Extensive contribution of embryonic stem cells to the development of an evolutionarily divergent host

Abstract

The full potential of embryonic stem (ES) cells to generate precise cell lineages and complex tissues can be best realized when they are differentiated in vivo—i.e. in developing blastocysts. Owing to various practical and ethical constraints, however, it is impossible to introduce ES cells of certain species into blastocysts of the same species. One solution is to introduce ES cells into blastocysts of a different species. However, it is not known whether ES cells can contribute extensively to chimerism when placed into blastocysts of a distantly related species. Here, we address this question using two divergent species, Apodemus sylvaticus and Mus musculus, whose genome sequence differs by ∼18% from each other. Despite this considerable evolutionary distance, injection of Apodemus ES cells into Mus blastocysts led to viable chimeras bearing extensive Apodemus contributions to all major organs, including the germline, with Apodemus contribution reaching ∼40% in some tissues. Immunostaining showed that Apodemus ES cells have differentiated into a wide range of cell types in the chimeras. Our results thus provide a proof of principle for the feasibility of differentiating ES cells into a wide range of cell types and perhaps even complex tissues by allowing them to develop in vivo in an evolutionarily divergent host—a strategy that may have important applications in research and therapy. Furthermore, our study demonstrates that mammalian evolution can proceed at two starkly contrasting levels: significant divergence in genome and proteome sequence, yet striking conservation in developmental programs.

INTRODUCTION

Pluripotent ES cells are capable of indefinite self-renewal as well as differentiation into all derivative lineages of the three embryonic germ layers (1). Their potential application in cell-replacement therapy has generated considerable excitement in recent years and has spurred intense research aimed at finding appropriate in vitro conditions for differentiating them into desired cell types (2). However, in vitro differentiation has several major limitations. First, in vitro conditions are a far cry from the enormously complex in vivo microenvironment that cells encounter in the course of development. As such, many cell types may be very difficult to derive in large quantities in vitro, and even for the cell types that can be derived, it is uncertain whether they are truly identical to their in vivo counterparts. Second, the differentiation of ES cells in vitro always stands the possibility that some cells remain undifferentiated or have differentiated into unintended cell types, which may pose significant problems to both research and therapy (3). Finally, it seems unlikely that in vitro conditions could lead to the generation of highly complex tissues and organs from ES cells, a situation that would limit their therapeutic potential only to diseases that could benefit from the transplantation of unstructured cells.

One solution to these limitations is to differentiate ES cells in an in vivo environment—i.e. placing them into blastocysts where they can differentiate through the normal course of embryonic development. However, due to various ethical and practical constraints, ES cells from some species cannot be routinely placed into blastocysts of that same species. For example, it is ethically impossible to differentiate human ES cells in vivo in the context of human embryos, though in vivo differentiation should in theory provide many opportunities for research and the therapeutic use of human ES cells. Similarly, although it may be feasible to obtain ES cells from certain wild and/or endangered species (such as the giant panda or the chimpanzee) for the purposes of basic research and conservation, it is highly impractical to obtain large quantities of blastocysts from these species, and it is also impractical to use these species as surrogates for carrying manipulated blastocysts to term. We therefore decided to explore methods of differentiating ES cells in vivo by introducing them into evolutionarily divergent host blastocysts. Such methods could facilitate many studies of the basic biology and therapeutic potential of ES cells in ways not accessible to in vitro means.

Several reports have demonstrated the feasibility of creating interspecies chimeras by aggregating early embryos from two species. Techniques employed in these studies include morula–morula aggregation, inner cell mass (ICM) and morula aggregation and ICM injection into blastocysts (4–13). Of these, viable chimeras have been obtained for only three pairs of species: between Mus musculus and Mus caroli (two laboratory mouse species), between Ovis aries (sheep) and Capra hircus (goat) and between Bos taurus (cow) and Bos indicus (zebu) (9–13). For each of these studies, the two species in the pair are rather closely related in terms of evolutionary distance, differing in DNA sequence by no more than 3% (Fig. 1A). Furthermore, for each pair, the two species can still mate and produce viable interspecies hybrids, though only rarely between sheep and goat (14–18). Thus, it is unclear whether viable chimeras from two species separated by a much greater evolutionary distance could be made by embryo aggregation. Furthermore, these studies did not involve ES cells, and therefore do not address whether ES cells could be used to create viable interspecies chimeras. Although human ES cells have been injected into mouse blastocysts, it did not result in viable chimeras and there was no demonstration of proper differentiation of the ES cells (19). Thus, it remains an open question as to whether the injection of ES cells from one species into the blastocysts of a distantly related species could lead to the development of viable chimeras.

Figure 1.

