 µl of cDNA template were used in an final volume of 20 µl. After an initial denaturation/activation step of 3 minutes at 95°C followed by 45 cycles of 20 seconds at 95°C, 20 seconds at 60°C, and 20 seconds at 72°C, the detection of fluorescent signal was performed at 72°C in each cycle. Cycle threshold (Ct) values were calculated using Rotorgene software. Quantification of gene expression was based on the Ct value for each sample. The Ct values were calculated as the average of duplicate measurements. A mathematical model has previously described this in detail [30]. Line SA001, SCED-passaged with either TrypLE Select (SCED passages 24 and 35) or trypsin-EDTA (SCED passages 25 and 35), as well as SA002 (SCED passage 5) and SA167 (SCED passage 5), were analyzed. Since the hESC colonies grow on top of the HFFs, the feeder cells were included with the hESC colonies for sample preparation. As controls, manually cut material from lines SA001 and SA002 from traditional cultures and pure HFF samples were used. As expected, HFF controls did not express any of the selected genes (data not shown).

Genetic Characterization
All hESC lines were genetically characterized prior to the onset of this study [3, 6, 24–29]. For karyotype analysis, the hESCs were incubated in the presence of colcemid, trypsinized, fixed, and mounted on glass slides. The chromosomes were visualized by DAPI staining, arranged, and documented using an inverted microscope equipped with appropriate filters and software (CytoVision; Applied Imaging, Santa Clara, CA, http://www.appliedimagingcorp.com). SA001 (TrypLE Select) was analyzed at SCED passages 27 and 39 (17 and 14 metaphases screened). SA001 (trypsin-EDTA) was analyzed after 25 and 39 SCED passages (17 and 13 metaphases screened). SA121 (TrypLE Select) and SA121 (trypsin-EDTA) were both analyzed after 21 SCED passages (16 metaphases per group screened).

For fluorescence in situ hybridization (FISH) analysis, commercially available preimplantation genetic testing multiprobe panel and chromosome enumerating probe kits (Vysis, Downers Grove, IL, http://www.vysis.com) containing probes for chromosomes 12, 13, 17, 18, 20, 21, X, and Y were used according to the manufacturer's instructions, with minor modifications. The slides were analyzed in an inverted microscope equipped with appropriate filters and software (CytoVision). For each probe, a minimum of 100 nuclei were analyzed. Analyses of the different groups are presented in Table 1.


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  		Table 1. Validation of human ESCs after SCED passage

 
Analysis of Pluripotency In Vivo
In vivo pluripotency of SA001 and SA121 cultured in the SCED system, using either TrypLE Select or trypsin-EDTA, for 20 consecutive passages, were assessed by teratoma formation in SCID mice as described earlier [3, 29]. In brief, undifferentiated hESC colonies were mechanically cut into 200 x 200-µm pieces and surgically placed under the kidney capsule of severe combined immunodeficient (SCID) mice (C.B-17/lcrCrl-scidBR; Charles River Laboratories). The mice were sacrificed after 8 weeks, and tumors were excised and fixed in 4% paraformaldehyde. Hematoxylin and eosin-stained paraffin sections were evaluated histologically for the presence of differentiated human tissue derived from all three embryonic germ layers (i.e., neuroectoderm, cartilage, and gut-like epithelium).

Colony Formation Assay
To test the supportive capacity of the SCED system, traditionally cultured hESCs (on MEFs using mechanical passaging) were dissociated to single cells by using either TrypLE Select or trypsin-EDTA for 3–5 minutes at 37°C. The hESCs were diluted and seeded into IVF dishes at two densities on either HFFs (70,000 HFFs per cm2) or MEFs (45,000–60,000 MEFs per cm2), resulting in approximately 350 and 700 hESCs per cm2.