Phylogenetic relationship between A. sylvaticus and other mammalian species. Horizontal lengths of the solid branches are scaled to DNA sequence divergence, whereas lengths of the dashed branches are arbitrary. Scale bar represents 5% DNA sequence divergence. (A) Evolutionary distance between mammalian species for which viable interspecies chimeras have been obtained by aggregating early embryos as done in previous studies or injecting ES cells into blastocysts as done in this study. The ability of these species to interbreed and produce interspecies hybrids through mating is also indicated. Abbreviations for methods of chimera production are as follows: ES-B, ES cell-to-blastocyst injection; M–M, morula–morula aggregation; ICM-B: ICM-to-blastocyst injection. (B) Evolutionary distance among A. sylvaticus, M. musculus, human, chimpanzee and rhesus macaque.

Here, we address this question by constructing interspecies chimeras between A. sylvaticus and M. musculus, two species separated by a rather large evolutionary distance. The sequence divergence between Apodemus and Mus is ∼18% (see Results), whereas it is no more than 3% for previously reported viable chimeras made by embryo aggregation (9–13). Importantly, the two species in our study are far too divergent to produce hybrids through mating. This is in contrast to the species employed in previous reports of viable chimeras where the two species used for embryo aggregation are sufficiently related that they can still mate and produce viable hybrids, at least in some cases. We show that the injection of Apodemus ES cells into Mus blastocysts could produce healthy chimeras bearing extensive contributions from both species in all major organs, including the germline. Our results demonstrate the feasibility of properly differentiating ES cells in vivo by introducing them into evolutionarily highly divergent host blastocysts. Our data also highlight the rather striking conservation of developmental programs between some mammalian species, despite their considerable divergence in DNA sequence.

RESULTS

Phylogenetic relationship between Apodemus and Mus

Apodemus sylvaticus, known by its common name as wood mouse, belongs to an entirely different genus as the house mouse M. musculus. The large evolutionary distance between these two species is immediately evident at the karyotypic level: A. sylvaticus has 48 metacentric or acrocentric chromosomes, whereas M. musculus has 40 telocentric chromosomes, and there are numerous chromosomal rearrangements between these two species (20). At the DNA sequence level, Apodemus and Mus differ by ∼18% as estimated from neutral sites of genes (see Materials and Methods for details). This is far greater than the 1–3% sequence divergence separating the species used in previous reports of viable interspecies chimeras made with embryo aggregation (Fig. 1A). In comparison, genome divergence is ∼1.5% between human and chimpanzee, ∼8% between human and the Old World monkeys (such as macaque or baboon), ∼20% between rat and mouse and ∼45% between human and mouse (Fig. 1B).

Derivation of Apodemus ES cells

To derive ES cells from A. sylvaticus, we retrieved 68 blastocyst-stage embryos from 17 Apodemus females 3.5 days post coitum. Of these, 34 embryos were cultured on mouse embryonic fibroblast (MEF) feeder cells for 1–2 days until hatching. After 3–4 days, the outgrowth from the ICM of each embryo was picked, dissociated and transferred onto fresh feeder. After 4–7 days, colonies became visible, which were dissociated and plated again onto fresh feeder. For two embryos from two separate matings, numerous colonies were observed several days later. These colonies resembled mouse ES cell colonies: they were dome-shaped with smooth and clearly defined borders, and cells within the colonies had a high nucleus-to-cytoplasm ratio, prominent nucleoli and a compact morphology (Fig. 2A, left panel). We report results from one of these two independent lines, named AS-ES1, where AS denotes A. sylvaticus.

Figure 2.

Images of Apodemus ES cells and Apodemus–Mus chimeras. (A) Left: phase-contrast image of AS-ES1 colonies; middle panel: green fluorescence image of AS-ES1-hrGFP1 colonies; right: red fluorescence image of AS-ES1-DsRed colonies. (B) Images of A. sylvaticus, M. musculus B6, and four different chimeras. (C) Green fluorescence images of two different chimeras created with AS-ES1-hrGFP1 cells.

Characterization of Apodemus ES cells

Similar to mouse ES cells, both leukemia inhibitory factor (LIF) and feeder cells promote the undifferentiated state of AS-ES1. In the absence of LIF but presence of feeder, AS-ES1 grew well and remained undifferentiated. When cells were grown in the presence of LIF but absence of feeder, they could form colonies, but the colonies were smaller and the morphology was more variable. In the absence of both LIF and feeder, cells differentiated completely.

Under our culture conditions, population doubling time was ∼12 h for AS-ES1 and about 17 h for a number of commonly used mouse ES cell lines in our possession such as E14Tg2a (data not shown). Thus, AS-ES1 appears to proliferate somewhat faster than the typical mouse ES cell lines.

Karyotype analysis was performed at several time points between passages 24 and 80, all revealing the presence of 48 chromosomes—the correct number for euploid Apodemus cells (Supplementary Material, Fig. S1A). PCR assays specific for the Y-linked Apodemus gene Tspy showed the presence of the Y chromosome in AS-ES1 (Supplementary Material, Fig. S1B), indicating that it is a 48 (X,Y) male line.

We examined AS-ES1 for the expression of several stem cell markers (Supplementary Material, Fig. S2). It is positive for alkaline phosphatase (ALP), Oct4, Nanog and Rex1, markers found to be positive in all previously examined ES cells, but negative for SSEA-1, SSEA-3, SSEA-4, Tra-1–60 and GCTM-2, markers associated with some but not all previously described ES cell lines (21). We note that the negativity of the immunohistochemistry-based SSEA markers may be due to species differences in the orthologous epitopes in Apodemus rather than the absence of the epitopes per se. Supplementary Material, Table S1 compares the expression status of the various markers in AS-ES1 with that found previously in mouse and human ES cells.

Under appropriate culture conditions, AS-ES1 cells readily produced both simple and cystic embryoid bodies (Supplementary Material, Fig. S3A). When allowed to further differentiate, these embryoid bodies gave rise to heterogeneous populations of cells that expressed markers of all three germ layers (Supplementary Material, Fig. S3B). When injected into immunodeficient nude mice, AS-ES1 cells produced teratomas containing derivatives of all three germ layers (Supplementary Material, Fig. S4).

AS-ES1 was passaged continuously for over 6 months (about 80 passages) without any detectable change in morphology, rate of proliferation, karyotype, expression of key markers or the potential of the cells to differentiate in vitro or in vivo (data not shown).

We stably transfected AS-ES1 cells with two plasmids, one expressing the green fluorescent protein hrGFP and the other expressing the red fluorescent protein DsRed, both under the control of a ubiquitous promoter. This resulted in two clonal lines, AS-ES1-hrGFP1 and AS-ES1-DsRed1 (Fig. 2A, middle and right panels), which were subjected to additional characterization. These transgenic lines grew normally and exhibited typical undifferentiated morphology in long-term culture (over 20 passages). Furthermore, they can produce embryoid bodies and teratomas (data not shown). However, teratomas made from AS-ES1-DsRed1 showed silencing of DsRed expression in ∼10% of the cells (this phenomenon was not seen in AS-ES1-hrGFP1 teratomas). This may be related to the previously reported cytotoxicity of DsRed (22). We also tested whether Apodemus ES cells were conducive to RNA interference (RNAi). AS-ES1-hrGFP1 cells were treated with small interfering RNA (siRNA) that targets the hrGFP coding region. One day after treatment, the level of hrGFP fluorescence was dramatically reduced (Supplementary Material, Fig. S5), indicating that the cells responded well to RNAi.

Generation of Apodemus–Mus chimeras

The original unmodified AS-ES1 cell line and the derivative AS-ES1-hrGFP1 and AS-ES1-DsRed1 sublines were all used for the generation of chimeras. Cells were in passages 40–60 at the time of injection. Recipient embryos were 3.5-day-old blastocysts from M. musculus. The Apodemus animals used to derive ES cells have light brown fur, whereas the recipient blastocysts should grow black fur (Fig. 2B). A total of 1250 embryos were injected with about 15 ES cells each and transferred into 44 surrogate ICR mice (Table 1). From these, 220 pups were born and 16 of them, or 7.3%, showed chimerism based on patchworks of light brown fur mixed with black fur (Fig. 2B). Of the 16 chimeras, three were from AS-ES1, four from AS-ES1-hrGFP1 and nine from AS-ES1-DsRed1 (Table 1). The Apodemus contribution to fur color ranged from ∼10 to ∼40% in the chimeras. All the chimeras appeared overtly healthy and thrived alongside their littermates. We confirmed the presence of Apodemus cells in chimeras by PCR assays that specifically amplified Apodemus but not Mus DNA (data not shown). Morphologically and behaviorally, the chimeras appeared to resemble both species to varying degrees. For example, Apodemus has much larger eyes than Mus, and some chimeras had eyes that seemed intermediate in size between the two species (Fig. 2B). The level of nervousness displayed by some chimeras toward handling also seemed intermediate between the very jumpy Apodemus and the much tamer Mus. However, it is difficult to statistically quantitate these phenotypes due to the limited number of chimeras and the variation in the extent of chimerism from animal to animal.

Table 1.

Summary of Apodemus–Mus chimera production

ES cell line	Number of blastocysts injected	Number of surrogates	Number of pregnant surrogates	Number of pups	Number of chimeras	Percent of chimeric pups
AS-ES1	434	16	10	68	3	4.40
AS-ES1-GFP1	383	14	10	67	4	6.00
AS-ES1-DsRed1	403	14	13	85	9	10.60
Total	1220	44	33	220	16	7.30

Chimerism is determined by coat color.