Medium was changed every 2–3 days, as described previously. Approximately 1 week later, the medium was removed, and the cells were washed with 1x PBS followed by alkaline phosphatase staining according to the manufacturer's description (Sigma-Aldrich). The number of hESC colonies obtained from all four test groups was determined. To allow semiquantitative evaluation of the numbers of undifferentiated colonies, a morphological grading system was used [25] in which a colony was scored positive if more than 50% of the colony was undifferentiated. The experiments were performed in duplicate and repeated three times. hESC line SA002.5 was used for these experiments.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 
Transfer of hESCs to the SCED Culture System
The derivation and characterization of all examined hESC lines, SA001, SA002, SA002.5, SA121, and SA167, was previously reported [3, 6, 24–29]. Preceding this study, the hESC lines were maintained on MEFs and mechanically passaged every 4–5 days. All hESC lines had been cultured on MEFs and mechanically passaged for <50 passages except for SA121, which had been cultured for >100 passages. Regardless of differences in passage number, these hESC lines responded to the enzymatic dissociation culture protocol in a similar manner. Following the first SCED passage, both homogeneous, undifferentiated colonies (Fig. 1A) and heterogeneous colonies, containing differentiated cells, were found (Fig. 1B). However, after 1–2 enzymatic passages, the hESCs had adjusted to their new culture environment, resulting in the disappearance of differentiated hESC colonies. No qualitative differences were observed between the two enzymes (TrypLE Select and trypsin-EDTA) used. However, consistently more hESC colonies were found in the groups passaged with TrypLE Select. This issue was further investigated in the colony-forming assay (as described below).


Figure 1
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  		Figure 1. Morphology of human ESCs (hESCs) cultured in the single-cell enzymatic dissociation (SCED) culture system. (A): Adjusting hESC colonies after their first enzymatic dissociation. Shown are areas with heterogeneous (undifferentiated + differentiated) cell populations observed during early adjustment. (B): Single hESCs in suspension during SCED passage. (C): Single-cell suspension of hESCs on human foreskin fibroblast feeder. (D): Adjusted hESCs, 2 days after enzymatic dissociation. (E): Adjusted hESC colonies. (F): Homogeneous hESC colony. (G): Culture well with hESC colonies, overview. Scale bar = 100 µm (A, B, G), 25 µm (C), 200 µm (D), 250 µm (E, F), 1.66 mm (H).

 
Culture and Expansion of hESCs Using SCED
At each SCED passage, all cells were enzymatically dissociated into single cells (Fig. 1C, 1D). If clusters remained, they were eliminated by trituration because cell clusters tended to give rise to heterogeneous/differentiated hESC colonies after seeding. Two to 3 days after enzymatic passaging, numerous small hESC colonies became visible in the culture dish (Fig. 1E). Unlike hESCs on MEF, the hESC colonies growing on HFF layers did not push the feeder cells aside but grew on top of the feeder layer (Fig. 1F, 1G). Colonies also appeared thinner than manually passaged hESC colonies grown on MEF feeders [29, 31–33]. All tested hESC lines grew as distinct undifferentiated colonies with clearly defined borders, and cells exhibited the characteristic morphology of hESCs, that is, densely packed cells with a high nucleus to cytoplasm ratio (Fig. 1F–1H). For all cell lines tested, a confluence of 25%–50% appeared to be optimal prior to SCED passaging. The split ratio at passage normally ranged from 1:4 to 1:40.

SCED-Passaged Cells Express Characteristic Markers for Undifferentiated, Pluripotent hESCs
To confirm the undifferentiated state of the hESCs, the cultures were evaluated immunohistochemically at approximately passages 5, 10, and 20. SA001 was also analyzed after 30 and at 40 enzymatic passages. At all time points, the cultures homogeneously expressed the expected markers of undifferentiated hESCs (Oct-4, TRA1-60, TRA1-81, SSEA-3, and SSEA-4), whereas they were completely negative for SSEA-1 (Figs. 2, 3; Table 1). The hESC colonies also displayed strong alkaline phosphatase activity (Figs. 2, 3; Table 1).


Figure 2
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  		Figure 2. Immunohistochemical staining of SA001 after 30 passages in the single-cell enzymatic dissociation culture system using TrypLE Select and trypsin EDTA. (A, C, E, G): SA001 TrypLE Select stained for Oct-4 (A), TRA-1-81 (C), SSEA-4 (E), and alkaline phosphatase (G). (B, D, F, H): SA001 trypsin-EDTA stained for Oct-4 (B), TRA-1-81 (D), SSEA-4 (F), and alkaline phosphatase (H). Scale bar = 100 µm (A–G), 250 µm (H).