Extensive chimerism in all major organs

For chimeras made with AS-ES1-hrGFP1 or AS-ES1-DsRed1 cells, fluorescence of the skin prior to the growth of fur allowed for a visually much more striking presentation of the chimerism (Fig. 2C). Fluorescence of Apodemus cells also allowed us to examine the extent of chimerism in internal organs. Given the silencing of DsRed transgene noted earlier, we will only present fluorescence data from chimeras made with AS-ES1-hrGFP1 cells. Figure 3 shows the green fluorescence in all the major organs prior to sectioning. Examination of sections from these tissues at the cellular level further confirmed the abundance of fluorescent cells (Supplementary Material, Fig. S6). The contributions of Apodemus-derived fluorescent cells were estimated to range from a few percent to ∼40% in the various tissues (excluding skeletal muscle). For skeletal muscle, the fraction of fluorescent cells was nearly 100% in some muscles and 50–70% throughout the body. This is presumably due to the fact that Apodemus- and Mus-derived myocytes have fused into single myotubes in many cases.

Figure 3.

Green fluorescence images of unsectioned organs in the chimera. Two images are presented for each organ, the left from chimera and the right from negative control B6 animal.

The 16 chimeras included 13 males, two females and one intersex individual. Sexing of the animals using Mus-specific PCR primers showed that all 13 males had a Mus-derived Y chromosome, whereas the two females did not. It is not clear why there is a bias for males, though this may simply be a coincidence. The one intersex animal does not carry a Mus-derived Y chromosome and is therefore likely the result of a female blastocyst that was partially masculinized during development by the injected Apodemus male ES cells.

Cellular identity of Apodemus-derived cells

We next performed immunofluorescence (IF) staining to examine the identity of Apodemus-derived cells in a wide range of tissues (Fig. 4). We first examined the three major types of neural cells: neurons, oligodendrocytes and astrocytes. For neurons, we performed IF staining of cortical sections with antibodies against MAP2 and NeuN, markers for processes and cell bodies of mature neurons, respectively. Both showed co-expression with hrGFP in many cells. For oligodendrocytes, we stained both the cortex and the corpus callosum with an anti-oligodendrocytes antibody and detected numerous cells positive for both this marker and hrGFP. For astrocytes, we stained against GFAP in the cortex and found hrGFP-positive cells that also expressed this marker, though the frequency of such cells is lower than hrGFP-positive neurons or oligodendrocytes, at least in the sections we examined. To investigate whether Apodemus ES cells had differentiated into dopaminergic neurons, we stained against tyrosine hydroxylase (TH) in the basal ganglia. Co-expression of TH and hrGFP was seen in some axon terminals, indicating the presence of Apodemus-derived dopaminergic neurons. We also stained cortical sections with antibody against PAX6, a marker for adult neural stem cells. Co-expression of PAX6 and hrGFP could be seen in isolated cells of the subventricular zone. In heart sections, the cardiomyocyte marker troponin 1 showed co-expression with hrGFP in patches of cells. In skin sections, staining of K14, a marker for stratifying epithelial cells, showed co-expression with hrGFP in many cells. Finally, we stained against the endothelium marker vWF in small intestines, which revealed co-expression with hrGFP in the vasculature. Thus, Apodemus ES cells have contributed abundantly to all the tissues examined, where they differentiated into a wide range of cell types appropriate for the tissue.

Figure 4.

IF characterization of the cellular identity of Apodemus-derived cells in chimeras. Each section has four images in a row, which are (from left to right) IF staining using the antibody indicated, GFP fluorescence marking Apodemus-derived cells, DAPI marking nuclei and merged image. The co-localization of IF signal and GFP indicates that Apodemus-derived cells have differentiated into cell types identified by IF. For merged images where co-localization between IF signal and GFP is not immediately obvious, arrows are used to indicate such co-localization. MAP2 and NeuN antibodies stain processes and cell bodies of neurons, respectively; anti-oligodendrocytes antibody stains oligodendrocytes and their myelin sheaths; GFAP antibody stains cell bodies of astrocytes; TH antibody stains axon terminals of dopaminergic neurons; PAX6 antibody stains cell bodies of adult neural stem cells; troponin 1 antibody stains cell bodies of cardiomyocytes; K14 stains cell bodies of epithelial cells; and vWF antibody stains cell bodies of endothelium. MAP2, NeuN, anti-oligodendrocytes, GFAP and PAX6 staining was performed on cortical sections. TH, Troponin 1, K14 and vWF staining was performed on basal ganglia, heart, skin and small intestine sections, respectively.

Chimerism in the germline

To examine whether there is chimerism in the germline, we purified sperm samples from two male chimeras, extracted DNA from the samples, and performed PCR using primers specific for either Apodemus or Mus (see Materials and Methods). Mus-specific PCR produced positive results for both animals. Apodemus-specific PCR produced positive result for one of the two animals, indicating that there is Apodemus contribution to the sperm sample. To gauge the proportion of Apodemus contribution, we performed competitive PCR using primers common to both Apodemus and Mus, followed by digestion with a restriction enzyme that only cuts the Apodemus-derived product due to sequence differences between the two species (see Materials and Methods). Quantitation of the digested PCR product showed that Apodemus contribution is ∼5% (Fig. 5). At this level of contribution, it is highly unlikely to be the result of contamination by somatic cells (see Materials and Methods). This result was confirmed two more times with independent sperm samples. Microscopic examination of the sperm showed that they all appeared grossly normal, and it was not possible to differentiate Apodemus sperm from Mus sperm by morphology. As yet, we do not know if the Apodemus contribution to the chimera's sperm sample represents fully functional sperm capable of fertilization.

Figure 5.

Quantitation of Apodemus contribution in chimera sperm sample. PCR was performed on DNA from purified chimera sperm sample, as well as a series of control samples (which were mixtures of A. sylvaticus and M. musculus genomic DNA with the percentage of Apodemus contribution ranging from 0 to 100%). PCR primers were designed to amplify both Apodemus and Mus DNA. PCR products were digested with a restriction enzyme specific for Apodemus sequence.

Apodemus ES cells form teratomas in chimeras

In chimeras, the contribution of Apodemus ES cells to various tissues should, in theory, create immune tolerance in these animals toward Apodemus cells. We found that, consistent with this prediction, Apodemus ES cells readily formed teratomas in the chimeras but not in non-chimeric control mice. Histological examination of the teratomas revealed derivatives of all three germ layers (data not shown). Thus, the chimeras indeed show immune tolerance towards many, and perhaps all, Apodemus cell types. This further supports the notion that Apodemus ES cells have differentiated into a wide range of cell types in the chimeras and elicited immune tolerance in the chimeras to Apodemus-derived cells.

DISCUSSION

Our results are notable in several regards. The first is the ability of Apodemus ES cells to fully integrate with the ICM of Mus blastocysts to produce viable chimeras, despite the considerable evolutionary distance separating these two species. The second is the extensiveness of chimerism in all the organs examined—as high as ∼40% Apodemus contribution in some tissues. The third is the correct differentiation of the ES cells into a wide range of cell types intimately integrated into the host tissues. The fourth is the relatively high frequency of chimeras (judging from coat color) among liveborn animals. These results demonstrate the feasibility of differentiating ES cells into a wide range of cell types in vivo by introducing them into an evolutionarily divergent host. This interspecies approach may be the only way to study ES cells of some species, such as human ES cells, in an in vivo context. One caveat in our results is the possibility that the Apodemus cells might have fused with Mus cells, and that this fusion is responsible for the observed chimerism. Although, we have not formally ruled out this possibility, we believe that it is highly unlikely in light of the extensiveness of the Apodemus contribution to the chimeras.

Further genetic manipulation of either the ES cells or the host blastocysts could in theory increase or decrease the degree to which ES cells contribute to a particular tissue or organ. For example, if the host blastocysts are engineered to carry genetic defects that block the development of a particular tissue (e.g. Pdx-1 mutation which leads to the agenesis of the pancreas), the percentage contribution of the ES cells to the affected tissue may increase dramatically. Conversely, if the ES cells are engineered to carry genetic defects that prevent their differentiation down a particular lineage, then the ES cells would not contribute to that lineage in the chimeras. For example, contribution of ES cells to the brain could be minimized by knocking out a gene in ES cells essential for neural differentiation. The Apodemus–Mus chimera system thus provides an informative model for investigating the means to manipulate the degree to which ES cells contribute to a particular tissue in interspecies chimeras, which may have important research and therapeutic implications.

Besides relevance to the study and therapeutic application of ES cells, our results also offer important evolutionary insights. As two species descend from a common ancestor and evolve apart from each other, their genomes become increasingly divergent due to the accumulation of mutations along the two lineages. However, it remains unclear how divergence in genome sequence translates into divergence in developmental programs. For the two species employed in this study, their genomes differ by ∼18% in DNA sequence, which means that protein sequences differ by ∼12% between them. This translates into ∼5.5×10⁸ nucleotide differences and ∼1.5×10⁶ amino acid differences between the two species. Yet, Apodemus ES cells and Mus blastocysts can combine and develop into one functionally coherent organism without any overt defect. Given that cells from the two species are combined at a very early stage of embryogenesis, both populations of pluripotent cells are completely unstructured when combined and must progress through the entire schedule of embryogenesis, including gastrulation, axial patterning, organogenesis and histogenesis, while remaining tightly integrated with each other. This means that the two populations of pluripotent cells, despite their dissimilarities in genome and proteome sequence, can properly engage each other in the myriad cell–cell interactions and cell–cell signaling events needed to specify cell lineages and regulate cell proliferation, migration, differentiation and apoptosis. As such, the developmental programs driving embryogenesis must be highly conserved between the two species. The Apodemus–Mus chimera system thus provides an informative platform for studying the evolution of development.

We note that our study involves the construction of primary chimeras, which are fundamentally different from secondary chimeras. Primary chimeras are created when early embryos containing pluripotent stem cells (or their pluripotent derivative cell lines such as ES cells) from two individuals are merged before any structure has formed in the embryo and then allowed to develop into a fully structured organism. Secondary chimeras, in contrast, are the result of tissue transplants performed during much later stages of embryonic development or in adults. One common example of secondary chimeras is chick–quail chimeras, which are typically made by grafting post-gastrulation fetal tissues from quail onto developing chick fetus (chick and quail genomes differ by ∼6% in DNA sequence). At these later stages of development, the embryos are already highly structured and the cells of the embryos have already gone a long way in their proliferation, differentiation and migration. The ability to graft tissues at these stages is therefore a much weaker indicator of developmental conservation between species.