 


Figure 3
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  		Figure 3. Characterization of SA121 after more than 20 passages in the single-cell enzymatic dissociation (SCED) culture system using TrypLE Select. (A): Oct-4 immunohistochemical staining; (B): TRA-1-60; (C): SSEA-3; (D): SSEA-4; (E): alkaline phosphatase. (F): Diploid normal karyotype of SA121 TrypLE after 20 SCED passages. (G–I): Teratomas derived from SA121 after 23 SCED passages. (G): Neuroectoderm (ectoderm); (H): cartilage (mesoderm); (I): secretory epithelium (endoderm). Scale bar = 100 µm (A–E), 50 µm (G–I).

 
Moreover, Oct-4, Nanog, and Cripto, but not AFP, mRNAs were expressed in SA001 after 24 and 35 passages with TrypLE Select or trypsin-EDTA, as well as in SA002 and SA167 after 15 passages with TrypLE Select (Table 1). In summary, SCED-passaged hESCs expressed the expected markers for undifferentiated, pluripotent hESCs, suggesting that SCED passaging supported maintenance of pluripotent hESCs.

SCED Passaging Did Not Result in Major Chromosomal Changes
Karyotype analyses revealed that all analyzed groups maintained a stable diploid normal karyotype. SA001 was passaged with TrypLE Select or trypsin-EDTA for more than 15 months (39 passages) in a diploid normal state (Fig. 4; Table 1). Consistently, SA121 was analyzed after 21 SCED passages (using both TrypLE Select and trypsin-EDTA), demonstrating that the cells remained diploid normal (Fig. 3; Table 1). FISH analyses on selected chromosomes (12, 13, 17, 18, 20 and 21, X, and Y) confirmed these findings (Fig. 4; Table 1).


Figure 4
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  		Figure 4. Karyotypes and fluorescence in situ hybridization (FISH) of SA001 after 25 passages in the single-cell enzymatic dissociation culture system using TrypLE Select (TrypLE) and trypsin-EDTA (TE). (A): The chromosomes from SA001 TrypLE were diploid normal. Shown is a representative karyotype. (B): SA001 TE, diploid normal. (C, D): FISH analysis of selected chromosomes from SA001 TrypLE (C) and SA001 TE (D) demonstrated that the cells were XY and diploid normal for chromosomes X (blue), Y (gold), 13 (red), 18 (aqua), and 21 (green).

 
SCED-Passaged hESCs Remained Pluripotent When Tested In Vivo
After more than 20 SCED passages of SA001 and SA121 with TrypLE Select or trypsin-EDTA, in vivo pluripotency was tested by transplanting clusters of undifferentiated hESCs under the kidney capsule of SCID mice (Table 1). Teratomas from all four SCED groups consisted of highly differentiated cells and tissues deriving from all three germ layers, possessing features undistinguishable from teratomas derived from their manually passaged counterparts [3, 29]. The appearance of ectodermal (neuroectoderm; Figs. 5A, 3G) mesodermal (cartilage; Figs. 5D, 3H), and endodermal (gut-like epithelium; Figs. 5E, 3I) tissues within the teratomas demonstrated that SCED-passaged hESCs exhibit the characteristic in vivo differentiation capacity of pluripotent hESCs. Comparable results were obtained with all four SCED groups (Table 1).


Figure 5
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  		Figure 5. In vivo pluripotency test of SA001. Teratomas were produced after more than 20 passages in the single-cell enzymatic dissociation culture system using TrypLE Select (TrypLE) and trypsin-EDTA (TE). Histological analysis of teratomas from SA001 TrypLE (A, C, E) and SA001 TE (B, D, F) after 27 respectively 22 enzymatic passages. (A, B): Neuroectoderm (ectoderm). (C, D): Cartilage (mesoderm). (E, F): Secretory epithelium (endoderm). Scale bar = 25 µm (A, C, E), 50 µm (B, D, F).