In their classic 1975 paper, ‘Evolution at two levels in humans and chimpanzees’, King and Wilson (23) reported the remarkable genetic similarities between humans and chimpanzees (only ∼1.5% DNA sequence divergence), despite their apparent phenotypic dissimilarities. This finding has since been cited as a prime example of selection driving the rapid divergence of developmental programs in mammals. Our results demonstrate that there is another perspective to interpret ‘evolution at two levels’—i.e. significant genetic divergence and yet striking developmental conservation between at least some mammalian species. As such, our study reveals important principles underlying the evolution of developmental programs in mammals in addition to its relevance to stem cell biology.

MATERIALS AND METHODS

Animal husbandry

Apodemus sylvaticus is a native rodent species to Europe that is found commonly in forests, grasslands and cultivated areas. An outbred colony was originally established at Oxford University, UK, from wild-caught animals, and animals from there were used to establish a colony at the University of Liverpool, UK, in 1992. About 20 animals were transferred to the University of Chicago in 2003 to establish another colony with which the present experiments were performed. Animals were maintained in a standard animal facility on a 14/10 light/dark cycle to mimic the summer time to promote breeding. They were fed the standard diet as used for laboratory mice. In captivity, the animals breed year round with a gestation time of 19–21 days and an average litter size of four to five pups. Sexual maturity occurs 7–9 weeks after birth for males and 6–8 weeks for females. Like mice, pregnancy can be timed by the presence of copulatory plugs.

Analysis of evolutionary divergence

Previous phylogenetic studies have shown that the evolutionary distance between Apodemus and Mus is similar to the distance separating Rattus and Mus (24–26), with the Rattus–Mus genome divergence at neutral sites being ∼20% (27). To confirm this, we obtained A. sylvaticus gene sequences by cDNA sequencing and searching GenBank. Orthologs in species of interest were obtained from ENSEMBL and GenBank using reciprocal best BLAST hits. Synonymous substitution rate (K_s) between orthologs, which approximates the substitution rate at neutral sites, was calculated for each gene (Supplementary Material, Table S2). The calculation of K_s, which corrects for multiple hits, is based on the standard Li method (28) as implemented by the Diverge function in the Wisconsin Package 10.2 (Accelrys Inc.). The average K_s from our data is 0.16±0.016. When further corrected for local insertions and deletions, this produced a sequence divergence between Apodemus and Mus at ∼18%, consistent with previous estimates (24–26). Although our method is designed to estimate the sequence divergence at neutral sites, the resulting value should be a reasonable approximation for the entire genome because the great majority of the genome evolves neutrally. This estimate does not consider large-scale genomic rearrangements such as translocations, interstitial deletions and segmental duplications. Such large-scale rearrangements must be numerous between Apodemus and Mus given the very different chromosome number and karyotypic structure between them. Our estimate of ∼18% sequence divergence between Apodemus and Mus is therefore conservative.

Derivation of Apodemus ES cells

Apodemus mating pairs were set up between young adults and pregnancy was ascertained by the presence of copulatory plugs. We did not use superovulation because the animals did not seem to respond to superovulation drugs. Blastocyst-stage embryos were obtained by flushing the uteri of females 3.5 days post-coitum with M2 medium. Embryos were then plated on a 24-well dish in ES cell medium, with the bottom of wells covered with a feeder layer of radiation-arrested MEF carrying multiple antibiotic resistance including puromycin (Cyagen Biosciences). The ES cell medium was high-glucose MEM-supplemented with 10% fetal bovine serum, 100 U/ml recombinant murine LIF, 0.05 mm β-mercaptoethanol, 1 mm sodium pyruvate, 1× non-essential amino acids and penicillin/streptomycin (Cyagen Biosciences). Embryos were kept in ES cell medium for 1–2 days till they hatched from the zona pellucida and attached to the feeder layers. Shortly thereafter, the trophectoderm of the embryos flattened and the ICM grew out. After 3–4 days of culture, the outgrowth from the ICM was picked up, away from the flat trophoblast cells, dissociated into clumps by trituration and replated onto fresh feeder in ES cell medium. Within 4–7 days, colonies with the morphology of undifferentiated ES cell (i.e. dome-shaped with a smooth surface) were observed. They were dissociated with trypsin and reseeded to fresh feeder layers in ES cell medium again. Culture dishes were kept at 37°C in humidified atmosphere with 5% CO₂. The ES cells were passaged every 2 days by trypsin and a change of culture medium every day with fresh ES cell medium.