 
SCED Passaging with TrypLE Select Results in Increased Clonal Survival
To investigate whether the SCED culture system affects clonal survival of hESCs, we performed colony-forming assays. Traditionally cultured hESCs (cultured on MEFs and mechanically passaged) were enzymatically dissociated by either TrypLE Select or trypsin-EDTA to single-cell suspensions and plated into IVF dishes at 350 and 700 hESCs per cm2 on either HFFs or MEFs. Regardless of whether the cells were transferred to MEFs or HFFs, TrypLE Select treatment resulted in a threefold increase in the number of hESC colonies formed compared with trypsin-EDTA treatment (Fig. 6). After 1 week in culture, twice as many colonies formed when cells were seeded at 700 hESCs per cm2 compared with a seeding density of 350 hESCs per cm2, suggesting that these results were independent of the seeding density of hESCs.


Figure 6
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  		Figure 6. Single-cell enzymatic dissociation passaging with TrypLE Select resulted in increased clonal survival. When cells were transferred to HFFs after TrypLE Select treatment, the result was a threefold increase in the number of human ESC (hESC) colonies formed compared with trypsin-EDTA treatment (p = .001). If dissociated hESCs were plated on HFFs, significantly increased numbers of good colonies were obtained (p = .2) compared with plating them on mouse embryonic fibroblasts. The data are presented as the mean + SE (n = 3). Abbreviations: HFF, human foreskin fibroblast feeder; MEf, mouse embryonic fibroblast; TE, trypsin-EDTA.

 
Regardless of which enzyme was used, approximately 90% of the colonies on HFFs were graded as undifferentiated (Fig. 6). For comparison, if dissociated hESCs were plated on MEFs, significantly decreased numbers of undifferentiated colonies were obtained; only approximately 60% of the colonies were graded as undifferentiated (Fig. 6). The qualitative differences were similar in both dilutions tested. Thus, using HFFs as feeders and TrypLE Select for dissociation appears to be the most favorable combination to facilitate clonal survival of undifferentiated, pluripotent hESCs.


   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosure of Potential...
 Acknowledgments
 References
 
hESCs have traditionally been cultured on mitotically inactivated MEFs [1, 3]. At each passage, the hESC colonies are manually microdissected into pieces and transferred to new culture dishes. This culture method has been proven to be reliable and very stable and is used by many groups [1, 18, 19, 28]. Propagation for an extended time using splitting ratios up to 1:8 and a 7-day passage interval has been reported [34]. Furthermore, efforts have recently been made to partly automate the mechanical passaging procedure [35]. Nevertheless, this culture method is very labor-intensive and requires manual micromanipulation, and therefore it lacks a realistic capacity for up-scaling.

It is therefore not surprising that an increasing number of groups have started to use enzymatic passaging methods, which are superior to mechanical passaging with respect to speed and simplicity [6, 8, 9, 14–16, 36, 37]. Limitations are associated with these methods as well; for example, several groups use protocols that require dissociation of hESCs into small clusters [6, 9, 16, 36, 38]. This causes problems in standardizing and automating expansion and limits the applicability of these methods to several important experimental techniques, such as transfection by electroporation, clonal selection, fluorescence-activated cell sorting, and various high-throughput screens. Another important issue is the need to up-scale hESC cultures. The achievable split ratios reported for the different enzymatic passaging protocols are 1:2 every 3–7 days [16, 37], 1:3 every 4–7 days [14], and 1:5 to 1:10 [36] approximately every 7 days. Given the increasing demand for hESCs for different nonclinical applications alone, a protocol that allows the use of high splitting ratios when necessary would be highly beneficial. Recently, Hasegawa et al. [15] reported a method for selection of hESC sublines with high replating efficiency. They report split ratios of 1:50 to 1:100 every 7 days using MEFs as feeder. However, this method demanded a lengthy adjustment time (more than five passages) and appeared not to be applicable on all lines tested. Furthermore, abnormalities in karyotypes were noted in several of the sublines.