Marker expression in Apodemus ES cells

To detect alkaline phosphatase activity, ES cell colonies were fixed in 4% paraformaldehyde (PFA) and stained with the alkaline phosphatase substrate BCIP/NBT. For immunocytochemistry, ES cell colonies were fixed with 4% PFA, permeabilized with 0.1% Triton X-100 and blocked with 10% goat serum. Cells were first incubated with primary antibodies against SSEA-1 (Developmental Studies Hybridoma Bank; 20× dilution), SSEA-3 (Developmental Studies Hybridoma Bank; 20× dilution), SSEA-4 (Developmental Studies Hybridoma Bank; 20× dilution), Tra-1-60 (Developmental Studies Hybridoma Bank; 20× dilution), GCTM-2 (from Dr. Martin Pera; 20× dilution), or OCT4 (Santa Cruz Biotechnology; 400× dilution), followed by incubation with FITC-conjugated secondary antibodies (Jackson ImmunoResearch). Specimens were mounted in Vectashield with DAPI (Vector Laboratories) for examination. Staining with the secondary antibody alone was used as negative controls.

To study gene expression by RT–PCR, cDNA was made from ES cells, followed by PCR amplification. The Apodemus genes targeted and their primer sequences were as follows: Oct4 (GAGGGAACCTCCTCTGAGCCCTGTGC and TCCTCCACCCACTTCTCCAGCAGGG), Nanog (AAGCAGAAGATGCGGACTGTGTTC and CTTCCAGATGCGTTCACCAGATAGC), Rex1 (GTGTTGTCCCCAAATACCACTGACC and ACTCACCTCGTATGATGCACTCTAGG) and Gapdh (AGAAGGTGGTGAAGCAGGCATC and GGGTGGTCCAGGGTTTCTTACTC).

Karyotype and PCR analysis of Apodemus cells

To make metaphase chromosome preparations, ES cells in growth phase were arrested with colcemid, dissociated with trypsin, fixed in methanol/acetic acid (3:1) and spread onto pre-chilled glass slides. The chromosome spreads were stained with Giemsa solution and photographed. At least 20 metaphase spreads were analyzed. The sex of Apodemus ES cells was determined by standard PCR amplification of the Y-linked gene Tspy (CTCCTGCTGGGGACAATC and CTCATTCTTGACATCCACCAC) from genomic DNA. As a control, PCR was also performed to amply the X-link gene PolA1 (TCCAGAAGATGAGCAGGAG and GCATCCAGCCAATAAAACTGAA) and the autosomal gene Gapdh (GGCATTGCTCTCAATGACAA and CTTGCTCAGTGTCCTTGCTG). PCR products were sequenced and shown to be distinct from M. musculus sequences. To confirm the presence of Apodemus cells in chimeras by PCR, two pairs of primers specific to Tspy in Apodemus were used (CAGATGGTGAAGCCTCTGGT and CCTGCTTGTGCTCCTTTACC, and GAGGTTGCTAAGGGGGAGTT and GTCCCACCACCTGACTGTG). Two pairs of primers specific to Tspy in Mus were used as PCR control (GCCCCAATATAGGCATCAGA and AACTTTAAATAGCAGCCTTGTCC, and GGGGACAAGGCTGCTATTTA and CATTCCACAGGCCTACCATC). To obtain sperm sample from the chimeras, they were mated with wild-type B6 females. Uteri were removed from plugged animals and sperm samples flushed. The recovered sample was briefly centrifuged to remove tissue debris and contaminating somatic cells before DNA extraction. To quantify the amount of Apodemus contribution in the sperm sample, we amplified a region of the GPR33 gene using a pair of primers whose sequences are common to both Apodemus and Mus orthologs (GGCCACCAAGATGAAAGAGA and TCAGGGTTTGTGTCCTTTCTG). We then digested the PCR product with the restriction enzyme PmlI, which only cuts Apodemus PCR product into two fragments. A set of control samples containing mixtures of Apodemus and Mus genomic DNA (with Apodemus contribution ranging from 0 to 100%) were also amplified and digested. The control samples allowed the quantitation of Apodemus contribution in the chimera sample using Image J software.

Genetic manipulation of Apodemus ES cells

We used two plasmids to transfect Apodemus ES cells: pTP6-hrGFP and pTP6-DsRed2, which express hrGFP and DsRed, respectively, under the control of the ubiquitous CAGG promoter as described previously (29). For stable transfection, 5×10⁶ undifferentiated ES cells were trypsinized and washed once in ES cell media and twice with PBS. The two plasmids were linearized by ScaI. Cells were then electroporated with 30 µg of linearized DNA at 800 V, 3 µF and a time constant of 0.4 s, using the Gene Pulser II System (Bio-Rad). These constructs also confer puromycin resistance. After growing the electroporated cells in ES cell medium for 48 h without selection, puromycin (2 µg/ml) was added. The MEF feeder cells were not affected by the drug because they already carry a puromycin-resistant transgene (Cyagen Biosciences). ES cell colonies surviving drug selection were picked after 7 days of selection, trypsinized and replated onto fresh feeder layers in ES cell medium. For RNAi, AS-ES1-hrGFP1 cells were passaged onto six-well plates at low density. Cells were transfected with double-stranded siRNA oligonucleotides specific for hrGFP (target mRNA sequence: gcgacaucaaccugaucga; siRNA sequence: gcgacaucaaccugaucga-dCdA and ucgaucagguugaugucgc-dCdA). A negative control was also included (siRNA sequence: uucuccgaacgugucacgu-dTdT and acgugacacguucggagaa-dTdT). Transfection of siRNA was performed by Lipofectamine 2000 (Invitrogen) and cells were re-transfected after 24 h. Fluorescence microscope analysis was performed 48 h after the second transfection.