A problem that may arise when hESCs are adjusted to new culture environments is that the cultures may over time become populated with culture-adapted cells, that is, cells that gain a proliferate advantage under the prevailing conditions [9, 17, 19, 39, 40]. For example, stress caused by enzymatic dissociation under suboptimal culture conditions may lead to selection of abnormal cells or a gradual transformation to an in vitro-adapted cell line [19]. Such adaptive changes may be the results of genetic and epigenetic changes. However, causes for genetic changes of hESCs, which may arise under various growth conditions, are poorly understood [18, 41]. Importantly, the introduction of the SCED passaging method has so far not resulted in any obvious karyotypic abnormalities. This includes analysis of hESCs (SA001) cultured for more than 15 months in the SCED system using either TrypLE Select or trypsin-EDTA. Our genetic analyze comprises karyotyping and FISH of selected chromosomes. These methods would reveal a gain of cf. chromosomes 17q and 12, genetic abnormalities previously reported in relation to enzymatically dissociated hESCs [17]. Minor genomic alterations, however, would not be detected.

Whereas the stability of maintenance of euploid hESCs was independent of the enzyme preparation used, dissociation with TrypLE Select increased hESC colony formation by a factor of 3 compared with dissociation with trypsin-EDTA. Importantly, this occurred without compromising the maintenance of an undifferentiated, pluripotent phenotype. One possible explanation for the obtained result is that the increased enzyme purity of recombinant protein-based TrypLE Select compared with the porcine trypsin may benefit the clonal survival and growth of hESCs. Another possible explanation for the differences between the two tested enzymes could be that a relatively high concentration of trypsin-EDTA had been used. However, colony-forming assays comparing 1x trypsin/EDTA (0.25% trypsin) with that of 0.2x trypsin/EDTA (0.05% trypsin) did not reveal any differences among the different concentrations (data not shown).

We also found that the efficiency of the SCED culture system was enhanced using HFFs feeders compared to MEFs. HFFs have previously been shown to support undifferentiated growth of hESCs [23, 29, 31–33, 36]. It has also been shown that hESCs previously grown on MEFs easily adjust to growth on HFFs [29, 32, 36], and that HFFs tend to allow longer passage intervals without background differentiation [29, 32, 36]. These characteristics may be of extra importance in the SCED system, since colony formation per se starts from single cells at each passage, and therefore the feeders have to support the hESCs for a longer interval. Here, we show that the SCED system efficiently supports consecutive enzymatic dissociation of hESCs in a highly efficient manner and that the hESC population remains homogeneously undifferentiated.

By transferring several hESC lines to the SCED culture system, we confirmed that the adjustment of manually passaged hESCs to this new culture environment is a general phenomenon and is not restricted to selected hESC lines or history of mechanical passages (SA121, >100; all other lines, 40–50). Thus, the method should be widely applicable and of great value for the scientific community.

We routinely use the SCED system and cultivate hESCs in six-well dishes with approximately 1–2 x 106 hESCs per well. Based on the hESC line used and the need of expansion, we routinely split the cells 1:4–1:40 every 7–11 days. However, when necessary, the SCED system allows further scale-up, using split ratios of 1:100 to 1:200. When high split ratios were used, the passage interval was extended to 12–14 days without cell loss due to excessive spontaneous differentiation or cell death. The results from colony-forming assays performed directly at the transfer step from traditional culture to the SCED culture system suggest that the possibility of using high split ratios is not a result of a gradual adjustment, but more likely the result of a very supportive culture environment. In summary, the short time needed to adjust hESC lines from the traditional microdissection culture, the ability of the SCED system to support very high split ratios, and the efficient dissociation of hESCs to single cells will increase the versatility of hESCs as a basic research tool by facilitating scale-up, gene targeting (electroporation, cloning efficiency), multiwell plate assays, and cell sorting.


   DISCLOSURE OF POTENTIAL CONFLICTS OF INTEREST
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
Acknowledgments
 References
 
H.S. owns stock in, has acted as a consultant for, and has served as an officer or member of the Board for Cellartis.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
References
 
This work was supported by the Juvenile Diabetes Research Foundation, the Swedish Research Council, BetaCellTherapy EU sixth FP Integrated Project, Invitrogen, and Cellartis AB. The work related to hESC lines SA001 and SA002 (listed on the NIH Human Embryonic Stem Cell Registry) was partly supported by NIH Grant R24RR019514–01 awarded to Cellartis AB. We thank G. Caisander, K. Emanuelsson, and J. Synnergren for their expertise and assistance.


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosure of Potential...
 Acknowledgments
 References
 

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