Formation of embryoid bodies and teratomas

For embryoid body formation, ES cells were dissociated to single cells and plated onto non-adherent Petri dishes in ES cell medium without LIF for 2 weeks to form embryoid bodies. Embryoid bodies were then plated onto the adherent surface of tissue culture dish in ES cell medium without LIF to induce differentiation. The resulting differentiated cells were probed with antibodies against albumin (Dako, A0001; 50× dilution), desmin (Neomarkers, MS-376-R7; 100× dilution), nestin (Chemicon, MAB353; 100× dilution), GFAP (Dako, Z0334; 500× dilution) and beta-III Tubulin (Tuj1; R&D Systems, MAB1195; 100× dilution) and followed by incubation with Cy3-conjugated goat anti-mouse secondary antibodies (Jackson ImmunoResearch; 200× dilution) or R-phycoerythrin (R-PE)-conjugated goat anti-rabbit secondary antibodies (SouthernBiotech; 200× dilution). For teratoma formation, ∼1–2×10⁶ undifferentiated ES cells were injected subcutaneously into 4–6-week-old nude mice. About 8 weeks later, teratoma was removed and fixed in 4% PFA in PBS. Paraffin sections were prepared using standard protocols and examined histologically after hematoxylin and eosin staining.

Generation of chimeras

ES cells were grown to near confluency in a 35 mm Petri dish and fed with fresh ES cell medium 3–4 h prior to injection. They were trypsinized into single cells. The suspension was then plated onto a dish coated with gelatin. The dish was returned to the incubator for 1 h to allow feeder fibroblasts to attach and viable ES cells to begin to adhere to the surface. Non-adherent cells were gently aspirated off, and loosely adhering ES cells (which were viable) were gently washed off with a pipette using culture medium. Cells were then centrifuged, suspended in 150 µl M2 medium and placed on ice ready for injection. B6 female mice 4–5 weeks old were injected with PMS between 1 and 2 p.m. and with hCG 46–48 h later. After the administration of hCG, one female was placed in a cage with one stud B6/DBA F1 male. Blastocysts were flushed using M2 medium from plugged females 3.5 days post-coitum. About 20 blastocysts were introduced into a microdrop of 20 µl M2 medium under oil in the injection chamber, followed by the introduction of 1 µl of ES cell suspension and 1 µl of DNase I (1 mg/ml). About 15 ES cells with good morphology (small, round and with sharp boundary) were injected into each blastocyst. About 20–25 injected blastocysts were transferred into the oviduct of each ICR recipient female mice 0.5 day post-coitum. Birth was expected 16–18 days after the operation.

IF analysis of chimeras

The chimera was perfused transcardially with 4% PFA. Organs were post-fixed in 4% PFA, cryoprotected with 30% sucrose, embedded in OCT and cut into 10 µm sections on a cryostat. Sections were permeabilized and blocked with 0.2% Triton X-100 and 3% normal goat serum, then incubated with the following primary antibodies: MAP2 (Sigma-Aldrich, M4403; 100× dilution), NeuN (Chemicon, MAB377; 100X dilution), anti-oligodendrocytes (Chemicon, MAB1580; 1000× dilution), GFAP (Dako, Z0334; 400× dilution), TH (Chemicon, AB152; 100× dilution), PAX6 (DSHB; 50X dilution), troponin I (Chemicon, MAB338; 200× dilution), vWF (Dako, A0082; 300× dilution), K14 (NeoMarkers, RB-9020-P0; 300× dilution). The sections were visualized by incubation with the appropriate secondary antibodies: Cy3-conjugated donkey anti-goat, Cy3-conjugated goat anti-mouse (Jackson ImmunoResearch; 200× dilution) or R-PE-conjugated goat anti-rabbit (Southernbiotech; 200× dilution). Slides were counterstained with DAPI and examined. Staining with the secondary antibody alone was used as negative controls.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at HMG Online.

FUNDING

This work was supported by the Key Scientific and Technological Projects of Guangdong Province (2003A3020103), the Key Scientific and Technological Projects of Guangzhou (2002U13E0011) and National Natural Science Foundation of China (30671023).

Conflict of Interest statement. None declared.

Author notes

†

The authors wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors.