Genetic Analysis of the Roles of BMP2, BMP4, and BMP7 in Limb Patterning and Skeletogenesis

Abstract

Bone morphogenetic protein (BMP) family members, including BMP2, BMP4, and BMP7, are expressed throughout limb development. BMPs have been implicated in early limb patterning as well as in the process of skeletogenesis. However, due to complications associated with early embryonic lethality, particularly for Bmp2 and Bmp4, and with functional redundancy among BMP molecules, it has been difficult to decipher the specific roles of these BMP molecules during different stages of limb development. To circumvent these issues, we have constructed a series of mouse strains lacking one or more of these BMPs, using conditional alleles in the case of Bmp2 and Bmp4 to remove them specifically from the limb bud mesenchyme. Contrary to earlier suggestions, our results indicate that BMPs neither act as secondary signals downstream of Sonic Hedghog (SHH) in patterning the anteroposterior axis nor as signals from the interdigital mesenchyme in specifying digit identity. We do find that a threshold level of BMP signaling is required for the onset of chondrogenesis, and hence some chondrogenic condensations fail to form in limbs deficient in both BMP2 and BMP4. However, in the condensations that do form, subsequent chondrogenic differentiation proceeds normally even in the absence of BMP2 and BMP7 or BMP2 and BMP4. In contrast, we find that the loss of both BMP2 and BMP4 results in a severe impairment of osteogenesis.

Synopsis

A group of related signaling molecules called bone morphogenetic proteins (BMPs) are known to play important roles in the formation of the structures such as the limbs. However, because different members of this group often have similar effects on target cells and are produced in overlapping regions of the embryo and hence can be redundant with one another, removal of any single member of the BMP family may not reveal the full extent of the roles they play during development. We have therefore improved on this type of analysis by removing pairs of these factors (BMP2 and BMP4 or BMP2 and BMP7) specifically from the developing limb. Although some have speculated that these signals play an early role in organizing or “patterning” the different tissues of the limb, we find no evidence for such a role. We do find, however, that a minimal amount of BMP signal is required to form cartilage, and hence some cartilaginous elements fail to form in limbs deficient in both BMP2 and BMP4. Moreover, in the absence of these two BMP family members, there is a severe impairment in the development of bone tissue, resulting in severely deformed limbs. This study gives important new insight into the roles of these BMP signals in making skeletal tissues in the embryo.

Introduction

Bone morphogenetic proteins (BMPs) are secreted signaling molecules belonging to the transforming growth factor β superfamily, originally identified on the basis of their ability to induce ectopic bone formation when implanted within soft tissue in vivo [1–3]. BMP family members are now known to play an extremely diverse set of roles in a wide variety of developmental processes [4]. Even in the context of the morphogenesis of a single structure, these molecules can play a series of quite divergent roles. For example, during limb development, BMPs have been postulated to act sequentially in multiple distinct aspects of patterning, cell type specification, and differentiation of various tissues, particularly of the skeleton.

The earliest step of limb development in which BMP signaling has been implicated is the establishment of the anterior-posterior limb axis. Differences in anterior-posterior pattern are instructed as a graded response to Sonic Hedghog (SHH) signaling emanating from the posterior margin of the limb bud [5]. It has remained controversial, however, whether this response is direct or indirect. If indeed the long-range effects of SHH are indirectly mediated by local production of secondary signals, the leading candidates have been two members of the BMP family, BMP2 and BMP7. Both are expressed in a broader domain than SHH in the early posterior limb bud mesenchyme [6,7], although BMP7 also has a second, weaker domain of expression in the anterior limb bud mesenchyme. BMP2 [8] and BMP7 [7] can be induced by ectopic SHH and their expression is greatly diminished in the absence of SHH activity [9]. BMP2 and BMP7 are thus secondary signals produced in response to SHH activity. Moreover, BMP signaling has a weak ability to posteriorly polarize the limb in ectopic grafting experiments [10], an activity enhanced by prior low-level exposure to SHH [11]. It remains unclear, however, whether BMP2 and BMP7 activity is required endogenously for anterior-posterior limb patterning by SHH. Bmp2 mutant embryos die too early to assess their limb phenotypes. A targeted deletion of Bmp7 has been made, and Bmp7-deficient embryos display hindlimb polydactyly with incomplete penetrance but otherwise phenotypically normal limbs [12,13]. Nonetheless, Bmp7 knockout mice do not show any defect in limb polarity. However, a redundant function in anterior-posterior patterning with BMP2 remains a possibility. In addition to BMP2 and BMP7, a third member of this family that is closely related to BMP2, BMP4, is also expressed in the early limb bud. Like BMP7, it is expressed in both the anterior and posterior margins of the limb bud mesenchyme [4,14]; however, it does not appear to be induced by SHH signaling, nor does its expression change in SHH-deficient limb buds. Thus, BMP4 is not a candidate for a secondary signal downstream of SHH in early patterning. However, all three of these molecules, BMP2, BMP4, and BMP7, have been suggested to act in a second distinct phase of limb patterning, when digit identities are established downstream of earlier patterning events. In the vertebrate limb, each digit can be uniquely identified based on its size, length, number of phalanges, and location within the autopod. As a consequence of the initial establishment of anterior-posterior positioned information within the limb by SHH and/or BMP signaling, the interdigital mesenchyme of the hand plate becomes specified in a graded manner. Grafting and extirpation experiments have shown that it is this polarized interdigital tissue which directs digit morphology [15]. BMP2, BMP4, and BMP7 are all expressed in the interdigital mesenchyme [4,14], and experiments inhibiting BMP signaling in this tissue suggested that differential levels of BMPs in the interdigital mesenchyme might be the relevant factor directing digit morphology [15]. As noted above, BMP7-deficient embryos produce mostly normal limbs, and mice harboring a conditional deletion of Bmp4 in the limb bud mesenchyme also do not show evidence of anterior digit transformation [16], as would have been predicted if BMP4 acted as an interdigital regulator of digit morphology. In the light of these observations, it would seem unlikely that quantitative differences in levels of interdigital BMP signal are responsible for establishing digit identity, although redundancy between BMP2, BMP4, and BMP7 in this role remains a possibility.

A third aspect of limb patterning in which BMP2, BMP4, and BMP7 have been implicated is in the apoptotic death of the interdigital tissue. For example, ectopic application of BMP antagonists demonstrates that BMP signaling is necessary for this process, and in its absence, webbing occurs [17–19]).

BMP activity also plays an indirect role in limb patterning as a key component of a feedback loop between SHH production in the posterior limb bud and fibroblast growth factor production in the overlying apical ectodermal ridge (AER). SHH activity leads to the upregulation of the BMP antagonist Gremlin in the mesenchyme [20,21]. This, in turn, prevents BMPs from downregulating fibroblast growth factor (FGF) production in the AER [17,22] and maintains the integrity of the AER itself [23]. FGF signaling feeds back to maintain SHH production in the posterior mesenchyme [8,24]. Thus, conditional removal of BMP4 activity from the posterior limb mesenchyme results in a persistence of the AER, expanded SHH signaling, and consequent preaxial and postaxial polydactyly [16].

At later stages of limb development, BMP activity is also believed to play critical roles in skeletogenesis. Activation and dominant-negative experiments in the chick have shown that signaling through the BMPR-IB receptor is necessary and sufficient for cartilage condensation in chick [25]. Likewise, conditional knockout of BmpR1A and BmpR1B blocks all chondrogenic differentiation in the mouse limb bud [26]. Similarly, when noggin, a potent BMP inhibitor, is introduced in early stage chick limb buds prior to skeletogenesis, mesenchymal condensation does not take place [27,28]. Taken together, these studies strongly implicate BMP signaling as necessary for mesenchymal condensations and the initiation of chondrogenesis. However, the phenotypes in these experiments are severe, causing either global chondrogenesis or absence of chondrogenesis, in the gain- and loss-of-function experiments, respectively. These severe defects during the early stages of skeletogenesis preclude investigation of any specific roles for BMPs in later events in cartilage differentiation, bone formation, and bone metabolism [29]. Conversely, removal of individual ligands such as BMP4 [16] or BMP7 [12,13] has not displayed defects in skeletal differentiation, presumably due to functional redundancy.

To gain further insights into the potential roles of BMP signaling in vertebrate limb patterning and skeletogenesis, we have produced mice simultaneously lacking BMP2 and BMP4 activity or BMP2 and BMP7 activity in the limb mesechyme during limb development. Since Bmp2 and Bmp4 mutants are both lethal early in embryogenesis [30,31], we used conditional alleles of both these genes. The conditional alleles were deleted early in limb development through the action of a cre-recombinase transgene expressed under the control of the Prx1 enhancer [32]. Neither limbs deficient in BMP2 and BMP7 nor those deficient in BMP2 and BMP4 had defects suggestive of loss of anterior-posterior patterning information, arguing against these factors acting as secondary signals downstream of SHH or acting as interdigital determinants of digit identity. The Bmp2/Bmp4 double mutants did display a loss of posterior digits with a broad posterior hand plate, suggestive of a block in the chondrogenic condensation of the posterior digit rays, rather than a change in anterior-posterior digit identity. At later stages, the combined activity of neither BMP2 and BMP4 nor of BMP2 and BMP7 is required for chondrogenesis. Osteogenesis is also initiated properly in the absence of these pairs of factors. However, at later stages, the combined loss of BMP2 and BMP4 evidently becomes limiting, and osteogenesis ceases. As all prior reported manipulations of the BMP pathway during skeletal development either result in a block at the first stage of chondrogenesis or allow complete skeletal formation, this is the first demonstration of a requirement for BMP activity in osteogenic differentiation.

Results

To investigate the roles of BMP signaling at various stages of limb patterning and skeletogenesis, we constructed a series of mice deficient singly or in combination in the ability to produce BMP2, BMP4, and BMP7. BMP7-deficient mice (kindly provided by Dr. Liz Robertson) survive until birth. However, BMP2 and BMP4 are both required for viability early in embryonic development [30,31]. We therefore constructed a conditional allele of Bmp2, introducing loxP sites flanking exon 3. We obtained a conditional allele of Bmp4, in which exon 4 is flanked by loxP sites, from Dr. Holger Kulessa and Dr. Brigid Hogan [16]. Both of these alleles would be expected to result in null alleles following recombination (see Materials and Methods for details). To conditionally inactivate Bmp2 and Bmp4 in the limb, we used a well-characterized transgene in which cre-recombinase is expressed under the control of the Prx1 limb enhancer [32]. This transgene expresses cre very early in limb development, resulting in complete recombination of floxed alleles at early limb bud stages. We verified the ability of Prx1::cre to recombine the conditional Bmp2 and Bmp4 alleles at earliest limb bud stages using in situ hybridization. Bmp2 is first expressed in the limb mesenchyme at embryonic day (E)10.5 in the mouse (Figure 1A, asterisk). By the time its expression is first detectable, the floxed Bmp2 allele appears to be completely recombined in the limb mesenchyme, as whole mount in situ hybridization does not detect any mesenchymal Bmp2 transcription (Figure 1B). At E10.5 Bmp4 is expressed in the mouse limb mesenchyme in two stripes at the anterior and posterior margins (Figure 1C, red arrows). These expression domains are completely lost by E10.5 in the presence of the Prx1::cre transgene (Figure 1D). Bmp2 and Bmp4 are also expressed in the AER, where Prx1::cre is inactive, and these domains of expression are not affected (Figure 1A–1D, black arrows).

An Allelic/Nonallelic Series of BMP-Deficient Limbs

Mice were generated with limbs deficient in BMP2, BMP4, or BMP7, both BMP2 and BMP4, or both BMP2 and BMP7. To obtain an initial indication of the range of phenotypes produced in these animals, we first examined the limb skeletons of newborn animals. BMP2-deficient limbs appeared remarkably normal, both in skeletal pattern and in gross aspects of chondrogenesis and osteogenesis indicated by Alcian blue/Alizarin red staining. The one notable defect in the appendicular skeleton of these animals is a characteristic malformation of the scapula (Figure 1, red arrow). BMP2-deficient limbs also exhibit 3/4 soft-tissue syndactyly with variable penetrance (Figure 2).

Mice with limbs deficient in BMP4 activity and the limbs of Bmp7 mutant mice have both been previously described. As reported [16], in the absence of BMP4 activity, limbs display a variable penetrance of preaxial and postaxial polydactyly, but otherwise, normal digit patterns and apparently normal skeletal differentiation (Figure 1G and 1O) take place. As previously described, mice homozygous for a null mutation in Bmp7 [12] have no defects in the formation of the normal appendicular skeletal elements (Figure 1H and 1P). We do occasionally observe preaxial polydactyly in these mutants.

Compound heterozygous mice, with one functional copy each of Bmp2 and Bmp4 in the limb (Bmp2+/C; Bmp4+/C; Prx1::cre), show no effect on either limb patterning or skeletogenesis (unpublished data). Similarly, limb skeletons of mice in which both copies of the Bmp4 gene and one copy of the Bmp2 gene have been conditionally removed in the limb (Bmp2+/C; Bmp4C/C; Prx1::cre) are phenotypically normal other than exhibiting the variable penetrance preaxial and postaxial polydactyly seen in mice deficient in BMP4 activity alone (Figure 1J and 1R). In contrast, mice in which both copies of Bmp2 and one copy of Bmp4 had been removed (Bmp2C/C; Bmp4+/C; Prx1::cre) showed more severe skeletal defects, including significantly thinner skeletal elements. However, the digit patterns of those animals are completely normal (Figure 1I and 1Q). Animals in which both copies of Bmp2 and Bmp4 were removed (Bmp2C/C; Bmp4C/C; Prx1::cre) had extremely malformed limbs (Figure 1K and 1S). They displayed severely short and malformed stylopods; one of the zeugopod elements was almost always missing, and the remaining one was so deformed that it was difficult to identify the element correctly. Moreover, the joint articulations are defective such that zeugopod and stylopod elements are fused (see below and Figure S1A–S1C). Interestingly, the autopods are less affected than the proximal elements. Nonetheless, the autopod elements are significantly reduced in size, and strikingly the two posterior-most digits are missing in the forelimbs of these animals.

Simultaneous removal of BMP2 and BMP7 had far less of an effect than did removal of BMP2 and BMP4 activity. Compound heterozygous removal of BMP2 and BMP7 (Bmp2+/C, Bmp7+/−; Prx1::cre) and heterozygous removal of BMP2 with complete removal of BMP7 (Bmp2+/C, Bmp7−/−; Prx1::cre) were both completely wild-type in skeletal pattern and differentiation (unpublished data). Mice homozygous for removal of Bmp2 and heterozygous for Bmp7 (Bmp2C/C, Bmp7+/−; Prx1::cre) displayed the same subtle scapular defect seen in Bmp2C/C; Prx1::cre mutants alone (unpublished data). In limbs developing in the complete absence of both BMP7 and BMP2 activity (Bmp2C/C, Bmp7−/−; Prx1::cre), the last phalanx was missing from digit III in the forelimb, and these limbs displayed the same phenotype in the hindlimb with variable penetrance (Figure 1L and 1T, black arrow). Additionally, the fibulae of these hindlimbs are malformed and do not articulate with the femur at the knee (Figure 1T, thick red arrow). They also display the same scapular defects seen in Bmp2C/C; Prx1::cre mice, and the overall size of the appendicular skeleton is slightly diminished. However, skeletal differentiation appears normal in the limbs of these animals.

Soft Tissue Syndactyly Observed in Limbs of Mice Devoid of BMP2 and BMP4 Activity

As noted above, Bmp2-deficient animals display a variably penetrant 3/4 soft tissue syndactyly phenotype (Figure 2B). In contrast, Bmp4-deficient mice show no evidence of syndactyly (unpublished data). However, one of the striking defects observed in the Bmp2C/C; Bmp4C/C; Prx1::cre mice is that in the newborn animals, the digits of both forelimb (unpublished data) and hindlimb show complete syndactyly (Figure 2D; compare to wild-type, Figure 2C). To determine the origin of this defect, we examined embryonic limbs at E15.5, the stage when interdigital separation is normally taking place (Figure 2E). In Bmp2C/C; Bmp4C/C; Prx1::cre limbs, only the very distal tips of each digit were separated, with the autopod adopting the shape of a notched pallet (Figure 2F). In wild-type development, the individual digits become free from one another through a process of interdigital apoptosis, observable by staining with acridine orange (Figure 2E and 2G, arrow). Interdigital apoptosis is dramatically reduced in the Bmp2C/C; Bmp4C/C; Prx1::cre limbs (Figure 2F and 2H, arrow). (Note that the strong acridine orange staining in Figure 2H is not in the interdigital mesenchyme but rather is in the remnant of the cells of the AER [red asterisk].)

Another region of very prominent apoptosis in the wild-type limb bud is within the AER. The AER is a specialized ridge of ectoderm running along the anterior-posterior axis at the distal margin of the early limb bud. One of the phenotypes characterizing the Bmp2C/C; Bmp4C/C; Prx1::cre mutant is expansion of the AER (see below). Moreover, in the Bmp2C/C; Bmp4C/C; Prx1::cre animals, the AER is maintained as late as E15.5, long after it has started to degrade in wild-type limbs. This can be seen morphologically (unpublished data) or by staining for expression of AER-specific marker such as Fgf8 (Figure 2I and 2J). The AER displays intense staining with acridine orange, TUNEL, or other markers for apoptosis at every stage when it is present, reflecting the rapid turnover of cells in this structure [33–35]. The Bmp2C/C; Bmp4C/C; Prx1::cre mutant limbs at E15.5 show strong acridine orange staining in the remnants of AER (Figure 2H, asterisk), in domains of the distal tip where the AER remains the longest (compare staining in Figure 2F and 2J). No significant difference was observed in the extent of apoptosis in the cells of the AER in earlier stage Bmp2C/C; Bmp4C/C; Prx1::cre mutant embryonic limbs (Figure S1D–S1G).

Patterning Defects in Limb Developing with Reduced Levels of BMP Signaling

BMP signaling has been proposed to play several distinct roles in limb patterning. The first of these is in establishing differences along the anterior-posterior axis of the limb downstream of sonic hedgehog (Shh). SHH signaling itself extends over a long range in the developing limb bud, based on both activation of target genes [36] and protein distribution [37]. Nonetheless, some models have suggested that the patterning effect of this morphogenic signal is mediated entirely [10] or in part [11] by BMP2 and BMP7 produced as secondary signals in response to SHH. We have removed both BMP2 and BMP7 activities in the early limb bud (Figure 1), and moreover, there is no apparent upregulation of BMP4 expression in the posterior of these limbs (Figure 3A–3D). The skeletal phenotypes of mouse limbs formed in the absence of BMP2, BMP7, and both BMP2 and BMP7 are illuminating, as none of these exhibit significant digit pattern defects (Figure 1). While a distal phalanx is missing in the middle digits of the double mutants (Bmp2C/C, Bmp7−/−; Prx1::cre), this is not a phenotype consistent with a decrease or loss of a posteriorly derived polarizing activity. Indeed, the terminal phalanx forms at the distal tip of each digit by a distinct mechanism, not dependent upon anterior-posterior patterning [38]. These data strongly indicate that BMP2 and BMP7 are not required secondary signals for SHH-mediated polarization of the digits along the anterior-posterior axis.

The lack of transformations in digit identity in the limb skeletons formed in the absence of BMP2 and BMP7 activity also argues against BMP signaling being the critical instructive signal from the interdigital mesenchyme to the forming digits. While a distal phalanx was lost from the middle digit of the forelimbs of these animals, these digits do not otherwise have a morphology consistent with their being transformed anteriorly into digit 1 (thumb). Indeed, in the hindlimbs, where the penetrance of the phenotype is variable, the mutant middle digits are otherwise indistinguishable from wild-type, other than the presence or absence of this variable skeletal element.

In contrast to the proposed roles for BMPs in anterior-posterior digit patterning, we obtained further support for the well-accepted idea that a threshold level of BMP signaling is necessary interdigitally for programmed cell death to occur. We observed syndactyly with variable penetrance in limbs developing without BMP2 activity (Figure 2B), and nearly complete soft tissue syndactyly of all digits in limbs deficient in both BMP2 and BMP4 (compare Figure 2C and Figure 2D).

The one dramatic patterning defect we observed in our BMP deficiency series was the loss of posterior digits in forelimbs deficient in both BMP2 and BMP4 (Figure 1K), also observed in earlier limb buds stained to show no expression of the condensation marker Sox9 (Figure 3). To understand this phenotype, we analyzed limb buds at various stages using a series of molecular markers. By E11.5, the Bmp2C/C; Bmp4C/C; Prx1::cre limb buds are noticeably broader, expanded in the posterior relative to wild-type. This correlates with an anterior expansion in Shh expression (Figure 3E and 3F, red bracket) and a concomitant expansion of the AER as visualized by Fgf8 expression (Figure 3G and 3H, red bracket). The expansion in Shh expression can first be detected at E10.5 and persists at least until E12.5, well after the time normal Shh expression disappears at E11.75 (Figure 3L and 3P). The expansion of the AER in the absence of BMP signaling may be understood on the basis of the feedback loop between the zone of polarizing activity (ZPA) and the AER. Expression of several Fgf genes [20,21] and maintenance of the AER itself [23] normally depend on BMP antagonism induced by SHH. Thus, a decrease in BMP signaling expands and maintains the AER. Because Shh expression is, reciprocally, supported by FGF signaling from the AER [8,24], this, in turn, feeds back on the ZPA, expanding Shh expression. While this explains the broader hand plate of the BMP2, BMP4–deficient limb buds, it is somewhat paradoxical in terms of the skeletal phenotype, as a broadened autopod would normally be expected to result in the formation of extra digits (polydactyly) (e.g., Figure 1G and 1O of this paper and [16]), not the loss of digits, as observed in the double mutant forelimbs.

To examine the loss of digits more closely, BMP2, BMP4–deficient limb buds were examined during cartilage condensation phase of the digit rays at E12.5. In wild-type limb buds the condensation of all five digits can be visualized at this stage by in situ hybridization with a probe directed against Sox9, the earliest known chondrogenic marker (Figure 3I). However, in Bmp2C/C; Bmp4C/C; Prx1::cre limb buds at E12.5, Sox9 only detects primordia of the anterior three digits of the forelimb, indicating a failure in the formation of posterior digits at this earliest stage (Figure 3J).

Genes such as Msx2 (Figure 3N) continue to be expressed throughout the posterior forelimb mesenchyme, indicating that the posterior forelimb tissue is viable at E12.5 in the Bmp2C/C; Bmp4C/C; Prx1::cre animals. Msx2 is normally expressed in the non-chondrogenic interdigital mesenchyme at this stage (Figure 3M), a continuation of an earlier expression domain throughout the distal mesenchyme. This expression of Msx2 in the posterior of the BMP2, BMP4–deficient forelimbs reflects a failure of chondrogenesis in this region. The loss of the posterior forelimb digits in these mice thus does not appear to represent a decrease in polarizing activity leading to a change in digit identity rather a consequence of the overall level of BMP signaling falling below a threshold for initiating chondrogenesis in the posterior of these forelimbs. The posterior of the hindlimb in the mutant, in contrast, appears to remain above this threshold, and more than five digit condensations form in the hindlimb bud (Figures 3K and 1S) and no ectopic expression of Msx2 is observed in the posterior (Figure 3O).

Skeletal Differentiation in Limbs Developing with Reduced Levels of BMP Signaling

BMPs were first discovered in the context of their skeletogenic activity. We therefore wanted to determine whether the loss of the specific BMP molecules under investigation here would perturb the skeletal differentiation pathways. Accordingly, skeletal preparations from each of the various mutant alleles were subjected to histological and molecular analyses. The skeletal elements that formed in the absence of BMP2, BMP4, or BMP7 or BMP2 and BMP7 all underwent normal chondrogenesis and osteogenesis by a series of criteria. All showed normal staining with Alcian blue (for cartilage) and Alizarin red (for mineralized tissue, see Figure 1). Most were also analyzed in histological sections stained with toluidine blue and showed osteoid formation and bone matrix deposition (compare Figure S2A and S2E; and unpublished data). The skeletal elements in these various BMP-deficient animals contained normal prehypertrophic chondrocytes expressing type II collagen, hypertrophic chondrocytes expressing type X collagen, as well as the osteoblasts expressing type I collagen, like their wild-type counterparts (Figure S2; compare Figure S2F–S2H with Figure S2B–S2D; and unpublished data). In contrast, however, limbs completely deficient in both BMP2 and BMP4 activity showed severe defects in skeletal differentiation.

BMP2 and BMP4 Are Not Critical for Chondrogenic Differentiation

As noted above, at E12.5, chondrogenic condensation of the digits could be visualized in Bmp2C/C; Bmp4C/C; Prx1::cre limb buds by in situ hybridization with a Sox9 probe. Similar analysis at E10.5 and E11.5 demonstrated a relatively normal process of condensation of the proximal skeletal elements (Figure S1H–S1M), despite the absence of detectable Bmp2 and Bmp4 transcripts. As shown in Figure S1N and S1O, expression of Col II, a chondrocyte marker, is observed in the BMP2, BMP4–deficient cartilage (Figure S1O) as early as E12.5, similar to a wild-type control (Figure S1N). Similarly, at E13.5, the entire limb skeleton could be visualized undergoing chondrogenesis by Alcian blue staining, albeit in a defective pattern, in the BMP2, BMP4–deficient limb buds (Figure 4A and 4B and unpublished data).

As the cartilage elements grow, a wave of hypertrophic differentiation occurs from the center toward the distal ends (reviewed in [39–41]). Histological examination of the humeri at E13.5 showed that while chondrogenic differentiation had taken place in the BMP2, BMP4–deficient humerus, hypertrophic differentiation was delayed (Figure 4C and 4D). However, by E15.5, hypertrophy was evident in these elements as seen by type X collagen expression (Figure 4E and 4F). The hypertrophic cells in BMP2, BMP4–deficient limb also express late hypertrophic markers, such as osteopontin [42] (unpublished data).

To examine whether the successful execution of chondrogenesis in the BMP2, BMP4–deficient limbs is a result of compensatory upregulation of other ostegenic BMPs, we carried out in situ hybridization for Bmp6 and Bmp7 in these limbs. At E13.5, we observed no increase in expression of Bmp6 (Figure S1P–S1S) or Bmp7 (compare Figure 4H with Figure 4G) mRNA in the mutant tissues compared to their wild-type counterparts

Osteogenesis Is Initiated Normally in BMP2, BMP4–Deficient Limb Buds

Bone formation first occurs at the interface between the late hypertrophic chondrocytes and the surrounding perichondrium, a region identified as the bone collar. We examined whether the osteogenesis program is initiated in the BMP2, BMP4–deficient limbs using several histological and molecular criteria, as described below (Figure 5). Adjacent to the hypertrophic cells, a bone collar is laid down normally in the absence of BMP2 and BMP4 (Figure 5A and 5B, red star). As can be seen in sagittal sections through an E15.5 humerus, the cells embedded in the mineralized matrix express the osteoblast marker Col I as normal (Figure 5C and 5D). By E17.5, the mineralized cartilage that makes up the late hypertrophic zone of the growth plate normally undergoes removal by osteoclasts (Figure 5G, brown-stained cells, red arrows), resulting in the formation of a bone marrow cavity (red arrow in Figure 5E and 5I, see also Figure 6). Osteoprogenitor cells from bone collar region now populate this site and deposit osteoid, forming the trabecular bone in the bone marrow cavity (Figure 6E, red arrow) and cortical bone at birth (Figure 5I, asterisk). In contrast, in E17.5 limbs deficient in BMP2 and BMP4, there is an overall delay in the normal endochondral process (Figure 5F). While vascularization occurs in Bmp2C/C; Bmp4C/C; Prx1::cre forelimbs, few osteoclasts are recruited to the mineralized cartilage (compare Figure 5G and Figure 5H, red arrows), resulting in a delay in bone marrow cavity formation in the absence of BMP2 and BMP4. In fact, bone marrow formation and trabecular bone formation have not yet started at birth in the absence of BMP2 and BMP4 (Figure 5J), and the morphology of Bmp2C/C; Bmp4C/C; Prx1::cre bones at birth is most similar to control embryos at E17.5 (Figure 5E). To differentiate between a simple delay in bone formation in the absence of BMP2 and BMP4 and the inability to form bone without these BMPs, we performed histological analyses in Bmp2C/C; Bmp4C/C; Prx1::cre mice up to 3 wks of age. As shown in Figure 6A through 6E, in the control mice, a bone collar is found adjacent to the zone of late hypertrophic chondrocytes at 1 wk (Figure 6A and 6B). Many osteoblasts line the newly formed cortical bone in the diaphyseal region (Figure 6C, blue arrow), and formation of a secondary ossification center has begun (Figure 6A, red star). At 3 wks of age, formation of the secondary ossification center is complete (Figure 6D, red star), along with robust bone marrow formation (Figure 6D, bm) and deposition of trabecular bone (Figure 6E, pale blue tissue marked by red arrow). In stark contrast, bone formation is not observed in the absence of BMP2 and BMP4 at 1 or 3 wks after birth (Figure 6F–6J). The appearance of the skeletal elements in the double mutants remains similar to E17.5 structures seen in wild-type limbs (Figure 5E), and although there is a bone collar at the site of hypertrophic cartilage (Figure 6G), no bone marrow cavity, trabecular bone, or cortical bone is present (Figure 6F). Many osteoclasts have been recruited to the remaining mineralized cartilage and are actively resorbing this tissue (red asterisks, Figure 6I), so that by 3 wks, all the mineralized cartilage in the diaphyseal region has disappeared, leaving a void where bone formation should have occurred (Figure 6J, also see Figures 6L and S3). This void is eventually invaded by soft tissues and the adjacent muscle (marked s and m, respectively, in Figure 6J). The resorption of bone at 3 wks of age is clearly visible in the proximal femur of mutant animals where a portion of that bone is missing (Figure 6L, black arrow). The inability to complete osteoblast differentiation in the Bmp2C/C; Bmp4C/C; Prx1::cre mice results in strikingly defective limb morphology at 1 wk of age (compare Figure S3G and S3H).

To determine if any osteoblast differentiation occurs in Bmp2C/C; Bmp4C/C; Prx1::cre forelimbs, we examined osteoblast-related gene expression (Figure 7). In the control mice, osteoblasts lining the surface of trabecular and cortical bone express type I collagen, runx2, and osterix genes at both 1 wk and 3 wks after birth (Figure 7A–7D and 7I–7L). In the absence of BMP2 and BMP4, cells which are fibroblastic in appearance (Figure 6H) are found adjacent to the mineralized cartilage surfaces. These cells express amounts of type I collagen and runx2 comparable to control osteoblasts, but their expression of osterix, while noticeable at 1 wk, becomes almost undetectable at 3 wks (Figure 7E–7H and 7M–7P). These results allow us to conclude that the fibroblast-like cells resident in the bone shaft (Figure 6H) may be osteoprogenitor cells unable to differentiate into mature osteoblasts in the absence of BMP2 and BMP4.

Discussion

Using a genetic approach to remove activity of different Bmp genes during limb development, we have found that BMP2, BMP4, and BMP7 either individually or in combination are not required for specification of normal digit identities. However, BMP2 and BMP4 are required in concert to promote condensation of the posterior digit anlagen. We have also shown that chondrogenesis can be initiated and chondrogenic differentiation will take place even in the absence of both BMP2 and BMP4 or BMP2 and BMP7. On the other hand, BMP2 and BMP4 together are required for completion of osteogenesis.

BMP Signaling and Pattern Formation

It has been proposed that BMP signaling may be involved in digit patterning as a secondary signal originating in the ZPA [7,11,43] or as a later mediator of interdigital patterning information [15]. In particular, BMP2, a secreted factor that is induced by SHH [8] in the posterior limb bud, has been suggested to act as a true morphogen secondary to SHH signaling to properly pattern the limb digits. However, inactivation of Bmp2 during early limb bud development clearly does not cause any digit patterning defects. BMPs are known, in many cases, to function redundantly [44]. In considering potential secondary signals mediating ZPA activity, the most relevant family member to consider is BMP7 as, like BMP2 (and unlike BMP4), it is positively regulated by SHH signaling [7]. However, Bmp7 expression does not overlap with Bmp2 in the early mouse limb bud [45], and in any case, mice in which we conditionally knocked out Bmp2 in the Bmp7−/− background do not show any obvious digit patterning defect other than the missing final phalanx in digit III. Although still poorly understood, the final phalanges form at the tip of every digit through a process distinct from the more proximal phalanges [38]. Our data implicate BMP signaling in this process.

At later stages, BMP signaling has been suggested to play a role in mediating the transfer of positional information from the interdigital mesenchyme to specify the identity of the adjacent digits [15]. Application of the BMP antagonist noggin to the interdigital tissue at the time of digit condensation results in loss of phalanges and a resultant pattern that can be interpreted as an anterior homeotic transformation in digit identity. However, increasing BMP concentrations interdigitally with exogenous protein does not result in the reciprocal posterior transformations, and the noggin experiments can also be interpreted simply as a block in the condensation of the primordia of distal phalanges when BMP activity is completely abolished [38]. Decreasing the level of interdigital BMP signaling, without abolishing it, provides an opportunity for differentiating between these models. If BMP signaling were only required as a permissive factor for chondrogenesis, partial reduction in the level of signaling might have no effect. However, if levels of BMP signaling were instructive in terms of digit identities, then a substantial decrease in the amount of BMP ligand present should cause transformations in digit identity. This is not what is observed, either in limbs deficient in both BMP2 and BMP4 or in those deficient in BMP2 and BMP7. It should be noted that the interdigital expression domains form several days after recombination of the floxed BMP alleles is complete. Moreover, there is no detectable compensatory upregulation of BMP7 in the interdigital mesenchyme of the BMP2, BMP4–deficient limbs (compare Figure 4G and Figure 4H). Thus, we conclude that BMP signaling is not likely to be responsible in a quantitative fashion for the instructive role of interdigital mesenchyme in establishing digit identities. However, in the Bmp2C/C; Bmp4C/C; Prx1::cre forelimbs, there is a complete loss of posterior digit ray condensations, presumably because the total level of BMP signaling falls below the threshold for initiation of condensation, as previously seen in cases where BMP antagonists have been used [27,28].

It is worth noting that the defects in chondrogenesis seen in the BMP2, BMP4–deficient limbs are much more severe proximally than distally. This is particularly significant because proximal elements condense and differentiate before distal ones. Thus, if the conditional alleles were partially recombined at early stages and only completely recombined at later stages, it is the distal elements, formed after greater recombination has taken place, that should be more severely affected. Since it is the proximal elements which are more severely affected, the explanation for the difference in severity must lie elsewhere. This is most likely explained by compensation in the distal limb bud by BMP2, BMP4, and BMP7 produced in the AER.

Compensation by BMPs in the AER also likely explains a difference seen between the Bmp4C/C; Prx1::cre mice in our experiments and a recently published analysis [16], which used the identical Bmp4 conditional allele and Prx1::cre driver. The earlier report, like ours, described preaxial and postaxial polydactyly. However, they observed these phenotypes with less variability and, moreover, described multiple preaxial and postaxial ectopic digits, while we saw, at most, a single preaxial and postaxial digit. The difference is that the mice analyzed in the previous report had one null allele and one conditional allele, while ours had two conditional alleles. Their mice, therefore, lost a half-dose of BMP4 from the AER in addition to complete loss in the mesenchyme, while our conditional alleles were only recombined in the mesenchyme by the Prx1::cre driver. It is also possible that the difference in severity relates to the time required to recombine one as opposed to two floxed alleles; however, we do not favor this explanation as all detectable BMP4 expression is lost prior to any morphological differences between the mutant and wild-type limb buds. Finally, differences in genetic background could contribute to the differences as the Prx1::cre line is outbred.

A final aspect of patterning previously associated with BMP signaling is interdigital apoptosis. Consistent with this, we see some syndactyly in the BMP2-deficient limbs. The fact that this is variable, and limited to the 3/4 interdigit, implies that the threshold for induction of apoptosis is low enough that loss of BMP2, BMP4 or BMP7 or BMP2 and BMP7 still leaves sufficient BMP signaling for most interdigital cell death to occur normally. However, loss of both BMP2 and BMP4 results in complete soft tissue syndactyly, i.e., webbing between all the digits. Although BMP activity has been previously implicated in regulating interdigital apoptosis, this is the first genetic verification of their requirement for this process.

BMP Signaling and Skeletal Development

As discussed above, our data indicate that threshold levels of BMP signaling are required for initiating chondrogenic condensation, consistent with prior results [26–28]. However, once chondrogenesis begins, cartilage differentiation can be sustained, albeit with some detectable delay, in the absence of BMP2 and BMP4 or in the absence of BMP2 and BMP7. Thus, the expression of either BMP5, BMP6, and BMP7 (in the absence of BMP2 and BMP4) or the expression of BMP4, BMP5, and BMP6 (in the absence of BMP2 and BMP7) is sufficient to support mesenchymal condensation and chondrocyte differentiation in the developing limb.

In one set of previous studies implicating BMP signaling in chondrogenesis, Yoon et al. [26] showed that if mice lose both Alk3 (BmprIa) and Alk6 (BmpIb), two of the three type I receptors used in BMP signal transduction, there is a dramatic decrease in the size of skeletal primordial due to a reduction of proliferation and increase in apoptosis. The size of the skeletal elements in Bmp2C/C; Bmp4C/C; Prx1::cre limbs are also decreased compared to those of control mice, although not to the extent seen in the double receptor mutant. We do not know if reduced size of the skeletal elements in our study is similarly due to reduced proliferation and increase in apoptosis, lack of replacement by bone, or a combination of these. It is possible that a triple mutant removing BMP2, BMP4, and BMP7 activities would have a similar phenotype as the BmprIa, BmprIb double mutant, although these could also be compensated from other BMPs. We do know, however, that osteogenesis is defective in these mice, although bone formation proceeds normally in siblings retaining one functional copy of either the Bmp2 or Bmp4 gene (Figure S4), as it does in mice completely deficient in BMP2 and BMP7 (Figures 1 and S2).

The phenotype we observe in limbs of Bmp2C/C; Bmp4C/C; Prx1::cre mice is similar to the phenotype reported for osterix knockout mice [46], which exhibited severe defects in osteoblastic differentiation. Since we observe fibroblastic cells adjacent to mineralized cartilage that have characteristics of osteoprogenitor cells but fail to express osterix in the absence of both BMP2 and BMP4, one important role of BMP2 and BMP4 during endochondral ossification may be to induce osterix gene expression in osteoprogenitors. Strikingly, however, we do observe osterix expression in the bone collar of E16.5 mice missing functional copies of both the Bmp2 and Bmp4 genes (K. Tsuji and A. Bandyopadhyay, unpublished data). This suggests that varying levels of expression from different BMP genes during preaxial and postaxial development result in fluctuations in total levels of BMP signaling, which at distinct times are above or below the threshold for supporting osteogenesis. While many osteogenic BMP molecules, apart from BMP2 and BMP4, such as BMP5, BMP6, and BMP7, are expressed during the endochondral process, our data suggest that these BMP molecules cannot compensate for the combined loss of BMP2 and BMP4 in bone formation. Recent biochemical and genetic experiments have suggested that individual BMPs have identical functions. BMP2, BMP4, BMP5, BMP6, and BMP7 can utilize the same type I (Alk2, Alk3, and Alk6) and type II receptors (BMP RII, ActRII, and ActRIIb) [47]. Once the complex between ligand and receptors is formed, BMP2, BMP4, BMP5, BMP6, and BMP7 direct the phosphorylation of the same set of BMP receptor–specific Smads (1, 5, or 8), and the signal that is transduced by each BMP appears to be identical in the skeletal target cells [48–51]. BMPs have also been shown to activate MAPK and AKT pathways and, based on current published information, each of the osteogenic BMPs appears to have the same capacity to activate these signaling pathways [52–54]. When this information is considered along with our observation that one allele of either Bmp2 or Bmp4 can rescue the osteoblast differentiation phenotype observed in Bmp2C/C; Bmp4C/C; Prx1::cre mice (see Figures 1I, 1J, 1Q, and 1R and S4), we favor the hypothesis that bone formation requires a threshold amount of BMP signaling which is not met when both BMP2 and BMP4 are completely absent. We cannot, however, exclude the possibility that BMP2 and BMP4 have distinct functions in osteoblastogenesis. Our analysis of skeletogenesis indicates that BMP2 and BMP4 are prerequisite for osteoblastogenesis while less important for chondrogenesis. These studies provide the first evidence linking BMP2 and BMP4 with bone formation in an in vivo setting where the preceding events of chondrogenesis are not compromised.

Materials and Methods

Reagents.

Xylene, formamide, and acetic anhydride were purchased from Fisher Scientific (http://www.fishersci.com). Alizarin red S, toluidine blue, paraformaldehyde, polyvinylpyrrolidone, proteinase K, RNase A, tRNA, and dithiothreitol were purchased from Sigma (http://www.sigmaaldrich.com). Alcian blue 8GX was purchased from Electron Microscopy Science (http://www.emsdiasum.com). Dextran sulfate was purchased from Pharmacia (http://www.pharmacia.com).

Generation of Bmp2 and Bmp4 conditional null allele.

Mice carrying floxed Bmp2 allele were available through a material transfer agreement between Harvard University and Wyeth Pharmaceuticals. In these mice, LoxP sites were integrated to excise the entire protein-coding region in exon 3 of the Bmp2 gene. Generation of Bmp4 conditional allele has been described [16,55]. The Prx1::cre transgene has been described [32].

Generation of Bmp2, Bmp4 double conditional mice and Bmp2 conditional, Bmp7 mutant mice.

Bmp2C/C animals were crossed with Bmp4C/C animals to generate Bmp2C/+; Bmp4C/+ animals. These animals were crossed with Bmp2C/C or Bmp4C/C animals to generate Bmp2C/C; Bmp4C/+ or Bmp2C/+; Bmp4C/C animals, respectively. Bmp2C/C; Bmp4C/+ or Bmp2C/+; Bmp4C/C animals were crossed with each other to generate Bmp2C/C; Bmp4C/C animals. Bmp2C/C; Bmp4C/C females were crossed with males bearing Prx1::cre transgene to generate Bmp2C/+; Bmp4C/+; Prx1::cre animals. Bmp2C/+; Bmp4C/+; Prx1::cre males were crossed with Bmp2C/C; Bmp4C/C females to generate Bmp2C/C; Bmp4C/C; Prx1::cre animals. Bmp2C/C; Bmp4C/C mice carrying floxed alleles of the Bmp genes without Prx1::cre are wild-type (WT). Genotyping of animals were done by PCR analyses with the primers (5′–3′) GTGTGGTCCACCGCATCAC (AHP2–9) and GGCAGACATTGTATCTCTAGG (AHP2–35) for Bmp2 and AGACTCTTTAGTGAGCATTTTCAAC (No. 79 [order of primers tried]) and AGCCCAATTTCCACAACTTC (No. 80) for Bmp4 following extraction of genomic DNA from embryonic or adult tail. For Bmp2, the floxed allele amplifies as a 545-bp product, while the wild-type allele amplifies as a 474-bp product. For Bmp4, the floxed allele amplifies as a 220-bp product, while the wild-type allele amplifies as a 180-bp product. Protocols utilized for mouse experiments were approved by the Harvard University Institutional Animal Care and Use Committee (Cliff Tabin #02735).

The strategy for generating a Bmp2 conditional null in the BMP7−/− background is similar to the strategy used for generating the Bmp2, Bmp4 double conditional allele with three important differences. First, the Bmp7−/− (strain kindly donated by Dr. Liz Robertson [12]) is not a conditional allele but a null mutation. Second, Bmp7−/− mice do not survive past birth due to kidney failure [12] and hence cannot be maintained as homozygous for the null allele. Third, Bmp2 and Bmp7 are on the same chromosome (Chromosome 2). To obtain Bmp2C/C, Bmp7−/−; Prx1::cre we crossed Bmp2C/C, Bmp7−/+ females with Bmp2C/+, Bmp7−/+; Prx1::cre males. Bmp2C/C, Bmp7−/−; Prx1::cre progeny were obtained at a rate slightly higher than 1 in 16. For genotyping of Bmp7 locus we used the following set of primers: Bmp7_ExI_5′ (5′–3′) TCGCCTGCAGCAAGTGACCTCGGGTC; Bmp7_ExI_3′ (5′–3′) TAGGGGTAGGAGAAGCCCTGTCCGTCC and Beta-geo 5′ of 3′ (5′–3′) CTGCATACGCTTGATCCGGCTACCTGC. From a wild-type chromosome Bmp7_ExI_5′ and Bmp7_ExI_3′ amplifies an approximately 423-bp PCR product, while from the lacZ insertion mutant this pair fails to amplify any product. Bmp7_ExI_5′ and Beta-geo 5′ of 3′ (this primer is designed from the Beta-geo cassette) amplifies an approximately 1-kb PCR product from the lacZ inserted chromosome.

Skeletal analysis.

Our protocol is an adaptation of the procedure described [56] previously. Newborn and older animals were killed using carbon dioxide. Skin, viscera, and adipose tissues were removed within 2 h after death, and samples were fixed in 95% ethanol for 5 d. Samples were then placed in acetone to remove residual fat for 2 d. For E13.5 embryos, skin was not removed. The dehydrated animals were stained with 0.015% w/v Alcian blue and 0.005% w/v Alizarin red in a 1:19 mixture of glacial acetic acid and 70% ethanol. The younger animals were stained for 5 h at 37 °C and then overnight at room temperature, while the newborns were stained overnight at 37 °C followed by 2 d at room temperature. The stained skeletons were kept in 1% KOH, with occasional change of solution, until cleared. The cleared skeletons were transferred into 100% glycerol and photographed.

Acridine orange staining for apoptotic cells.

A working stock of 5 mg/ml acridine orange was diluted 1:10,000 in PBS. Dissected fresh embryos were transferred to the working solution of acridine orange and incubated for 30 min at 37 °C in the dark. Embryos were then washed twice in PBS for 5 min and viewed with a fluorescence microscope.

Whole mount in situ hybridization.

In situ hybridization staining on whole embryos was performed as described [57]. For section in situ hybridizations, please see below.

Histology, section in situ hybridizations, and immunohistochemistry.

For histology and section in situ hybridization, samples were fixed in 4% paraformaldehyde in PBS at 4 °C overnight. Newborn and older animals were then decalcified in Tris buffer containing 10% EDTA and 7.5% polyvinylpyrrolidone (pH 7.5) at 4 °C for 3 wks. Samples were then dehydrated through a graded ethanol series, cleared in xylene, and embedded in paraffin. 8μm sections were collected and stained with hematoxylin and eosin (H & E) or toluidine blue following standard procedure.

Section in situ hybridization with digoxigenin labeled probes was performed as described [57]. Section in situ hybridization with radiolabeled probes was performed as described [58] with a small modification. Briefly, sections were deparaffinized in xylene and rehydrated through a graded ethanol series. Sections were postfixed in 4% paraformaldehyde in PBS at room temperature for 15 min, digested with 10 μg/ml proteinase K at room temperature for 10 min, and again fixed in 4% paraformaldehyde at room temperature for 10 min and acetylated in the solution containing 0.2% hydrochloric acid, 0.1 M triethanol amine, and 0.25% acetic anhydride at room temperature for 10 min. Sections were then dehydrated in increasing concentrations of ethanol and air-dried. Hybridization was performed in a humidified chamber in a solution containing 50% formamide, 10% dextran sulfate, 1× Denhardt's solution, 0.6 M sodium chloride, 0.01 M Tris buffer (pH 7.5), 0.001 M EDTA, 0.05 M dithiothreitol, 0.25% SDS, 200 μg tRNA, and 35S-labeled cRNA probe at the final concentration of 5 × 106 cpm/ml at 55 °C for 16 h. After hybridization, sections were washed with a solution containing 5× SSC and 10 mM dithiothreitol at 50 °C for 30 min, incubated in a solution containing 50% formamide, 2× SSC, and 10 mM dithiothreitol at 65 °C for 30 min, treated in a solution containing 10 μg/ml RNase A in TNE [0.01 M Tris buffer (pH 7.6), 0.5 M sodium chloride, and 0.001 M EDTA] at 37 °C for 30 min and then washed with 50% formamide, 2× SSC, and 10 mM dithiothreitol at 65 °C for an additional 30 min followed by two washes with 2× SSC/10 mM dithiothreitol at 65 °C for 30 min, and 0.1× SSC/10 mM dithiothreitol at 65 °C for 30 min.

To visualize the signal, sections were dipped into NTB-2 emulsion (Kodak, http://www.Kodak.com) and placed at 4 °C. After developing, sections were counterstained with H&E. The 35S-labeled cRNA probes were transcribed from plasmids encoding type I collagen [59], runx2 [60], osteopontin [61], and osterix [46]. Sense and antisense probes were synthesized from linearized plasmids using a Riboprobe Combination System (Promega, http://www.promega.com).

Supporting Information

Figure S1

Chondrogenic Differentiation in Bmp2C/C; Bmp4C/C; Prx1::cre Animals

(A–C) In BMP2, BMP4–deficient limbs, there is only one zeugopod present, and it is often fused with the stylopod. All panels are from E17.5 Bmp2C/C; Bmp4C/C; Prx1::cre embryos. (A) Hematoxylin and eosin–stained section of the forelimb. Alcian blue– and Alizarin red–stained skeletons shown in (B) (forelimb) and (C) (hindlimb). Black arrow shows the fusion of the zeugopod and stylopod in BMP2, BMP4–deficient limbs.

(D and E) Acridine orange–stained limbs from E11.5 wild-type (D) and Bmp2C/C; Bmp4C/C; Prx1::cre embryos (E).

(F and G) Acridine orange–stained hindlimbs from E13.5 wild-type (F) and Bmp2C/C; Bmp4C/C; Prx1::cre (G) embryos.

(H–M) In the absence of BMP2 and BMP4, condensation begins in the limb normally. Whole mount in situ staining of Sox9 mRNA in embryonic limb buds from E10.5 (H and I) and E11.5 (J–M) embryos. (H, J, and L) From wild-type embryos, (I, K, and M) from Bmp2C/C; Bmp4C/C; Prx1::cre embryos. (H, I, J, and K) Forelimbs, (L and M) hindlimbs.

(N and O) In situ hybridizations with Col II mRNA probes of sections derived from E12.5 wild-type (N) and Bmp2C/C; Bmp4C/C; Prx1::cre (O) embryos.

(P–S) Bmp6 mRNA expression is not increased in E13.5 Bmp2C/C; Bmp4C/C; Prx1::cre cartilage. Bmp6 in situ on sagittal sections from wild-type (Q) and Bmp2C/C; Bmp4C/C; Prx1::cre (S) embryonic forelimbs. (P and R) Bright field images of (Q) and (S), respectively.

(23 MB TIF)

Click here for additional data file.

Figure S2

Skeletal Differentiation Occurs Normally in Absence of BMP2

(A) Toluidine blue–stained section of newborn femurs from Bmp2C/C animals.

(B–D) Marker analyses. Sections of femurs from Bmp2C/C animals hybridized with Col II (B), Col X (C), and Col I (D) mRNA probes.

(E) Toluidine blue–stained section of newborn femurs from Bmp2C/C; Prx1::cre animals.

(F–H) Marker analyses. Sections of femurs from Bmp2C/C; Prx1::cre animals hybridized with Col II (F), Col X (G), and Col I (H) mRNA probes.

(15 MB TIF)

Click here for additional data file.

Figure S3

BMP2, BMP4–Deficient Limb Skeleton Is Resorbed

Alcian blue– and Alizarin red–stained skeletons of forelimbs (A–D) and hindlimbs (E–F) from 1-wk-old (A, B, E, and F) and 3-wk-old (C and D) animals. (A, C, and E) Wild-type animals, (B, D, and F) Bmp2C/C; Bmp4C/C; Prx1::cre animals. The limbs of Bmp2C/C; Bmp4C/C; Prx1::cre (H) animals are severely defective compared to a wild-type (G) animal at 1 wk of age.

Please note that the proximal part of the femur is present at 1 wk of age but missing at 3 wks (refer to Figure 6L).

(12 MB TIF)

Click here for additional data file.

Figure S4

Osteogenesis Occurs in Mice Lacking Three of Four Alleles of Bmp2 and Bmp4 Combined

Toluidine blue–stained sections of femurs from adult control, Bmp2C/+; Bmp4C/C; Prx1::cre, and Bmp2C/C; Bmp4C/+; Prx1::cre animals.

(11 MB TIF)

Click here for additional data file.

Acknowledgements

We thank Dr. Andy Dudley for his help in designing mouse genotyping strategies. We also thank Jose Rivera-Feliciano and Dr. Douglas Kim for helpful discussions. We are immensely grateful to Drs. Brigid Hogan and Holger Kulessa for their kind gift of the Bmp4 conditional allele and Drs. Liz Robertson and Andy Dudley for the Bmp7 knockout allele. We would like to acknowledge the help of Dr. Ernestina Schipani and Jan Saxton for help with histology.

This manuscript is dedicated to the memory of Dr. Holger Kulessa, whose efforts contributed to the groundwork for this study and whose dedication and intense commitment to excellence in science are greatly missed.

Abbreviations

AER - apical ectodermal ridge

BMP - bone morphogenetic protein

E - embryonic day

FGF - fibroblast growth factor

SHH - Sonic Hedgehog

Shh - sonic hedgehog

ZPA - zone of polarizing activity

Figures and Tables

Figure 1

An Allelic/Nonallelic Series of BMP-Deficient Limbs

(A–D) Prx1::cre efficiently recombines Bmp2 and Bmp4 conditional alleles in the limbs. Bmp2 (A and B) and Bmp4 (C and D) whole mount mRNA in situ hybridization in the limb. Wild-type (A) and Bmp2C/C; Prx1::cre (B) are forelimbs from E10.5 mouse embryos. Mesenchymal expression [asterisk (A)] of Bmp2 is abolished in Bmp2C/C; Prx1::cre embryo while the AER expression [black arrow (A and B)] of Bmp2 persists. Note that pink staining in the central region of the limb bud in (B) is nonspecific background. Wild-type (C) and Bmp4C/C; Prx1::cre (D) are forelimbs from E10.5 mouse embryos. Mesenchymal expression [red arrow (C)] of Bmp4 is abolished in Bmp4C/C; Prx1::cre embryo while the AER expression [black arrow (C and D)] of Bmp4 persists.

(E–T) Depletion of BMP2 and BMP4 together causes severe limb skeletal defects. (E–T) Whole mount skeletons from newborn animals stained with Alcian blue and Alizarin red. (E–L) Forelimbs, (M–T) hindlimbs. (E and M) Wild-type, (F and N) Bmp2C/C; Prx1::cre, (G and O) Bmp4C/C; Prx1::cre, (H and P) Bmp7 −/−, (I and Q) Bmp2C/C; Bmp4+/C; Prx1::cre, (J and R) Bmp2+/C; Bmp4C/C; Prx1::cre, (K and S) Bmp2C/C; Bmp4C/C; Prx1::cre, (L and T) Bmp2C/C, Bmp7 −/−; Prx1::cre. Thin red arrow in (F), (I), and (L), defective scapula; thick red arrow in (T), failure of fibula to articulate with knee, and thick black arrow in (L) and (T), missing phalanx in digit III.

Figure 2

Depletion of BMP Signaling Causes Interdigital Syndactyly

(A–D) Forelimb of adult wild-type (A) and Bmp2C/C; Prx1::cre mouse (B) and hindlimbs of newborn wild-type (C) mouse and newborn Bmp2C/C; Bmp4C/C; Prx1::cre (D) mouse. The black arrow in (B) shows soft tissue syndactyly in Bmp2C/C; Prx1::cre mouse.

(E and F) Wild-type and Bmp2C/C; Bmp4C/C; Prx1::cre, respectively, showing acridine orange–stained hindlimbs of E15.5 mouse embryos. Acridine orange stain is in yellow.

(G and H) Enlarged views of selected regions from (E) and (F), respectively. Black arrow in (G) and (H) show acridine orange–stained apoptotic cells in the interdigital mesenchyme, and asterisk in (H) shows the remnant of the AER.

(I and J) Fgf8 mRNA expression in the hindlimbs of E13.5 wild-type (I) and Bmp2C/C; Bmp4C/C; Prx1::cre (J) embryos. The thick black arrows in (J) show Fgf8 mRNA expression.

Figure 3

Patterning Defects in Limbs Deficient of Different Combinations of BMP Molecules

(A–D) Bmp4 expression in limb buds from E11.5 wild-type (A and C) and Bmp2C/C, Bmp7 −/−; Prx1::cre (B and D) mouse embryos. (A and B) Forelimbs, (C and D) hindlimbs.

(E–H) Shh (E and F) and Fgf8 (G and H) expression in E11.5 forelimbs and hindlimbs, respectively. (E and G) Wild-type embryos, (F and H) Bmp2C/C; Bmp4C/C; Prx1::cre embryos.

(I–P) Sox9 (I–K) and Msx2 (M–O) expression in E12.5 wild-type (I and M) and Bmp2C/C; Bmp4C/C; Prx1::cre mouse embryonic forelimbs (J and N) and hindlimbs (K and O). (L and P) Shh expression in E12.5 wild-type (L) and Bmp2C/C; Bmp4C/C; Prx1::cre (P) embryonic hindlimbs.

The red brackets in (F) and (H) show the broadened domains of expressions of Shh and Fgf8, respectively.

Figure 4

Chondrogenesis Starts and Proceeds Normally Even in the Absence of BMP2 and BMP4

(A and B) Whole mount skeletons from E13.5 embryos that are stained with alcian blue. (A) Wild-type embryo, (B) Bmp2C/C; Bmp4C/C; Prx1::cre embryo. Black arrow in (B) shows the fusion of the zeugopod and stylopod.

(C and D) Hematoxylin and eosin–stained sagittal sections of humeri from E13.5 wild-type (C) and Bmp2C/C; Bmp4C/C; Prx1::cre (D) embryos. Thick red arrow in (C) shows the hypertrophic region.

(E and F) Sagittal sections of wild-type (E) and Bmp2C/C; Bmp4C/C; Prx1::cre (F) humeri from E15.5 embryos were hybridized with digoxigenin-labeled antisense rioprobes for ColX.

(G and H) Sagittal sections of forelimbs from E13.5 wild-type and BMP2, BMP4–deficient embryos, respectively, stained with radioactive riboprobes for Bmp7 mRNA. (G′) and (H′) show the bright field views of (G) and (H), respectively.

Figure 5

In the Absence of BMP2 and BMP4, Osteogenesis Begins During Early Embryonic Development

(A–D) Sagittal sections of forelimb from E15.5 embryos. (A and B) Stained with toluidine blue. (A) Distal ulna from wild-type embryo, (B) distal ulna/radius from Bmp2C/C; Bmp4C/C; Prx1::cre embryo. Mineralized cartilage is shown by dark purple and osteoid is shown by light blue (red asterisk). (C and D) Humeri sections were hybridized with digoxigenin labeled riboprobe for ColI. (C) Section of wild-type humerus from E15.5 embryo, (D) from Bmp2C/C; Bmp4C/C; Prx1::cre E15.5 embryo.

(E–H) Sagittal sections of zeugopod from E17.5 embryos. (E and F) Stained with toluidine blue while G and H are stained for TRAP (brown stain, red arrow). (E and G) E17.5 proximal radius from wild-type embryo, (F and H) E17.5 proximal ulna/radius from Bmp2C/C; Bmp4C/C; Prx1::cre embryo. Although vascularization occurs in the absence of BMP2 and BMP4, few osteoclasts are observed in the mineralized cartilage in Bmp2C/C; Bmp4C/C; Prx1::cre mice. Please note all the panels other than (G) and (H) are photographed at ×20, while (G) and (H) were photographed at ×40.

(I and J) Sections of newborn proximal radius (I) from control animal and newborn proximal ulna/radius (J) from Bmp2C/C; Bmp4C/C; Prx1::cre animal were stained with toluidine blue. Red arrow shows the bone marrow cavity in (E) and (I). Note that bone collar (red *) forms at the right time and place in the absence of BMP2 and BMP4. Also note that since Bmp2C/C; Bmp4C/C; Prx1::cre mice occasionally have only one bone in zeugopod and the elbow joint is occasionally fused, it is difficult to distinguish between the ulna and the radius.

Figure 6

Defects in Bone Formation in the Absence of BMP2 and BMP4

Toluidine blue staining of sagittal sections of distal femurs.

(A–E) Control femurs at 1 wk (A) and 3 wks (D) of age. Boxed areas in (A) are enlarged in (B) and (C). Boxed area in (D) is enlarged in (E). Blue arrows in (C) point to osteoblast cells lining the surface of cortical bone. Pink arrows in (G) show similar cells to those in (C) in Bmp2C/C; Bmp4C/C; Prx1::cre femurs. Red star in (A) and (D) marks the secondary ossification center. Pale blue–stained tissue in (E), marked by red arrows, is trabecular bone. bm, marks the bone marrow cavity in (D).

(F–J) Bmp2C/C; Bmp4C/C; Prx1::cre femurs at 1 wk (F) and 3 wks (J) of age. Boxed areas in (F) are enlarged in (G), (H), and (I). (A), (D), (F), and (J) are shown with the same magnification. Note that there are defects in bone formation but no defect in osteoclast mediated bone resorption. Mineralized tissue in the mid shaft of femur is almost resorbed at 3 wks. (H) Fibroblast-like cells present in the bone shaft of Bmp2C/C; Bmp4C/C; Prx1::cre mouse. Red star in (I) shows the osteoclasts invading the Bmp2C/C; Bmp4C/C; Prx1::cre femurs at 1 wk of age. s and m mark soft tissues and muscle in (J), respectively.

(K and L) Three-wk-old wild-type (K) and Bmp2C/C; Bmp4C/C; Prx1::cre (L) hindlimb skeletons stained with Alcian blue and Alizarin red. The black arrow shows the missing part of proximal femur.

Figure 7

Osteoblast Maturation Is Inhibited in the Absence of BMP2 and BMP4

(A–D) Distal femurs from control mice at 1 wk of age.

(E–H) Distal femurs from Bmp2C/C; Bmp4C/C; Prx1::cre mice at 1 wk of age.

(I–L) Distal femurs from control mice at 3 wks of age.

(M–P) Distal femurs from Bmp2C/C; Bmp4C/C; Prx1::cre mice at 3 wks of age.

In situ hybridization of early osteoblast differentiation-related marker genes Col I (B, F, J, and N), runx2 (C, G, K, and O) and osterix (D, H, L, and P).

Footnotes

Competing interests. The authors have declared that no competing interests exist.

A previous version of this article appeared as an Early Online Release on November 6, 2006 (doi:10.1371/journal.pgen.0020216.eor).

Author contributions. AB, KT, BDH, VR, and CJT conceived and designed the experiments. AB, KT, KC, and BDH performed the experiments. AB, KT, VR, and CJT analyzed the data. AB and KT wrote the paper.

Funding. This work was supported by a grant from the National Institutes of Health (P01 DK56246 to CJT) and by funds from the Forsyth Institute and Harvard School of Dental Medicine (to VR).


Combining global genome and transcriptome approaches to identify the candidate genes of small-effect quantitative trait loci in collagen-induced arthritis

Abstract

Quantitative traits such as complex diseases are controlled by many small-effect genes that are difficult to identify. Here we present a novel strategy to identify the candidate genes for small-effect quantitative trait loci (QTL) in collagen induced arthritis (CIA) using global genome and transcriptome approaches. First, we performed genome linkage analysis in F2 progeny of the CIA susceptible and resistant strains to search for small-effect QTL. Second, we detected gene expression patterns of both strains during CIA. The candidate genes were identified using three criteria: they are located in a genomic region linked to CIA; they are disease-specific differentially expressed during CIA; and they are strain-specific differentially expressed regarding the two parental strains. Eight small-effect QTL controlling CIA severity were identified. Of 22,000 screened genes, 117 were both strain-specific and disease-specific differentially expressed during CIA. Of these 117 genes, 21 were located inside the support intervals of the 8 small-effect QTL and thus were considered as candidate genes.

Introduction

Susceptibility to most complex diseases is controlled by many genes, each having a small effect on the disease. One example is rheumatoid arthritis (RA), a common complex multifactorial autoimmune disease. Several studies have been carried out to detect the genetic basis of RA, and more than 30 genomic regions have shown evidence of linkage to the disease. Most of these genomic regions did not reach a genome-wide significant threshold value of linkage, with P values between 0.05 and 0.001 [1-5]. Thus, these loci only have a small effect on RA. Small genetic contributions could also be seen from the susceptibility genes of RA identified so far, including HLA-DR4, PADI4, PTPN22 and FCRL3 [6-9]. Except for HLA-DR4, which is strongly associated with RA, all the other susceptibility genes have only a small effect on the disease. In the mouse model of RA, small genetic contributions are also often observed. For example, in a previous study, we carried out a genome screen to identify the quantitative trait loci (QTL) in collagen-induced arthritis (CIA), which is a widely used animal model of RA. Only one QTL, Cia2, was identified for the phenotype of CIA severity, but this QTL contributes to only 16% of the phenotype variations for CIA susceptibility in F2 progeny [10]. This suggests that there must be other susceptibility genes whose contributions were not big enough to reach the stringent significance threshold value of linkage analysis.

One aim of using animal models for complex diseases is to detect the genetic basis of these diseases. With controllable environmental factors as well as the known genetic background, animal models are powerful tools to search for susceptibility genes for complex diseases, and have been intensively employed for that purpose. More than 27,000 QTL have been identified in the mouse genome since the first QTL was identified at the beginning of the 1990s [11]. By 2005, approximately 20 quantitative trait genes (QTGs) in the mouse genome had been identified [12,13]. Interestingly, most QTGs identified in animal models have the causal polymorphisms in the protein-coding region [14], which provoke protein structure changes or protein deficiency. This suggests, on the one hand, that small-effect QTL are difficult to identify with traditional strategies and, on the other hand, that the polymorphisms regulating gene expression might only slightly affect the quantitative traits, and thus are more difficult to identify.

Microarray-based global gene expression is a powerful technique for investigating complex diseases. During disease development, genes involved in the disease are likely to be differentially regulated. Therefore, signature genes of the diseases could be identified by detecting the expression patterns of the disease-related cells/tissues and their ideal controls. In the past decade, many studies applied this technique to study both RA and its animal models [15-22]. Indeed, genes involved in arthritis show distinct expression patterns in certain tissues and pathological stages of the disease. Genes involved in immunoinflammatory responses were differentially expressed in the blood cells in RA patients [18]. Chemokines and adhesion molecules were upregulated in the joint at the initiation phase of arthritis in animal models [21,22], while genes involved in cartilage destruction and bone erosion were differentially expressed at the late phase of arthritis in animal models of RA [15,16]. Besides detecting genes involved in complex diseases, microarrays could also be used to detect the genetic polymorphisms regulating gene expression because differential expressions between two strains might be the result of a polymorphism located in regulatory elements.

To identify the small-effect QTL of CIA as well as the potential candidate genes inside them, we investigated CIA genetically susceptible and resistant strains at both the genome and transcriptome levels. At the genome level, F2 progeny of the CIA susceptible (DBA/1) and resistant (FVB/N) strains were generated and a genome-wide linkage analysis was performed to identify small-effect QTL. At the transcriptome level, we detected the gene expression patterns of both the DBA/1 and FVB/N strains at four different phases of CIA. The potential candidate genes were identified based on three criteria: they are located within the genomic region linked to CIA; they are disease-specific differentially expressed during CIA; and they are strain-specific differentially expressed between the two parental strains during CIA.

Materials and methods

Animals, immunisation and assessment of arthritis

Both DBA/1 and FVB/N mice used in this study were obtained from the Jackson Laboratory and kept in a climate-controlled environment with 12 hour light/dark cycles in the animal facility at the University of Rostock. All animal experiments were pre-approved by the State Animal Care Committee. CIA was induced in control and experimental animals according to established protocols described previously [10]. In brief, DBA/1J, FVB/N and (DBA/1J × FVB/N)F2 progeny were immunised at 8 to 12 weeks at the base of the tail with 125 μg of bovine Collagen II (Chondrex, Redmond, WA, USA) emulsified in CFA (DIFCO, Detroit, MI, USA). The clinical scoring of arthritis commenced 18 days after immunisation, and animals were monitored three times weekly for signs of CIA. Arthritis development was monitored in all four limbs using a three-score system per limb as described previously [10].

Eight-week old FVB/N and DBA/1J mice were used for the detection of gene expression. They were divided into four experimental groups according to the different phases of CIA, namely naive control (NC), post-immunisation (PI), onset of arthritis (OA) and chronic arthritis (CA) (Table 1). The NC group contained non-immunised mice that were sacrificed at the age of 8 weeks. The mice in the PI group were sacrificed on day 10 after immunisation. The mice in the OA group were sacrificed on day 35 after immunisation. Three FVB/N non-arthritic mice and three DBA/1 mice that showed signs of arthritis on day 33 or 34 after immunisation were sacrificed on day 35 after immunisation. The mice in the CA group were sacrificed on day 95 after immunisation. Three non-arthritic FVB/N mice and three DBA/1 mice that had developed arthritis for at least two months were sacrificed on day 95 after immunisation.

Linkage analysis

Detailed information on genotyping of the genome screen has been described previously [10]. In short, we genotyped 290 F2 mice using 126 informative microsatellite markers covering the genome with an average inter-marker distance of 11.5 cM for 290 F2 progeny. All linkage analyses were performed with QTX Map manager software [23]. The main clinical phenotype of CIA, arthritis severity, was taken as phenotype. To detect the small-effect QTL, the threshold value of linkage was set as P = 0.05 (Chi-square test).

Sample preparation and microarray hybridisation

Lymph nodes (LNs) draining the immunisation site were used for total RNA preparation. The total RNA was extracted from the tissue homogenates using a commercial kit in accordance with the provided protocol (QIGEN, Hilden, Germany). Analysis of gene expression was conducted using a U430A array (Affymetrix, Santa Clara, CA, USA) interrogating more than 22,000 genes. RNA probes were labelled in accordance with the manufacturer's instructions. Samples from individual mice were hybridised onto individual arrays. Hybridisation and washing of gene chips were done as previously described [16]. Fluorescent signals were collected by laser scan (Hewlett-Packard Gene Scanner).

Microarray analysis

Normalisation of the expression level was done using Affymetrix software MAS 5, which is based on global scaling of total gene expression level per microarray. The normalised expression values were imported to and analysed by dCHIP [24]. Differentially expressed genes were identified by defining the appropriate filtering criteria in the dCHIP software as: lower 90% confidence boundary of fold change between the group means exceeded twofold; the absolute difference between the two groups exceeded 100; the P value threshold of the unpaired t-test was 0.05. The false discovery rate was established with a permutation test for each pairwise comparison to estimate the proportion of false-positive genes.

Hierarchical gene clustering was performed with dCHIP to characterise the gene expression patterns during CIA. The default clustering algorithm of genes was as follows: the distance between two genes is defined as 1 – r, where r is the Pearson correlation coefficient between the standardised expression values of the two genes across the samples used. To characterise the functional relationship between differentially expressed genes, Gene Ontology (GO) term classification incorporated in DNA-Chip Analyzer was performed. The significant level for a function cluster was set at P < 0.005, and the minimum size of a cluster was three genes.

Results

Small-effect QTL of CIA in (DBA/1 × FVB/N) F2 progeny

In a previous study, we carried out a genome screen to identify QTL controlling CIA susceptibility in (DBA/1 × FVB/N) F2 progeny. For the phenotype of arthritis severity, only one QTL, Cia2, was identified, with a highly significant logarithm of the odds (LOD) score of 12 [10]. However, Cia2 contributed to only 16% of the phenotype variations, indicating that there should be some small-effect QTL whose contributions to CIA were not big enough to reach the significant threshold value of linkage. To identify these potential small-effect QTL, we reanalyzed the data using a lower threshold value of linkage (P = 0.05). We reasoned that since the main candidate gene of Cia2, complement component C5 (Hc), was proven to be essential for CIA development and because the FVB/N strain is C5 deficient [10,25], some small-effect QTL might be masked by Cia2. To exclude the masking effect of C5, we performed linkage analysis with 3 datasets, the first containing all 290 F2 progeny, the second 77 C5+/+ F2 progeny and the third 133 C5+/- F2 progeny. Eight genomic regions were linked to the phenotype of CIA severity (loci 1 to 8, Table 2), with P values varying between 0.043 and 0.003. These eight small-effect QTL were located on chromosomes 5, 6, 7, 10, 11, 16, and 17. Five loci were identified in at least two datasets. Of the eight loci, five had DBA/1 as the susceptibility allele, and three had FVB/N as the susceptibility allele.

Lander and Botstein [26] suggested a LOD score of between 2 and 3 to ensure an overall false positive rate of 5%, which means that using a lower threshold value will prevent false negative QTL at the expense of increasing false positive QTL. Being aware of this, we examined these genomic regions to search whether they, or their syntenic genomic regions on the human genome, have been previously linked to arthritis. Four small-effect QTL overlapped with arthritis QTL on the mouse genome identified previously. Locus 1 and 2 overlap with Cia13 and Cia14, which control severity of CIA in (DBA/1 × BALB/C) F2 progeny [27]. Locus 5 located on chromosome 10 overlaps with Cia8, which was identified in (DBA/1 × B10.Q) F2 progeny [28]. Locus 6 overlaps with Pgia7, which controls susceptibility to proteoglycan-induced arthritis (PGIA) and was identified in (BALB/C × DBA/2) F2 progeny [29]. The syntenic genomic regions of five small-effect QTL on the human genome have been reported to be linked to RA. These are genomic regions 22q11 and 12p13-q24 on chromosome 22 and 12 (the counterparts of locus 2), 12p13-pter on chromosome 12 (the counterpart of locus 3), 21q22-qter and 10q22-23 on chromosome 21 and 10 (the counterparts of locus 5), 17q21-25 on chromosome 17 (the counterpart of locus 6) and 3q29-qter on chromosome 3 (the counterpart of locus 7) [2,4].

Strain-specific differentially expressed genes

We detected the gene expression profiles using three mice per group, which is a small number. Being aware of the importance of data reproducibility, we determined the coefficient of variation (CV) to measure data variability. The CV for each gene on the chip and the mean CV for the entire probe set were calculated. The mean CV ranged between 18.4% and 25.8% for all experimental groups, and this relatively low CV indicated that these data could be used for further analysis (Table 1).

To search for strain-specific differentially expressed genes, we performed comparisons of gene expression between the DBA/1 and FVB/N strains at all four phases of CIA, including NC, PI, OA and CA. For the naive mice without immunisation, 361 genes were differentially expressed between the two strains. On day 10 after immunisation, when both strains did not show any sign of the disease, 141 genes were differentially expressed. After DBA/1 mice developed CIA, 184 and 85 differentially expressed genes were identified between these two strains at the onset and chronic phases, respectively. When the lists of the differentially expressed genes at the four phases were merged and overlapping genes were excluded, 509 genes were identified (Additional file 1). Twenty-one genes consistently showed differential expression between the two strains at all phases. Besides these 21 genes, only 3 additional genes were strain-specific differentially expressed during the 3 phases after CIA induction (PI, OA and CA; Figure 1).

Disease-specific differentially expressed genes

To identify the disease-specific differentially expressed genes in CIA, we detected the genes that were differentially expressed in LNs during CIA in the susceptible strain. Three experimental conditions, PI, OA and CA, were compared with the NC group. On day 10 after immunisation, 102 genes were differentially expressed – most of them were upregulated (78 out of 102) – while at the onset phase of the disease, only 26 genes were differentially expressed. At the chronic phase of the disease, 184 differentially expressed genes were identified, with 156 downregulated genes. Only one gene was differentially expressed at all three phases of CIA. Besides this gene, five, one and six differentially expressed genes were shared by PI with OA, PI with CA and OA with CA, respectively (Figure 2a). Taken together, 310 disease-specific differentially expressed genes were differentially regulated during CIA in DBA/1 mice (Additional file 2).

To further characterise the gene expression pattern during CIA, we performed hierarchical cluster analysis for these 310 genes. Six gene clusters were identified (clusters I to VI, Figure 2b), each with a distinct gene expression pattern during CIA. Cluster I contains 16 genes, representing genes that were upregulated after induction of CIA. The expression of these genes reached a peak at the onset phase of the disease and functional clustering results revealed that they are related to the immune response. Cluster II contains 12 genes whose expression was gradually upregulated and reached a peak at the chronic phase of CIA. These genes are mainly related to the immune response, organelle membrane and extracellular region and space. Cluster III contains 78 genes that were only upregulated at the PI phase. These genes are related to the intercellular junction. More than half of the genes (156 of 310) belong to cluster IV and represent genes specifically downregultaed at the chronic phase. These genes are functionally related to lymphocyte proliferation, T cell activation, protein binding as well as the notch signal pathway. Cluster V contains eight genes downregulated at the PI phase. Cluster VI contains 18 genes downregulated at the OA phase. The GO term classification showed no functional cluster that was significantly enriched in these two gene clusters.

Candidate genes for the small-effect QTL of CIA

To identify candidate susceptibility genes for the CIA small-effect QTL, we compared the list of strain-specific differentially expressed genes with the list of disease-specific differentially expressed genes; 117 genes were shared by both lists (Additional file 3). Figure 3 visualises positions of the 117 genes retrieved from Ensembl [30] in relation to the 8 small-effect QTL. The eight loci were located on 7 chromosomes, 5, 6, 7, 10, 11, 16 and 17. Since the confidence intervals of QTL in F2 progeny are around 20 cM [26], we used 40 Mb as the confidence intervals for all loci. Twenty-one genes were located in the confidence intervals of six of the eight QTL. We located 5, 4, 2, 1, 3 and 6 potential candidate genes within the confidence intervals of loci 1, 2, 3, 5, 6 and 8, respectively, while no candidate gene was identified for loci 4 and 7. Table 3 summarises the 21 candidate genes identified in this study. Two genes, hspa1a and Oas1a, were upregulated at the OA phase of CIA and Oas1a was also upregulated at the PI phase. Except for these two genes, all other 19 genes were downregulated at the chronic phase of CIA. All genes, except hspa1a, showed expression differences between the two strains at the NC phase. Five genes were differentially expressed at all phases of CIA, including H2-Q10, Mapk14, Pscd1, Kpnb1 and Wdr1. Among these five genes, H2-Q10 had a consistently higher expression in the DBA/1 than the FVB/N strain in all CIA phases, while the other four genes had a higher expression in the DBA/1 strain at the early stages, including NC, PI and OA, but a lower expression at the chronic phase. GO term classification analysis revealed that the functional cluster of protein kinase cascade was significantly enriched in the 21 candidate genes. This functional cluster contained four genes, including Mapk14, Mapk8ip3, Stat5a and Gna12.

Discussion

In this study, we attempted for the first time to identify small-effect QTL in an F2 progeny. Small-effect QTL are defined as those reaching the threshold value of P = 0.05 but that did not reach the significant threshold value suggested by Lander and Botstein [26]. Although not significant, there is evidence that most of the eight small-effect QTL likely contain susceptibility genes for CIA. First, we performed the linkage analysis in three datasets, including all 290 F progeny, 76 C5+/+ F2 progeny and 133 C5+/- F2 progeny. Five of the eight small-effect QTL were identified in at least two datasets, suggesting that these QTL are reproducible. Second, many QTL identified in the present study overlap with arthritis QTL previously identified, including loci 1, 2, 5 and 6. In addition, syntenic analysis revealed that the counterpart genomic regions on the human genome of many of these eight QTL are linked to RA.

For five of the eight small-effect QTL the DBA/1 alleles are the arthritis-enhancing alleles, while the FVBN alleles are the arthritis-enhancing alleles in the other three QTL, indicating that some susceptibility genes could come from the resistant strain. Interestingly, loci 2 and 7 partially overlap with two arthritis-related QTL identified by us in the same F2 progeny [10]. Locus 2 was located at the same genomic region as Cia27, a QTL controlling IgG2a antibody levels to collagen II. Recently, we have refined this QTL into a 4.1 Mb genomic region and showed that a gene within this region regulates CIA severity by controlling the IgG2a antibody levels to collagen II [31]. Locus 7 on chromosome 16 overlaps with Lp1, which controls lymphocyte proliferation. Furthermore, according to our unpublished data, loci 8 on chromosome 17 controls lymphocyte adherence during development of CIA. Therefore, the gene within the small-effect QTL could affect CIA severity through controlling arthritis-related phenotypes.

Several studies have been carried out to detect gene expression during CIA, all of which used joints as the target tissue [15,16,21,22]. This study, for the first time, detected gene expression in LNs during CIA. We present an extensive study of gene expression patterns in LNs of both genetically susceptible and resistant strains at four different phases of CIA. In both strains, differentially regulated genes were highly concentrated at the PI and CA phases, and only a small number of genes were differentially expressed in two or three phases. This indicates that biological responses in LNs were stronger in the PI and CA phases than in the OA phase, and the responses at different phases were different. When comparing the susceptible to the resistant strain, the biggest difference was found in one cluster of genes (cluster IV, Figure 2). These genes had a higher expression in DBA/1 than in FVB/N at the early phases of CIA (NC, PI and OA) and the opposite expression pattern in the CA phase. GO term classification revealed that these genes were related to lymphocyte proliferation and activation, suggesting that lymphocytes in the DBA/1 strain are more activated than those in the FVB/N strain. However, this difference is not CIA specific because the expression difference between the two strains existed in mice without immunisation. Additionally, some genes related to the immune response were upregulated in the DBA/1 strain but not in the FVB/N strain during CIA. These differences could explain why a higher antibody response to collagen II occurred in the DBA/1 strain compared to the FVB/N strain, and might partially explain the difference of the susceptibility to CIA between both strains.

Twenty-one genes were identified as potential candidate genes for six of the eight small-effect QTL according to their gene expression patterns during CIA and their genomic locations. No candidate genes were located in QTL 4 and 7, suggesting that QTG polymorphisms of the susceptibility genes inside these two QTL might not affect the phenotype by regulating gene expression. Two of the 21 candidate genes were reported to be involved in arthritis. Mapk14, a candidate gene for locus 8 and also called p38 mitogen-activated protein kinase (MAPK) alpha, regulates the production of arthritis-essential cytokines, such as tumour necrosis factor and interleukin-1 [32]. Moreover, inhibitors of p38 MAPK could attenuate CIA in rats [33], and p38 MAPK is becoming a potential therapeutic target in RA [32]. Stat5a, a candidate gene for loci 6, is an essential molecule for lymphoid development and differentiation [34]. Stat5a-deficient mice were reported to lose tolerance, resulting in the development of autoimmune diseases. Stat5a is suggested to contribute to tolerance through maintenance of the CD4+CD25+ regulatory T cell population [35].

Therefore, we have presented a strategy to identify small-effect QTL and search for potential candidate genes within them. However, it is noteworthy that the low statistical threshold and small number of animals per group could lead to some false positive results. On the genome level, some of the eight small-effect QTL identified using a very low threshold value (P < 0.05) could be false positives. For example, locus 4 was identified with a low P value and does not overlap with any previously identified arthritis QTL. On the transcriptome level, the small number of animals per group and the low threshold used to detect gene expression could also result in false positives in the differentially expressed genes. Furthermore, the differential expression of a gene could result not only from allele difference between two strains, but also from other factors. Therefore, our findings should be confirmed in future studies.

Conclusion

We present a strategy to search candidate genes for small-effect QTL. With this strategy, we identified 21 candidate genes for 8 small-effect QTL regulating CIA susceptibility. Our future studies will be carried out using two approaches. The first is generating congenic animals for promising small-effect QTL that have relatively high P values and overlap with previously-identified arthritis QTL. The second approach is investigating candidate genes using both mouse and human studies. Candidate genes will be selected according to their function and polymorphism between the two strains. Thereafter, we will generate knock-out mice to investigate the role of the genes in CIA. For the loci whose counterparts on the human genome are linked to RA, we will investigate the candidate genes using case-control association studies in RA cohorts.

Abbreviations

CA = chronic arthritis; CIA = collagen-induced arthritis; CV = coefficiant of variation; GO = Gene Ontology; LN = lymph node; LOD = logarithm of the odds; MAPK = mitogen-activated protein kinase; NC = naive control; OA = onset of arthritis; PI = post-immunisation; QTG = quantitative trait gene; QTL = quantitative trait loci; RA = rheumatoid arthritis.

Competing interests

The authors declare that they have no competing interests.

Authors' contributions

XY and KB performed the animal experiments. DK and HJT performed gene expression profiling experiments. XY and DK performed the bioinformatic analysis. SI conceived and designed the experiment. XY and SI drafted the manuscript. All authors read and approved the final manuscript.

Supplementary Material

Additional file 1

Summary information on the 509 genes differentially expressed between DBA/1 and FVB/N strains. Comparison was performed between two strains at four stages of CIA, NA, PI, OA and CA. Three criteria were applied for selecting the differentially expressed genes: the lower 90% confidence bound of fold change between the group means exceeded twofold; the absolute difference between the two groups exceeded 100; the P value threshold of the unpaired t-test was 0.05

Click here for file

Additional file 2

Summary information on the 311 genes differentially expressed in joints in the DBA/1 strain during CIA. Pairwise comparisons were performed by comparing PI, OA and CA with NA. Three criteria were applied for selecting the differentially expressed genes: the lower 90% confidence bound of fold change between the group means exceeded twofold; the absolute difference between the two groups exceeded 100; the P value threshold of the unpaired t-test was 0.05

Click here for file

Additional file 3

Summary information on the 117 genes that were strain-specific differentially expressed and were dysregulated in the DBA/1 strain during CIA. The physical positions of the genes were retrieved from Ensembl [30]

Click here for file

Acknowledgements

The authors wish to thank Ilona Klamfuss for animal care. This work was supported by a grant from the EU FP6 (MRTN-CT-2004-005693, EURO-RA).

Figures and Tables

Figure 1

Differentially expressed genes between collagen-induced arthritis (CIA) susceptible and resistant strains. Comparison of the gene expression between the DBA/1 and FVB/N strains was performed at four phases of CIA. Of the 22,000 screened genes, 509 were differentially expressed between both strains at one or more phases of CIA, including 361 genes at the naive control (NC) phase, 141 genes at post-immunisation (PI) phase, 184 genes at the onset of arthritis (OA) phase and 85 genes at the chronic arthritis (CA) phase. The Venn diagram indicates the number of overlapping genes differentially expressed at different phases of CIA.

Figure 2

Differentially expressed genes in the DBA/1 strain during collagen-induced arthritis (CIA). Three experimental conditions, post-immunisation (PI), onset of arthritis (OA) and chronic arthritis (CA), were compared with naïve control (NC) to search for differentially expressed genes. (a) Venn diagram indicating the number of overlapping genes differentially expressed at three phases of CIA. (b) Hierarchical clustering of the 310 differentially expressed genes. The left panel shows the distribution of relative gene expression across the hierarchical tree structure. Rows represent individual genes; columns represent individual value of triplicate samples for each experimental group. Each cell in the matrix represents the expression level of a gene, with red and green indicating transcription levels above and below the normal values for that gene, respectively. Four sample groups are indicated above the expression matrix. Six basic gene clusters (clusters I to VI) were yielded by the analysis according to the gene expression pattern during CIA. The expression patterns of the gene cluster are graphed and the major biological activities for each cluster that were examined by functional clustering analysis are indicated on the right.

Figure 3

Visualisation of all the chromosomal locations of the small-effect QTL identified in this study (blue bar) as well as 120 genes of interest from gene expression profiling (black letters). The positions of the 120 genes and the peak markers of the QTL were retrieved from Ensembl [30]. Confidence intervals of all the QTL were set as 40 Mb.

Table 1

Experimental groups used for gene expression profiling

CA, chronic arthritis; CFA; complete Freund's adjuvant; CV, coefficient of variation; LN, lymph node; NC, naive control; OA, onset of arthritis; PI, post-immunisation.

Table 2

Summary of the small-effect QTL identified in this study

aIdentified in all F2 290 progeny. bIdentified in 76 C5+/+ F2 progeny. cIdentified in 133 C5+/- F2 progeny. QTL, quantitative trait loci; RA, rheumatoid arthritis.

Table 3

Summary of the small-effect QTL candidate genes

aFold change calculated by comparing DBA/1 with FVB/N. bFold change calculated by comparing post-immunisation (PI), onset of arthritis (OA) and chronic arthritis (CA) with naive control (NC), respectively. Chr., chromosome; CIA, collagen-induced arthritis.


Pygo1 and Pygo2 roles in Wnt signaling in mammalian kidney development

Abstract

Background

The pygopus gene of Drosophila encodes an essential component of the Armadillo (β-catenin) transcription factor complex of canonical Wnt signaling. To better understand the functions of Pygopus-mediated canonical Wnt signaling in kidney development, targeted mutations were made in the two mammalian orthologs, Pygo1 and Pygo2.

Results

Each mutation deleted >80% of the coding sequence, including the critical PHD domain, and almost certainly resulted in null function. Pygo2 homozygous mutants, with rare exception, died shortly after birth, with a phenotype including lens agenesis, growth retardation, altered kidney development, and in some cases exencephaly and cleft palate. Pygo1 homozygous mutants, however, were viable and fertile, with no detectable developmental defects. Double Pygo1/Pygo2 homozygous mutants showed no apparent synergy in phenotype severity. The BAT-gal transgene reporter of canonical Wnt signaling showed reduced levels of expression in Pygo1-/-/Pygo2-/- mutants, with tissue-specific variation in degree of diminution. The Pygo1 and Pygo2 genes both showed widespread expression in the developing kidney, with raised levels in the stromal cell compartment. Confocal analysis of the double mutant kidneys showed disturbance of both the ureteric bud and metanephric mesenchyme-derived compartments. Branching morphogenesis of the ureteric bud was altered, with expanded tips and reduced tip density, probably contributing to the smaller size of the mutant kidney. In addition, there was an expansion of the zone of condensed mesenchyme capping the ureteric bud. Nephron formation, however, proceeded normally. Microarray analysis showed changed expression of several genes, including Cxcl13, Slc5a2, Klk5, Ren2 and Timeless, which represent candidate Wnt targets in kidney development.

Conclusion

The mammalian Pygopus genes are required for normal branching morphogenesis of the ureteric bud during kidney development. Nevertheless, the relatively mild phenotype observed in the kidney, as well as other organ systems, indicates a striking evolutionary divergence of Pygopus function between mammals and Drosophila. In mammals, the Pygo1/Pygo2 genes are not absolutely required for canonical Wnt signaling in most developing systems, but rather function as quantitative transducers, or modulators, of Wnt signal intensity.

Background

Wnt signaling is of critical importance in several stages of kidney development. Mutual inductive interactions between the ureteric bud and metanephric mesenchyme drive nephrogenesis [1]. The ureteric bud synthesizes Wnt9b, which is essential for induction of the mesenchyme to form nephrons [2]. Wnt4 is made by the induced metanephric mesenchyme and is also required for nephrogenesis [3]. Furthermore, Wnt11, secreted by the ureteric bud tips, participates in a positive feedback loop promoting glial cell line-derived neurotrophic factor (GDNF) expression by the metanephric mesenchyme [4]. Mutations in Wnt9b or Wnt4 result in a dramatic block in nephron formation, while Wnt11 mutants show a significant reduction in nephron number. It is interesting to note that Wnt4 and Wnt11 have been shown to signal, at least in some cases, through noncanonical pathways [5-7], while there is evidence indicating that Wnt9b activates canonical Wnt signaling in the kidney [2].

Genetic studies in Drosophila have identified the Pygopus (Pygo) gene as a critical component of canonical Wnt signaling [8-11]. Pygo and Lgs interact with β-catenin during the formation of the canonical transcriptional complex and are required for accumulation of β-catenin in the nucleus [12]. Lgs binds the central armadillo repeats of β-catenin, while Pygo interacts with Lgs, mediating activation of Wnt targets [9,13]. The N-terminal domain of Pygo is required for Wnt transcriptional activation, while the PHD motif is required for the association of Pygo with Lgs [9,13]. Additionally, a putative nuclear localization signal (NLS) was identified within the N-terminal domain of Pygo, suggesting a possible role of nuclear importation of β-catenin [8,11]. Analyses of multiple aspects of the pygopus mutant phenotype indicate that this gene is dedicated to, and required for, canonical Wnt signaling during Drosophila development [9].

The mammalian genome carries two, and only two, orthologs of Drosophila Pygo, Pygo1 and Pygo2 [8,9,14]. In this report, we generated targeted mutations of Pygo1 and Pygo2 to determine their functions, with a particular interest in the contributions of these genes to canonical Wnt signaling during kidney development. The resulting double-homozygous mutant embryos showed a context-dependent reduction in canonical Wnt signaling as measured by Wnt reporter transgene expression. Development remained, however, surprisingly normal, with survival to birth and few apparent defects in most organ systems. Our phenotypic analysis focused on the kidney, which showed altered branching morphogenesis of the ureteric bud, and expansion of the zone of condensed mesenchyme surrounding the ureteric bud, yet relatively normal nephron formation, as measured by histology, confocal analysis, in situ hybridization and microarray analysis. The obvious conclusion is that in mammals, unlike Drosophila, Pygopus-mediated canonical Wnt signaling is not absolutely necessary in most developing organ systems.

Results

Pygo1 and Pygo2 targeted mutations

We targeted both the Pygo1 and Pygo2 genes by inserting LoxP sequences to flank critical coding regions including the PHD domains. The resulting targeted mice were mated with transgenic CMV-Cre mice [15] to drive germline LoxP recombination, resulting in the null mutant alleles that were used for this study. PCR confirmed the deletion of the bulk of the coding sequences for both genes, including 89% of coding sequence for Pygo1 and 87% of coding sequence for Pygo2. Previous studies in Drosophila have shown that even a single missense mutation in the PHD domain can eliminate Pygo function in Wnt signaling [8].

Pygo1 and Pygo2 mutant phenotypes

Pygo1 homozygous null mice were viable and fertile with no developmental defects detectable. This was surprising given the importance of the pygopus gene in Wnt signaling in Drosophila, and the reported expression during development of the mouse Pygo1 gene in, for example, the brain, limbs, kidney and branchial arches [14,16] (Yu J, Valerius MT, McMahon AP, contribution to GUDMAP, ).

The Pygo2 homozygous null mice survived to birth, but with rare exceptions, died shortly afterwards. The gut, heart and limbs developed without detectable abnormality (data not shown) despite known requirements for Wnt signaling. The Pygo2 mutants did, however, show growth retardation, lens agenesis and a kidney phenotype with high penetrance, exencephaly, and cleft palate with incomplete penetrance.

Double-homozygous mutant Pygo1 and Pygo2 mice had a phenotype similar to that of single Pygo2 nulls (Figure 1, Table 1). There was no significant synergism of developmental abnormalities in the double mutants. Together these results suggest that Pygo2 is required for the proper development of a limited number of structures, whereas Pygo1 is not necessary for normal development.

Figure 1

Pygo1/Pygo2 mutant embryos. (A) E18.5 Pygo1-/-/Pygo2+/- embryos, with only one wild-type Pygo2 allele, appeared normal. (B, C) Double homozygous mutant Pygo1-/-/Pygo2-/- embryos were smaller, with eye defects including absent or rudimentary lens and folded pigmented retina (arrows). (C) A small percentage of Pygo1/Pygo2 null embryos also displayed exencephaly.

Table 1

Phenotypes of Pygo2 wild-type, heterozyogous, and null E18.5 embryos on a Pygo1 null background.

* Denotes a significant change (p < 0.01) of the average weight of the Pygo2 null group compared with both the Pygo wild-type and heterozygous groups.

Expression of Pygo1 and Pygo2 in the developing kidney

In situ hybridization has been used previously to characterize expression of Pygo1 [14,16](Yu J, Valerius MT, McMahon AP, contribution to GUDMAP, ). In this report, we used immunofluorescence to better define the expression patterns of the Pygo1 and Pygo2 genes in the developing kidney. Both genes were widely expressed, showing nuclear localization of encoded proteins in the ureteric bud, capping mesenchyme, and stromal cells (Figure 2). Raised expression of Pygo1, and to a lesser degree of Pygo2, was seen in stromal cells, and significant nuclear staining was detected in essentially all cells of the developing kidney for both proteins.

Figure 2

Pygo1 and Pygo2 expression in the E18.5 developing kidney. Immunofluorescence was used to determine expression patterns of the Pygo1 and Pygo2 proteins in the cortex of the E18.5 kidney. Both Pygo1 and Pygo2 (red) were localized in the nucleus, as expected. Both genes showed widespread expression, with signal detected in all components of the developing kidney, but with elevated levels in the stromal cell compartment (arrows). Epithelial cells, primarily ureteric bud in these sections, were labeled blue using E-cadherin antibody. Original magnification × 200.

Confocal analysis of Pygo1/Pygo2 mutant kidneys

Histological examination of Pygo2 null and Pygo1/Pygo2 double-null mutant kidneys did not reveal any abnormalities in nephrogenesis (data not shown). Confocal analysis was therefore performed to characterize renal development in E18.5 Pygo1/Pygo2 null kidneys more precisely (Figure 3). Wt1 (red) and Cited1 (red) antibodies both stain the capping metanephric mesenchyme around the ureteric bud tips [17]. WT1 antibody also stains renal vesicles and glomerular anlage. Antibodies to Cdh1, also known as E-cadherin (blue), were used to identify epithelial structures, including the branching ureteric bud and nascent nephrons of the developing kidney [18]. Dolichos biflorus (DBA) lectin [16] was used to selectively stain ureteric bud-derived structures [19].

Figure 3

Confocal analysis of Pygo1/Pygo2 mutant E18.5 kidneys. (B, D) Pygo1-/-/Pygo2-/- mutant E18.5 kidneys and (A, C) kidneys of normal littermates were stained using antibodies to (C, D) Cited1 or (A, B) Wt1, both expressed in the condensing metanephric mesenchyme, and colored as red, Cdh1, a general marker of epithelia (blue), and DBA lectin staining the ureteric tree [16]. Confocal Z-sections were obtained every 5 μm for 75–80 μm. A reduced number of ureteric tips per area was observed in (B, D) the Pygo1/Pygo2 null kidneys compared with (A, C) control littermates. In addition, ureteric bud tips were often more dilated in Pygo1/2 null mutants compared with controls. The condensed mesenchyme surrounding the ureteric bud was also significantly expanded in the mutants. Original magnification × 200.

Wt1 and Cited1 (red) staining revealed an increase of approximately 30% in the thickness of the capping mesenchyme surrounding the mutant ureteric buds (Figure 3, Table 2). Nevertheless, the mutant metanephric mesenchyme underwent relatively normal nephrogenesis. Cdh1-staining nephrons (blue) were identified connecting to the ureteric bud tips [16] and extending into the medulla of the kidney, as normal. Intermediate structures of nephrogenesis, including renal vesicles, comma-shaped bodies, and S-shaped bodies appeared normal.

Table 2

Confocal analysis of Pygo2 wild-type, heterozygous, and null E18.5 kidneys on the Pygo1 null background.

*Significant change (P < 0.01) between the three groups.

SD, Standard deviation; n, number

Confocal analysis also showed that the ureteric bud tips [16] of Pygo1/Pygo2 null kidneys were dilated and misshaped compared with those of littermates with at least one wild-type Pygo2 allele (Figure 3). In addition, the Pygo1/Pygo2 double-homozygous mutant kidneys also had a decrease of approximately 25% in the number of ureteric bud tips per area compared with littermates with at least one wild-type copy of Pygo2 (Table 2). No significant difference in ureteric bud-tip density was seen between Pygo2+/- and Pygo2+/+ embryos (P = 0.33).

In situ hybridization

We examined expression of three Wnt genes in Pygo mutants. Wnt7b is expressed in the stalks [20], Wnt11 in the tips [4], and Wnt9b in the stalks and weakly in the tips of the branching ureteric bud [2]. All three genes showed similar expression patterns in Pygo2+/+ and Pygo1/Pygo2 double-mutant kidneys (Figure 4). In addition, these in situ hybridization patterns confirmed the confocal microscopy results, showing a reduced number of tips (Wnt11-positive) per surface area in the Pygo1/Pygo2 double-homozygous mutants.

Figure 4

Expression of Wnt7b, Wnt9b, and Wnt11 in E18.5 Pygo1/Pygo2 compound null kidneys. Whole-mount in situ of hybridizations of (A-C) E18.5 wild-type kidneys and (D-F) Pygo1-/-/Pygo2-/- mutant kidneys with the ureteric bud derivative markers: (A, D) Wnt7b, (B, E) Wnt9b, and (C, F) Wnt11. The mutant kidneys showed normal expression patterns for the ureteric stalk markers Wnt7b and Wnt9b, and reduced density of ureteric tips as measured by Wnt11. Original magnification × 32.

Reduced canonical Wnt signaling in Pygo1/Pygo2 mutant mice

The BAT-gal transgene reporter of canonical Wnt signaling [21] was used to examine changes in Wnt signaling in the mutant mice. Both Pygo2-/- and Pygo1/Pygo2 double-null E10.5 embryos showed a decrease in canonical Wnt signaling (Figure 5). There was, however, tissue-specific variability in the degree of reduction, with the somites, for example, showing strong reduction, while the mutant telencephalon still showed robust Wnt signaling. These results suggest that the mammalian Pygo genes are significant modulators of canonical Wnt signaling in some, but not in all developing systems.

Figure 5

Reduced canonical Wnt signaling in Pygo2 and Pygo1/Pygo2 mutant embryos. E10.5 embryos, all with the BAT-gal transgene reporter of canonical Wnt signaling. (A) Pygo1+/+/Pygo2+/- embryos showed normal X-gal staining in the developing brain, pharyngeal pouches, otic vesicle, apical ectodermal ridges of the fore and hind limb buds, and in somites. (B) Pygo1+/-/Pygo2-/- embryos, with loss of both Pygo2 alleles, showed reduced but not absent BAT-gal reporter expression in many developing structures, including pharyngeal pouches, otic vesicle, and somites. (C) Pygo1-/-/Pygo2+/- embryo, with mutation of both Pygo1 alleles, but one wild-type Pygo2 allele, showed normal BAT-gal expression. (D) Double-homozygous mutant Pygo1-/-/Pygo2-/- embryos still showed some remaining BAT-gal expression, suggesting residual canonical Wnt signaling. Embryos in panels (A) and (B) were from the same litter and were processed in parallel, while embryos in (C) and (D) were from a separate litter, also processed in parallel, and were slightly more developmentally advanced. Original magnification (A, B) × 20; (C, D) × 16.

We also examined BAT-gal reporter expression in more detail in the developing urogenital system of Pygo mutants. The results suggested that the Pygo2 gene is required for canonical Wnt signaling in the nephric duct. Both Pygo2 null and Pygo1/Pygo2 double-null E10.5 embryos showed an absence of reporter expression in the nephric duct, while control littermates showed strong expression (Figure 6A–C). In Pygo1/Pygo2 double-null mutants, the nephric duct did form, however, and give rise to the ureteric bud outgrowth, which showed reduced but not absent BAT-gal reporter expression (Figure 6A–C).

Figure 6

Pygo2 is required for BAT-gal reporter expression in ureteric bud-derived structures of the developing kidney. X-Gal staining of BAT-gal transgenic (A-C) E10.5, and (E-L) E13.5 urogenital tracts, and (M-O) E18.5 kidneys. (A) Pygo1-/-/Pygo2+/+ (left) and Pygo1+/-/Pygo2-/- (right). Note the loss of reporter activity in the nephric duct (black arrowhead) and reduction of staining in the ureteric bud (white arrow) of the Pygo2 null embryo (Right). (B) Pygo1-/-/Pygo2+/- E10.5 embryo with BAT-gal reporter activity in the nephric duct (black arrowhead) and ureteric bud (white arrow). (C) Pygo1-/-/Pygo-/- embryo, with reporter expression lost in the nephric duct and reduced in the ureteric bud (white arrow). (D) Control background X-gal staining of a urogenital tract from an E13.5 embryo without the BAT-gal transgene (Non-Tg), showing absence of endogenous beta-galactosidase activity. (E) Pygo1+/-/Pygo2+/+, with BAT-gal reporter activity in the ureteric compartment of the developing kidney, including the ureteric tips, ureteric tree, and ureter. (F) Pygo1+/-/Pygo2-/-, with marked reduction of BAT-gal reporter activity in the ureteric compartment. (G) Pygo1+/-/Pygo2+/-, with reporter expression in the paramesonephric duct (white arrowhead) and ureteric tree. (H, I) Pygo1-/-/Pygo2+/-, with (H) ventral view showing reporter expression in the paramesonephric duct (white arrowhead), and (I) dorsal view showing ureteric tree expression in the kidney. (J)Pygo1+/-/Pygo2+/-, a control processed in parallel with (K) and (L), with expression in ureteric tree and paramesonephric duct. (K, L)Pygo1-/-/Pygo2-/-, reporter activity was lost in the paramesonephric duct (white arrowhead), and in the ureter and ureteric compartment of the developing kidneys (dashed circles), except for (K) a few faintly staining cells. (M, N) Pygo1+/-/Pygo2+/- (left) and Pygo1-/-/Pygo2-/- (right) E18.5 kidneys. The kidneys in (N) were bisected. BAT-gal reporter expression was seen in the ureteric tree components of the cortex and medulla of the double heterozygotes but was almost completely lost in the double-homozygous mutants. (O) Pygo1+/-/Pygo2+/+ (left), Pygo1-/-/Pygo2+/- (middle), and Pygo1-/-/Pygo2-/- (right) E18.5 kidneys. Reporter activity was present in the ureteric compartments of the Pygo1+/-/Pygo2+/+ (left) and Pygo1-/-/Pygo2+/- (middle) kidneys, but lost in the Pygo1-/-/Pygo2-/- (right) kidney. Original magnification: (A-C) × 32, (D-F) × 63, (G-I) × 40, (J-L) × 50, (M, N) × 10, and (O) × 12.5.

At E13.5, the Pygo2 gene appeared to play a major role, and the Pygo1 gene a minor role, in canonical Wnt signaling in the ureteric tree, as measured by BAT-gal expression. A negative control kidney, without the BAT-gal transgene, showed minimal background X-gal staining (Figure 6D), whereas a BAT-gal transgenic kidney with at least one wild-type Pygo2 gene showed strong X-gal staining in the ureteric tree (Figure 6E). In contrast, Pygo1+/-/Pygo2-/- E13.5 kidneys showed very weak reporter expression (Figure 6F), suggesting a significant loss of canonical Wnt signaling. Homozygous loss of the Pygo1 gene alone, however, had a small effect on reporter expression (Figure 6G–H). Homozygous mutation of both the Pygo1 and Pygo2 genes gave a more dramatic reduction of BAT-gal expression than loss of Pygo2 alone (Figure 6F, L).

The Pygo1 and Pygo2 genes were also required for BAT-gal reporter expression in the paramesonephric (Mullerian) ducts. In Pygo2-/- mice, there was a significant loss of reporter expression (data not shown), and double-homozygous mutants showed loss of X-gal staining in the paramesonephric ducts (Figure 6K), whereas Pygo1-/-/Pygo2+/- and Pygo1+/-/Pygo2+/- mice showed normal levels of BAT-gal expression (Figure 6G, H, J).

BAT-gal reporter analysis of the Pygo mutants at a later developmental stage, E18.5, also identified a significant decrease in canonical Wnt signaling in the cortical ureteric branches and renal pelvis of the developing kidney (Figure 6M–O). Cortical X-gal staining was seen in the ureteric branches of a Pygo1+/-/Pygo2+/- kidney (Figure 6M, left), but was completely absent in the cortex of a Pygo1+/-/Pygo2-/- kidney (Figure 6M, right). Bisection revealed a significant loss of X-gal staining cells in the collecting ducts and renal pelvis of the Pygo2 null kidney compared with control littermates (Figure 6N). Side by side comparison of Pygo1+/+/Pygo2+/- (Figure 6O, left), Pygo1-/-/Pygo2+/- (Figure 6O, middle), and Pygo1-/-/Pygo2-/- (Figure 6O, right) E18.5 kidneys suggested a significant role for the Pygo2 gene in canonical Wnt signaling in the ureteric tree and its derivatives.

In order to validate and quantify the BAT-gal reporter expression changes in the Pygo2 null and Pygo1/Pygo2 nulls, we performed ELISA measurements of transgene specific β-galactosidase levels in E18.5 kidneys (Figure 7). Loss of the Pygo2 gene (Pygo1+/-/Pygo2-/- and Pygo1-/-/Pygo2-/-) gave greater than 90% reduction in BAT-gal expression. Loss of Pygo1 alone (Pygo1-/-/Pygo2+/+) did not result in a significant change. Interestingly, however, the Pygo1-/-/Pygo2+/- showed only 50% of wild-type BAT-gal expression, suggesting a minor contribution by Pygo1 in canonical Wnt signaling. Although BAT-gal expression was decreased in the E18.5 Pygo1-/-/Pygo2+/- kidney, confocal analysis of kidneys with this genotype revealed no dilated ureteric tips or significant changes in ureteric tip number per area compared with Pygo1-/-/Pygo2+/+ kidneys (data not shown). Collectively, these BAT-gal reporter results suggest a significant role for the Pygo1 and Pygo2 genes in canonical Wnt signaling during development of the ureteric tree of the kidney.

Figure 7

Quantitative analysis of BAT-gal reporter expression in Pygo1/Pygo2 E18.5 kidney extracts. Transgene-specific beta-galactosidase was quantified by ELISA analysis. Pygo1+/-/Pygo2+/+, Pygo1+/-/Pygo2+/- and Pygo1+/-/Pygo2+/+ kidneys all showed similar levels of BAT-gal expression. In the Pygo2 hetereozygote, Pygo1 homozygous mutants, however, reporter expression was reduced by about 50%. In Pygo2 homozygous mutants BAT-gal expression was uniformly low, with or without wild-type Pygo1 alleles. Each experimental group included a sample size of at least four.

Interestingly, however, even in wild-type mice the BAT-gal reporter showed no expression in the developing metanephric mesenchyme, or metanephric mesenchyme-derived structures, such as renal vesicles, S-shaped bodies, tubules, and glomeruli. This has been reported previously, and was interpreted to indicate the absence of canonical Wnt signaling in the metanephric mesenchyme [21]. Results from other studies, however, argue for the presence of canonical Wnt signaling in the metanephric mesenchyme. For example, ureteric bud expression of Wnt1, thought to act through canonical Wnt signaling, can rescue Wnt9b mutants and induce nephrogenesis of the metanephric mesenchyme [2]. This suggests that the BAT-gal transgene might not accurately report canonical Wnt signaling in the metanephric mesenchyme. To address this question, we incubated E11.5 metanephric mesenchyme in lithium chloride (LiCl), which activates canonical Wnt signaling through inhibition of GSK3, and also functions as an inducer of nephrogenesis in kidney organ culture [22]. We observed that LiCl-treated metanephric mesenchyme did undergo nephrogenesis, as expected, but failed to show BAT-gal expression (data not shown), suggesting that this transgene is not an accurate reporter of canonical Wnt signaling in the metanephric mesenchyme of the developing kidney.

Microarray analysis of Pygo1/Pygo2 null kidneys

To further examine possible disturbance of gene expression in the Pygo1/Pygo2 mutant kidneys, we used microarrays to perform a global analysis of gene expression changes. Whereas the BAT-gal transgene reporter monitors the response of one promoter to Wnt signaling, microarrays can be used to follow expression changes of all genes, including all known Wnt targets. E18.5 wild-type and Pygo1-/-/Pygo2-/- kidneys were examined in biological triplicate. In total, 45 genes were identified as significantly changed, using a relatively low stringency screen of the data, with a p-value cutoff of <0.05, and fold change >2 (Table 3). Notably, both Pygo1 and Pygo2 were identified as downregulated (- 5.3-fold and - 3.9-fold, respectively) in the mutant kidneys. Other genes with significant decreases in mutant kidneys included Cxcl13 (- 2.3-fold), Slc5a2 (- 2.8-fold), and Slco1a4 (- 2.1-fold). Slc5a2 is expressed in the proximal tubules of the adult nephron and has been implicated in autosomal recessive renal glucosuria, characterized by loss of glucose uptake by the nephron [23]. The organic anion transporter, Slco1a4, is also strongly expressed in the tubules of the adult kidney [24]. These results suggest that the Pygo1/Pygo2 genes might play a role in nephron maturation and subsequent function.

Table 3

Genes differentially expressed (p < 0.05, two-fold change or greater) in the Pygo1/Pygo2 null E18.5 kidney normalized to wild-type samples.

Noteworthy genes upregulated in the Pygo1/Pygo2 null kidney included Klk5 (2.8-fold), Klk6 (2.7-fold), Ren2 (2.2-fold), and Timeless (2.4-fold). Klk5 and Klk6 are members of the kallikrein family of trypsin-like serine proteases. Kallikreins have diverse functions in cancer, tissue remodeling, and regulation of blood pressure [25]. Ren2, a homolog of the endopeptidase Renin 1, activates the renin-angiotensin system, increasing blood pressure [26]. Disruption of the renin-angiotensin system during development results in congenital abnormalities of the ureter and collecting-duct system [27]. Timeless, a transcription factor involved in the regulation of circadian rhythms, and expressed in the branching ureteric tips, has also been shown to regulate ureteric branching morphogenesis [28].

We were interested in the possible altered expression of previously known Wnt targets in the Pygo1-/-/Pygo2-/- mutant kidneys. A list of known Wnt targets was compiled from . Both Pygo1 and Pygo2 probe sets were included as references to illustrate significant down regulation. Remarkably, the known Wnt target genes showed very few differences in expression between wild-type and mutant kidneys (Figure 8). Only the expression of Ccnd1 encoding cyclin D1 [29], and Wisp1 [30] were significantly changed, with expression level changes of <1.5-fold in Pygo1/Pygo2 null kidneys. These results indicate an absence of dramatic changes in expression of previously known Wnt signaling target genes in the Pygo1/Pygo2 mutant kidneys.

Figure 8

Gene expression changes of common Wnt signaling targets in the E18.5 Pygo1/Pygo2 null kidney. Microarray analysis was performed in triplicate on wild-type and Pygo1/2 compound null E18.5 kidneys. Possible Wnt targets were selected from those compiled at [39]. An initial gene list of 82 Wnt targets was created and then reduced to a total of 40 genes using an expression level restriction requiring the raw expression intensity to be >100 in at least 3 samples. Pygo1 and Pygo2 probes were included to demonstrate significant changes in expression levels.

We used quantitative real-time PCR, with independent biological samples, to validate the microarray results. Nine genes were tested, and for seven we did observe a significant fold change in the same direction predicted by the microarray results (Table 4). It is not surprising that two of the nine genes could not be validated, considering the relatively low stringency used in screening the microarray data.

Table 4

Validation of gene expression changes in the E18.5 Pygo1/Pygo2 null kidneys normalized to E18.5 wild-type kidneys.

Discussion

In Drosophila, the Pygopus gene is a key mediator of canonical Wnt signaling. In one study, 12 distinct measures of Wnt signaling in Pygo mutants were performed, including analysis of leg, wing, and eye imaginal discs. In 2 cases there was a significant reduction of Wnt signaling and in 10 cases a complete block [10]. A second study in Drosophila examined the effects of Pygo mutation on cuticle patterning, midgut constriction, central nervous system, and cardiac development, and concluded that "Pygo is an essential component in the Wg signal transduction pathway". [8].

Given these results in Drosophila, the relatively mild phenotypes of mice with targeted Pygo1, Pygo2, or double Pygo1/Pygo2 mutations were striking. Pygo2 mutants developed to birth and showed limited abnormalities, while Pygo1 homozygous mutants were normal and fertile. Furthermore, there was no detected synergism in the phenotype of the double Pygo1/Pygo2 mutant, although in several tissues the BAT-gal Wnt reporter did show more a severe reduction in expression. One possible explanation of these unexpected results is a failure of the gene targeting to eliminate functioning of the Pygo1 and Pygo2 genes. The Pygopus deletion alleles described in this report, however, are almost certainly functional nulls. In Drosophila, it has been shown that the PHD domain is absolutely necessary for Pygopus function in Wg signaling. The PHD domain is 60 amino acids with seven cysteines and a histidine, predicted to chelate two zinc ions. PHD domains are found in diverse proteins, including transcription factors, and have been implicated in chromatin remodeling and protein-protein interactions [8]. In Drosophila the pygoF107 allele, with a single missense mutation converting amino acid 802 in the PHD domain from cysteine to tyrosine, loses Wnt signaling function in both embryogenesis and imaginal disc development [8]. The Pygo1/Pygo2 mutant alleles made in this report carried deletions of the entire PHD domains, as well as most other coding sequences. For the Pygo1 gene, the coding region for 372 of 417 total amino acids was deleted, and for the Pygo2 gene, we deleted coding for 354 of 405 amino acids. It is therefore very unlikely that the relatively mild phenotypes observed were the result of residual function of the targeted alleles.

We focused our analysis on the developing kidney, in which Wnt signaling has been shown to be of critical importance in several stages of nephrogenesis. Wnt9b is made by the ureteric bud and induces the metanephric mesenchyme to undergo nephrogenesis [2]. Downstream of Wnt9b is Wnt4 [2], which is made by the metanephric mesenchyme, and is also required for nephrogenesis [3]. In addition, Wnt11 is produced by the ureteric bud tips and induces GDNF expression in the metanephric mesenchyme [4].

In this report we describe a novel Wnt function in kidney development. The BAT-gal transgene reporter indicated the presence of canonical Wnt signaling in the ureteric bud and its derivatives in the developing kidney. Further, in the Pygo2 mutants this signal was lost, suggesting significant reduction of Wnt signaling. In addition, we observed a resulting decrease in ureteric tip density, reduced kidney size and altered morphology of the ureteric tree in mutants, indicating a role for canonical Wnt signaling in branching morphogenesis of the ureteric bud. While the simplest interpretation is direct Wnt signaling to the ureteric bud, it remains possible that the observed abnormalities are the result of indirect effects, with altered Wnt signaling to the metanephric mesenchyme then affecting mesenchyme to ureteric bud signaling.

The Pygo1/Pygo2 mutant phenotype of reduced branching morphogenesis of the ureteric bud is surprisingly similar to that previously reported for Wnt11 mutants [4]. The underlying mechanisms, however, are likely to be distinct. Wnt11 is generally thought to act through a noncanonical pathway, (although exceptions have been noted [31]), whereas the Pygopus genes promote canonical Wnt signaling. Further, the Wnt11 mutants showed an altered feedback loop between Wnt signaling from the ureteric bud to the mesenchyme and GDNF signaling from the mesenchyme to the bud, whereas in the Pygo1/Pygo2 mutants, we observed disrupted Wnt signaling in the ureteric bud.

The Pygo1/2 mutants also showed an expansion of the zone of thickened mesenchyme that caps the ureteric bud. Nevertheless, the mutant metanephric mesenchyme formed nephrons normally. This was particularly interesting, as Wnt9b signaling from the ureteric bud to the metanephric mesenchyme has been shown to induce nephrogenesis via canonical Wnt signaling [2].

The results in this report indicate a striking and surprising evolutionary divergence of Pygopus function between Drosophila and mammals. In Drosophila, the Pygopus gene is often required for canonical Wnt signaling, while in mammals the Pygo1/Pygo2 genes appear to play a smaller role in canonical Wnt signaling. The BAT-gal transgene reporter of canonical Wnt signaling showed reduced but generally not absent signal in Pygo1/Pygo2 mutant embryos, with tissue-specific variation in level of diminution. In addition, the kidneys were not unique in showing surprisingly normal development in Pygo1/Pygo2 mutants. Indeed, organogenesis generally proceeded without detectable abnormality, with few exceptions. These results suggest that the proteins encoded by mammalian Pygopus genes are often mere modulators of canonical Wnt signaling intensity, and not essential components.

The microarray results further support this conclusion. Whereas the BAT-gal transgene reporter monitors the expression level of only one Wnt responsive promoter, the microarray allows the analysis of the activity levels of promoters of all genes. Interestingly, the mutant kidneys showed gene-expression profiles surprisingly similar to wild type. Some genes did, however, show expression differences, and a high percentage of these differences could be validated by real-time PCR with independent biological samples. Assuming that the Pygopus genes in mammals are dedicated to canonical Wnt signaling, as has been previously shown in Drosophila, the genes with expression differences represent candidate Wnt targets (direct or indirect) in kidney development. We would predict these genes to show greater changes in expression level in a developing kidney with a more complete removal of canonical Wnt signaling.

Conclusion

In conclusion, the mammalian Pygo2 gene is required for normal branching morphogenesis of the ureteric bud, with mutants showing dilated tips and reduced numbers of tips. In addition, in Pygo2 mutants there was an expansion of the zone of metanephric mesenchyme that caps the ureteric buds. Nevertheless, nephron formation proceeded remarkably normally, even in Pygo1/Pygo2 double-homozygous mutants. This was surprising considering the importance of the Pygopus gene in canonical Wnt signaling in Drosophila, and the importance of canonical Wnt signaling in nephrogenesis. The results argue that the mammalian Pygopus genes are, in most developing systems, only quantitative transducers of Wnt signaling. Previous cell culture studies [32,33] and the reduced BAT-gal transgene reporter expression in the Pygo1/Pygo2 knockout mice described in this report, do confirm an evolutionarily conserved function in canonical Wnt signaling. In mammals, however, the phenotypic effects of Pygopus mutation are much milder than in Drosophila. The degree of importance of the Pyg1/2 genes in Wnt signaling was context-dependent, but in general, mammalian organogenesis remained intact in Pygopus mutants. Perhaps the simplest explanation is that in mammals, other genes show partial functional redundancy with the two Pygopus genes. The β-catenin transcription-factor complex includes a large and growing number of proteins , some of which may share the nuclear localization and/or transcription activation functions of Pygo1/Pygo2 in mammals. The identities and roles of these Pygopus redundant genes remain to be determined.

Methods

Targeting of Pygo1 and Pygo2

Pygo1 and Pygo2 genes were each targeted by flanking the critical coding regions of the third exon containing the PHD motif with LoxP sequences. Targeting constructs were made using the Flp/Cre vector previously described [34], which carries a neomycin-resistance gene flanked by FRT sequences, and three unique restriction sites for subcloning the two blocks of homology for driving recombination, and the critical region to be flanked by LoxP sequences. For each construct, the three genomic segments required for subcloning were made by PCR from RI ES cell DNA. The resulting targeting constructs were confirmed by sequencing.

For Pygo1, the genomic sequences used for PCR were: 5' forward, GTGAAGGAGAGATGGATAAGTATG; 5' back, TAGACCCTAACCACCTACAAG; exon forward, GGTTAGGGTCTATGTGCTGG; exon back, TCACCAAATCTCTGTTCTACAC; 3' forward, TGTGTAGAACAGAGATTTGGTG; 3' back, CAGTGAAGAAAGAGGGTCAG. For Pygo2, the genomic sequences used for PCR were: GCCTGGGTTGCTTGTCTTCTG and CCACCTTACTTGTGTGTGAGGATACATAC, CCAAGTCCCAGCATCTCTTAC and CCAGTCATACCAGCAACAAG, and exon sequences TGGGTGCTGGGAACAGAAC and CAACAACAACAGAAGACAAGC.

Linearized constructs were electroporated into RI ES cells, and resulting targeted ES cells used for C57/Bl6 blastocyst injections according to standard protocols. Resulting chimeras were mated with Swiss Black mice, and the targeted stocks maintained on a mixed 129/Swiss Black background.

Germline null alleles of both Pygo1 and Pygo2 were generated by mating heterozygous floxed mice with the CMV-Cre mice [15]. The sequences of the primers used for genotyping PCR were: Pygo2 null allele, forward (F) CCTGGATTCTTGTTGCTGGTATG; reverse (R) AAGGTATTTGGTGCTCCGAGGG; Pygo2 WT or floxed allele, F TGTCTTGATGACAGCGTTTAGCC, R AGATTCAGTAAGCTGAGCCTGGTG; Pygo1 null allele, F AGTTTGAAATAGCGACGAGTTTGAG, R 5'-CACTTCTGCCCCTCTCTTTGC; Pygo1 WT or floxed allele, F AAGCGTGCCTCATCTCCATCCCTAAG, R GCCCTCCCCGACGTTTATATTG.

The noon of the day that vaginal plugs were observed was designated E0.5.

Confocal microscopy

Kidneys were dissected, fixed in paraformaldehyde, treated with methanol, and washed with PBS containing Tween-20 (PBS-T) prior to treatment with lectins and antibodies. PBS-T was used for incubations of the tissues with fluoroscein-conjugated Dilichos biflorus aggutinin (DBA, Vector; Burlingame, USA), and PBS-T plus 2% goat serum was used for incubations with the antibodies. The primary antibodies were anti-WT1 (c-19; Santa Cruz Biotechnology, Santa Cruz, CA, USA), anti-uvomorulin (e-cadherin, Cdh1, Sigma-Aldrich St. Louis, MO, USA), and anti-Cited1 (Neomarkers-Lab Vision Corporation, Fremont, CA, USA). The secondary antibodies were Alexa 555-conjugated anti-rabbit and Alexa 633-conjugated anti-rat antibodies (Molecular Probes, Eugene, OR, USA).

The tissues were imaged with a laser scanning microscope (Carl Zeiss, Thornwood, New York, USA) equipped with an argon (488 nm) and two HeNe lasers (543 nm and 633 nm). Optical sections approximately 2 μm thick were obtained every 5 μm to a depth of at least 65 μm. The sections began at the surface of the kidney and were on a plane tangential to it. Two Z-stack series were obtained, one from each of the two kidneys of each embryo. Ureteric bud tips identified by section tracing were counted within a defined area of the confocal image.

In situ hybridization

Whole-mount in situ hybridization was performed as previously described [35]. Riboprobes to Wnt11 and Wnt7b were described previously [35]. The Wnt9b riboprobe was provided by T. Carroll [2].

Pygo1 and Pygo2 antibody production

To generate anti-human Pygo2 (anti-hPygo2) and anti-mouse Pygo1 (anti-mPygo1) polyclonal antibodies, we subcloned cDNA by PCR corresponding to amino-acid residues 80–327 of human Pygo 2 or amino-acid residues 76–263 of mouse Pygo1 into pGEX4T1 (Amersham Health, Piscataway, NJ, USA). The PCR fragments of hPygo 2 and mPygo 1 lack both NHD and PHD conserved regions of hPygo2 and mPygo1. GST-hPygo2 and GST-mPygo1 proteins were expressed in bacteria, purified, and injected into rabbits for antibody production by (Proteintech Group Inc., Chicago, IL, USA). The rabbit antisera of anti-mPygo1 and anti-hPygo2 were initially allowed to bind to the GST affinity matrix to remove any antibodies against GST. The anti-hPygo2 and anti-mPygo1 antisera were then separated from the GST affinity matrix and allowed to bind to the GST-hPygo2 or GST-mPygo1 affinity columns, respectively. The bound antibodes were eluted with elution buffer. To further ensure antibody specficity, the purified antibodies were extensively incubated with Pygo1/2 double-homozygous mutant embryo extract before use.

BAT-gal transgene reporter assay of canonical Wnt signaling

X-gal staining of both embryos and developing kidneys was performed as previously described [21]. Care was taken to reduce endogenous β-galactosidase activity within the developing kidney by increasing pH of the X-gal staining solution to 8.0. Changes in transgene β-Gal expression were quantitated using a β-Gal ELISA (Enzyme-linked immunoassay) kit (Roche, Indianapolis, IN), normalizing according to total protein. Each genotype was represented by a sample size of 4 except the non-transgenic (n = 5), Pygo1+/- ; Pygo2+/+ (n = 6), Pygo1-/- ; Pygo2+/- (n = 8), and Pygo1-/- ; Pygo2-/- (n = 6) groups.

Microarray analysis

Total RNA was isolated from E18.5 kidneys dissected from normal and Pygo1/Pygo2 compound null embryos using a commercial kit (Stratagene Absolutely RNA Microprep Kit; La Jolla, CA, USA). An aliquot (300 ng) of total RNA was processed and labeled using a commercial kit (TargetAmp 1-Round Aminoallyl-aRNA Ki; Epicentre, Madison, WI, USA). Labeled RNA was hybridized to microarrays (Sentrix Mouse-6 expression Beadchip; Illumina, San Diego, CA) providing coverage of over 47000 genes and expressed sequence tags as previously described [36]. Microarray analysis of Pygo1/Pygo2 null and normal wild-type kidneys was performed in biological triplicate. Raw signal intensities of each probe were obtained from data analysis software (Beadstudio ;Illumina) and imported into GeneSpring GX 7.3 (Agilent Technologies, Palo Alto, CA, USA). Genes were selected on the basis of greater than two-fold average fold change and statistical significance (p-value < 0.05). Previously described Wnt target genes were obtained from .

Quantitative PCR validation of microarray results

Total RNA from E18.5 Pygo1/Pygo2 null and control kidneys (both represented in duplicate and distinct from the kidneys used for microrray analysis) was purified using a commercial kit (Absolutely RNA Microprep Kit; Stratagene, La Jolla, CA USA) including DNAse1 treatment. cDNA was generated using random hexamers according to conventional protocols (Invitrogen, Carlsbad, CA, USA). The following primers were generated to include the sequence obtained from the Illumina probe: Actb (TTGCTGACAGGATGCAGAAG, ACATCTGCTGGAAGGTGGAC); Aldh1a7 (CCAGAAAGTGGTGTTTTGCT, GAGTTACAGAGAGCTTGCACCTT); Col8a1 (GCAGACAGGCATCTTCACCT, TGTGTACATCATGGGCTCGT); Csrp1 (CAGCATAGCCCAGGGTAGAG, TGGGCAAGGTAGTGAAGGTT); Klk5 (GCAGCATTACACCCGTCATA, TTGCCTCCATCCATATCTCC); Picalm (GGGAGGGAACAGAAATCCTT, GCACCGATCAACAGTGCAG); Pygo2 (TTCTGGGAACTTGTGCACTG, AACTTCCTCCAGCCCATTTT); Ren2 (TTGTGAGCTGTAGCCAGGTG, TGTGCACAGCTTGTCTCTCC) and Tia1 (TGATTGAAGGGCTACTAGAGTGGT, AGCCCCAGAGGACAATTTTT) using Primer3 software [37]. Relative quantitative PCR was performed according to the conventional SYBR Green protocol (Stratagene) using a quantitative PCR system (Mx3000p; Stratagene). Dissociation curve and agarose-gel analysis of each primer set were used to insure specificity of the amplicon. All data were normalized to an internal housekeeping control (Actb) and analyzed using the 2(-Delta Delta C(T)) method [38].

Authors' contributions

SP designed and did most of the work for targeting the Pygo genes, provided oversight for the project and helped write the paper. KS did most of the experiments, including confocal analysis with LP, in situ hybridizations with HH, and helped write the paper. XL made and tested the Pygo antibodies. NS and RL provided useful suggestions and helped with the immunostaining and Bat-Gal analyses.

Acknowledgements

L. McClain provided essential technical assistance and we thank P. Groen and D. Witte for initial histology and embryological analyses. D. Ash helped make targeting constructs. We thank T. Carroll for providing a Wnt9b riboprobe construct. We thank H. Liang for generation of microarray data. This work was supported by grant DK61916 from the National Institutes of Health (S.S.P.).


A Mammal-Specific Doublesex Homolog Associates with Male Sex Chromatin and Is Required for Male Meiosis

Abstract

Gametogenesis is a sexually dimorphic process requiring profound differences in germ cell differentiation between the sexes. In mammals, the presence of heteromorphic sex chromosomes in males creates additional sex-specific challenges, including incomplete X and Y pairing during meiotic prophase. This triggers formation of a heterochromatin domain, the XY body. The XY body disassembles after prophase, but specialized sex chromatin persists, with further modification, through meiosis. Here, we investigate the function of DMRT7, a mammal-specific protein related to the invertebrate sexual regulators Doublesex and MAB-3. We find that DMRT7 preferentially localizes to the XY body in the pachytene stage of meiotic prophase and is required for male meiosis. In Dmrt7 mutants, meiotic pairing and recombination appear normal, and a transcriptionally silenced XY body with appropriate chromatin marks is formed, but most germ cells undergo apoptosis during pachynema. A minority of mutant cells can progress to diplonema, but many of these escaping cells have abnormal sex chromatin lacking histone H3K9 di- and trimethylation and heterochromatin protein 1β accumulation, modifications that normally occur between pachynema and diplonema. Based on the localization of DMRT7 to the XY body and the sex chromatin defects observed in Dmrt7 mutants, we conclude that DMRT7 plays a role in the sex chromatin transformation that occurs between pachynema and diplonema. We suggest that DMRT7 may help control the transition from meiotic sex chromosome inactivation to postmeiotic sex chromatin in males. In addition, because it is found in all branches of mammals, but not in other vertebrates, Dmrt7 may shed light on evolution of meiosis and of sex chromatin.

Author Summary



Genes related to the sexual regulator Doublesex of Drosophila have been found to control sexual development in a wide variety of animals, ranging from roundworms to mammals. In this paper, we investigate the function of the Dmrt7 gene, one of seven related genes in the mouse. Female mammals are XX and males are XY, a chromosomal difference that presents specific challenges during the meiotic phase of male germ cell development. Some of these are thought to be overcome by incorporating the X and Y chromosomes into a specialized structure called the XY body. We find that DMRT7 protein is present in germ cells, localizes to the male XY body during meiosis, and is essential for male but not female fertility. The XY body normally is altered by recruitment of additional proteins and by specific modifications to histone proteins between the pachytene and diplotene stages of meiosis, but modification of the “sex chromatin” in Dmrt7 mutant cells is abnormal during this period. Because Dmrt7 is found in all branches of mammals, but not in other vertebrates, these results may indicate some commonality in regulation of sex chromatin among the mammals.

Introduction

Sexual differentiation generates anatomical, physiological, and behavioral dimorphisms that are essential for sexual reproduction. Many of these dimorphisms affect somatic cells, but the sexual dimorphisms that most directly mediate sexual reproduction are those of the gametes themselves. Gametes differ between the sexes in size and morphology, sometimes dramatically so, reflecting their very different roles in zygote formation. Indeed, the morphology of the gametes is what defines sex: females are the sex that produces the larger gametes and males produce the smaller ones.

Mammalian meiosis is regulated sex-specifically starting in embryogenesis and continuing through much of adult life (reviewed in [1]). For example, the timing and synchrony of meiosis are very different in the two sexes. In females, germ cells synchronously initiate meiosis in the embryo and arrest during meiotic prophase I. After puberty, oocytes are selectively recruited for ovulation, when they proceed to metaphase II and then complete meiosis after fertilization occurs [2]. In contrast, male meiosis occurs entirely postnatally, without the arrest periods found in females. In females, each meiosis can produce a single haploid oocyte (and two extruded polar bodies), whereas each male meiosis can produce four haploid spermatocytes.

Other meiotic processes, such as recombination and chromosome pairing (synapsis), occur in both sexes but operate somewhat differently. For example, there is a higher failure rate for meiosis in females, with human oocyte aneuploidy rates up to 25% versus about 2% in human sperm [3], and this may indicate that the checkpoints controlling and monitoring the events of meiotic progression in males are more stringent. Consistent with this idea, genetic analysis of a number of meiotic regulatory genes in the mouse has demonstrated a much stronger requirement in males than in females [1,4].

The existence of heteromorphic sex chromosomes, such as the XX/XY system of mammals, creates sex-specific challenges. One is the need for mechanisms to balance expression of sex-linked genes between the sexes, which in mammals is accomplished by X chromosome inactivation in females [5,6]. In male germ cells there is another sex-specific consideration during meiosis. In prophase I, when the homologous chromosomes synapse and homologous recombination occurs, X and Y chromosome pairing is limited to a region termed the pseudoautosomal region, leaving large portions of each chromosome unpaired. In eutherian and marsupial mammals, these unpaired chromosome regions are associated with a specialized chromatin domain termed the XY body or sex body. The function of the XY body is uncertain [7–11], but there is evidence that it is essential for male meiotic progression [12].

Several proteins are reported to localize to the XY body, including BRCA1, ATR, the histone variant H3.3, and modified histones such as ubiquitinated H2A (Ub-H2A) and phosphorylated H2AX (γH2AX) [12–15]. In the XY body, the sex chromosomes are transcriptionally silenced in a process termed meiotic sex chromosome inactivation (MSCI). The XY body disappears after pachynema; however, many sex-linked genes remain transcriptionally silent into spermiogenesis [16]. This maintenance of silencing is associated with a distinct set of chromatin marks that define a sex chromatin domain termed postmeiotic sex chromatin (PMSC) [16,17].

Regulators of sexual differentiation have been identified in a number of organisms, but in contrast to many other developmental processes, such as axial patterning or development of many body parts, the molecular mechanisms that regulate sexual differentiation are highly variable between phyla. A notable exception involves genes related to doublesex (dsx) of Drosophila, which share a Doublesex/MAB-3 DNA-binding motif called the DM domain [18,19]. DM domain–encoding genes have been shown to regulate various aspects of sexual differentiation in insects, nematodes, and mammals [20]. The mab-3 gene of Caenorhabditis elegans has been shown to function analogously to DSX in several respects and can be functionally replaced by the male isoform of DSX, suggesting that the similarity in the sequence of these genes may stem from conservation of an ancestral DM domain sexual regulator [18,21,22].

Vertebrates also have DM domain genes, and analysis to date, although limited, has shown that these genes also control sexual differentiation. Mammals have seven DM domain genes (Dmrt genes), several of which exhibit sexually dimorphic mRNA expression [23,24]. The best studied of these genes, Dmrt1, is expressed in the differentiating male genital ridges and adult testis of mammals, birds, fish, and reptiles, and a recently duplicated Dmrt1 gene functions as the Y-linked testis-determining gene in the Medaka fish [25–29]. Human DMRT1 maps to an autosomal locus, which, when hemizygous, is associated with defective testicular development and consequent XY feminization [30]. Similarly, mice homozygous for a null mutation in Dmrt1 have severe defects in testis differentiation involving both germ cells and Sertoli cells [31]. Female mice mutant in Dmrt4 have polyovular follicles, indicating that this gene also plays a role in gonadal development [32]. It appears from these studies that the involvement of DM domain genes in sexual differentiation is ancient and conserved. However, vertebrate Dmrt gene function is not limited to sexual differentiation: Dmrt2 is required in both sexes for segmentation in mice and fish [33–35].

Here, we have investigated the expression and function of the Dmrt7 gene in the mouse. Dmrt7 is expressed only in the gonad, and, unlike the other Dmrt genes, appears to be present exclusively in mammals and not in nonmammalian vertebrates [23,36]. We find that DMRT7 protein is expressed only in germ cells and is selectively localized to the XY body of male pachytene germ cells. To test its function, we generated a conditional null mutation of Dmrt7 in the mouse. We find that Dmrt7 is required in males for progression beyond the pachytene stage of meiotic prophase but is not required in females. In rare mutant cells that survive to diplonema, we observed sex chromatin abnormalities. Based on these observations, we suggest that Dmrt7 plays a critical role in a male-specific chromatin transition between pachynema and diplonema during meiotic prophase.

Results

Expression of DMRT7 Proteins in Testis

Our previous mRNA expression analysis suggested a possible meiotic function for Dmrt7, based on the expression of Dmrt7 mRNA in the fetal gonads of the two sexes [23]. In the fetal ovary, Dmrt7 mRNA was detected primarily from E13.5 to E15.5, the time during which meiosis progresses from pre-meiotic replication to the pachytene stage [4], whereas Dmrt7 expression in the non-meiotic fetal testis was very low. Because this earlier work did not examine adult Dmrt7 expression, we first performed reverse transcriptase (RT)-PCR on mRNA from ten adult organs and detected strong Dmrt7 mRNA expression in the testis and a trace of expression in heart, but not in any other tissue tested (Figure 1A). We examined the timing of Dmrt7 mRNA expression during postnatal testis development and detected strong expression beginning at 2 wk, which roughly coincides with the onset of the pachytene stage during the first synchronous wave of spermatogenesis (Figure 1B) [37].

Figure 1

Expression of Dmrt7 mRNA and Protein

(A) RT-PCR analysis of Dmrt7 mRNA from ten organs of adult mouse. cDNA from each organ was amplified with primers specific for Dmrt7 (top row) and β-actin (bottom row).

(B) Dmrt7 mRNA expression during the first round of spermatogenesis. cDNAs obtained from testis at the indicated days after birth were amplified as in (A).

(C) DMRT7 protein expression. Immunofluorescence of testis sections from 6-wk-old male stained with antibody to DMRT7 (green) and DAPI (blue).

(D) DMRT7 subcellular localization to XY body. Testis sections from 6-wk-old male stained with antibodies to DMRT7 (red) and SUMO-1 (green). SUMO-1 is localized to the XY body. Right-most panel shows merge of other two panels. Inserts show higher magnification of pachytene spermatocytes with XY bodies.

To investigate DMRT7 protein expression, we generated an antibody against the C-terminal portion of the protein. The antibody was raised against a unique region lacking the DM domain in order to avoid cross-reaction with other DM domain proteins. Immunofluorescent staining with purified DMRT7 antisera showed that DMRT7 protein is expressed predominantly in mid- to late-pachytene spermatocytes (Figure 1C), as well as in sperm, and is not detectable in other germ cell types including spermatogonia and round spermatids. We did not detect DMRT7 protein in somatic cells such as Sertoli cells, peritubular myoid cells, or Leydig cells. To more precisely determine the pachytene stages of DMRT7 expression, we double-stained with an antibody to GATA1, which is expressed in Sertoli cells from stages VII to IX [38]. This confirmed that DMRT7 is expressed in mid- to late-pachytene spermatocytes, starting slightly earlier than stage VII and extending through stage IX (unpublished data).

Within pachytene spermatocytes, DMRT7 is concentrated in the XY body, or sex body, a densely staining chromatin domain that harbors the sex chromosomes. These undergo transcriptional inactivation and heterochromatinization as a result of their incomplete pairing during prophase of mammalian male meiosis [17]. To verify DMRT7 protein expression in the XY body, we double-stained mouse testis sections for DMRT7 and small ubiquitin-related modifier 1 (SUMO-1), which is concentrated in the XY body during pachynema [39,40]. DMRT7 and SUMO-1 were colocalized, confirming that DMRT7 protein is preferentially localized to the XY body (Figure 1D). We also confirmed XY body localization of DMRT7 by double staining for other markers including Ub-H2A and γH2AX (unpublished data). DMRT7 is not preferentially localized to the XY body at all stages but instead is dynamic. Based on epithelial staging, it appears that DMRT7 localizes to the XY body from mid- to late-pachynema, becomes diffusely distributed in late-pachynema, and disappears in diplonema (unpublished data). This localization was confirmed by staining of meiotic spreads (Figure S1). DMRT7 also is specifically localized in sperm, with antibody staining mainly in the perinuclear ring of the sperm head manchette. This staining coincided with the epithelial stages in which DMRT7 localizes to the XY body in spermatocytes (Figure 1C and 1D).

Targeted Deletion of Dmrt7

To establish the functional requirement for Dmrt7, we generated Dmrt7−/− mice by targeted disruption in embryonic stem (ES) cells using a strategy diagrammed in Figure S2A. The Dmrt7 gene has nine exons with the DM domain encoded by the second and third exons. Because the DM domain is essential for function of other genes, including mab-3, mab-23, and dsx [18,19,41], we generated a conditionally targeted “floxed” allele in which the DM domain–containing exons of Dmrt7 are flanked by recognition sites for the Cre recombinase (loxP sites). The targeting vector also contained a neomycin resistance cassette (neo) flanked by Flpe recognition sites. The removal of these sequences by Cre-mediated recombination eliminates the DM domain and the translational start site, thus generating a putative null allele. We identified three homologously targeted ES cell clones by Southern blotting (Figure S2B) and injected cells from two clones into C57BL/6 blastocysts. Chimeric animals from both cell lines transmitted the targeted allele through the germ line. Targeted animals were bred to β-actin Cre mice to delete the DM domain–encoding exons, generating the Dmrt7− allele, or to Flpe transgenic mice to delete the neo cassette, generating the Dmrt7flox allele. Dmrt7+/− mice were interbred to generate Dmrt7−/− mice. To confirm the lack of functional DMRT7 protein in Dmrt7−/−testes, we stained meiotic spreads from Dmrt7 mutants (Figure S1) and sections from mutant testes (Figure S2C) and carried out western blot analysis (unpublished data). In each case, we detected no DMRT7 protein in the mutant testes.

Dmrt7 Is Required for Male but Not Female Gametogenesis

Breeding of Dmrt7 heterozygotes produced homozygous mutant progeny of both sexes at the expected frequency (63/264; 23%). Male and female homozygous mutants were viable, grew to adulthood normally, and exhibited normal sexual behavior. Female homozygotes were fertile, produced litters of normal size, and had no obvious ovarian abnormalities as judged by histological analysis (unpublished data). In contrast, Dmrt7 homozygous mutant males were completely infertile and had testes about one-third the weight of those of heterozygous or wild-type adult littermates (Figure 2). To determine when defective testis development begins in Dmrt7 mutants, we compared the testes of wild-type and mutant littermates during the first wave of spermatogenesis. Prior to postnatal day 14 (P14), mutant testes appeared histologically normal and the testis weights were similar to those of heterozygous and wild-type littermates, indicating that spermatogonia and early meiotic germ cells form normally (Figure 2B; unpublished data). Thereafter, the testes of the Dmrt7 mutant mice ceased to grow and the weight difference was significant. Microscopic examination of P21 and P42 Dmrt7 mutant testes revealed that germ cells arrest in pachynema, and later stages of germ cells are largely missing (Figure 2C and 2D). Dmrt7 mutant mice are deficient in postmeiotic spermatids and lack epididymal spermatozoa, although a few cells develop to the round spermatid stage. These meiotic defects are in agreement with a recent preliminary analysis of another Dmrt7 mutation [42]. While some Dmrt7 mutant tubules are highly vacuolated and contain primarily Sertoli cells and spermatogonia, others have abundant primary spermatocytes. In addition, some tubules contain multinucleated cells and cells with darkly stained nuclei that are typical of apoptotic cells (Figure 2D).

Figure 2

Reduced Testis Size and Germ Cell Apoptosis in Mice with Targeted Deletion in Dmrt7

(A) Testes from a 6-wk-old wild-type (+/+) mouse and a homozygous (−/−) Dmrt7 mutant littermate.

(B–D) Sections of testes from 14-d-old (B), 21-d-old (C), and 42-d-old mice (D) stained with hematoxylin and eosin. Wild-type is in left column and mutant in right. No significant difference is observed at 14 d (B), but by 21 d some tubules are lacking abundant spermatocytes (C, asterisk) or cells with typical apoptotic morphology are present (open arrowhead, C and D). Mutant tubules contain multinucleate cells (closed arrowhead, D).

(E and F) TUNEL labeling of Dmrt7-deficient mouse testes. Testes from wild-type and homozygous mutant littermates were analyzed by TUNEL labeling to detect apoptotic cells. Testis sections from 21-d-old (E) and 6-wk-old mice (F). Apoptotic cells (brown) are much more abundant in seminiferous tubules of homozygous Dmrt7 mutant mice relative to wild-type. Bars in (B–F) represent 100 μm.

Since Dmrt7 mutant testes lack most post-pachytene cells, we used TUNEL analysis to test whether the missing cells are eliminated by apoptosis. At 3 wk, Dmrt7 mutant testes contain significantly more apoptotic cells than those of wild-type controls. The percentage of tubule sections with five or more apoptotic nuclei was about three times higher in Dmrt7 mutants compared with wild-type (20% versus 7%; Figure 2E). A similar elevation of apoptosis was apparent in mutant testes at 7 wk (Figure 2F). In mutants, many apoptotic cells were in the middle of the tubules, whereas the apoptotic cells in wild-type occur mainly near the seminiferous tubule periphery. The numbers of Sertoli cells were not significantly different between wild-type and mutant testes, and we observed no difference in somatic cell apoptosis in mutants (unpublished data). From these results, we conclude that loss of Dmrt7 causes a block in meiotic progression, mainly in pachynema, leading to the elimination, by apoptosis, of the arrested cells.

Pachytene Arrest of Dmrt7 Mutant Germ Cells

To better define the spermatogenic stage at which Dmrt7−/− male germ cells arrest and die, we used antibodies against several stage-specific germ cell markers. TRA98 is expressed in PGCs and spermatogonia [43]. In the wild-type adult testis, strongly staining TRA98-positive cells form a layer one cell deep; however, in the mutant TRA98, strongly positive cells were abnormally organized, and some tubules had a layer several cells deep (Figure 3A). The BC7 antibody recognizes spermatocytes in the leptotene to early-pachytene stages [44]. Dmrt7 mutant testes had BC7-positive cells in approximately normal numbers, but again abnormally organized, with many positive cells in the center rather than the periphery of the tubules (Figure 3B). The TRA369 antibody recognizes a calmegin protein expressed in pachytene spermatocytes through elongated spermatids [45]. In contrast to the earlier stages, far fewer TRA369-positive cells were present in mutant testes relative to wild-type (Figure 3C). We also quantitated the number of cells at each meiotic stage using spermatocyte spreads, assaying chromosome-pairing status by staining for SYCP3, a component of the synaptonemal complex (Figure 4). We found that Dmrt7 mutants accumulate pachytene cells but have greatly reduced numbers of cells in late-pachynema and beyond. Together, these results confirm that the meiotic arrest in Dmrt7 mutants occurs primarily during pachynema and results in efficient elimination of arrested cells.

Figure 3

Prophase I Arrest of Dmrt7 Mutant Germ Cells

Testis sections from 6-wk-old wild-type (+/+) and Dmrt7 mutant (−/−) littermates stained with stage-specific antibodies specific to spermatogenic cells.

(A) TRA98 antibody strongly stains spermatogonia, which are present in wild-type and mutant.

(B) BC7 antibody stains spermatocytes, which are present in wild-type and mutant.

(C) TRA369 antibody stains pachytene and later germ cells, which are severely deficient in the mutant testis.

Figure 4

Profile of Meiotic Arrest in Dmrt7 Mutant Testis

Graph of distribution of meiotic stages of germ cells from heterozygous (n = 1,970) and homozygous mutant (n = 2,084) P26 testes, spread and stained for the synaptonemal complex protein SYCP3 to permit precise staging, which was performed as described [64,65].

Normal Meiotic Prophase in Dmrt7 Mutant Germ Cells

Defects in chromosome pairing, synapsis, or recombination can trigger pachytene arrest and apoptosis [46]. We therefore examined these events in Dmrt7 mutant testes. To assess homolog synapsis, we used antibodies to SYCP1, a synaptonemal complex transverse element component, and SYCP3, a component of the axial element, which remains on the desynapsed axes during diplonema [47,48]. Formation of synaptonemal complexes in the mutant was indistinguishable from that in wild-type, as indicated by the proper accumulation of SYCP1 (unpublished data) and SYCP3 (Figure 5A). Likewise, the Dmrt7 mutant zygotene spermatocytes showed normal accumulation of the early recombination repair marker RAD51, suggesting that early meiotic recombination is not significantly affected (Figure 5B). Dmrt7 mutant spermatocytes exhibited the expected decline in the presence of RAD51 foci associated with the autosomal synaptonemal complexes (Figure 5B; unpublished data) [49]. The few surviving cells that progressed beyond pachynema also underwent apparently normal desynapsis during diplonema (Figure 5A). From these results, we conclude that chromosomal pairing, synapsis, recombination, and desynapsis during prophase I in Dmrt7 mutant males are grossly normal.

Figure 5

Normal Synapsis and Recombination in Dmrt7 Mutant Germ Cells

(A) Testicular cells from Dmrt7 heterozygous (+/−) or homozygous (−/−) mutant mice were spread and stained with antibody to SYCP3 (red). The developmental stages of meiotic prophase based on SYCP3 organization are indicated in each panel. By pachynema, all autosomes are fully synapsed, and the XY bivalent is synapsed only at the pseudoautosomal region in both wild-type and mutant cells.

(B) Testicular cells in zygonema and pachynema spread and stained with antibodies to RAD51 (green) and SYCP3 (red). Size and distribution of RAD51 foci are similar between wild-type and mutant spermatocytes.

Abnormal Cellular Organization and the Role of Sertoli Cells

Sertoli cells interact with germ cells during spermatogenesis and the interaction is critical for germ cell maturation [50]. Although we did not detect DMRT7 expression in Sertoli cells by antibody staining, we nevertheless considered the possibility that Sertoli cell defects might contribute to the male-specific germ line failure of Dmrt7 mutants. To characterize Sertoli cell differentiation, we examined expression of the Sertoli cell markers GATA4 (a marker of immature postnatal Sertoli cells) and GATA1 (a mature Sertoli cell marker). The levels of these proteins appeared normal relative to wild-type at P14 and P42 (Figure 6A–6C), as did the androgen receptor (Figure S3; unpublished data). However, the organization of Sertoli cells in Dmrt7 mutant testes was abnormal: in some tubules GATA1-positive Sertoli cell nuclei were displaced from their usual close apposition with the basement membrane (Figure 6C). In such tubules, nuclei of pre-meiotic germ cells and spermatocytes were packed close to the basal membrane and few germ cells were found in the adlumenal region.

Figure 6

Abnormal Sertoli Cell Organization in Dmrt7 Mutant Testes

(A) Testes from P14 wild-type and Dmrt7 mutant mice were sectioned and stained with antibody to GATA4.

(B) P14 testis sections stained with antibody to GATA1. At P14, GATA4 and GATA1 levels are similar in wild-type and mutant Sertoli cell.

(C) Wild-type and mutant testis sections double-stained with antibody to GATA1 (red) and DAPI (blue). Most Sertoli cell nuclei were adjacent to the basal membrane in wild-type, but mutant Sertoli cells were displaced in some tubules (arrowhead). White dotted line indicates position of basal membranes.

(D) Testis sections from 10-wk-old wild-type and Sertoli cell-specific Dmrt7 mutant (SC-Dmrt7KO) mice stained with hematoxylin and eosin. Spermatogenesis and spermiogenesis are normal in SC-Dmrt7KO testis.

(E) Testis sections from 10-wk-old wild-type and SC-Dmrt7KO mice stained with antibodies to GATA1 (red) and smooth muscle actin (to outline seminiferous tubules; green). Sertoli cell nuclei are positioned normally near the basal membrane in SC-Dmrt7KO mice.

The aberrant Sertoli cell organization in Dmrt7 mutant testes raised the possibility that the germ cell phenotype might indirectly result from defects in Sertoli cell function. To test this possibility, we deleted Dmrt7 just in the Sertoli cell lineage by crossing mice carrying the floxed Dmrt7 allele with Dhh-Cre transgenic mice [51]. The Desert hedgehog (Dhh) promoter is active starting at about E12.5 in pre-Sertoli cells but not in germ cells, allowing deletion of Dmrt7 in Sertoli cells well before any likely requirement for its function [52]. Testicular size in Sertoli-targeted (SC-Dmrt7KO) animals was slightly reduced from that of wild-type, but histological analysis revealed no obvious difference between wild-type and SC-Dmrt7KO testes (Figure 6D). Spermatogenesis appeared normal, mature sperm were present, and SC-Dmrt7KO mice were fertile. In addition, GATA1 staining showed that Sertoli cell nuclei were located adjacent to the basement membrane as in wild-type (Figure 6E). These results suggest the germ cell defects of Dmrt7 mutants are not caused by lack of Dmrt7 in Sertoli cells. Rather, the abnormal organization of Sertoli cells appears to result from lack of Dmrt7 in the germ line.

XY Bodies Form Normally in Dmrt7 Mutant Cells

The data presented so far indicate that Dmrt7 mutant germ cells undergo apparently normal early meiosis and then arrest during pachynema due to a strict requirement for Dmrt7 in the germ line. To better understand the basis of the meiotic arrest, we more closely examined meiotic germ cells in the mutant. We focused on the XY body, which is thought to be essential for meiotic progression and is the site of preferential DMRT7 localization. Condensation of the X and Y chromosomes begins in late-zygotene cells, and, by mid-pachynema (when homologous chromosome pairs are fully aligned) the sex chromatin forms a microscopically visible spherical structure near the nuclear periphery [53].

We first asked whether DMRT7 is required for XY body formation by evaluating several characteristic XY body chromatin features. First, we tested γH2AX expression by immunofluorescent staining. H2AX is a variant of H2A that is crucial for XY body formation and MSCI [12]. γH2AX localized normally to the XY body of DMRT7 mutant cells in meiotic spreads (Figure 7A), and many γH2AX-positive puncta were present in germ cells of Dmrt7 mutant testes (Figure 7B). Next, we examined SUMO-1 localization in the mutant testis. SUMO-1 expression normally increases in the XY body of early- to mid-pachytene spermatocytes at the time of sex chromosome condensation. Prior to the completion of the first meiotic division, SUMO-1 disappears from the XY body as the X and Y chromosomes desynapse [40]. Punctate SUMO-1 localization was present in Dmrt7 mutant germ cells, again consistent with formation of a correctly marked XY body (Figure 7C). However, some tubules in mutants had multiple layers of cells with SUMO-1-condensed spots (Figure 7C), rather than the normal single layer of cells. This accumulation of XY body–containing cells also was apparent with γH2AX staining and is consistent with a developmental arrest of mutant cells in mid- to late-pachytene. We also examined Ub-H2A localization in Dmrt7 mutant testes. In early-pachytene, Ub-H2A is concentrated in the XY body; by mid-pachytene Ub-H2A is observed throughout the entire nucleus, but it again becomes limited to the XY body in late-pachytene spermatocytes [13]. Analysis of nuclear spreads revealed that Ub-H2A localizes normally to the XY body in Dmrt7 mutants (Figure S4). Collectively, these results indicate that Dmrt7 mutant germ cells can establish an XY body with at least some of the normal chromatin marks.

Figure 7

XY Body Forms Normally during Pachynema in Dmrt7 Mutant Mice

(A) Testicular cells spread and stained with antibodies to γH2AX (red) and SYCP3 (green). In pachytene cells, γH2AX is concentrated in the XY body both in wild-type and Dmrt7 mutant.

(B) Testes from 6-wk-old mice sectioned and stained with antibody to γH2AX (red).

(C) Testes from 6-wk-old mice sectioned and stained with antibody to SUMO-1 (green). Abundant pachytene cells with XY bodies are present in wild-type and mutant testes.

Abnormal Sex Chromatin in Dmrt7 Mutant Germ Cells

Although the XY body can form during pachynema in Dmrt7 mutants, we considered the possibility that transcriptional silencing might not be properly established. This would be consistent with the Dmrt7 phenotype: pachytene cells that escape from MSCI normally are eliminated prior to late-pachytene [17]. Recently, MSCI has been shown to continue into meiosis II and spermiogenesis, apparently mediated by a distinct chromatin compartment termed postmeiotic sex chromatin (PMSC) that is established starting in diplonema [16]. We therefore asked whether the pachytene germ cell death in Dmrt7 mutants is associated with a failure either to initiate or to maintain sex chromosome inactivation.

First, we examined the mid-pachytene XY body. To examine XY transcriptional status, we carried out Cot-1 RNA fluorescence in situ hybridization (FISH) to detect nascent RNA polymerase II transcription, combined with DAPI staining to locate the XY body on spreads of seminiferous tubules (Figure 8A and 8B). In Dmrt7 mutants, the XY body was formed and excluded Cot-1 hybridization (Figure 8B), indicating that transcriptional silencing is established normally in mutant pachytene cells. We also examined expression of the Y-linked gene Rbmy, which normally is inactivated during pachytene and reactivated after secondary meiosis begins [54,55]. Rbmy was inactivated normally in pachytene cells of Dmrt7 mutants, based on immunofluorescent staining with an anti-RBMY antibody (Figure S5). We also examined heterochromatin protein 1 beta (HP1β), which normally localizes to the X centromere at mid-pachynema and then spreads through the XY body as it internalizes during diplonema [56]. We found that HP1β localization is normal in DMRT7 mutant cells in mid-pachynema (Figure 8C and 8D). These results suggest that XY body formation and initiation of MSCI both occur normally in Dmrt7 mutant germ cells.

Figure 8

Chromatin Abnormalities in Dmrt7 Mutant Diplotene Germ Cells

(A and B) MSCI occurs normally in mid-pachytene spermatocyte of Dmrt7 mutant. Arrows indicate XY body in all panels. Cot-1 RNA FISH (red) reveals normal silencing of sex chromosome transcription in XY body. This is consistent with wild-type mid-pachynema as described previously [16].

(C and D) HP1β localization in mid-pachytene spermatocytes. HP1β localizes to X chromosome centromere (arrowhead) in wild-type (C) and mutant (D).

(E and F) Cot-1 hybridization in diplotene. Presumptive XY body, based on DAPI and γH2AX localization, is indicated by arrow in (F).

(G and H) HP1β in diplotene. Wild-type cell (E) has HP1β throughout XY body, whereas mutant cell (H) only has localization to X chromosome centromere (arrowhead).

(I and J) H3-2meK9 localization in diplotene cells. Mutant cell (J) lacks strong concentration of this mark to the XY body seen in wild-type (I).

(K and L) H3-3meK9 localization in diplotene cells. Mutant cell (L) has no localization of this mark to the XY body. γH2AX localizes to autosomes in mutant cell, possibly indicating onset of apoptosis.

(M) Example of mutant diplotene cell with normal HP1β accumulation to the XY body.

All images except those in (M) are single Z sections.

We next considered the possibility that sex chromatin is established normally but is not properly modified as cells exit pachynema and begin to form PMSC. Although most Dmrt7 mutant cells are eliminated by apoptosis prior to diplonema, we were able to examine epigenetic markers of PMSC in rare Dmrt7 mutant spermatocytes that escaped pachytene arrest and progressed into diplonema. First, we examined nascent transcription by Cot-1 hybridization. Although heterochromatic regions generally showed lower Cot-1 signal than euchromatic regions (Figure 8E and 8F), in some mutant cells the sex chromatin appeared to be incompletely silenced relative to wild-type (Figure 8F). We also examined three epigenetic signatures of PMSC: histone H3 dimethylated or trimethylated at lysine-9 (H3-2meK9, H3-3meK9) and spreading of HP1β through the XY body [16,57,58] (S. H. Namekawa, unpublished data). We observed defects in sex chromatin localization of all three markers in Dmrt7 diplotene cells. Although HP1β localization to the X chromosome centromere initially appeared normal at mid-pachynema, we observed Dmrt7 mutant diplotene cells that failed to show spreading of HP1β to the entire XY body (Figure 8G and 8H). Similarly, we found Dmrt7 mutant diplotene cells lacking accumulation of H3-2meK9 and H3-3meK9 marks onto the sex chromatin (Figure 8I–8L).

Not all Dmrt7 mutant diplotene cells showed abnormal localization of HP1β to the sex chromatin (Figure 8M). In one experiment, 11/27 mutant cells in diplonema lacked HP1β on the XY body, as compared with 2/22 wild-type cells. We hypothesize that the mutant cells with normal HP1β may be those that can complete meiosis (Figures 4 and 5). Some of the mutant diplotene cells showing abnormal sex chromatin also had abnormal autosomal γH2AX staining (Figure 8L). γH2AX localizes to double-strand DNA breaks, so this staining may indicate that some diplotene mutant cells are approaching or entering apoptosis [59]. We did not observe sex chromatin defects prior to diplonema, but we cannot exclude the possibility that earlier defects exist and the affected cells are rapidly eliminated.

In the preceding experiments, we staged cells based on XY body internalization. Because this process could be abnormal in the mutant cells, we also staged mutant cells by chromosome morphology using an antibody to SYCP3 (Figure 9). This independently identified Dmrt7 mutant diplotene cells lacking HP1β accumulation in the XY body, such as the example in Figure 9B. From these results, we conclude that Dmrt7 mutant cells establish a normal XY body in mid-pachynema, but then have multiple epigenetic defects in the sex chromatin transition from pachynema to diplonema.

Figure 9

Abnormal Sex Chromatin in Cells Staged by Chromosome Pairing Status

(A) Spread of wild-type germ cell stained with DAPI, anti-SYCP3, and anti-HP1β showing chromosome morphology typical of diplonema and internalized XY body with HP1β accumulation.

(B) Spread of Dmrt7 mutant germ cell showing normal diplotene chromosome morphology and internalized XY body, but no HP1β accumulation in the XY body.

XY chromosome pairs are indicated by arrow.

Discussion

In this study, we find that the DM domain protein DMRT7 is required for male germ cells to complete meiotic prophase but is dispensable in the female germ line. In males, DMRT7 expression is highest in pachytene spermatocytes, and the protein preferentially localizes to the XY body. Consistent with this expression, we found that most mutant male germ cells arrest in pachynema and undergo apoptosis, although a small proportion can progress to diplonema and sometimes beyond. Examination of chromatin and nascent transcription in mutant cells that progressed to diplonema revealed sex chromatin abnormalities, as discussed below.

The pachytene stage of prophase involves tremendous chromosomal changes as the homologs align, synapse, and recombine. During this period, at least one pachytene surveillance system exists to monitor key events of meiotic progression. Cells in which any of these events is anomalous are efficiently eliminated by apoptosis [46]. Another key event of pachynema in male mammals is the packaging of the sex chromosomes into the XY body and the establishment of MSCI. Examination of male meiosis in XYY mice and mice carrying a sex-chromosome-to-autosome translocation showed that cells in which a sex chromosome escapes MSCI are eliminated prior to late-pachynema [17]. This indicates that the establishment of MSCI also is subject to surveillance.

Since the arrest and apoptosis of Dmrt7 mutant spermatocytes could result from perturbation of any of the critical pachytene events mentioned above, we tested whether they occur abnormally in the mutant cells. We found that chromosomal synapsis and recombination appear normal in Dmrt7 mutant cells. We therefore focused on the XY body, the most prominent site of DMRT7 accumulation. First, we tested whether the XY body forms and MSCI is established in Dmrt7 mutant cells. Surprisingly, we found that these cells form an XY body with normal morphology and proper accumulation of all the chromatin marks we examined. Moreover, Cot-1 hybridization and analysis of RBMY expression demonstrated that MSCI initiates normally in the XY body of mid-pachytene Dmrt7 mutant cells.

We did, however, observe three specific defects in the sex chromatin of Dmrt7 mutant germ cells that avoided arrest in pachynema and were able to enter diplonema. Normally cells accumulate H3-2meK9 and H3-3meK9 marks and HP1β protein on the sex chromatin as they progress to diplonema, but we observed mutant diplotene cells lacking these features. Thus, although a minority of Dmrt7 mutant germ cells can progress from pachynema to diplonema, there are defects in sex chromatin modification during the transition. A function in male sex chromatin can reconcile the findings that DMRT7 is required for meiosis, but only in males, and is present only in mammals. A proportion of mutant diplotene cells have apparently normal sex chromatin (for example, Figure 8M); these are likely to be the cells that can progress beyond diplonema.

Because most Dmrt7 mutant germ cells are eliminated by apoptosis around the time at which we observed sex chromatin defects, a simple model is that the apoptosis is a consequence of the sex chromatin defects. The reciprocal situation (sex chromatin defects caused by apoptosis) is possible, but seems unlikely, because we observed mutant cells with sex chromatin defects but no indications of apoptosis. Alternatively, apoptosis and abnormal sex chromatin may be two independent consequences of Dmrt7 loss. This question cannot be answered definitively until we know the detailed molecular mechanism of DMRT7.

A number of other proteins have been identified that interact with the XY body, including histone variants and modified histones, a testis-specific histone methyl transferase, chromobox proteins, an orphan receptor germ-cell nuclear factor, and recombination-related proteins [60]. A common feature of these proteins is involvement with heterochromatin and/or transcriptional repression. DMRT7 is unusual among XY body proteins in being related to highly site-specific transcriptional regulators. An attractive speculation is that DMRT7 may provide sequence specificity in recruiting other proteins, such as chromatin modifiers, to the XY body as part of the transition to PMSC. Chromatin regulation may be a common mechanism for DM domain proteins, as we find that other DM domain proteins associate with chromatin modifying complexes (M. W. Murphy, D. Zarkower, and V. J. Bardwell, unpublished data).

The finding that Dmrt7 is essential for mammalian meiosis expands the known functions of this gene family. Invertebrate DM domain genes so far have only been found to function in somatic cells. Two other DM domain genes, Dmrt1 and Dmrt4, do affect germ cell development in the mouse. Dmrt1 is required in pre-meiotic male germ cells for differentiation of gonocytes into spermatogonia, as well as in Sertoli cells, but it is not expressed in meiotic cells [31] (S. Kim and D. Zarkower, unpublished data). The requirement for DMRT1 in pre-meiotic germ cells and DMRT7 in meiotic germ cells demonstrates that DM domain proteins act at multiple critical points of male germ cell development. Ovaries of Dmrt4 mutant females have polyovular follicles (follicles containing multiple oocytes), but it is unknown whether this reflects a defect in the germ line or the soma. It is notable that at least three mammalian DM domain genes affect gonadal development only in one sex, given the similar roles of related proteins in directing sex-specific somatic development in other phyla.

Strikingly, Dmrt7 is present, not only in placental mammals, but also in marsupials and a monotreme (egg-laying mammal), the platypus, which has a clear Dmrt7 ortholog [36]. However, no close Dmrt7 ortholog has been reported in nonmammalian vertebrates, and our database searches did not reveal one. Thus, Dmrt7 likely arose, presumably by duplication and divergence of another Dmrt gene, shortly before or coincident with the mammalian radiation. Monotremes have five X and five Y chromosomes, which form an extended pairing chain during meiosis and appear unrelated to the sex chromosomes of the other mammals [61]. The presence of Dmrt7 in both lineages may support a common origin for either the sex chromosomes or the sex chromatin of monotremes and other mammals. A plausible model is that Dmrt7 evolved during the establishment of mammalian sex determination to help cope with ancestral differences in gene dosage, chromosome pairing, recombination, or other meiotic issues arising from sex chromosome heteromorphy. In this regard, we speculate that the recruitment of Dmrt7 during mammalian evolution may be analogous to the recruitment of chromatin regulatory complexes to achieve somatic dosage compensation during evolution of heteromorphic sex chromosomes in several phyla (reviewed in [62]). It will be of interest to determine whether DMRT7 localizes to sex chromosomes during monotreme meiosis.

In summary, we have found that the mammal-specific DM domain protein DMRT7 is essential for meiotic prophase progression in males. DMRT7 localizes to the sex chromosomes after they are assembled into specialized heterochromatin, and many Dmrt7 mutant cells have epigenetic defects in the modification of the sex chromatin between pachytene and diplotene. Although Dmrt7 belongs to an ancient and conserved gene family, it is found only in mammals, and to our knowledge DMRT7 is the only example of a mammal-specific protein that is essential for meiosis. It will be important to determine the precise mechanism by which DMRT7 affects sex chromatin regulation during male meiosis.

Materials and Methods

Generation of Dmrt7 mice.

A mouse Dmrt7 cDNA fragment containing sequences from exon 8 was used to screen a mouse BAC library from the 129/SvJ strain (Stratagene, http://www.stratagene.com), and clones containing promoter sequences were isolated and sequenced to obtain Dmrt7 genomic sequence. The targeting vector pDZ169 (diagrammed in Figure S2) was constructed by the following scheme: The vector pDZ157 was used as a backbone vector [31]. 3′ to Pgk-neo and the loxP site, we inserted, as a XmaI/XmaI DNA fragment, the third intron of Dmrt7 (from 366 bp to 2,773 bp downstream of exon 3) generated by PCR. 5′ to Pgk-neo, we inserted an EcoRI/NotI PCR fragment extending from 4,107 bp to 336 bp 5′ of the Dmrt7 translational start. Finally, we inserted a loxP site and NotI site 336 bp 5′ of the Dmrt7 translational start. In the resulting vector, the second and third exons of Dmrt7 are flanked by loxP sites (floxed). The Dmrt7-containing portions of pDZ169 were completely sequenced.

pDZ169 was linearized with PmeI and electroporated into CJ7 ES cells (originally derived from the 129S1 strain). Three homologous recombinants were identified from 296 G418-resistant colonies by Southern hybridization by use of a DNA probe from the sequences upstream of exon 1 to screen genomic DNA digested with EcoRI. Homologous recombination was confirmed on both ends of the targeted region by Southern hybridization. Probes for Southern hybridization were made by PCR using primers DM5S10/DM5S11 (5′ probe) and DM5PR1/DM5PR2 (3′ probe), listed below. Two targeted ES cell clones containing the floxed allele Dmrt7neo were injected into C57Bl/6 blastocysts to generate chimeras. Chimeric males were bred with C57Bl/6 females to generate heterozygotes carrying Dmrt7neo. Dmrt7+/Dmrt7neo females were bred with male β-actin-Cre transgenic mice to delete the floxed sequences and generate heterozygous Dmrt7−/+ deletion mutants, which were interbred to generate homozygous Dmrt7−/− mutants.

Genotyping and RT-PCR.

For genotyping, tail-clip DNA was amplified for 35 cycles. Chromosomal sex was determined by PCR with primers to the Y chromosome gene Zfy (below). The wild-type Dmrt7 allele Dmrt7+ was detected by PCR with DM5S4/DM5S5, with an annealing temperature of 60 °C. The Dmrt7flox allele was detected by PCR with DM5S5F/DM7KO7R with an annealing temperature of 62 °C. The deleted Dmrt7 allele Dmrt7− was detected with DM5S3/DM7KO7R with an annealing temperature of 62 °C. RT-PCR for Dmrt7 expression analysis was as described [23] using primers SK111/SK112 with an annealing temperature of 62 °C.

Primers.

DM5S3: 5′-AGAGTGGATTGAATCGGATAGCTC-3′

DM5S4: 5′-AGGATCTTAGTGTCGAATGAATAC-3′

DM5S5: 5′-CCCTTATCCTCCTGCATCCAGATC-3′

DM5S10: 5′-CAGGCTATGGTTAGACTTGAGCAC-3′

DM5S11: 5′-CATCACTCGCGGACAAAGATGCAG-3′

DM5PR1: 5′-CTTCTGCTACAGCCACAGGTCTGG-3′

DM5PR2: 5′-GAATTCAACTAGTATCTGTCCC-3′

DM7KO7R: 5′-CGAGGATCAAGCTCAGGTCACTAGG-3′

DM5S5F: 5′-GATCTGGATGCAGGAGGATAAGGG-3′

ZFYF: 5′-CCTATTGCATGGACAGCAGTCTTATG-3′

ZFYR: 5′-GACTAGACATGTTCTTAACATCTGTCC-3′

SK111: 5′-CCCTTCTGGAAAAGAGAACATAGC-3′

SK112: 5′-GCTCCAGGGGCCTGTGGCTGTAGC-3′

β-actinF: 5′-TGCGTGACATCAAAGAGAAG-3′

β-actinR: 5′-GATGCCACAGGATTCCATA-3′

Histological analysis and TUNEL assay.

Dissected testes were fixed in Bouin's fixative or phosphate-buffered formalin overnight at 4 °C, progressively dehydrated in a graded ethanol series, and embedded in paraffin wax. Sections (6 μm) were deparaffinized, rehydrated, and stained with hematoxylin and eosin. For TUNEL analyses, deparaffinized sections were treated with proteinase K for 15 min and quenched in 3.0% hydrogen peroxide in PBS for 5 min at room temperature. Subsequently, nuclear staining in apoptotic cells was detected using ApopTag kit (Chemicon, http://www.chemicon.com) according to the manufacturer's instructions.

Tissue immunofluorescent staining.

Slides with paraffin sections were washed in PBT (0.1% Tween 20 in PBS) and autoclaved in 10 mM citric acid (pH 6.0) to retrieve antigenicity. Slides were blocked in 5% serum (matched to the species of the secondary antibody) in PBS for 1 h at room temperature and incubated with primary antibodies overnight at 4 °C prior to detection with secondary antibodies.

Antibodies.

Rabbit polyclonal antibodies to DMRT7 were raised against a purified fusion protein containing glutathione-S-transferase (GST) fused to the C-terminal 279 amino acids of DMRT7. Antibodies to GST were removed by GST-affigel 10 chromatography and the antiserum was then purified by GST-DMRT7-affigel 15 chromatography. DMRT7 antibody was used at 1:200 dilution with a goat anti-rabbit secondary antibody (Molecular Probes, http://www.invitrogen.com) at 1:200 dilution. Other primary antibodies used for immunofluorescence were rat anti-GATA1 (1:200, Santa Cruz Biotechnology, http://www.scbt.com, sc-265), goat anti-GATA4 (1:200, Santa Cruz Biotechnology, sc-1237), rat anti-TRA98 (1:200, gift of H. Tanaka and Y. Nishimune), rat anti-BC7 (1:50, gift of H. Tanaka and Y. Nishimune), rat anti-TRA369 (1:200, gift of H. Tanaka and Y. Nishimune), rabbit anti-RAD51 (1:600 Calbiochem, http://www.calbiochem.com, PC130), mouse anti-GMP-1/SUMO-1 (1:200, Zymed, http://invitrogen.com, 33–2400), rabbit anti-phospho-H2AX (Ser139) (1:200, Upstate, http://www.millipore.com, 01–164), mouse anti-phospho-H2AX (1:200, Upstate, 05–636), mouse anti-SYCP3 (1:200, Abcam, http://www.abcam.com, ab12452), rabbit anti-HP1β (1:100, Abcam, ab10478), rabbit anti-H3-2meK9 (1:100, Upstate, 07–441), rabbit anti-H3-3meK9 (1:200, Upstate, 07–442), rabbit anti-AR (N-20) (1:200, Santa Cruz Biotechnology, sc-816), and mouse anti-αSMA clone 1A4 (1:800, Sigma, http://www.sigmaaldrich.com, A2547). Secondary antibodies used were goat anti-rabbit Alexa 488, goat anti-rabbit Alexa 594, goat anti-rat Alexa 594, and goat anti-mouse Alexa 488 (Molecular Probes) used at 1:250. Donkey anti-goat FITC (Jackson ImmunoResearch Laboratories, http://www.jacksonimmuno.com) and donkey anti-rabbit Texas Red (Jackson) were used at 1:50 according to the manufacturer's instructions.

Meiotic chromosome spread preparations and FISH.

Meiotic chromosome spread preparations were made from 3-wk-old mice, prepared as described by Reinholdt et al. [63]. For analysis of PMSC and Cot-1 RNA FISH, meiotic slides were prepared as previously described [16]. Slides containing chromosome spreads or meiotic spermatocytes were subjected to immunofluorescent staining or RNA FISH, as previously described [16,63]. For combined RNA FISH/immunostaining, we carried out RNA FISH first, followed by immunofluorescence. DNA FISH was performed using chromosome painting (Cambio, http://www.cambio.co.uk). Z-sections were captured by Zeiss Axioplan microscope (Zeiss, http://www.zeiss.com) and processed by Openlab (Improvision, http://www.improvision.com).

Supporting Information

Figure S1

Wild-Type and Dmrt7 Mutant Meiotic Spreads

Arrows indicate the location of XY body. SYCP3 antibody staining (green) shows normal synapsis of homologous chromosome synapse in Dmrt7 mutant cell. DMRT7 protein (red) is localized to the XY body in wild-type (top row) and is not detected in Dmrt7 mutant (bottom row).

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Figure S2

Targeted Disruption of Dmrt7

(A) Diagram of targeting strategy. Homologous recombination of the targeting vector with the wild-type Dmrt7 allele (Dmrt7+) resulted in the targeted allele Dmrtneo. This allele contains loxP sites flanking the translational start and the second and third exons (containing the DM domain, gray), as well as a neomycine-resistance cassette flanked by Flp recombinase recognition sites (frt sites). Mice heterozygous for the Dmrt1neo allele were mated with transgenic mice expressing Cre recombinase, resulting in deletion of the sequence between the two loxP sites, including the neo cassette. The resulting deletion allele is called Dmrt7−. Mating of Dmrt7neo mice with transgenic mice expressing the Flpe recombinase excised the neo cassette, generating the Dmrt7flox allele.

(B) Southern blots of targeted and wild-type ES cell clones. Genomic DNAs from ES cell clones were digested with NsiI/NotI and EcoRI. The 5′ external probe hybridizes to 14.2-kb (wild-type) and 6-kb (targeted) NsiI/NotI fragments. The 3′ probe hybridizes to 14.6-kb (wild-type) and 4.7-kb (targeted) EcoRI fragments.

(C) Testis sections from adult mice. SUMO-1 (green) on XY body is detected in both wild-type (top) and Dmrt7 mutant (bottom) section. DMRT7 is detected in wild-type but not detected in Dmrt7 mutant in SUMO-1-positive cells. (Wild-type images are the same as in Figure 1D.)

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Figure S3

Normal Androgen Receptor Expression in Dmrt7 Mutant Sertoli Cells

Testis sections from adult mice stained with antibody to androgen receptor (red) and counterstained with DAPI (blue). In both wild-type (top) and mutant (bottom) testis sections, androgen receptor protein is expressed in Sertoli cell and peritubular myoid cell. Mutant testis section shows abnormal localization of Sertoli cell nuclei, which are displaced from basement membrane.

AR, androgen receptor.

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Figure S4

Ub-H2A Localization to XY Body in Dmrt7 Mutant Spermatocyte

Spread pachytene spermatocytes from wild-type (top) and Dmrt7 mutant (bottom) mice, stained with antibody to Ub-H2A (green) and DAPI (blue). Arrows indicate XY body. Both wild-type and mutant pachytene cells have Ub-H2A properly localized to XY body.

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Figure S5

RBMY Silencing in Dmrt7 Mutant Spermatocytes

(A) SYCP3 (green) and RBMY (red) immunostaining of spread wild-type and Dmrt7 mutant testicular cells. In wild-type and mutant cells, RBMY is expressed at low levels in leptotene stage. In pachytene spermatocytes, RBMY has fallen to background levels in both wild-type and mutant.

(B) SUMO-1 (green) and RBMY (red) immunostaining of adult testis sections from wild-type and Dmrt7 mutant. Pre-meiotic cells near the basal membrane express high level of RBMY whereas SUMO-1-positive pachytene cells do not express RBMY, in both wild-type and mutant, indicating proper silencing.

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Click here for additional data file.

Acknowledgements

We thank members of the Zarkower and Bardwell labs for many helpful discussions; Dan Camerini-Otero, John Logsdon, and Scott Hawley for insights into evolution of meiotic regulators; Hiromitsu Tanaka, Yoshitake Nishimune, and David Elliot for generously sharing antibodies; and Dies Meijer for sharing Dhh-Cre mice. We thank the University of Minnesota Mouse Genetics Laboratory for ES cell injection. Some images were obtained using the Middlebury Biology Imaging Facility, equipped in part through the National Science Foundation (Course, Curriculum, and Laboratory Improvement, 0088412). SHN is a research fellow of the Japan Society for Promotion of Science.

Abbreviations

DM - Doublesex/MAB-3 domain

ES - embryonic stem

FISH - fluorescence in situ hybridization

GST - glutathione-S-transferase

HP1β - heterochromatin protein 1 beta

γH2AX - phosphorylated H2AX

MSCI - meiotic sex chromosome inactivation

P14 - postnatal day 14

PMSC - postmeiotic sex chromatin

RT-PCR - reverse transcriptase-PCR

SUMO-1 - small ubiquitin-related modifier 1

Ub-H2A - ubiquitinated H2A

Footnotes

Competing interests. The authors have declared that no competing interests exist.

A previous version of this article appeared as an Early Online Release on March 7, 2007 (doi:10.1371/journal.pgen.0030062.eor).

Author contributions. All authors conceived and designed the experiments and analyzed the data. SK, SHN, LMN, and JOW performed the experiments. JOW contributed reagents/materials/analysis tools. SK, SHN, JOW, JTL, VJB, and DZ wrote the paper.

Funding. This work was supported by the US National Institutes of Health (GM059152) and the Minnesota Medical Foundation. LMN was supported by the Bicentennial Fund of Middlebury College.


Deletion at ITPR1 Underlies Ataxia in Mice and Spinocerebellar Ataxia 15 in Humans

Abstract

We observed a severe autosomal recessive movement disorder in mice used within our laboratory. We pursued a series of experiments to define the genetic lesion underlying this disorder and to identify a cognate disease in humans with mutation at the same locus. Through linkage and sequence analysis we show here that this disorder is caused by a homozygous in-frame 18-bp deletion in Itpr1 (Itpr1Δ18/Δ18), encoding inositol 1,4,5-triphosphate receptor 1. A previously reported spontaneous Itpr1 mutation in mice causes a phenotype identical to that observed here. In both models in-frame deletion within Itpr1 leads to a decrease in the normally high level of Itpr1 expression in cerebellar Purkinje cells. Spinocerebellar ataxia 15 (SCA15), a human autosomal dominant disorder, maps to the genomic region containing ITPR1; however, to date no causal mutations had been identified. Because ataxia is a prominent feature in Itpr1 mutant mice, we performed a series of experiments to test the hypothesis that mutation at ITPR1 may be the cause of SCA15. We show here that heterozygous deletion of the 5′ part of the ITPR1 gene, encompassing exons 1–10, 1–40, and 1–44 in three studied families, underlies SCA15 in humans.

Author Summary



We have identified a spontaneous in-frame deletion mutation in the gene Itpr1 that causes a recessive movement disorder in mice. In an attempt to define whether any similar disease occurs in humans we performed a literature search for diseases linked to the human chromosomal region containing ITPR1. We identified the disease spinocerebellar ataxia 15 as linked to this region. High-density genomic analysis of affected members from three families revealed that disease in these patients was caused by deletion of a large portion of the region containing ITPR1. We show here that this mutation results in a dramatic reduction in ITPR1 in cells from these patients. These data show convincingly that ITPR1 deletion underlies spinocerebellar ataxia 15 in humans.

Introduction

The use of forward genetics to define novel loci of interest in human disease has become increasingly viable with the implementation of large-scale mutagenesis programs. Prior to these initiatives this work was carried out in part by the investigation of spontaneous mutations that cause disorders in mouse breeding colonies. Careful observation of these serendipitous events has led to the establishment and study of many in vivo disease models [3].

During the generation of a knockout line of mice we noted an early movement disorder that was inherited independently of targeting vector transmission. We embarked on a series of experiments to identify the genetic lesion underlying this movement disorder and to identify a cognate disease and corresponding mutation in humans. Here we describe this effort and the discovery of deletion at the ITPR1 locus as a cause of this disorder in mice and of spinocerebellar ataxia 15 (SCA15) in humans.

Results/Discussion

During the generation of a line of mice with knockout of the gene Park7 we noted an early movement disorder that was inherited independently of targeting vector transmission. Our initial observations suggested the affected mice suffered from an apparently paroxysmal movement disorder, often induced by touch. The abnormal movements occurred predominantly below the cervical level, and the disorder appeared progressive. At initial examination, a human movement disorder specialist (K. G.-H.) likened the disorder to episodic intermittent ataxia or kinesiogenic paroxysmal dystonia and predicted the involvement of an ion channel mutation in the etiology. Affected mice presented at approximately postnatal day 14, and survival time without weaning was on average 4 wk after onset.

Breeding experiments suggested that the observed disorder was inherited in an autosomal recessive manner. To map the location of the disease-causing lesion, we performed genome-wide linkage analysis using strain-specific single nucleotide polymorphisms (SNPs) at 120 loci across the mouse genome. Analysis of these data showed a single genomic region with significant linkage to disease, providing a two-point LOD score of 5.13 at marker 20.MMHAP85FLG2 on Chromosome 6qE1. The linked haplotype suggested the mutation had occurred on the 129x1/SvJ background (Figure S1).

Literature searches revealed that among disease lines mapped to 6qE1, the spontaneous mutant opt mouse displays a strikingly similar presentation to that described here [1]. The underlying genetic lesion causing the opt phenotype is a homozygous in-frame deletion of exons 43 and 44 of the gene Itpr1 (Itpr1opt/opt), encoding inositol 1,4,5-triphosphate receptor 1 (Itpr1). Sequencing of all exons and intron–exon boundaries of Itpr1 in affected mice from the current study revealed a single mutation within Itpr1: a novel in-frame deletion of 18 bp within exon 36 (Itpr1Δ18/Δ18). To confirm the pathogenicity of this mutation we crossed heterozygous mice from the current study (Itpr1wt/Δ18) with mice heterozygous for the opt mutation (Itpr1wt/opt). This resulted in two litters of mice with a total of four affected Itpr1opt/Δ18 pups (from a total of 15) with a phenotype indistinguishable from that of the Itpr1Δ18/Δ18 and Itpr1opt/opt mice [1]. Furthermore, this phenotype was similar, although less severe, to that described in a mouse line with targeted deletion of Itpr1, where ataxia was described as a prominent feature [4]. As with the Itpr1opt/opt mice, where the deletion of exons 43 and 44 is also predicted to leave the translational reading frame unaffected, the in-frame Itpr1Δ18/Δ18 deletion mutation results in markedly decreased levels of Itpr1 in cerebellar Purkinje cells. In these two spontaneous mutants [1] and in the Itpr1-deficient mouse [4] generated by gene targeting, decreased Itpr1 expression is associated with the same autosomal recessive movement disorder (Figure 1).

Given our interest in human neurological disease we sought to identify any cognate human disorders where linkage had been established to the syntenic region of the human genome, but where no causal mutation had been identified. SCA15, an adult-onset autosomal dominant progressive ataxia is linked to this locus [5]. Although missense mutation of ITPR1 had previously been ruled out [2] and the mode of inheritance was inconsistent with that seen in the Itpr1Δ18 and Itpr1opt mice, the phenotypic presence of ataxia in the mice led us to reexamine this candidate gene as a possible cause of SCA15.

We obtained genomic DNA from three affected family members and one family member with unknown disease status from the kindred originally used to define and map SCA15 (family AUS1, of Australian Anglo-Celtic origin) [2]. We performed two experiments concurrently in three affected members of this family: sequence analysis of the coding exons of ITPR1 and high-density genome-wide SNP genotyping. Sequence analysis failed to show any coding alterations segregating with disease or any alterations that were inconsistent with Mendelian patterns of inheritance within the family. However, visualization of log R ratio and B allele frequency metrics from the genome-wide SNP genotyping experiments clearly showed data consistent with a heterozygous genomic deletion across the first one-third of ITPR1 and across the first half of a neighboring gene, SUMF1 (Figure 2). This deletion was apparent in all three affected family members studied and absent from the family member with unknown affection status (Figure 3). The SNP data showed a deletion of between 188 kb and 210 kb in size; examination of SNPs at the flanking unknown regions of this deletion allowed us to delimit the borders of the deletion to 7.5 kb on the telomeric side of the deletion (between rs12634249 and rs793396) and ~14.4 kb on the centromeric side of the deletion (between rs4073665 and rs17709863). In an attempt to define whether this variation was a benign polymorphism we analyzed genome-wide SNP data at this locus, produced using the same genotyping chip, from 577 individuals of European descent who were either controls or individuals with an unrelated neurological disorder. We failed to find any deletions affecting the coding sequence of either gene, ITPR1 or SUMF1; we did, however, identify a single individual with a possible heterozygous deletion approximately 6 kb in size within intron 40–41 of ITPR1, at least 5 kb away from exon 40. Given the location of this alteration it is unlikely to effect the expression or splicing of ITPR1.

In an attempt to fine-map the breakpoints of the disease-causing deletion we performed a series of experiments designed to refine the unknown intervals at the edges between definite deleted and definite diploid sequences. These data narrowed the unknown borders to ~4 kb on the telomeric side and ~7 kb on the centromeric side. We used all possible combinations of forward orientation primers designed within the newly defined telomeric boundary and of reverse orientation primers designed within the newly defined centromeric boundary in PCR assays in an attempt to amplify across the deletion in affected family members. Using PCR primers T3F and C11R, which should be more than 200 kb apart, we were able to amplify a fragment 953 bp in size using DNA from each of the three affected family members as template. Sequencing of this fragment revealed a deletion of 201,509 bp (Figure S3), removing the first three of the nine exons of SUMF1 and the first ten of the 58 exons of ITPR1. We were unable to amplify the deletion-specific fragment in the family member of unknown affection status, or in 275 neurologically normal controls.

To further establish genetic deletion at ITPR1 as the cause of SCA15 we analyzed two additional families with an inherited cerebellar ataxia similar to that described in the AUS1 family, ascertained through neurology clinics in London, United Kingdom. DNA extracted from probands from these two families (family H33 and family H27) were also analyzed using Illumina Infinium HumanHap550 genotyping chips. These experiments showed deletion at the SCA15 locus in all affected members assayed, from SUMF1 through ITPR1. These mutations segregated with disease in these two families (Figure S3). A strategy similar to the one outlined above enabled us to sequence over the breakpoint in family H27 but not family H33. In the former, the deletion spans 344,408 bp, removing exons 1–3 of SUMF1 and 1–44 of ITPR1; in the latter, we estimate that the deletion is 310 kb in length and that it removes exons 1–3 of SUMF1 and exons 1–40 of ITPR1. The site of mutation is of interest, particularly the fact that in each of the three families the telomeric end of the deletion is anchored between exons 3 and 4 of SUMF1; sequence searches failed to identify any repeat sequences that might explain this phenomenon. With three cerebellar ataxia families segregating a SUMF1–ITPR1 deletion, and this deletion not observed in a control population, we may reasonably conclude that the association is causal, and that the deletion is indeed the genetic basis of the disease, with SCA15 the diagnosis in the two British families as well as the original Australian family.

It is improbable that heterozygosity for the deletion of SUMF1, encoding sulfatase modifying factor 1, of itself causes or contributes to SCA15. Homozygous mutation of SUMF1 results in autosomal recessive multiple sulfatase deficiency, a metabolic disorder characterized by hepatosplenomegaly, deafness, and developmental delay [6,7]. No co-occurrence of ataxia has been described in (heterozygous) parents of patients with multiple sulfatase deficiency. Conversely, mutation of ITPR1 is biologically plausible as a cause of ataxia: the protein is highly expressed in Purkinje cells; as we have shown here, mice with mutation at this locus present with ataxia; and perturbed Ca2+ signaling has previously been implicated in the etiology of ataxia, notably in episodic ataxia type 2 and SCA6 [8]. In further support of this conclusion, analysis of protein levels of ITPR1 in Epstein-Barr virus (EBV) immortalized lymphocytes from affected and unaffected AUS1 family members revealed that all affected members showed a dramatic decrease in ITPR1 levels when compared with the family member without the deletion (Figure 4).

Itpr1 contains three domains, an N-terminal inositol triphosphate binding domain, a coupling domain, and a C-terminal transmembrane domain; it also contains two protein kinase A phosphorylation sites and an ATP-binding site. Itpr1 is coupled to Ca2+ channels and facilitates Ca2+ release from the endoplasmic reticulum after binding by the intracellular second messenger inositol 1,4,5-triphosphate [9]. Itpr1 is enriched in the Purkinje cells of the cerebellum [4]. ITPR1 mutations have more than one potential pathogenic mechanism. First, the disease may be a result of haploinsufficiency at ITPR1; this concept is consistent with the observation that heterozygous deletion leads to a later onset disorder in humans, whereas homozygous deletion in mice leads to an early onset disorder, able to be expressed within the much shorter life span of the mouse. Second, we cannot rule out the existence of an alternate start site for ITPR1 that may result in a product that confers a pathogenic gain of function to the protein; however, Western blot analysis of cells derived from affected AUS1 family members, which was performed using an antibody raised against the C-terminal portion of ITPR1, failed to identify any disease-specific truncated protein products. Clearly, the identification of distinct ITPR1 mutations underlying SCA15 will help elucidate the pathogenic mechanism of this disorder.

We show here the utility of investigating spontaneous mouse mutations in understanding human disease. Currently, the small number of aged Itpr1wt/Δ18 animals precludes us from examining these mice for subtle signs and symptoms similar to those seen in SCA15 patients; however, these mice are clearly of interest to us as a potential model of SCA15. These data also demonstrate that genome-wide SNP assay can facilitate rapid detection of structural genomic mutations that may underlie disease. The data provided by these approaches provide compelling evidence that heterozygous deletion of ITPR1 causes SCA15. Clearly, sequence analysis of ITPR1 in potential SCA15 cases may provide additional insight into the disease, particularly if a stop mutation were to be identified; however, the mutational mechanism noted here means that standard sequencing approaches alone are insufficient to confidently rule out ITPR1 mutation as a cause of disease: a comprehensive gene dosage approach is also required. Given that SCA16 and autosomal dominant congenital nonprogressive ataxia have both recently been mapped to regions overlapping with the SCA15 locus [10,11], ITPR1 is a gene of importance for screening in these families. These data add weight to a role for aberrant intracellular Ca2+ signaling in Purkinje cells in the pathogenesis of spinocerebellar ataxia.

Materials and Methods

Genome-wide linkage in mice.

One hundred and twenty DNA fragments were amplified across the genome, each selected to contain one or more strain-specific SNPs that would differentiate between C57BL/6J and 129x1/SvJ inbred strains [12]. Each fragment was initially amplified in 11 affected mice and nine unaffected mice; genotype calling was performed by dye-terminator sequencing of these fragments. Linkage analysis using these data was performed using mlink [13], which revealed a positive linkage at Chromosome 6qE1, on the 129x1/SvJ background (two-point LOD score 5.13 at marker 20.MMHAP85FLG2). In an attempt to narrow the disease interval we performed backcross experiments that resulted in the generation of three additional affected mice. Genotyping of all affected mice across the disease-segregating interval revealed flanking recombinants and a candidate region of ~5 Mb, between markers D6Mit37 and 44.MMHAP85FLG5 (Figure S1). This region contains 16 genes and predicted transcripts.

Identification of the underlying genetic lesion in mice.

Identification of similar phenotypes in mice linked to the 6qE1 interval was performed by literature searches. This revealed the Itpr1opt/opt mouse, in which disease is caused by homozygous deletion mutation of exons 43 and 44 of Itpr1. Primer pairs were designed to sequence each of the coding exons and at least 50 bp of each flanking intronic sequence of Itpr1. PCR amplification of each exon was performed using DNA from two affected mice as templates. The Itpr1Δ18/Δ18 mutation was confirmed by sequencing in all affected mice (Figure S2).

Breeding experiments were performed between two female mice heterozygous for the current mutation (Itpr1wt/Δ18) and a male mouse heterozygous for the Itpr1opt mutation (Itpr1wt/opt). This resulted in two litters of mice with a total of four affected Itpr1opt/Δ18 pups (from a total of 15; two of seven from first mating; two of eight from the second mating) with a phenotype indistinguishable from that of the Itpr1Δ18/Δ18 and Itpr1opt/opt mice.

Analysis of Itpr1 protein in mice.

We performed Western blot analyses using standard techniques with ECL detection kits (Amersham, http://www.amersham.com). Briefly, dissected whole brains from postnatal day 21 littermates were homogenized in a buffer containing 50 mM Tris-HCl, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 1% sodium deoxycholate, 0.1% SDS, and a cocktail of protease inhibitors (Roche, http://www.roche.com). Homogenates were diluted appropriately, mixed with 4× reducing sample buffer, and loaded onto 4%–12% precast gradient gels (Novex, http://www.invitrogen.com) for SDS-PAGE and immunoblotting. The antibodies to Itpr1 (1:2,000) and Actb (1:5,000) were used as recommended by manufacturers.

Immunohistochemistry.

Brains were isolated from 21-d-old mice, perfused with 4% paraformaldehyde in PBS, and post-fixed overnight in the same fixative. Brains were embedded in gelatin, and 35-μm sagittal sections were cut using a sliding microtome (NeuroScience Associates, http://www.neuroscienceassociates.com). Sections from wild-type, heterozygote, and homozygous brains were placed in the MultiBrain template. Sections were washed in 1× PBS prior to 1 h of incubation in block solution containing 1× PBS with 20% normal goat serum and 0.3% Triton X-100 (pH 7.4). Sections were incubated overnight at 4 °C in primary antibodies: affinity purified polyclonal Itpr1 antibody (1:2,000, Chemicon International, http://www.chemicon.com) and monoclonal anti-Calb1 antibody (1:6,000, Sigma-Aldrich, http://www.sigmaaldrich.com) diluted in carrier solution. Following extensive washes (in 6.0 ml of PBS, three times), sections were incubated with appropriate secondary antibodies (Alexa Fluor 555 goat anti-rabbit IgG and Alexa Fluor 488 goat anti-mouse IgG [Invitrogen, http://www.invitrogen.com]) for 1 h at room temperature. Sections were washed and mounted on glass slides in a buffered medium containing Mowiol (Calbiochem, http://www.emdbiosciences.com) as described earlier [14]. Sections were imaged using a laser scanning confocal microscope (LSM 510; Zeiss, http://www.zeiss.com). Imaging parameters (pinhole, detector gain, laser power) were optimized, and were kept constant for the wild-type, heterozygous, and homozygous mutant brains. Specificity of the Itpr1 antibodies was verified by preabsorption control experiments. Antibody dilutions were incubated for 24 h at 4 °C with the immunizing peptide. Tissue sections were incubated with the preabsorbed antibodies and processed as described above. Under these conditions, no staining above autofluorescence was detected.

Analysis of ITPR1 in SCA15 patients.

DNA was extracted from EBV immortalized lymphocytes, derived from family members. The coding exons and at least 50 bp of flanking introns of ITPR1 were PCR amplified and sequenced using dye-terminator sequencing (BigDye version 3.1; Applied Biosystems, http://www.appliedbiosystems.com). Sequence reactions were run on an ABI3730XP automated sequencer as per the manufacturer's instructions (Applied Biosystems). This analysis was performed in all three affected family members for whom genomic DNA was available (members 6, 7, and 19). Primer sequences and conditions are available upon request. Sequence data were analyzed using Sequencher (Gene Codes Corporation, http://www.genecodes.com). Genome-wide SNP genotyping was performed using Infinium HumanHap550 SNP genotyping chips as per the manufacturer's protocol (Illumina, http://www.illumina.com). This product assays 555,352 unique SNPs. Data were collected using the Illumina BeadStation scanner and data collection software. Genotypes were produced using the genotyping module of BeadStudio (version 2.3.25; Illumina), and log R ratio and B allele frequency were visualized using the genome viewer tool within this package. In order to rule out the possibility that the observed deletion within ITPR1 was a benign copy number variant we examined log R ratio and B allele frequency metrics of HumanHap550 genotyping data at this locus from 577 individuals of Northern European descent from North America and Europe, produced by us as a part of an ongoing study.

In an attempt to narrow the unknown intervals flanking the deletion observed in family AUS1, we designed primers for 30 PCR amplifications that would generate overlapping fragments across the two bordering regions (primer sequence and conditions available upon request). There were ten primer pairs in the telomeric flanking region and 20 pairs in the centromeric flanking region (Figure S3). On average each product was ~750 bp in size, and amplifications were performed using genomic DNA from each of the three affected individuals (family members 6, 7, and 19). Dye-terminator sequencing of each product was performed using the forward and reverse primers designed for amplification; running and analysis of each fragment was performed as described above. Amplification of a fragment from a normal diploid genome was denoted by the presence of a heterozygous polymorphism; amplification of a fragment from a region of the genome harboring a heterozygous genomic deletion was inferred when homozygosity for the major allele and the minor allele were noted among the three affected family members (i.e., this is inconsistent with Mendelian inheritance in related individuals known to share a common haplotype).

Using the data from the experiments described above we were able to limit the size of unknown regions flanking the deletion to ~4 kb on the telomeric side and 7 kb on the centromeric side. All combinations of forward primers from the newly defined region flanking the deletion on the telomeric side with reverse primers from the newly defined region flanking the deletion on the centromeric side were used in PCR amplification reactions performed with DNA from the three affected family members and single unaffected family members. This experiment was performed in an attempt to amplify across the deleted fragment and define the exact breakpoint. A single fragment was obtained from the third forward primer from the telomeric side (T3f 5′-TGAATGCTCAATTTTCCAGC-3′) with the 11th reverse primer from the centromeric side (C11r 5′-GGGAAAATGGATAGAGGGTG-3′). The fragment, which is 953 bp in size, was sequenced as described above and compared to the current build of the human genome. A similar series of experiments was performed to identify the deletion breakpoints in families H27 and H33; we were able to amplify a 369-bp PCR product across the breakpoint found in affected members of family H27 using primer pair H27-11F 5′-GACCTCAAGAAGGCATGAATAC-3′ and H27-3R 5′-ATGGTGGCCAGGTACACAAG-3′ (Figure S4), but to date we have been unable to identify the breakpoint in family H33.

Western blot analysis in SCA15 patients.

EBV immortalized lymphoblasts from three affected family members who carry the deletion and one family member without the mutation were used as a readily accessible source of protein; all samples came from members of family AUS1. Protein extraction was performed using lysis buffer containing 1× TBS, 1% Triton X-100, and a cocktail of protease inhibitors (Roche) with overnight lysis at −80 °C. Homogenates were diluted appropriately, mixed with 4× reducing sample buffer, and loaded onto 4%–12% precast gradient gels (NuPAGE, Invitrogen) for SDS-PAGE and immunoblotting. The antibodies to ITPR1 (1:1,000) and ACTB (1:5,000) were used as recommended by manufacturers.

Supporting Information

Figure S1

Schematic of Genotyping Results across Mouse Chromosome 6 in Affected Mice

Black squares are indicative of a C57BL/6J homozygous genotype; light grey squares, a 129x1/SvJ homozygous genotype; grey squares, a C57BL/6J 129x1/SvJ heterozygous genotype; white squares, undetermined genotype. The black box bounds a region of homozygous 129x1/SvJ genotypes that segregate with disease; thus, the critical region was determined to be between markers D6Mit37 and 44.MMHAP85FLG5.

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Figure S2

Sequence of Exon 36 of Itpr1 from Four Mice

A wild-type homozygous C57BL/6J mouse (A), a wild-type homozygous 129x1/SvJ mouse (B), an affected 129x1B6 mice homozygous for the 18-bp deletion mutation (C), and an unaffected mouse heterozygous for the 18-bp deletion mutation (D). The deleted nucleotides are bounded by a green box.

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Figure S3

Additional Families Harboring Deletion at the SCA15 Locus

(A) Family H33; (B) family H27. Upper panel shows log R ratio and B allele frequency metrics generated from Infinium HumanHap550 arrays for an affected family member from each family. Log R ratio is the ratio of normalized, observed R to expected R for each SNP (each SNP is a blue dot) and thus serves as a surrogate of copy number at each locus. B allele frequency is a measure of the number of times the A or B alleles are detected at each locus (each SNP is denoted by a blue dot). Thus, SNPs with a B allele frequency of one are apparent B/B homozygotes, SNPs with a B allele frequency of 0.5 are apparent A/B heterozygotes, and those with a B allele frequency of zero are apparent A/A homozygotes. These plots show a contiguous region ~310 kb long (family H33) and ~350 kb long (family H27) with decreased copy number and apparent homozygosity indicative of a genomic deletion (shaded grey). The pedigrees below show the available family members assayed for these deletions, all of whom were affected and all of whom carried a deletion at this locus.

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Figure S4

Deleted Regions Identified in Families AUS1 and H27

(Top) Family AUS1; sequence from the PCR product generated using primers T3f and C11r from genomic DNA from an affected family member. Red arrowhead denotes the deletion breakpoint; the deletion is 201,510 bp in length.

(Bottom) Family H27; sequence flanking deleted region. Green font indicates nucleotides telomeric to the deletion; blue font indicates nucleotides centromeric to the deletion. The deletion is 344,408 bp in length. Basepair positions are based on NCBI genome build 36 reference assembly.

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Click here for additional data file.

Accession Numbers

The OMIM (http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=OMIM) accession number for SUMF1 is 607939.

Acknowledgements

We thank the SCA15 family members for participating in this study.

Abbreviations

EBV - Epstein-Barr virus

SCA[number] - spinocerebellar ataxia [number]

SNP - single nucleotide polymorphism

Figures and Tables

Figure 1

Immunohistochemistry and Western Blot Analysis of ITPR1 Protein Levels in Mouse Cerebellum

(A–F) Immunohistochemistry of cerebellum from a wild-type mouse (A and D), a mouse heterozygous for the Itpr1 18-bp deletion (B and E), and a mouse homozygous for the 18-bp Itpr1 deletion (C and F). (A–C) Immunohistochemistry using polyclonal Itpr1 anti-rabbit antibody (1:2,000; Alexa Fluor 555); (D–F) immunohistochemistry using monoclonal Calb1 anti-mouse antibody (1:6,000; Alexa Fluor 488). Scale bars denote 100 μm. As previously described, Iptr1 is highly expressed in the Purkinje cells. Notably, there appears to be decreased immunoreactivity to Itpr1 in the heterozygous and homozygous mutant mice.

(G) Western blot performed to examine Itpr1 levels in whole brain from wild-type, Itpr1wt/Δ18, and Itpr1Δ18/Δ18 mice; this clearly shows a reduction of Itpr1 in brain tissue from Itpr1wt/Δ18 mice and a greater reduction of Itpr1 in Itpr1Δ18/Δ18 mice.

Figure 2

Metrics Derived from Analysis of DNA from Affected Family Member 7 Using Illumina Infinium HumanHap550 Genotyping Chips

The upper and lower plots are log R ratio and B allele frequency, respectively, at an ~800-kb segment on the p arm of Chromosome 3. Log R ratio is the ratio of normalized, observed R to expected R for each SNP (each SNP is a blue dot) and thus serves as a surrogate of copy number at each locus. B allele frequency is a measure of the number of times the A or B alleles are detected at each locus (each SNP is denoted by a blue dot). Thus, SNPs with a B allele frequency of one are apparent B/B homozygotes, SNPs with a B allele frequency of 0.5 are apparent A/B heterozygotes, and those with a B allele frequency of zero are apparent A/A homozygotes. Clearly, these plots show a contiguous region ~200 kb long with decreased copy number and apparent homozygosity (bounded by a red box). As we have demonstrated previously, this is indicative of a heterozygous genomic deletion [15]. Below these plots is a schematic of the two known genes affected by this deletion, ITPR1 and SUMF1.

Figure 3

Mutation Analysis in the Australian SCA15 Family

(Top) Pedigree of kindred. Filled symbols denote affected individuals; open symbols, unaffected individuals; grey symbol denotes unknown disease status; bulls-eye symbol denotes obligate carrier. w/w, wild-type at ITPR1; w/m, heterozygous carrier of the ITPR1 deletion.

(Middle) Schematic of primer pairs used to narrow the unknown regions between known deleted sequence and known diploid sequence at the SCA15 locus. Nine primer pairs (T1–T9) were used to amplify across the unknown region telomeric to the known deleted region; 19 primer pairs (C1–C19) were used to amplify across the unknown region centromeric to the known deleted region. All PCRs were carried out in the three affected family members. Analysis of these data narrowed the unknown region, and ultimately we were able to use primer T3f and C11r to amplify across the deletion breakpoint in the three affected family members, producing a fragment of 953 bp in affected individuals.

(Bottom) Gel showing amplification product using primer pair T3f and C11r from affected pedigree members 6, 7, and 19; in pedigree member 23, with unknown disease affection status; in a neurologically normal control (C); and in a no template control (NC).

Figure 4

Western Blot Analysis of ITPR1 Protein Levels in EBV Immortalized Lymphoblasts from AUS1 Family Members

Western blot performed to examine ITPR1 levels in EBV immortalized lymphocytes from AUS1 affected family members carrying the ITPR1 deletion and from an AUS1 family member of unknown disease status who does not carry the deletion. Notably the samples from patients with ITPR1 deletion show a dramatic decrease in ITPR1 levels. To demonstrate equal loading, these samples were diluted one in five, and the Western blot was repeated using an antibody against ACTB.

Footnotes

A previous version of this article appeared as an Early Online Release on May 16, 2007 (doi:10.1371/journal.pgen.0030108.eor).

Author contributions. EMCF, JTR, and ABS conceived and designed the experiments. JvdL, JC, MAK, LAH, SS, MRC, HCF, XL, DH, JSS, IR, and HC performed the experiments. JvdL, JC, MAK, SS, MRC, KGH, HCF, JSS, JTR, and ABS analyzed the data. HH, NWW, PG, JH, ES, RJMG, SMF, HC, and ABS contributed reagents/materials/analysis tools. RJMG, EMCF, and ABS wrote the paper.

Funding. This research was funded in part by the intramural programs of the National Institute on Aging and the National Institute on Neurological Disorders and Stroke (NINDS), both of the National Institutes of Health, Department of Health and Human Services, United States of America. MAK was supported by a NINDS Competitive Postdoctoral Fellowship. HH is supported by the Medical Research Council, United Kingdom.

Competing interests. The authors have declared that no competing interests exist.


Rb-Mediated Neuronal Differentiation through Cell-Cycle–Independent Regulation of E2f3a

Abstract

It has long been known that loss of the retinoblastoma protein (Rb) perturbs neural differentiation, but the underlying mechanism has never been solved. Rb absence impairs cell cycle exit and triggers death of some neurons, so differentiation defects may well be indirect. Indeed, we show that abnormalities in both differentiation and light-evoked electrophysiological responses in Rb-deficient retinal cells are rescued when ectopic division and apoptosis are blocked specifically by deleting E2f transcription factor (E2f) 1. However, comprehensive cell-type analysis of the rescued double-null retina exposed cell-cycle–independent differentiation defects specifically in starburst amacrine cells (SACs), cholinergic interneurons critical in direction selectivity and developmentally important rhythmic bursts. Typically, Rb is thought to block division by repressing E2fs, but to promote differentiation by potentiating tissue-specific factors. Remarkably, however, Rb promotes SAC differentiation by inhibiting E2f3 activity. Two E2f3 isoforms exist, and we find both in the developing retina, although intriguingly they show distinct subcellular distribution. E2f3b is thought to mediate Rb function in quiescent cells. However, in what is to our knowledge the first work to dissect E2f isoform function in vivo we show that Rb promotes SAC differentiation through E2f3a. These data reveal a mechanism through which Rb regulates neural differentiation directly, and, unexpectedly, it involves inhibition of E2f3a, not potentiation of tissue-specific factors.

Author Summary



The retinoblastoma protein (Rb), an important tumor suppressor, blocks division and death by inhibiting the E2f transcription factor family. In contrast, Rb is thought to promote differentiation by potentiating tissue-specific transcription factors, although differentiation defects in Rb null cells could be an indirect consequence of E2f-driven division and death. Here, we resolve different mechanisms by which Rb controls division, death, and differentiation in the retina. Removing E2f1 rescues aberrant division of differentiating Rb-deficient retinal neurons, as well as death in cells prone to apoptosis, and restores both normal differentiation and function of major cell types, such as photoreceptors. However, Rb-deficient starburst amacrine neurons differentiate abnormally even when E2f1 is removed, providing an unequivocal example of a direct role for Rb in neuronal differentiation. Rather than potentiating a cell-specific factor, Rb promotes starburst cell differentiation by inhibiting another E2f, E2f3a. This cell-cycle–independent activity broadens the importance of the Rb–E2f pathway, and suggests we should reassess its role in the differentiation of other cell types.

Introduction

The simplicity of the retina makes it an ideal tissue to study neurogenesis. Its development proceeds through three overlapping steps starting with retinal progenitor cell (RPC) proliferation, followed by birth of post-mitotic retinal transition cells (RTCs, also referred to as precursors), and ending with terminal differentiation of seven major cell types (Figure 1A) [1]. RPCs are multipotent and exit the cell cycle to generate different RTCs at specific time periods in development [1]. This process of RTC “birth” requires coupling of differentiation and cell cycle exit. Once born, post-mitotic RTCs migrate and form different retinal layers. Rods and cones make up the outer nuclear layer (ONL); horizontal, bipolar, and amacrine cells, as well as Müller glia cell bodies, reside in the inner nuclear layer (INL); and ganglion and displaced amacrine cells form the ganglion cell layer (GCL) (Figure 1A). The outer plexiform layer (OPL) and inner plexiform layer (IPL) house synaptic connections separating the ONL/INL and INL/GCL, respectively.

Figure 1

E2f1, but Not E2f2 or E2f3, Loss Rescues Ectopic Division and Cell Death in the Rb KO Retina

(A) Retinal development. At E11 the retina is a NBL of dividing RPCs (white circle, green nuclei). RPC cell bodies oscillate along processes as they progress through the cell cycle. By P0 the NBL contains both RPCs and post-mitotic RTCs (coloured circles, red nuclei) and is separated from the GCL by the IPL. By P8 there are no RPCs, fewer RTCs, an OPL, and more differentiated rods (r) and cones (c) in the ONL; horizontal (h), bipolar (b), Müller (m), and amacrine (a) cells in the INL; and ganglion (g) and displaced amacrine cells in the GCL. Development is complete by ~P18.

(B) Rb is thought to regulate cell cycle and apoptosis by repressing E2fs, but to promote differentiation by potentiating tissue-specific transcription factors. However, Rb loss could also perturb differentiation through the indirect effects of abnormal division or death, and/or through direct regulation of differentiation genes by E2fs.

(C and D) Horizontal retinal sections of the indicated genotypes and ages were stained for nuclei (DAPI, blue), and (C) S-phase (anti-BrdU, red) or (D) apoptosis (TUNEL, red). Scale bars are 50 μm.

(E–G) Quantification of (E) all BrdU+ cells, (F) ectopic BrdU+ cells in GCL at P0, and (G) total TUNEL+ cells.

(H) Real-time RT-PCR analysis of E2fs and E2f target genes in P8 retinas of the indicated genotypes.

Error bars represent SD of measurements from three animals, and asterisks indicate a significant difference between the WT and indicated genotypes (*, p <0.05; **; p <0.01; ANOVA and Tukey HSD test for [E–G] and Fisher test for [H]).

The retinoblastoma protein (Rb) is critical for cell cycle exit during retinal transition cell birth. Rb knockout (KO) RTCs continue to proliferate inappropriately and some (rod, ganglion, and bipolar cells) die by apoptosis [2,3]. Rb controls the cell cycle by binding and inhibiting E2f transcription factors (E2fs) (Figure 1B), first defined as transcription factors that bind adenoviral E2 regulatory elements and subsequently shown to be critical cell cycle regulators [4,5]. E2fs bind to DNA as heterodimers with proteins of the related Tfdp family. E2f1, E2f2, and E2f3a are “activating E2fs” that are required for fibroblast division. They are strong transcriptional activators that can drive G0 fibroblasts into cycle, and are inhibited when bound to Rb [4,5]. Ectopic division in Rb KO embryos can be rescued to various extents in different tissues by knocking out E2f1, E2f2, or E2f3 [6–9], but which member(s) drive division in Rb KO RTCs is unknown. Other members of the family, such as E2f4 and E2f5, are known as “repressive E2fs” because they are weak activators and appear to be primarily involved in gene silencing in quiescent or differentiated cells.

Activating E2fs may also promote apoptosis in the Rb KO retina (Figure 1B). Originally, E2f1 was considered the primary pro-apoptotic member of the family [10]. However, this view was reevaluated when it was shown that either E2f1 or E2f3 deletion rescues apoptosis in the developing central nervous system (CNS) of Rb KO embryos [6,11]. Subsequently, CNS apoptosis was shown to be an indirect result of placental defects and probable hypoxia [12–14]. Indeed, E2f3-induced apoptosis in fibroblasts has recently been shown to require E2f1 [15]. Thus, it is controversial whether E2f3 is required for apoptosis of any Rb KO cell type. Determining which activating E2fs promote death in distinct Rb KO tissues requires conditional rather than germ line models of Rb deletion to avoid secondary indirect effects (such as hypoxia).

E2f family diversity is expanded by E2f3 isoforms. Alternative promoters generate two forms (a and b) that are identical except for distinct first exons [16]. E2f3a is a strong activator, and, like other activating E2fs, its expression is induced when quiescent cells are stimulated to divide [16]. E2f3b, like repressive E2fs, is present in both quiescent and dividing cells, and in quiescent fibroblasts it associates primarily with Rb, suggesting that it mediates repression [16–18]. Indeed, silencing the Cdkn2d (p19Arf) locus in unstressed cells relies on E2f3b [19]. Other E2fs may also exist in isoforms since at least two mRNA species have been detected for E2f1 and E2f2 [16]. The roles of E2f isoforms in vivo are unknown.

E2fs are also regulated by subcellular localization. Although this feature has been best characterized for repressive E2fs [20–22], it also affects activating E2fs [23–25]. The distribution of E2f isoforms has never been assessed.

It has been known for many years that Rb loss perturbs neuronal differentiation [26–29]. However, prior work could not exclude the possibility that differentiation defects are simply an indirect consequence of abnormal division and death. If Rb does regulate differentiation directly it is unclear whether it does so in all or a subset of neurons. Moreover, the mechanism has never been solved. In other cell types where Rb may promote differentiation directly, such as muscle and bone, it seems to do so through E2f-independent means by potentiating tissue-specific transcription factors (Figure 1B) [30–33]. In the retina, others have noted abnormally shaped Rb KO rods and have suggested Rb may directly promote their morphogenesis by activating retina-specific factors [29]. However, differentiation defects in any Rb KO neuron could be an indirect effect of ectopic division and/or apoptosis (Figure 1B). Thus, it is critical to study differentiation of Rb KO cells in the absence of ectopic proliferation and death.

Here, we establish that Rb suppresses RTC division and death by inhibiting E2f1, not E2f2 or E2f3. When these defects were rescued, most retinal neurons, including rods, survived, differentiated, and functioned normally. Thus, unexpectedly, retina-specific differentiation factors function independently of Rb. However, comprehensive assessment of the Rb/E2f1 double-null rescued retina revealed a differentiation defect in cholinergic starburst amacrine cells (SACs). Recent breakthroughs have revealed that these interneurons are critical for direction selectivity and developmentally important rhythmic bursts [34–36]. However, their differentiation is poorly understood. Contrary to the prevailing view that Rb promotes differentiation through E2f-independent tissue-specific transcription factors, we show that Rb facilitates SAC development through E2f3. Defects in Rb null SACs correlated with specific E2f3 expression in these cells, and E2f3 expression was absent in neurons that differentiated without Rb. E2f3 is also present in a specific subset of other CNS neurons, implying that this may be a general mechanism by which Rb facilitates neurogenesis. To define the mechanism in even more detail, we determined which E2f3 isoform Rb targets to control SAC differentiation. E2f3b mediates Rb function in quiescent fibroblasts [19], yet no prior studies to our knowledge have dissected E2f3a or E2f3b functions in vivo. Using an isoform-specific null mouse we show that Rb drives SAC differentiation through E2f3a. Thus, independent of E2f1-mediated effects on division and death, Rb does regulate neuronal differentiation, but only in specific neurons and, unexpectedly, through E2f3a, not tissue-specific differentiation factors.

Results

Rb Regulates Division and Death through E2f1

We used the α-Cre transgene to delete floxed Rb exon 19 at embryonic day (E) 10 in peripheral retina [2]. RbloxP/loxP;α-Cre mice were bred with strains lacking E2f1 or E2f2 in the germ line, or a strain carrying a floxed E2f3 allele [5]. RbloxP/loxP;E2f1+/− and RbloxP/loxP;E2f1+/−;α-Cre mice were bred to produce RbloxP/loxP;E2f1−/−;α-Cre mice at a frequency of 1/8 and littermate controls at the same or higher (1/4) frequency. For simplicity we will refer to the RbloxP/loxP;E2f1−/−;α-Cre peripheral retina as the Rb/E2f1 double knockout (DKO) retina. Similar strategies were employed in the case of E2f2 or E2f3. Cre-mediated excision of Rb and E2f3 alleles in the retina was confirmed by PCR as described previously [2,5].

To measure ectopic cell division, mice were pulse-labelled with bromodeoxyuridine (BrdU) 2 h before sacrifice and the peripheral retina analyzed for BrdU incorporation by immunofluorescence. As reported before [2,3], Rb KO retinas exhibited both spatial and temporal ectopic DNA synthesis (Figures 1C and S1A). This is easily detected at E14, E16, and postnatal day (P) 0 in the inner retina where abnormal BrdU+ ganglion and amacrine RTCs are located, or on the outermost region of the P0 retina, where BrdU+ photoreceptor RTCs reside (Figures S1A and S2, arrows) [2]. Ectopic RTC division in Rb KO retinas is even more obvious at P8 or P18, when division is completed in wild-type (WT) retina (Figures 1C and S1A). Strikingly, the ectopically positioned S-phase cells at E14, E16, and P0 and all the abnormal division at P8 and P18 were completely suppressed in the Rb/E2f1 DKO retina (Figures 1C, 1E, 1F, S1A, and S2). In contrast, deletion of E2f2 or E2f3 had no effect at any stage of development. Analysis of mitotic cells with anti–phosphohistone 3 (PH3)–specific antibodies confirmed that loss of E2f1, but not E2f2 or E2f3, suppressed ectopic division (Figure S3). Deleting one E2f1 allele partially suppressed ectopic S-phase and mitosis in Rb KO RTCs (Figures 1C, 1E, 1F, S1A, S2, and S3), suggesting that E2f1 drives ectopic division in Rb KO RTCs in a dose-dependent fashion. These data contrast with previous findings in the lens and CNS of Rb KO embryos, where deletion of any activating E2f suppresses ectopic division to some extent [6–9].

Loss of Rb in the retina results in considerable RTC apoptosis, eliminating most bipolar and ganglion cells as well as many rods (Figure 2A–2D) [2,3]. The loss of Rb KO rods is evident from the thinner ONL, and the death of these cells as well as bipolar and ganglion neurons can be detected directly by double labelling for apoptotic and cell-type-specific markers [2] (M. P. and R. B., unpublished data). Loss of peripheral Rb KO ganglion cells is also evident from thinning of the optic nerve (D. C. and R. B., unpublished data). Deleting E2f1, but not E2f2 or E2f3, blocked this ectopic cell death in a dose-dependent fashion (Figures 1D, 1G, and S1B).

Figure 2

E2f1 Deletion Rescues Ganglion, Rod, and Bipolar Cells in the Rb KO Retina

(A) Horizontal retinal sections from mice of the indicated ages and genotypes were stained for nuclei (DAPI, blue) and markers that detect ganglion cells (Pou4f2, red), rods and cones (Sag [rod arrestin], green), and rod bipolar cells (Prkca, green, and Cabp5, red). Scale bars are 50 μm.

(B) Quantification of Pou4f2+ ganglion cells.

(C) Quantification of Prkca+ and Cabp5+ bipolar cells.

(D) Thickness of the ONL, which represents the number of rods.

Error bars represent SD of measurements from three animals, and asterisks indicate a significant difference between retinas of WT and the indicated genotypes, unless indicated otherwise by connecting lines (*, p < 0.05; **; p < 0.01; ANOVA and Tukey HSD test).

(E and F) ERGs were recorded from the indicated genotypes under dark-adapted (scotopic) conditions, and (E) intensity series and (F) b-wave amplitudes as a function of the logarithm of the flash intensity were determined. (F) Further illustrates that the relative influence of the mutations on the photoreceptors (indicated by the saturated a-wave amplitude, right graph) was not substantially different from their effect on the b-wave response (dominated by the bipolars, left graph) at the same intensity of 10 cd·s/m2.

To investigate the molecular mechanism that underlies the unique role of E2f1, we assessed the expression of known E2f targets as well as other genes that regulate the cell cycle and apoptosis. Numerous positive and negative cell cycle and apoptotic regulators were up-regulated in the Rb KO retina (Figure 1H). Among the E2f family, E2f1, E2f2, E2f3a, and E2f7 were induced following Rb loss, but E2f3b, E2f4, and E2f5 were unaffected. Consistent with the BrdU and terminal dUTP nick-end labelling (TUNEL) analyses, E2f1 deletion specifically reversed all these molecular defects, but E2f3 deletion had no effect (Figure 1H).

Normal Differentiation in the Rb/E2f1 DKO Retina

Because E2f1 deletion blocks abnormal division and death in the Rb KO retina, the Rb/E2f1 DKO retina provided a unique opportunity to evaluate whether Rb controls differentiation independent of cell cycle effects. The Rb/E2f1 DKO retina had many Sag+ (S-antigen/rod arrestin) photoreceptors, Pou4f2+ (Brn3b) ganglion cells, and numerous Prkca+/Cabp5+ bipolar neurons (Figure 2A–2D). In contrast, there was no such rescue of cell types in Rb/E2f2 or Rb/E2f3 DKO retinas (Figure S4). Analysis with general neuronal markers Mtap2 (MAP2) and Snap25, as well as other markers expressed in bipolar cells (Chx10, Rcvrn, Vsx1, Tacr3, and Atp2b1) and rod photoreceptors (Rho and Rcvrn) confirmed rescue of the Rb/E2f1 DKO retina (Table S1). Moreover, neurons exhibited the same complex morphology as in WT retina. Bipolar cell bodies were located in the INL and had ascending and descending processes ending in the OPL and IPL, respectively (Figure 2A). In addition, the Rb/E2f1 DKO retina had a healthy appearing ONL consisting of morphologically normal rods with descending processes ending in the OPL and ascending processes that terminated in inner and outer segments (Figure 2A). It was suggested that Rb might regulate photoreceptor differentiation, possibly through rod-specific transcription factors (Figure 1B) [29]. However, if Rb does regulate photoreceptor differentiation, it does so by inhibiting E2f1, not by potentiating rod differentiation factors, such as Otx2, Crx, or Nrl. It is impossible to tell whether E2f1 perturbs differentiation directly, by affecting the expression of genes that modulate maturation, and/or indirectly through its effects on proliferation and survival (Figure 1B).

As with ectopic division and apoptosis (Figure 1C and 1D), the rescue of Rb KO retinal bipolar, ganglion, and rod cells was dependent on E2f1 dose (Figure 2A–2D). Separate from its role in driving ectopic division of Rb KO RTCs, E2f1 also promotes normal RPC division since in its absence RPC proliferation drops ~2-fold (D. C. and R. B., unpublished data). This modest reduction of RPC numbers in the absence of E2f1 accounts for the slight reduction in the number of ganglion cells at P0, in the number of bipolar cells at P18 or P30, and in the thickness of the ONL at P18 or P30 in the E2f1 KO and Rb/E2f1 DKO retina (Figure 2B–2D). The morphology of E2f1 KO neurons was WT (Figure 2A). Despite a slight drop in absolute cell numbers, the proportion of Rb/E2f1 DKO and E2f1 KO bipolar cells was the same as WT (data not shown). Slightly reduced cell numbers were not due to residual RTC death since we have not observed ectopic apoptosis at any embryonic or postnatal stage in the developing Rb/E2f1 DKO retina (Figures 1D, 1G, and S2). Moreover, deleting Ccnd1, which acts upstream of Rb proteins, also reduces RPC number, but does not suppress any defect in the Rb KO retina (D. C. and R. B., unpublished data). Thus, slightly reduced RPC division and dramatic rescue of severe defects in Rb KO RTCs are distinct effects stemming from the deletion of E2f1.

Normal Function of the Rb/E2f1 DKO Retina

The discovery that E2f1 loss rescues even the morphology of Rb KO neurons is surprising because Rb is thought to regulate differentiation primarily through E2f-independent pathways [30–33]. However, normal morphology may not equate to completely normal differentiation. Thus, we compared the electroretinograms (ERGs) of adult WT (α-Cre), E2f1−/−, α-Cre;RbloxP/loxP, and α-Cre;RbloxP/loxP;E2f1−/− mice. ERGs functionally assess visual signalling in the mammalian retina from photoreceptors to amacrine cells (but usually not gangion cells), and are dominated by rod and cone bipolar cells. Typically, an ERG signal begins with a negative deflection initiated by the photoreceptors (the a-wave), which is terminated by a large positive deflection due to the activation of ON bipolar cells (the b-wave).

Responses to dim light in dark-adapted (scotopic) conditions specifically assess the rod system, and were defective in the Rb KO retina (Figure 2E). The substantial reduction of both a- and b-waves is consistent with rod and bipolar cell apoptosis [2]. The sensitivity of the residual response appeared unchanged, suggesting it arose from the Cre-negative portions of the retina. Responses were about the same in the WT and E2f1 KO retina, and, most importantly, also the Rb/E2f1 DKO response median lay at the lower end of the normal range for most intensities (Figure 2F). Thus, E2f1 deletion almost completely rescued the rod system in the Rb KO retina.

Light-adapted (photopic) recordings to specifically assess the cone system yielded comparable results. Cones represent only 3% of photoreceptors and, unlike rods, develop without Rb, but they require rods for survival, and in the Rb KO retina, they have abnormal morphology and their synaptic targets, bipolar cells, are much depleted [2]. The photopic response, a product of cone and mainly bipolar activity, was much reduced by Rb loss, but was rescued considerably in the Rb/E2f1 DKO retina (Figure S5). Again, the median amplitude lay at the lower end of the E2f1 KO range. The photopic response in E2f1 KO mice was slightly reduced relative to WT (Figure S5B), possibly because E2f1 is required for maximal expansion of embryonic RPCs, and the E2f1 KO retina has, as noted earlier, slightly fewer cells than the WT retina, although cell type proportions are unaffected (D. C. and R. B., unpublished data). Thus, marginally subnormal photopic responses in the Rb/E2f1 DKO retina can be attributed to both a reduction of cone numbers in E2f1 KO mice alone, and a “genuine” slight reduction in cone function attributable to Rb loss relative to WT. This slight effect may relate to a true differentiation defect in a subset of amacrine cells discussed below. This discussion should not obscure the major outcome that E2f1 deletion recovers most of the ERG response. Thus, E2f1 deletion not only rescues morphology but also both rod and cone system function in the Rb KO retina.

Abnormal SAC Differentiation Independent of Cell Cycle and Survival Defects

ERGs primarily assess photoreceptor and bipolar cell function, but may miss differentiation defects in other cells. To test for subtle differences we stained the Rb/E2f1 DKO retina with 43 markers (Table S1). Thirty-two proteins displayed identical patterns in WT, E2f1 KO, and Rb/E2f1 DKO retina (Table S1). The other 11 markers revealed a cell-cycle– and apoptosis-independent differentiation defect in SACs. We first studied Calb2 (calretinin), which marks a subset of amacrine and ganglion cell bodies as well as three tracks corresponding to their processes in the IPL (Figure 3A). Normal Calb2 staining was seen in the E2f1 KO IPL (data not shown). However, only one Calb2+ track was evident in the Rb KO IPL, and this defect was not rescued in the Rb/E2f1 DKO retina (Figure 3A). We quantified Calb2+ cell bodies in the Rb KO INL (corresponding to amacrine cell staining only) and observed a reduction from P8 onwards (Figures 3C and S6).

Figure 3

Differentiation Defects in Rb KO SACs

(A) P18 horizontal sections of WT, Rb KO, and Rb/E2f1 DKO retina were stained for nuclei (DAPI, blue), Calb2 (green), and Slc18a3 (red).

(B) Confocal images of P30 horizontal sections of WT and Rb KO retina were stained for nuclei (DAPI, blue), Chat and Slc18a3 (both red), and Camk2a (green). In the Rb KO section, the red stain is Chat only, as Slc18a3 is missing (see [A]).

(C) Quantification of dense Calb2+ cell bodies in the INL, total Slc18a3+ cell bodies, and Camk2a+ cell bodies in the INL. Error bars represent SD of measurements from three animals, and asterisks indicate significant differences between retinas of WT and the indicated genotypes (*, p <0.05; **; p <0.01; ANOVA and Tukey HSD test).

Scale bars are 50 μm in (A) and 20 μm in (B).

Of the three Calb2+ tracks in the IPL, the two outer tracks are from SACs, named after their extensive dendritic-tree-like morphology [37]. SACs are cholinergic, represent ~5.2% of amacrine neurons [38], and are critical for both direction selectivity [34,35] and spontaneous rhythmic activity that occurs during normal retinal development [36]. SACs in the INL synapse in the OFF layer of the IPL that responds to decreasing light, while displaced SACs in the GCL have processes that synapse in the nearby ON layer of the IPL that responds to increasing light (reviewed in [39]). Mature SAC processes stain specifically for Slc18a3 (vesicular acetyl choline transporter, VAChT) [37], and, significantly, this marker was absent in the peripheral Rb KO or Rb/E2f1 DKO P18 retina (Figures 3A and S7B). Chat, expressed from the same locus, is also SAC specific, but marks both cell bodies and processes of mature SACs [37]. Chat was seen in fewer cells in the mature Rb KO retina, and was present in the soma but absent from processes (Figure 3B). We obtained similar results for Sv2c, a synaptic vesicle protein found in SACs [40]; Kcnc1b and Kcnc2, potassium channels expressed on SAC soma and dendrites as well as a very small number of ganglion cells [41]; gamma-aminobutyric acid (GABA), an inhibitory neurotransmitter present in about half of amacrine cells including SACs, as well as horizontal and some bipolar neurons [37]; and Calb1 (calbindin), which is expressed in many amacrine cells and labels SAC process faintly (Figure S7A and S7B; Table S1; and data not shown). Finally, we also examined the effect of Rb deletion on SAC differentiation using a Chx10-Cre transgene that is expressed in a mosaic pattern across the retina, generating patches of Cre-expressing cells [42]. Consistent with the mosaic deletion pattern, we observed markedly reduced Chat/Slc18a3 staining in the IPL of Chx10-Cre;RbloxP/loxP retina compared to WT (Figure S7C). Together, these results suggest a role for Rb in SAC differentiation.

The above findings could indicate a defect in SAC specification, SAC survival, or the levels and/or transport of the markers described above. Camk2a marks both SACs and ganglion cells [37], but because ganglion cells are eliminated in the Rb KO retina, Camk2a is a specific SAC marker in this context. Importantly, Camk2a+ tracks and dendrites were present in both the WT and Rb KO retina (Figure 3B), and the number of Camk2a+ soma was similar in WT and Rb KO retina at P30 and beyond, although fewer cells stained in Rb KO retina at P18, suggesting a delay in its appearance (Figures 3C and S6B). Thus, Rb is not required for SAC survival or for process outgrowth, but rather it seems to regulate the expression and/or stability of Calb2, Calb1, Chat, Slc18a3, Sv2c, Kcnc1b, Kcnc2, and GABA in SACs, but leaves Camk2a expression virtually unaffected. The presence of Chat in some cell bodies but never in processes (Figure 3B) also suggests a transport defect. The developmental pattern of Slc18a3 expression also supported this notion. In mature WT SACs Slc18a3 is only in processes, but in early postnatal SACs, it is found in the cell body, and moves into emerging processes at approximately P4–P6. As noted above, Slc18a3 was absent at P18 in the Rb KO retina (Figure 3A); at P4 or P5 it was in cell bodies, yet was rarely present in Rb KO processes (Figures 4A and S6). Slc18a3 became virtually undetectable in Rb KO SACs by P8 (Figures 3C and S6C). These data suggest that Rb affects both the synthesis/stability and transport of SAC markers.

Figure 4

E2f3 Loss Rescues Differentiation of Rb KO SACs

(A) Horizontal retinal sections of the indicated ages and genotypes were stained for nuclei (DAPI, blue), mitosis marker PH3 (green), and Slc18a3 (red), which marks SAC soma at early stages and processes from ~P5 onwards. Arrows show mitotic PH3+ nuclei in Rb KO, Rb/E2f2 DKO, and Rb/E2f3 DKO retinas. E2f1 loss rescues the ectopic mitosis and cell death defects, but not the SAC defect. E2f2 loss has no effect. E2f3 loss does not rescue the ectopic mitosis and cell loss defects, but rescues the SAC defect. Inactivating E2f1 and E2f3 together rescues the ectopic mitosis, cell death, and SAC defects.

(B) The fraction of Camk2a+ cells that are Chat + and Slc18a3+ in the P30 retina.

(C) Horizontal P5 retinal sections were stained for nuclei (DAPI, blue), Slc18a3 (green), and Isl1 (red). Arrows show double-labelled Isl1+/Slc18a3+ cells in the inner INL.

(D) Horizontal P0 retinal sections of the indicated genotypes were stained for nuclei (DAPI, blue), cell division marker Mki67 (green), and Isl1 (red). Arrows show double-labelled dividing SACs.

(E) The fraction of Isl1+ cells in the inner NBL (INBL) of P0 retinas that are dividing (Mki67+).

Error bars represent SD of measurements from three animals, and asterisks indicate significant differences between retinas of WT and the indicated genotypes (*, p <0.05; **; p <0.01; ANOVA and Tukey HSD test). Scale bars in (A), (C), and (D) are 50 μm.

Rb Regulates SAC Differentiation through E2f3

Rb binds more than 100 proteins [43] and in some non-neuronal cells, such as skeletal muscle, adipocytes, and bone, Rb is thought to bind and potentiate tissue-specific transcription factors that promote differentiation [31–33]. Thus, we expected that Rb might interact with retina-specific factors to facilitate SAC differentiation. A direct role for E2f in mediating Rb-dependent differentiation defects (independent of cell cycle or death defects) has to our knowledge not been described, but because E2f can regulate some differentiation genes [44–48], we first tested whether E2f2 or E2f3 might perturb Rb KO SAC maturation. At multiple time points, E2f1 deletion suppressed ectopic mitosis (PH3+ cells), but did not reverse the SAC defect, and E2f2 deletion had no effect on either defect (Figure 4A). Remarkably, although E2f3 deletion did not reverse ectopic mitosis, it rescued Calb2, Slc18a3, Chat, GABA, Kcnc1b, Kcnc2, and Sv2c staining at multiple times (Figure 4A and data not shown). Rb/E2f3 DKO SAC tracks were slightly more disordered than in WT retina, most likely because of the absence of synaptic partner cells, which are killed by E2f1. Indeed, this minor defect was rescued in the Rb/E2f1/E2f3 triple knockout retina, where bipolar and ganglion cell death was rescued and SAC differentiation was restored (Figure 4A). E2f3 deletion alone did not affect SAC differentiation (Figure 4A); thus, it is unleashed E2f3 activity that is detrimental, and the critical role for Rb is to inhibit E2f3.

We quantified the fraction of Camk2a+ SACs in different genotypes and found that 60% of WT P30 Camk2a+ cells expressed Chat and Slc18a3, which dropped to only 5.6% in the Rb KO retina, and remained low at 3.7% in the Rb/E2f1 DKO retina, but rose to 91% in the Rb/E2f3 DKO retina (Figure 4B). The latter fraction is higher than WT because ganglion cells, which normally make up ~40% of Camk2a+ cells, are killed by apoptosis.

To quantify the effect of different E2fs on ectopic division specifically in SACs, we exploited Isl1 (Islet1). This marker is expressed in both SACs and ganglion cells, thus Isl1+ cells in the INL are exclusively SACs [49]. We found that 98.2% ± 1.8% of Isl1+ cells in the forming inner INL at P5 were also Slc18a3+, confirming that Isl1 is an excellent SAC marker (Figure 4C). Moreover, Isl1, unlike Slc18a3, is nuclear, which facilitates scoring of Isl1+/Mki67+ cells. It is also expressed earlier than Slc18a3, permitting analysis of SACs soon after their birth at ~E15; thus, we could study retina at P0, a time when ectopic division is high in the inner retina and prior to Rb-independent cell cycle exit associated with terminal differentiation [2]. At P0, no WT Isl1+ cells in the inner neuroblastic layer (NBL) (which is the future INL) were dividing, but 57 ± 14 Isl1+/Mki67+ cells were detected in the Rb KO inner NBL (Figure 4D). Indeed, about one-third of all Isl1+ cells in the entire inner NBL were dividing in the Rb KO retina, or ~50% in the periphery where Cre is expressed (Figure 4E and data not shown). This defect was suppressed in the Rb/E2f1 DKO retina, where we detected only 1 ± 1 dividing SAC, but not the Rb/E2f3 DKO retina, where there were 53 ± 8 dividing SACs (Figure 4D and 4E). We observed similar effects at P0 with Calb2, which marks newborn SACs and other amacrine cells (data not shown). Thus, in Rb KO SACs, E2f1 deletion suppresses ectopic division but not aberrant differentiation, whereas E2f3 deletion suppresses aberrant differentiation but not ectopic division.

Specific Expression of E2f3 in SACs and Other Subsets of CNS Neurons

The unique effect of E2f3 in disrupting the differentiation of SACs but not other retinal neurons might be due to cell-type-specific expression or cell-type-specific activity of E2f3. Determining between these two possibilities is not easy, as E2f immunostaining in mouse tissues is problematic. We did not solve this issue for E2f1 or E2f2, but used a modified protocol [50] to successfully track E2f3 expression (Figure 5). At P0, E2f3 was detected in RPCs, consistent with a putative role in normal proliferation (Figure 5A). The signal was specific as it was absent in the E2f3 KO peripheral retina (Figure 5A). As the retina differentiated and RPC division diminished, the number of E2f3+ cells also dropped, and by P8, when division is virtually over, only a subset of post-mitotic cells in the inner retina expressed E2f3 (Figure 5A). By P18, E2f3 was also detected in two tracks in the IPL (Figure 5A and 5B), reminiscent of SAC markers such as Chat and Slc18a3 (c.f. Figures 3 and 4). This cytoplasmic E2f3 staining was also specific, as it was absent in the E2f3 KO peripheral retina of α-Cre;E2f3loxP/loxP mice (Figure 5A). Indeed, double labelling with E2f3 (red) and Chat plus Slc18a3 (green) confirmed that E2f3 is present in both SAC soma and dendrites (Figure 5B). Rb protein was also detected in the inner retina (Figure 5A), and showed a similar distribution as E2f3 in SACs (Figure 5B), and was also present in mature ganglion cells and Müller cells as reported [51]. Rb staining in SAC processes was specific as it was absent in the peripheral retina of αCre;RbloxP/loxP mice (Figure 5A). These data suggest that Rb and E2f3 colocalize in SACs and that E2f3 triggers defects in SAC differentiation because it is specifically expressed in these retinal neurons.

Figure 5

E2f3 and Rb Expression in SACs

(A) Left panels: horizontal P0, P8, and P18 retinal sections of the indicated genotypes were stained for E2f3 (red) and DAPI (blue). The arrow indicates the junction between the E2f3 null peripheral and WT central P0 retina. Note the absence of E2f3 protein in the peripheral E2f3 KO RPCs at P0 and in peripheral inner retinal neurons at P18. Far right panel: P18 retinal sections of the indicated genotypes were stained for Rb (red) and DAPI (blue). Note the absence of Rb protein in the peripheral Rb KO inner retinal neurons.

(B) WT P18 retinal sections were stained for nuclei (DAPI, blue), E2f3 (red) or Rb (red), and Chat plus Slc18a3 (green). Arrows indicate double-labelled soma. Note that the IPL processes are also double-labelled.

Scale bars are 50 μm.

We also found that E2f3 is present in a specific subset of mature neurons in various brain regions (data not shown). For example, in the P20 amygdala, E2f3 colocalized with the general neuronal markers Mtap2 and Mecp2 [52], but not with Calb2, which marks a subset of neurons, or with the glial marker Gfap (data not shown). Unlike in retinal SACs, E2f3 was not coexpressed in Chat+ or Slc18a3+ cholinergic neurons located in various regions of the brain and spinal cord (data not shown). In agreement, we could not detect defects in cholinergic Rb KO neurons in the developing forebrain, but other Rb KO neurons in this region showed differentiation defects that were rescued by deleting E2f3 [53]. Together, these results suggest that the common mechanism by which Rb promotes neural differentiation is through E2f3 inhibition.

Distinct Localization of E2f3 Isoforms

As noted above, E2f3 and Rb staining in SACs was both nuclear and cytoplasmic (Figure 5A and 5B). The antibody that worked in immunostaining recognizes a C-terminal region and thus, does not distinguish a/b isoforms. To our knowledge, the subcellular location of E2f3 isoforms has not been determined in any cell type. To verify the dual locations of E2f3 and to determine which isoforms were present in retina, we analyzed nuclear and cytoplasmic fractions by Western blot at different times during development. Analysis with the pan-E2f3 antibody (sc-878, Santa Cruz Biotechnology) detected a 55-kD E2f3a species and a 40-kD E2f3b polypeptide (Figure 6). To confirm that the upper species in our retinal lysates was E2f3a, we exploited novel mice that lack E2f3 exon 1a and thus express E2f3b exclusively (R. O. and G. L., unpublished data). The genotyping strategy is discussed in detail later and is outlined in Figure 7A. Western analysis confirmed that the upper band was absent in E2f3a−/− mice (Figures 6 and S8). Consistent with the drop in E2f3-expressing cells during WT retinal maturation (Figure 5A), the total amount of E2f3a was less at P18 compared to P0 (Figure 6). E2f3b was present in similar amounts at both time points. At P0 and P18, E2f3a was present in both nuclear and cytoplasmic fractions, but in marked contrast, E2f3b was exclusively nuclear at both times (Figure 6). Two closely migrating E2f3a bands were detected, more clearly evident at P18, of which the faster migrating species was dominant in nuclear and the slower species was dominant in cytoplasm (Figure 6). The identity of both as E2f3a species was confirmed by their absence in the P18 E2f3a KO retina (Figure S8). Analysis of Pou4f2, a nuclear transcription factor expressed in ganglion cells, showed that nuclear proteins had not contaminated the cytoplasmic fraction, and analysis of Slc18a3, a cytoplasmic SAC marker, confirmed that the reverse had also not occurred (Figure 6). These data show, to our knowledge for the first time, that E2f3a and E2f3b exhibit distinct patterns of subcellular distribution, and raise the possibility that E2f3a localization may be regulated by as yet unknown post-translational modifications.

Figure 6

Subcellular Distribution of E2f3 Isoforms and Other Cell Cycle Proteins in the Developing Retina

Nuclear and cytoplasmic extracts from an equivalent number of retinal cells from mice of the indicated genotypes and ages were analyzed by Western blotting to detect the indicated proteins. Lysate from E2f3a−/− mice was used as a control to confirm the location of E2f3a protein. C, cytoplasmic extracts; N, nuclear extracts.

Figure 7

The E2f3a Isoform Drives the Differentiation Defect in Rb KO SACs

(A) Schematic diagrams of the mouse WT, E2f3a−/−, and the Cre-recombined floxed E2f3 loci (indicated here as E2f3−/− for simplicity). E2f3a−/− mice lack most of E2f3 exon 1a and part of intron 1a (red dotted box). Arrows indicate PCR primers. Genotyping of an E2f3a+/− mouse is shown on the right.

(B) RT-PCR detection of E2f3a and E2f3b mRNA in the retina. The sequences of primers are 1aF (5′-GCCTCTACACCACGCCACAAG-3′), 1bF (5′-CGGAAATGCCCTTACAGC-3′), and 4R (5′-CTCAGTCACTTCTTTGGACAG-3′). WT retina expresses both E2f3a and E2f3b mRNA. As expected, E2f3a−/− retina lacks E2f3a mRNA and still expresses E2f3b mRNA. E2f3−/− retina lacks full-length E2f3a and E2f3b mRNAs, and instead expresses a truncated mRNA lacking exon 3.

(C) Real-time RT-PCR analysis of E2f genes in P8 retinas of the indicated genotypes. Error bars represent SD of measurements from three animals, and asterisks indicate a significant difference between WT and the indicated genotypes (*, p <0.05; **; p <0.01; ANOVA and Tukey HSD test).

(D) Rescue of Rb KO SACs by E2f3a deletion. Horizontal retinal sections of the indicated genotypes and ages were stained for nuclei (DAPI, blue), M-phase (PH3, green), and the SAC marker Slc18a3 (red). E2f3a deletion does not suppress ectopic division, but rescues the SAC defect. Scale bars are 50 μm.

M, molecular size marker.

We also examined the distribution of other cell cycle regulators during retinal development. Like E2f3a, Rb was present in both the WT cytoplasm and nucleus at P0, but at P18, when the levels of Rb had increased, it was primarily nuclear (Figure 6). A very faint cytoplasmic Rb signal was evident at P18, which is consistent with Rb staining of SAC processes (Figure 5B), and with the very small proportion of SACs in the retina [38]. E2f1 was also detected in both nuclear and cytoplasmic fractions, although unlike E2f3a it was predominantly nuclear both at P0 and P18 (Figure 6). The E2f dimerization partner, Tfdp1, which lacks a nuclear localization signal [54], was primarily cytoplasmic at both P0 and P18, and the Cdk inhibitors Cdkn1a and Cdkn1b showed a similar pattern of distribution (Figure 6). Thus, among the cell cycle regulators we examined, most showed bivalent distribution, and E2f3b was unusual in its solely nuclear compartmentalization.

Rb Regulates SAC Differentiation through E2f3a

To test which E2f3 isoform is responsible for aberrant Rb KO SAC differentiation we exploited E2f3a−/− mice (Figure 7A). The genotyping strategy outlined in Figure 7A was used to distinguish the E2f3a, WT, and null alleles. Reverse transcriptase PCR (RT-PCR) confirmed the presence of both E2f3a and E2f3b RNA species in the developing WT retina, and the specific absence of E2f3a RNA in the E2f3a−/− retina (Figure 7B). E2f3a protein was absent in E2f3a−/− retinal lysate (Figures 6 and S8). Importantly, the levels of E2f3b message were similar in the Rb KO and Rb/E2f3a DKO retina, ruling out the possibility that any effects of E2f3a deletion we might observe were due to down-regulation of E2f3b (Figure 7C). Also, the levels of other E2fs were the same in the Rb KO, Rb/E2f3 DKO, and Rb/E2f3a DKO retina, ruling out any cross-regulatory effects (Figure 7C) [55]. E2f3a can trigger cell cycle induction, but because SAC defects are not linked to cell cycle perturbation (Figures 3A and 4), and in view of the predominant association between E2f3b and Rb in quiescent cells [16,19], we suspected that E2f3b may perturb differentiation in Rb KO SACs. Unexpectedly, however, E2f3a deletion suppressed the Rb KO SAC defect (Figure 7D). Thus, separate from its role in cell cycle control, Rb regulation of E2f3a is critical to ensure proper neuronal differentiation.

Discussion

Rb Controls Retinal Cell Division and Death through E2f1

Work in the early 1990s showed that Rb loss triggers defects in neuronal cell cycle exit, survival, and differentiation [26–28]. Much of the death is an indirect consequence of probable hypoxia linked to placental defects [12–14]. However, targeted KO and chimeric studies reveal that Rb autonomously promotes cell cycle exit in newborn neurons, and is required for survival of a subset of neurons, particularly in the retina [2,3,13,14,56–59]. However, whether Rb also regulates differentiation is obscured by potentially indirect effects of ectopic division and death. Moreover, a mechanism though which Rb may regulate neuronal maturation has not been elucidated.

Here, deleting E2f1 specifically rescued ectopic division and death in the Rb KO retina. Importantly, major Rb/E2f1 DKO neurons differentiated normally, and ERGs revealed the rescue of rod- and cone-mediated function, implicating a regular signal flow from photoreceptors to bipolar and amacrine cells. Division and death genes were induced in Rb KO cells, and deleting E2f1, but not E2f2 or E2f3, reversed these molecular events. E2f1 may also regulate differentiation targets, but whether this contributes to defects in retinal cell maturation is impossible to separate from potentially indirect consequences of deregulated division and death. In any case, it is clear that in most retinal cells, including photoreceptors [29], transcription factors that promote differentiation function independently of Rb.

We have also found that E2f1 deletion rescues cell-autonomous ectopic division, death, and differentiation defects in sporadic Rb KO clones generated using a Cre retrovirus vector (M. P. and R. B., unpublished data). These data are consistent with the observation that E2f1 overexpression in newborn photoreceptors drives ectopic division and apoptosis [60], and add to the growing evidence indicating that E2f1 is the major, and perhaps only, member of the three activating E2fs required to induce apoptosis in Rb KO cells [10,15]. Thus, deregulated E2f1 activity in the retina, whether resulting from the inactivation of Rb or from overexpression, promotes unscheduled cell division and triggers apoptosis in susceptible RTCs. E2f1, rather than other E2fs, may be a potential target for novel therapeutics to prevent retinoblastoma in RB1+/− humans.

Our ERG studies revealed rescue of the Rb KO rod–bipolar system, and almost complete restoration of the cone–bipolar system following E2f1 deletion. There was a slightly lower response in the Rb/E2f1 DKO retina relative to the E2f1 KO control retina. This difference might reflect a role for Rb in the development of cones, bipolar cells, or other cells that may contribute to the photopic ERG, including potentially SACs, which do have a serious defect in the Rb/E2f1 DKO retina.

Rb Controls SAC Differentiation through E2f3a

Comprehensive marker analysis revealed that, in striking contrast to other retinal neurons, E2f1 deletion did not suppress defects in Rb KO cholinergic SACs. Instead, we observed E2f1-independent defects in the synthesis and transport of a large cohort of SAC proteins. These data expand insight into the development of these important interneurons, but more critically, provide to our knowledge the first unambiguous evidence that Rb regulates neurogenesis beyond terminal mitosis. Rb binds more than 100 factors [43], and in several non-neuronal cells, such as skeletal muscle, adipocytes, and bone, it binds and potentiates tissue-specific transcription factors that promote differentiation [31–33]. The idea that Rb promotes muscle differentiation by potentiating Myod1 activity was contested [61], and other mechanisms proposed [62,63], but not involving E2f repression. Strikingly, however, we discovered that Rb promotes SAC differentiation through E2f3 (Figure 8).

Figure 8

Rb Regulates Distinct Processes through E2f1 and E2f3a

Red text and arrows indicate Rb-dependent events. Black text and arrows indicate events for which there is no direct evidence of Rb involvement. Rb does not appear to temper RPC expansion and is not required for differentiation of RPCs into RTCs, but is essential to couple RTC birth to terminal mitosis, thus locking them out of cycle. Rb performs this function by inhibiting E2f1. Rb is also required for SAC differentiation, and in this case, acts by inhibiting E2f3a. There is no direct evidence that Rb is required for terminal differentiation of other cell types. Colour codes and abbreviations as in Figure 1A.

Rb regulation of SAC differentiation through E2f3 was independent of its role in controlling division or death: E2f3 deletion rescued Rb KO SAC defects but did not suppress aberrant proliferation or death, whereas E2f1 deletion reversed abnormal proliferation and death but did not rescue SAC differentiation. Double labelling confirmed that E2f1 but not E2f3 deletion reversed Rb KO SAC division. Moreover, deleting E2f1, but not E2f3, reversed deregulated expression of cell cycle and apoptotic genes in the Rb KO retina. E2f3 is expressed in a subset of CNS neurons (this work) and drives specific cell-cycle–independent defects in Rb KO forebrain neurons [53]. Thus, E2f3 inhibition is the first, and may be the only, mechanism by which Rb participates directly in neuronal differentiation.

To further dissect the mechanism of action of Rb in SACs we determined the E2f3 isoform it targets to promote differentiation. E2f3b was the primary candidate, since Rb and E2f3b collaborate to repress targets in quiescent cells in vitro [19]. However, in the first work to our knowledge to examine the function of any E2f protein isoform in vivo, we made the surprising observation that Rb regulates SAC differentiation through E2f3a (Figure 8). Formally, we cannot exclude the possibility that deleting E2f3b might also rescue SAC differentiation, but definitive proof will require analysis of E2f3b null mice. Nevertheless, our data prove that Rb definitely regulates SAC differentiation through the activating E2f3 isoform.

Distinct E2f3a and E2f3b Localization

The subcellular location of E2f isoforms has not to our knowledge been addressed before. E2f3a and E2f3b share 110 C-terminal amino acids that encode the NLS, DNA-binding, marked box, transactivation, and Rb-binding domains [16], yet they exhibit different subcellular distribution in developing retinal cells. E2f3a is both nuclear and cytoplasmic, but E2f3b is always nuclear. The unique 121- and six-residue N-termini of E2f3a and E2f3b, respectively, likely mediate this difference. This region in E2f1, E2f2, and E2f3a binds Ccna2, establishing a negative regulatory loop that deactivates E2fs in mid-late S-phase [64,65]. However, even E2f3b, which lacks this domain, binds and is regulated by Ccna2 [18], so the domain difference may not explain the unique distributions we observed. Rb family and Tfdp proteins can also determine E2f localization [20–22], and we found that a portion of both Rb and Tfdp1 proteins are cytoplasmic in retinal cells. Indeed, immunostaining revealed that Rb and E2f3 colocalize to SAC processes.

The nuclear localization of E2f3b contrasts with that of other repressive E2fs in differentiating muscle, where E2f5 switches from the nucleus to cytoplasm, while E2f4 remains in both compartments [23]. The distinct compartmentalization of E2f3a and E2f3b in the retina suggests temporally and functionally distinct activities. Rb distribution matches that of E2f3a, consistent with its critical role in supporting SAC differentiation through E2f3a.

Ectopic Division and Differentiation

Rb is critical to ensure that many types of terminally differentiating cells leave the cell cycle (e.g., neurons, gut and skin epithelia, muscle, and lens fibres) (reviewed in [66]). Early overexpression studies in vitro suggested Rb might temper expansion of cycling cells, but KO studies in vivo indicate that its major role is to block division in terminally differentiating cells. In its absence, many (but clearly not all) aspects of differentiation go ahead relatively unperturbed. In the retina, differentiating transition cells are born in the absence of Rb, migrate to the correct layer, and express appropriate markers ([2] and this work). In brain, Rb KO neurons migrate away from the ventricular zone and switch on Tubb3 (βIII-tubulin), but continue to incorporate BrdU [13], and in gut epithelia, differentiated enterocytes migrate up the villi and activate expression of serotonin, yet continue to incorporate BrdU [67]. In the case of SACs, the differentiation defects we observed (e.g., loss of Slc18a3 and Chat) were not due to aberrant division, but it is possible there are other problems with these cells that are caused by ectopic division. Nevertheless, it is clear that many aspects of differentiation in multiple cell types are compatible with ectopic division. However, division of terminally differentiating cells is dangerous, since it may facilitate transformation, as is the case in retinoblastoma (reviewed in [66]).

How Does E2f3a Perturb SAC Differentiation?

E2f3a could disrupt SAC differentiation through its well known role as a transcriptional activator, or, in view of the discovery that it is partially cytoplasmic, E2f3a may affect processes other than gene regulation. Both scenarios are feasible since E2fs regulate differentiation genes [44–48], and cell cycle regulators, such as Cdkn1b, have cytoplasmic activities that influence differentiation [68,69]. Many transcription factors shuttle between nucleus and cytoplasm during neurogenesis (e.g., [70] and references therein). It may be difficult to identify E2f3a-specific target genes or cytoplasmic proteins in SACs since these neurons are a small proportion (<1%) of the total retina and only ~5.2% of amacrine neurons [38].

Do E2fs Mediate All Rb Functions?

Others have suggested that Rb promotes differentiation in non-neuronal cells through E2f-independent means [31–33]. However, these studies did not assess whether these cell types differentiate normally if Rb is deleted along with one or more E2f family members. One study reported that Rb mutants that do not bind E2f still induce differentiation [30]. However, the binding assays were performed in solution, and we have found that several of these mutants do bind E2f, albeit weakly, on chromatin (T. Yu and R. B., unpublished data). It is possible that Rb-mediated potentiation of tissue-specific transcription factors may, at least in some cases, be a redundant activity, and that the only critical Rb function is to inhibit E2f. Our study is the first to our knowledge to assess comprehensively whether Rb KO cells can differentiate in the absence of different E2fs. In light of our findings, it will be important to reassess differentiation defects in other Rb KO tissues in the absence of individual and combined activating E2f family members.

Materials and Methods

Mouse strains and genotyping.

Mice were treated according to institutional and national guidelines. α-Cre mice (P. Gruss), Chx10-Cre mice (C. Cepko), RbloxP/loxP mice (A. Berns), E2f1–/– mice, E2f2–/– mice, E2f3loxP/loxP mice, and E2f3a−/− mice were maintained on a mixed (NMRI × C57/Bl × FVB/N × 129sv) background. A detailed description of E2f3a−/− mice will be published elsewhere. Mice of different genotypes were compared within the same litter and across a minimum of three litters. We have not noted any phenotypic differences in separate litters. Genotyping was performed as before [2,5], and the primers used for genotyping E2f3a−/− mice were E2f3a KL (5′-CTCCAGACCCCCGATTATTT-3′), E2f3a KR1 (5′-TCCAGTGCACTACTCCCTCC-3′), and E2f3a KM (5′-GCTAGCAGTGCCCTTTTGTC-3′).

Histology, immunofluorescence, and measurements.

Eyeballs were fixed in 4% paraformaldehyde for 1 h at 4 °C, embedded in OCT (TissueTek 4583, Sakura, http://www.sakuraeu.com), frozen on dry ice, and cut into 12-μm sections on Superfrost plus slides (VWR, http://www.vwr.com). For S-phase analysis, BrdU (100 μg/g of body weight) was injected intraperitoneally 2 h prior to sacrifice. BrdU+ cells were detected using a biotin-conjugated sheep polyclonal antibody (1:500, Maine Biotechnology Services, http://www.mainebiotechnology.com). All other antibodies are described in Table S1. For E2f3, Mki67, and Rb staining, antigen retrieval was performed by boiling sections in citric acid solution for 15 min according to Ino [50], except on frozen sections. TUNEL was performed as described [13]. Briefly, sections were incubated for 1 h at 37 °C with 75 μl of mixture solution consisting of 0.5 μl of terminal deoxynucleotide transferase, 1 μl of biotin-16-dUTP, 7.5 μl of CoCl2, 15 μl of 5× terminal deoxynucleotide transferase buffer, and 51 μl of distilled water. After three washes in 4× SSC buffer, sections were incubated with Alexa 488– or Alexa 568−streptavidin (1:1,000; Molecular Probes, http://probes.invitrogen.com) for 1 h at room temperature. Primary antibodies or labelled cells were visualized using donkey anti-mouse Alexa 488 or Alexa 568, donkey anti-rabbit Alexa 488 or Alexa 568, donkey anti-goat Alexa 488 or Alexa 568, and streptavidin Alexa 488 or Alexa 568 (1:1,000; Molecular Probes). Nuclei were counter-stained with 4,6-diamidino-2-phenyindole (DAPI; Sigma, http://www.sigmaaldrich.com). Labelled cells were visualized using a Zeiss (http://www.zeiss.com) Axioplan-2 microscope with Plan Neofluar objectives and images captured with a Zeiss AxionCam camera. For double-labelled samples, confocal images were obtained with a Zeiss LSM 5.0 laser scanning microscope.

The retina was separated into three bins by dividing the ventricular edge of the retina into equal parts and extending a line to the vitreal edge [2]. Bin 1 contains only cells that expressed Cre as progenitors; bin 3 is at central retina and contains cells derived from progenitors that did not express Cre. For cell counts or thickness measurement we used a region 0–100 μm peripheral to the boundary separating bins 1 and 2. Measurements were performed on an Axioplan-2 microscope using Axiovison software. Quantification of S-phase, M-phase, and apoptotic cells was performed on horizontal sections that included the optic nerve. Quantification of differentiated cell types was performed using horizontal sections at equal distances from the optic nerve. A minimum of three sections per eye and three eyes from different litters were counted.

RNA extraction, reverse transcription, and PCR.

Total RNA was isolated from dissected peripheral retina using TRIzol reagent (Invitrogen, http://www.invitrogen.com) followed by digestion with RNase-free DNase (DNA-free, Ambion, http://www.ambion.com) to remove DNA contamination. First-strand cDNA was synthesized from 0.2–0.5 μg of total RNA using the SuperScript II first-strand synthesis system (Invitrogen). PCR primers are listed in Table S2. Real-time quantitative PCR was performed using an Applied Biosystems (http://appliedbiosystems.com) PRISM 7900HT. Tests were run in duplicate on three separate biological samples with SYBR Green PCR Master Mix (Applied Biosystems) exactly as we described previously [71]. Briefly, master stocks were prepared such that each 10-μl reaction contained 5 μl of SYBR Green PCR Master Mix, 0.1 μl of each forward and reverse primer (stock 50 μM), 0.8 μl of blue H2O (0.73% Blue Food Colour; McCormick, http://www.mccormick.com), 2 μl of diluted cDNA template, and 2 μl of yellow H2O (0.73% Yellow Food Colour). PCR consisted of 40 cycles of denaturation at 95 °C for 15 s and annealing and extension at 55 °C for 30 s. An additional cycle (95 °C, 15 s, 60 °C) generated a dissociation curve to confirm a single product. The cycle quantity required to reach a threshold in the linear range was determined and compared to a standard curve for each primer set generated by five 3-fold dilutions of genomic DNA or cDNA samples of known concentration. Values obtained for test RNAs were normalized to Hprt1 mRNA levels.

Western blots.

Mouse retinas were homogenized by passing them through a 30-gauge BD 9 http://www.bd.com) needle 5–10 times in 1× PBS solution. Nuclear and cytoplasmic proteins were extracted using the NE-PER Nuclear and Cytoplasmic Extraction Kit (Product# 78833, Pierce Biotechnology, http://www.piercenet.com). Proteins were separated by 10% SDS-PAGE and transferred to nitrocellulose. After blocking overnight at 4 °C in 5% skim milk, membranes were incubated in the primary antibody for 2 h at room temperature. After three 10-min washes in TPBS (100 mM Na2HPO4, 100 mM NaH2PO4, 0.5 N NaCl, 0.1% Tween-20), membranes were incubated for 30 min at room temperature in the secondary horseradish peroxidase-conjugated antibody (Jackson ImmunoResearch Laboratories, http://www.jacksonimmuno.com). Blots were developed using the ECL-Plus chemiluminescent detection system (Amersham Pharmacia Biotech, http://www.pharmacia.ca), according to the manufacturer's instructions.

The following primary antibodies were used: E2f-1 (SC-193), E2f-3 (SC-878), Cdkn1a (p21, SC-471), Cdkn1b (p27, SC-528), Pou4f2 (Brn3b, SC-6062), and Tfdp1 (Dp1, SC-610) from Santa Cruz Biotechnology (http://www.scbt.com), pRB (554136) from BD Science-Pharmingen (http://www.bdbiosciences.com), and Slc18a3 (VAChT, G448A) from Promega (http://www.promega.com).

Electroretinography.

ERGs were recorded from dark-adapted mice as described [72]. Briefly, mice were dark-adapted overnight and anaesthetized by subcutaneous injection of ketamine (66.7 mg/kg body weight) and xylazine (11.7 mg/kg body weight). The pupils were dilated and single-flash ERG recordings were obtained under dark-adapted (scotopic) and light-adapted (photopic) conditions. Light adaptation was accomplished with a background illumination of 30 candela (cd) per square meter starting 10 min before recording. Single white-flash stimulation ranged from 10−4 to 25 cd·s/m2, divided into ten steps of 0.5 and 1 log cd·s/m2. Ten responses were averaged with an inter-stimulus interval of either 5 s (for 10−4, 10−3, 10−2, 3 × 10−2, 10−1, and 3 × 10−1 cd·s/m2) or 17 s (for 1, 3, 10, and 25 cd·s/m2). Band-pass filter cut-off frequencies were 0.1 and 3,000 Hz.

Statistics.

Different genotypes were evaluated using analysis of variance (ANOVA) followed by the Tukey honestly significant difference (HSD) test or Fisher test (XLSTAT program, http://www.xlstat.com).

Supporting Information

Figure S1

Deleting E2f1, but Not E2f2 or E2f3, Rescues Ectopic Division and Cell Death in P0 and P18 Rb KO Retina

Horizontal sections of the indicated genotypes and ages were stained for nuclei (DAPI, blue), and (A) S-phase (anti-BrdU, red) or (B) apoptosis (TUNEL, red). In Rb−/− retinas, BrdU+ cells extend beyond the normal boundaries at P0 (arrows), and ectopic DNA synthesis continues in multiple layers at later stages. Scale bar is 50 μm. The NBL is where dividing RPCs are located.

(815 KB PDF)

Click here for additional data file.

Figure S2

Deleting E2f1 Rescues Ectopic Division and Apoptosis in the Embryonic Rb KO Retina

Horizontal sections of the indicated genotypes and ages (E14 and E16, the period during which SACs are born) were stained for nuclei (DAPI, blue), and either S-phase (upper two panels, anti-BrdU, red) or apoptosis (lower two panels, TUNEL, red). In Rb−/− retinas, BrdU+ and TUNEL+ cells can be seen in the inner retina (arrows). Inactivation of E2f1 rescued these defects. Scale bar is 50 μm. The NBL is where dividing RPCs are located.

(754 KB PDF)

Click here for additional data file.

Figure S3

Deleting E2f1, but Not E2f2 or E2f3, Rescues Ectopic Mitosis in the Rb KO Retina

(A) Horizontal retinal sections of the indicated genotypes and ages were stained for nuclei (DAPI, blue) and M-phase (anti-PH3, red). Scale bar is 50 μm.

(B) Quantification of all PH3+ cells.

(C) Quantification of ectopic PH3+ cells.

Error bars represent standard deviation (SD), and asterisks indicate significant difference between retina of WT and indicated genotypes (*, p <0.05; **, p <0.01; ANOVA and Tukey HSD test).

(628 KB PDF)

Click here for additional data file.

Figure S4

Deleting E2f2 or E2f3 Does Not Rescue Ganglion, Rod, or Bipolar Cell Death in the Rb KO Retina

(A) Horizontal retinal sections from mice of the indicated ages and genotypes were stained for nuclei (DAPI, blue) and markers that detect ganglion cells (Pou4f2, red), rods and cones (Sag [rod arrestin], green), and rod bipolar cells (Prkca, green). Scale bar is 50 μm.

(B) Quantification of total ganglion (Pou4f2+) cells.

(C) Quantification of total rod bipolar (Prkca+) cells.

(D) Thickness of the ONL, which represents the number of rods.

Error bars represent SD, and asterisks indicate significant difference between retina of WT and indicated genotypes (**, p <0.01; ANOVA and Tukey HSD test).

(488 KB PDF)

Click here for additional data file.

Figure S5

E2f1 Deletion Rescues α-Cre;RbloxP/loxP Retinal Function

ERGs were recorded from the indicated genotypes under light adapted (photopic) conditions.

(A) Intensity series.

(B) The b-wave amplitudes as a function of the logarithm of the flash intensity.

(383 KB PDF)

Click here for additional data file.

Figure S6

Differentiation Defects in Rb KO SACs

Horizontal retinal sections of indicated genotypes and ages were stained for nuclei (DAPI, blue) and Calb2 ([A], red; only densely stained cells were counted for Figure 3C), Camk2a ([B], green), and Slc18a3 ([C], red). Scale bars are 50 μm.

(564 KB PDF)

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Figure S7

GABA Neurotransmitter in the Rb KO Retina and Abnormal SACs in Chx10-Cre;RbloxP/loxP Retina

Horizontal sections of the indicated genotypes and ages of retina were stained for nuclei (DAPI, blue), and (A and B) GABA (red) and Slc18a3 (green) or (C) Chat and Slc18a3 (red).

(A) In P18 WT retina, GABA labelled four IPL tracks, of which the two inner tracks co-stained with Slc18a3. The latter tracks disappeared in the Rb KO retina, and were rescued by E2f3 KO but not E2f1 KO.

(B) At the boundary of the WT (central) and Rb KO area (peripheral retina) the inner GABA+ SAC tracks can be seen disappearing towards the periphery (left).

(C) Slc18a3 staining in the IPL of Chx10-Cre;RbloxP/loxP retina is consistent with the mosaic pattern of Rb inactivation.

Scale bars are 50 μm.

(633 KB PDF)

Click here for additional data file.

Figure S8

Subcellular Distribution of E2f3a Isoform in the Developing Retina

Nuclear and cytoplasmic extracts from an equivalent number of retinal cells from mice of the indicated genotypes and ages were analyzed by Western blotting to detect the E2f3a protein. Lysates from E2f3a−/− mice of matched ages were used as a control to confirm the location of E2f3a protein. C, cytoplasmic extracts; N, nuclear extracts.

(115 KB PDF)

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Table S1

List of Antibodies and Marker Patterns in Rb/E2f1 DKO SACs

(97 KB DOC)

Click here for additional data file.

Table S2

Real-Time RT-PCR Primers

(49 KB DOC)

Click here for additional data file.

Accession Numbers

The GenBank (http://www.ncbi.nlm.nih.gov/genbank) accession numbers for the major genes and gene products discussed in this paper are Camk2a (NM_009792), Chat (NM_009891), E2f1 (NM_007891), E2f2 (NM_177733), E2f3 (NM_010093), Rb (NM_009029), and Slc18a3 (NM_021712).

Acknowledgements

We thank K. McClellan and R. Slack for sharing unpublished data, L. Galli-Resta for suggesting Isl1 (Islet1) as an early SAC marker, J. Eubanks for advice on immunostaining, A. Berns, C. Cepko, and P. Gruss for mice, and T. Edlund, F. Haeseleer, R. Janz, S. Sugita, P. A. Hargrave, C. M. Craft, X. Zhu, R. McInnes, R. L. Chow, and J. Saari for antibodies.

Abbreviations

ANOVA - analysis of variance

BrdU - bromodeoxyuridine

cd - candela

CNS - central nervous system

DAPI - 4,6-diamidino-2-phenyindole

DKO - double knockout

E[number] - embryonic day [number]

E2f - E2f transcription factor

ERG - electroretinogram

HSD - honestly significant difference

GABA - gamma-aminobutyric acid

GCL - ganglion cell layer

INL - inner nuclear layer

IPL - inner plexiform layer

KO - knockout

NBL - neuroblastic layer

ONL - outer nuclear layer

OPL - outer plexiform layer

P[number] - postnatal day [number]

PH3 - phosphohistone 3

Rb - retinoblastoma protein

RPC - retinal progenitor cell

RTC - retinal transition cell

RT-PCR - reverse transcriptase PCR

SAC - starburst amacrine cell

SD - standard deviation

TUNEL - terminal dUTP nick-end labelling

WT - wild-type

Footnotes

Author contributions. DC and RB conceived and designed the experiments. DC performed most of the experiments. RO, PW, and GL provided the E2f3 floxed and E2f3a KO mice. DC and RB analyzed the data. MP, NT, and MWS performed the ERG analysis. DC and RB wrote the paper.

Funding. This work was supported by grants from the Canadian Institute for Health Research and Foundation Fighting Blindness Canada to RB, and by the German Research Council (DFG Se837/4–1 and 5–1) and the European Union (IP “EVI-GenoRet” LSHG-CT-512036) to MWS.

Competing interests. The authors have declared that no competing interests exist.


Mouse Pachytene Checkpoint 2 (Trip13) Is Required for Completing Meiotic Recombination but Not Synapsis

Abstract

In mammalian meiosis, homologous chromosome synapsis is coupled with recombination. As in most eukaryotes, mammalian meiocytes have checkpoints that monitor the fidelity of these processes. We report that the mouse ortholog (Trip13) of pachytene checkpoint 2 (PCH2), an essential component of the synapsis checkpoint in Saccharomyces cerevisiae and Caenorhabditis elegans, is required for completion of meiosis in both sexes. TRIP13-deficient mice exhibit spermatocyte death in pachynema and loss of oocytes around birth. The chromosomes of mutant spermatocytes synapse fully, yet retain several markers of recombination intermediates, including RAD51, BLM, and RPA. These chromosomes also exhibited the chiasmata markers MLH1 and MLH3, and okadaic acid treatment of mutant spermatocytes caused progression to metaphase I with bivalent chromosomes. Double mutant analysis demonstrated that the recombination and synapsis genes Spo11, Mei1, Rec8, and Dmc1 are all epistatic to Trip13, suggesting that TRIP13 does not have meiotic checkpoint function in mice. Our data indicate that TRIP13 is required after strand invasion for completing a subset of recombination events, but possibly not those destined to be crossovers. To our knowledge, this is the first model to separate recombination defects from asynapsis in mammalian meiosis, and provides the first evidence that unrepaired DNA damage alone can trigger the pachytene checkpoint response in mice.

Author Summary



It is critical that the chromosomes carried by sperm and eggs contain faithful representations of the genome of the individual that produced them. During the process of meiosis, the maternal and paternal copies of each chromosome “synapse” with each other (become tightly associated), exchange genetic material via the process of recombination, then separate into daughter cells in the first of two meiotic cell divisions. The intricate chromosome behavior is subject to errors, so most organisms have evolved meiotic “checkpoints” that monitor fidelity of chromosome synapsis and repair of DNA damage. These checkpoints cause defective cells to self destruct rather than generate defective sperm or eggs. We studied the effects of deleting mouse Trip13, a gene that in distant organisms plays a key role in meiotic checkpoint control. These experiments revealed that instead of having a checkpoint role, Trip13 is required for one of the two major classes of recombination in meiosis that is required for repairing broken DNA molecules. The chromosomes still synapsed normally, but animals were sterile due to massive death of oocytes and spermatocytes. These results indicate that, in addition to a checkpoint that responds to failed synapsis, one exists to specifically detect unrepaired DNA damage that is due to failed recombination.

Introduction

The genesis of gametes containing an intact, haploid genome is critical for the prevention of birth defects, and is highly dependent upon the fidelity of chromosome dynamics before the first meiotic division. Homologous chromosomes must pair, synapse, undergo recombination, and segregate properly to opposite poles. Recombination, which repairs repair double strand breaks (DSBs) that are genetically induced in leptonema, is coupled with synapsis in budding yeast and mammals. While our knowledge of the assembly and nature of recombination machinery is extensive, little is known about the disassembly of recombination intermediates, recruitment of DNA replication machinery during recombinational repair, and how the choice between different repair pathways is made.

Defects in recombination can preclude homologous chromosome pairing, leave unrepaired chromosome breaks, and cause aneuploidy by abrogating crossing over. To avoid such deleterious outcomes, surveillance systems (“checkpoints”) exist to sense meiotic errors and eliminate cells containing unresolved defects. In many organisms, including S. cerevisiae, Drosophila melanogaster, C. elegans, and mice [1–4], meiocytes with defects in recombination and/or chromosome synapsis trigger meiotic arrest in the pachytene stage of meiotic prophase I. This response to meiotic defects is referred to as the “pachytene checkpoint” (reviewed in [5]). Genetic experiments in S. cerevisiae have identified elements of the pachytene checkpoint machinery (reviewed in [5]). In addition to meiosis-specific proteins, these include factors that play roles in DNA damage signaling in mitotic cells [6–10]. Arabidopsis thaliana does not appear to have a pachytene checkpoint akin to that in yeast [11], nor do male Drosophila.

The pachytene checkpoint is known to monitor two aspects of meiotic chromosome metabolism in S. cerevisiae and C. elegans: (1) DSB repair and (2) chromosome synapsis [2,12]. In mice, both spermatocytes and oocytes harboring mutations that disrupt DSB repair (such as Dmc1, Msh5, and Atm) are efficiently eliminated in pachynema, but spermatocytes are much more sensitive to DSB repair–independent synapsis defects than oocytes [13–15]. However, because recombination is required for synapsis in mice (mutations in recombination genes such as Dmc1 cause extensive asynapsis [16]), it has remained formally uncertain whether there is a distinct pachytene checkpoint that responds to defects in meiotic recombination, and if so, whether it would be identical to that used in somatic cells. The mechanisms of putative pachytene checkpoint control remain unknown in mammals, since no mutations have been identified that abolish it.

PCH2, encoding a nucleolar-localized AAA-ATPase that was originally identified in an S. cerevisiae genetic screen for mutants that relieve pachytene arrest of asynaptic zip1 mutants [8], was recently determined to be an essential component of the pachytene synapsis (but not DSB repair) checkpoint in yeast and worms [2,12]. PCH2 orthologs are present in organisms that undergo synaptic meiosis, but not asynaptic meiosis, prompting the suggestion that a Pch2-dependent checkpoint evolved to monitor synaptonemal complex (SC) defects from yeast to humans [12]. Here, we generated mice deficient for the Trip13, the ortholog of PCH2, and evaluated whether it also plays a role in the pachytene checkpoint. Surprisingly, while we found no evidence for checkpoint function, we did uncover a potential role for this protein in noncrossover (NCO) repair of meiotic DSBs.

Results

Trip13 Is a Widely Expressed Mammalian Ortholog of PCH2 with Unusual Phylogenetic Relationships

The mammalian ortholog of PCH2, Trip13 (thyroid hormone receptor interacting protein 13), encodes a protein with extensive amino acid homology in regions alignable to the yeast and worm orthologs (Figure S1) [12]). Interestingly, phylogenetic analysis of TRIP13/Pch2p shows that the mammalian protein clusters more closely to plants than it does to the evolutionarily more closely related worms and flies (Figure 1A; see Discussion). Semi-quantitative reverse-transcriptase PCR (RT-PCR) analysis showed Trip13 mRNA to be expressed in a variety of embryonic and adult tissues, including testis (Figure 1B), consistent with mouse and human EST data summarized in Unigene (http://www.ncbi.nlm.nih.gov/UniGene). It is also highly expressed in human and mouse oocytes [17].

Figure 1

The Mouse PCH2 Ortholog TRIP13 and Expression in Wild Type and Mutant

(A) Phylogenetic tree of presumed PCH2/TRIP13 orthologs. The database sequence accession number of each protein is presented in Table S1. Numbers shown are bootstrap values (see Materials and Methods). The clustering was the same regardless of whether the whole entire AA sequence or trimmed sequences (where regions showing little conservation were removed) were used for the analysis. Major eukaryotic groups are indicated in color, with deuterostomia in blue, plants in green, protostomia in purple, and fungi in maroon.

(B) Amplification products of cDNA from the following tissues: 1, heart; 2, brain; 3, spleen; 4, lung; 5, liver; 6, skeletal muscle; 7, kidney; 8, testis; 9, E7 embryo; 10, E11 embryo; 11, E15 embryo; and 12, E17 embryo.

(C) Intron–exon structure of TRIP13 and insertion site of gene-trap vector. See Materials and Methods for details on how the precise insertion site was identified.

(D) RT-PCR of Trip13 and a control gene Med31 from testis RNA. The Trip13 primers are situated in the first and last exons (see Materials and Methods).

(E) Western blot analysis of testis protein with anti-TRIP13 antibody. The blot was later probed with anti-alpha tubulin actin as a loading control. The expected TRIP13 protein is ∼48 KDa.

(F) Localization of TRIP13 in testes. Wild-type (top) and mutant (bottom) testis sections were probed with chicken anti-TRIP13, and detected with HRP-conjugated anti-chicken IgG (brown/red staining). Expression in WT was most prominent in the nuclei of Type B spermatogonia (Sg), leptotene spermatocytes (LS), and early pachytene spermatocytes (PS), but not late pachytene spermatocytes (LP). No nuclear staining was seen in mutant testis sections, although reddish cytoplasmic background is present. Identification of cell types was judged in part by estimating the epithelial stage of the tubules as described [67].

(G) TRIP13 localization in surface-spread spermatocytes. Preparations were immunolabeled with anti-SYCP3 (S) and TRIP13 (T). Both individual and merged images are shown for leptotene (Lep), zygotene (Zyg), and pachytene (Pac) spermatocytes. Nuclear staining was absent in the mutant.

Generation of Trip13 Mutant Mice

To explore the function of TRIP13 in mammals, we generated mice with a gene trap-disrupted allele, Trip13RRB047 (Figure 1C; abbreviated as Trip13Gt). Heterozygotes were normal in all respects, but homozygotes were present at ∼2/3 the expected ratio from intercrosses between heterozygotes (91 Trip13+/+, 183 Trip13Gt/+, and 61 Trip13Gt/Gt). Since >90% of prewean mice that died were mutant homozygotes, this discrepancy is apparently due to a partially penetrant lethality. Most surviving Trip13Gt/Gt animals were grossly normal. However, homozygotes that were semi-congenic (N4) on the C57BL/6J strain were often markedly smaller and/or had kinked or shorter tails (Figure 2A and 2B).

Figure 2

Developmental Phenotypes of Trip13 Mutant Mice

(A) Shown are 21-d-old littermates. Note the shortened tail in the mutant, but overall similar body size.

(B) Shown are 23-d-old littermates. The mutant is smaller in this case, but the tail is not as truncated as the mouse in (A).

(C) Wild-type (WT) and homozygous Trip13 mutant (MUT) testes.

(D) and (E) are cross sections through 17.5-d-old heterozygous (“WT”) and homozygous mutant Trip13 testes, respectively. Whereas the tubules in WT show coordinated spermatogenesis with pachytene spermatocytes present in all tubules (proximal to the lumen), developmental progression in the mutant is not synchronized between tubules. Some tubules have no pachytene spermatocyes (asterisks), while in others, development is somewhat disorganized (#).

RT-PCR analysis of Trip13Gt expression (Figure 1D) revealed a low level of normally spliced transcripts in testes of homozygotes that is presumably a consequence of incomplete usage of the gene trap's splice acceptor. Western blot analysis, using a polyclonal antibody raised against a peptide encoded by exon 3, revealed multiple species in wild-type and heterozygous testes, one of which corresponds to the expected size of 48 kDa (Figure 1E). This and three other species were undetectable in homozygous mutant testes, but a reduced amount of an intense ∼38 kDa smaller band was present. It is not clear if this corresponds to TRIP13. The greatly decreased Trip13 mRNA and predicted correct-length protein in mutants indicate that the Trip13RRB047 allele is severely hypomorphic.

To determine the germ cell types in which TRIP13 is expressed, and to assess possible expression in the mutant by means other than Western analysis, testis sections were immunolabeled for TRIP13 using a polyclonal chicken antipeptide antibody (see Materials and Methods). The most intensely labeled cells in control testes were Type B spermatogonia and leptotene spermatocytes (Figure 1F). Zygotene/pachytene spermatocytes stained less strongly, and there was no detectable staining in late pachytene spermatocytes. TRIP13 appeared to be nuclear localized. There was no such staining of nuclei in mutant seminiferous tubules (Figure 1F). To further assess the nuclear localization, TRIP13 was used to probe meiotic chromosomes prepared by surface spreading of spermatocyte nuclei. In wild type, there was diffuse nuclear staining, and no evidence of concentration on SC cores (marked by the axial element protein SYCP3) at any meiotic substage (Figure 1G). TRIP13 signal was noticeably absent in mutant meiotic nuclei.

Infertility Due to Meiotic Disruption in TRIP13-Deficient Meiocytes

Homozygotes of both sexes had small gonads (Figure 2C; see below) and were invariably sterile. Ovaries of adult Trip13Gt/Gt females were severely dysmorphic and had few or no follicles (Figure 3A and 3B). The majority of oocyte loss occurred in late embryogenesis or early in postnatal development, since 2 d postpartum ovaries were markedly smaller than those of control littermates, and were lacking oocytes or newly forming follicles (Figure 3C and 3D). Thus, oocytes failed to progress to the dictyate (resting) phase. Since we observed oocytes with pachytene stage chromosomes in 17.5 d Trip13Gt/Gt embryonic ovaries (unpublished data), this indicates that oocytes were eliminated somewhere between pachynema and dictyate.

Figure 3

Histology of Mutant Gonads

All are hematoxylin/eosin-stained paraffin sections. Testes are from 6-wk-old males, except as indicated below.

(A) Wild-type 25-d-old ovary.

(B) Trip13Gt/Gt 25-d-old ovary, showing dysgenesis from an absence of oocytes.

(C) Trip13Gt/+ 2-d-old control ovary. Arrows point to oocytes in newly forming follicles.

(D) Trip13Gt/Gt 2-d-old ovary, dysgenic due to lack of oocytes. Magnification is the same as its littermate in “C.”

(E) Wild-type testis.

(F) Trip13Gt/Gt testis with uniform pachytene arrest.

(G) Trip13Gt/Gt 3-mo-old testis with some postmeiotic spermatids (arrows).

(H) Spo11−/- testis. A tubule with spermatocytes at leptotene/zygotene transition is labeled ZP, and tubules with apoptotic spermatocytes are marked with an asterisk. The specimen was taken from a littermate of that in (I).

(I) Spo11−/− Trip13Gt/Gt testis. Labeling is the same as in (H). The inset contains a tubule with leptotene-zygotene spermatocytes.

(J) Mei1−/− Trip13Gt/+ testis. The specimen was taken from a littermate of that in (K).

(K) Mei1−/− Trip13Gt/Gt testis.

(L) Rec8Mei8/Rec8Mei8 Trip13Gt/+ testis. The Rec8Mei8 allele was described [39]. The specimen was taken from a littermate of that in (M).

(M) Rec8Mei8/Rec8Mei8 Trip13Gt/Gt testis.

(N) Dmc1−/− Trip13Gt/Gt testis.

(O) Spo11−/− Trip13Gt/+ 25-d-old ovary. The specimen was taken from a littermate of that in (P).

(P) Spo11−/− Trip13Gt/Gt 25-d-old ovary.

(Q) Mei1−/− Trip13Gt/+ 25-d-old ovary. The specimen was taken from a littermate of that in (R).

(R) Mei1−/− Trip13Gt/Gt 25-d-old ovary.

(S) Rec8Mei8/Rec8Mei8 Trip13Gt/+ 25-d-old ovary. The specimen was taken from a littermate of that in (T).

(T) Rec8Mei8/Rec8Mei8 Trip13Gt/Gt 25-d-old ovary.

Histological sections of mutant testes revealed a lack of postmeiotic cell types that are characteristic of wild-type seminiferous tubules (Figure 3E). The most developmentally advanced seminiferous tubules contained adluminal spermatocytes with condensed chromatin characteristic of pachynema (Figure 3F). The absence of coordinated spermatogenic progression beyond this stage is indicative of a pachytene arrest. This was revealed more clearly by chromosome analysis (see below). Some sections of adult seminiferous tubules contained postmeiotic spermatids (Figure 3G), although we saw no motile epididymal sperm. These drastic meiotic defects stand in contrast to yeast and C. elegans, in which deletion of Pch2 alone has minor effects on spore/gamete development [2,8].

TRIP13-Deficient Meiocytes Undergo Homologous Chromosome Synapsis Despite the Presence of Unrepaired DSBs in Pachynema

To better characterize the degree of meiotic progression in Trip13Gt/Gt spermatocytes, we immunostained chromosome spreads for SYCP3 and SYCP1, components of the axial/lateral elements and transverse filaments, respectively, of the synaptonemal complex (SC). Pachytene spermatocyte nuclei from postpubertal mutant testes could assemble normal SC cores and exhibited full synapsis of chromosomes as judged by colabeling of SYCP1 and SYCP3 along the full lengths of all autosomes (Figure 4A). Additionally, the X and Y chromosomes were normally synapsed at their pseudoautosomal region. More prepubertal (17.5 d postpartum) mutant spermatocytes contained asynaptic or terminally asynapsed chromosomes than age-matched controls (62.5% versus 25%, respectively; Figure 4B). We attribute this to a delay in the first wave of postnatal spermatogenesis (Figure 2D and 2E), likely related to systemic developmental retardation (Figure 2A and 2B). Nevertheless, since Trip13Gt/Gt spermatocytes progress to pachynema with no gross SC abnormalities, and oocytes were eliminated soon after birth (a characteristic of DNA repair mutants [13]), this suggested that unrepaired DSBs are responsible for eventual meiotic arrest and elimination.

Figure 4

Immunohistochemical Analysis of Pachytene Spermatocyte Chromosomes

Surface-spread chromosomes were immunolabeled with the indicated antibodies and fluorophores. As indicated in the upper right of each panel, cells were from wild type (WT, either +/+ or Trip13Gt/+) or Trip13Gt/Gt (Mut). There were no differences seen between heterozygotes and +/+ spermatocytes.

(A) A mutant pachytene nucleus with full synapsis. Areas of SYCP1/SYCP3 colabeling are yellow.

(B–E) Spermatocytes nucleus from 17.5 d postpartum mutant. Asynapsed chromosomes or regions of chromosomes are indicated by white and yellow arrows, respectively. Unlike the normal distribution in wild-type pachytene spermatocytes (C), BLM foci are present on synapsed pachytene chromosomes in the mutant (D). RAD51 foci, which are abundant earlier in prophase, disappear from autosomes in wild-type pachytene nuclei (E) and the bulk of staining is over the XY body (arrow).

(F) RAD51 persists on the synapsed mutant chromosomes (arrows).

(G) H2AX phosphorylation is restricted to the XY body in WT.

(H) In addition to a large area of γH2AX staining (arrow) over the XY body, there is extensive autosomal H2AX phosphorylation (arrows).

(I, J) Note that in wild-type pachytene spermatocytes, TOPBP1 is present only over the XY body (yellow arrow). In the mutant (J), an arrow denotes one area of intensive staining that may be over the sex chromosomes, but many other chromosome cores are positively stained.

(K, L) RPA persists along synapsed cores in the mutant, not WT.

(M, N) Arrows indicate examples of MLH3 foci on SCs.

(O) In WT late pachytene spermatocytes, RAD51 is present only at background levels.

(P) As in (F), extensive RAD51 staining delineates SCs in mutant pachytene nuclei (indicated by white arcs). MLH1 foci colocalize with these tracts (arrows) at the typical 1–2 foci per chromosome as in (M).

To elucidate the cause of meiotic arrest, we analyzed meiotic chromosomes with a variety of markers that are diagnostic of recombination and synapsis. Recombination in Trip13Gt/Gt spermatocytes appeared to initiate normally as judged by the presence of γH2AX in leptonema (Figure S2A and S2B), which reflects the presence of meiotically induced DSBs [18]. RAD51 and/or DMC1, components of early recombination nodules (ERNs), was also present as abundant foci in Trip13Gt/Gt zygotene spermatocytes (unpublished data; the anti-RAD51 antibody cross-reacts with DMC1), indicating that recombinational repair of DSBs is initiated. The cohesin complex, which is essential for completion and/or maintenance of synaptic associations, appeared to assemble normally as judged by immunolabeling for the meiosis-specific cohesins STAG3 (Figure S2C and S2D) and REC8 (unpublished data). Because yeast PCH2 localizes to telomeres in a Sir3p-dependent manner, we tested for possible telomere defects by immunolabeling for TRF2, a component of a protein complex that plays an essential role in telomere protection [19]. It was localized to telomeres of both fully synapsed and telomerically asynaptic mutant chromosomes (Figure S2E and S2F).

Defects in DSB repair became apparent in pachynema upon probing of mutant spermatocyte nuclei with antibodies against molecules involved in various stages of recombination. In >99% of Trip13Gt/Gt chromosome spreads, BLM helicase (Figure 4C and 4D), RAD51/DMC1 (Figure 4E and 4F), γH2AX (Figure 4G and 4H), and TOPBP1 (Figure 4I and 4J) all persisted abnormally on synapsed chromosomes. For RAD51/DMC1, mutant pachytene spermatocytes contained 138 ± 6 foci (compared to 11 ± 3 foci in wild type, most of which were on the XY body), down from 218 ± 13 in zygonema (compared to 220 ± 13 foci in wild type). TOPBP1 is a DNA damage–checkpoint protein involved in ATM protein–dependent activation of ATR protein [20,21]. It binds sites of DSBs and unsynapsed regions of meiotic chromosomes [22,23]. BLM has been reported to colocalize with markers (RPA and MSH4) of recombination at sites distinct from those that become resolved as crossovers (CO) [24]. We therefore assessed the distribution of RPA, the ssDNA binding protein, which is normally present at focal sites of synapsing meiotic chromosomes before disappearing in mid-pachynema [25]. It is thought to bind D-loops of recombination intermediates [26]. RPA also persisted on pachytene mutant chromosomes (Figure 4K and 4L). These data indicate that unrepaired DSBs, or unresolved recombination intermediates, remain in pachynema and activate a DNA damage checkpoint system.

It should be noted that chromosomes affected by meiotic sex chromosome inactivation (MSCI) and meiotic silencing of unpaired chromatin (MSUC) are heavily stained by antibodies for several DSB repair-associated molecules, including γH2AX. H2AX phosphorylation due to MSCI and MSUC is conducted by ATR, not ATM [27–29]. Since mutant chromosomes are fully synapsed, and MSUC is known to occur only as a result of asynapsis, the decoration of Trip13Gt/Gt chromosomes with DNA repair markers is probably attributable to incomplete DNA repair rather than transcriptional silencing.

Consistent with the presence of rare (<1%) Trip13Gt/Gt pachytene spermatocytes devoid of persistent DNA repair markers, and testis histology showing some degree of postmeiotic progression (Figure 3G), we observed both diplotene nuclei that lacked autosomal RAD51/DMC1 and γH2AX (Figure S3A–S3D), and also metaphase I spreads with 20 bivalents (Figure S3E–S3F). Since Trip13Gt may not be a complete null, these diplotene and metaphase I spermatocytes might arise by virtue of having sufficient wild-type TRIP13.

CO-Associated Markers Appear Normally in the Absence of TRIP13

The persistence of BLM on Trip13Gt/Gt spermatocyte chromosomes suggests that at least a subset of the unrepaired DSBs correspond to sites of defective NCO recombinational repair. To assess whether CO recombination occurs in the mutant, we examined the distribution of the mismatch repair proteins MLH1 and MLH3, which are normally detectable as foci in mid-late pachynema and mark the locations of chiasmata [30,31]. Remarkably, MLH1/3 foci were formed; we observed 1–2 foci/chromosome as in wild type and at typical overall levels (MLH3 = 23 ± 2, N = 10; [30,32]) on mid-late pachytene chromosomes (Figure 4M and 4N; MLH1 not shown). Since <1% of Trip13Gt/Gt pachytene nuclei had normal repair (as judged by absence of persistent DSB repair markers; see above), but most of the pachytene nuclei had MLH1/3 foci, it was unlikely that the MLH1/3 foci formed only on chromosomes with fully repaired DSBs. To test this directly, we conducted double staining for MLH1 and RAD51/DMC1. MLH1 foci were present on chromosomes that also contained numerous RAD51/DMC1 foci (Figure 4O and 4P).

To assess whether these MLH1/3 foci in Trip13Gt/Gt pachytene spermatocytes represent CO events completed to a point where they could maintain interhomolog attachments though the end of prophase I, we treated testicular cells from 17.5–20.5-d-old control (+/+), Trip13Gt/Gt, and Dmc1−/− mice with the protein phosphatase inhibitor okadaic acid (OA), a chemical that induces degradation of the SC, chromosome condensation, and premature progression to metaphase I [33]. Fifteen metaphase spreads were identified for each genotype. Whereas all of the Dmc1−/− spreads had ∼35 or more condensed chromosomes, all of the +/+ and Trip13Gt/Gt spreads had 20–25, suggesting that the MLH1/3 foci in Trip13Gt/Gt pachytene spermatocytes represent sites of completed, or near-completed, COs. Because the preparations were made from whole testes, it is possible that the univalent-containing metaphases from Dmc1−/− mice were from spermatogonia, not spermatocytes.

TRIP13 Deficiency Does Not Alleviate Meiotic Arrest Phenotypes of Mutants Defective in Synapsis

To determine if TRIP13 deficiency prevents apoptosis triggered by asynapsis as in C. elegans, we analyzed mice that were doubly mutant for Spo11 and Trip13. SPO11 is a transesterase that is essential for the creation of genetically programmed DSB during leptonema of many organisms, including mice [18]. In C. elegans, spo-11 mutant gametes have extensive asynapsis, which triggers PCH-2 dependent apoptosis in pachynema [2]. In mice, Spo11−/− spermatocytes are severely defective in homologous chromosome synapsis [34,35], and arrest with chromosomes in a state characteristic of the zygotene/pachytene transition (Figure 3H). Spermatocytes in Trip13Gt/Gt Spo11−/− testes progressed maximally to that point before undergoing death (Figs 3I), well before the spindle checkpoint that eliminates achiasmate spermatocytes [36]. There was no evidence of metaphase I spermatocytes or postmeiotic spermatids in these testes, unlike those seen in Trip13 single mutants (Figure 3G). In contrast to the complete synapsis in Trip13Gt/Gt pachytene spermatocytes (Figure 5A), in which SPO11 is available in leptonema to initiate (via DSB induction, Figure S2A and S2B) a recombination-driven homolog search, chromosome synapsis in doubly mutant spermatocytes was highly disrupted as in Spo11 single mutants (Figure 5B and 5C). Identical studies were performed with mice deficient for Mei1, a vertebrate-specific gene also required for DSB formation and chromosome synapsis [37], with similar results (Figure 3J and 3K; immunocytology not shown).

Figure 5

Immunocytological Analysis of Trip13 Compound Mutants

Surface-spread chromosomes were immunolabeled with the indicated antibodies and fluorophores. Genotypes are indicated, as are those panels in which dual staining patterns are merged. Note that (H) and (I) are at lower magnification to show multiple nuclei.

In yeast, deletion of PCH2 alleviates the pachytene arrest caused by asynaptic mutants zip1 and zip2 [8]. Although mouse SYCP1 might be a functional equivalent of Zip1p, because Sycp1 mutant spermatocytes arrest at approximately the same point as Trip13 mutants, there would be no opportunity to observe bypass of Sycp1−/−. Since Zip2p is present at sites of axial associations, even in zip1 mutants, it has been suggested that Zip2p promotes initiation of chromosome synapsis [38]. These observations raise the possibility that in yeast, Pch2p responds to synapsis polymerization rather than initiation. To test this, we performed epistasis analysis with a Rec8 allele (Rec8Mei8, abbreviated as Rec8−). Meiotic chromosomes of Rec8 mutant spermatocytes undergo apparent homolog pairing and interhomolog synaptic initiation, but are defective in DSB repair and fail to maintain interhomolog synapsis [39,40]. Rather, sister chromatids appear to synapse and are bound by SYCP1 along their axes. Rec8 mutants do not progress to diplonema or metaphase I. Double mutant analysis indicated that Rec8 is epistatic to Trip13. As in the Spo11 and Mei1 experiments, histology of testes deficient for both REC8 and TRIP13 resembled the Rec8 mutant, with no evidence of progression to metaphase I that occurs in Trip13Gt/Gt mice (Figure 3L and 3M). Immunocytological analysis of spread chromosomes showed a failure of homologous chromosome synapsis in both the Rec8−/− and Rec8−/− Trip13Gt/Gt spermatocytes, as previously reported for Rec8 mutants (Figure 5D and 5E) [39,40].

Although subsequent reports indicate otherwise [10,12], deletion of PCH2 in yeast was originally reported to alleviate meiotic arrest caused by deficiency for the meiosis-specific RecA homolog DMC1 [8]. To investigate this relationship in mice, we constructed animals doubly mutant for Trip13 and Dmc1. As in Dmc1−/− mice, in which spermatocytes undergo meiotic arrest from defective DSB repair and failed chromosome synapsis [16], spermatogenesis in Dmc1−/− Trip13Gt/Gt testes was uniformly arrested at the point where spermatocytes contained chromatin characteristic of zygonema/pachynema (Figure 3N). Immunocytological analysis indicated that both Dmc1−/− and Dmc1−/− Trip13Gt/Gt chromosomes had extensive asynapsis compared to Trip13Gt single mutants (Figure 5F–5H), and all had persistent RAD51/DMC1 foci and phosphorylated H2AX (γH2AX; Figure 5I–5L), confirming that Dmc1 is epistatic to Trip13. Doubly mutant females had residual ovaries, phenocopying Dmc1−/− and Trip13Gt/Gt single mutants (unpublished data).

Meiotic Defects in Trip13Gt/Gt Oocytes Are DSB-Dependent

Epistasis analysis of females was insightful with respect to the cause of arrest in Trip13 mutants. Both Mei1−/−/Trip13Gt/Gt and Spo11−/−/Trip13Gt/Gt females had ovaries with numerous follicles, identical to Mei1 and Spo11 single mutants (Figure 3O–3R). Thus, Spo11 and Mei1 are epistatic to Trip13 in oogenesis, just as they are to Dmc1 [13,41]. This demonstrates that oocyte loss in Trip13Gt/Gt females is dependent on DSB formation. In conjunction with the immunohistochemical data, these data provide strong evidence that meiotic arrest in Trip13 mutant mice is due to defects in DSB repair. As expected, ovaries of Rec8 Trip13 double mutants were devoid of oocytes as were those from either single mutant (Figure 3B, 3S, and 3T).

Discussion

Genetic experiments in S. cerevisiae provided evidence that the pachytene checkpoint monitors and responds to recombinational DSB repair and synapsis independently. Wu and Burgess concluded that the repair checkpoint is RAD17-SAE2 dependent, while the synapsis checkpoint is PCH2-ZIP1 dependent [12]. Of these four genes, SAE2 and ZIP1 do not have clear mammalian orthologs (although SYCP1 may be a functional ortholog of ZIP1), and mutation of the mouse RAD17 ortholog, Rad1, presumably causes embryonic lethality [42]. Thus, mutational analysis of mouse Pch2 (Trip13), which is also critical for the synapsis checkpoint in C. elegans [2], was the best remaining option to evaluate potential functional conservation in mammalian meiotic checkpoint control.

Our results demonstrate that in mice, the primary meiotic function of TRIP13 is in recombination itself. We found no evidence that it is involved in pachytene checkpoint control. Our data suggest that while recombination events destined to be resolved as COs can proceed normally in Trip13 mutants, DSBs that enter the NCO repair pathway are incompletely resolved or processed inefficiently. This hypothesis is compatible with current knowledge of meiotic recombination pathways. In S. cerevisiae, CO and NCO pathways are distinct [43]; they have different recombination intermediates, and are dependent upon different proteins [44,45]. Mice also appear to have independent CO versus NCO recombination pathways [46]. As in yeast, both require SPO11-induced breaks, but only the CO pathway requires MLH1. Both types of recombinant products are formed by mid-late pachynema. Another possibility is that the recombination defects are a result of defective intersister recombination. However, this type of DSB repair is suppressed in meiotic cells. Ablation of RAD54, which mediates intersister recombination in yeast, does not significantly disrupt meiosis in either yeast or mice [47,48]. Interestingly, RAD54-deficient spermatocytes display abnormal persistence of RAD51 foci on pachytene chromosomes, similar to those in TRIP13 mice, but there are no deleterious effects on meiotic progression or fertility [49].

Data from budding yeast also indicate that Pch2p functions in recombination. Deletion of PCH2 delays meiotic progression by ∼2 h in SK1 yeast, and causes a minor decrease in ascus formation [50]. DSBs persist >2 h longer in pch2Δ yeast than in wild type, and hyperresection of DSBs in dmc1Δ pch2Δ double mutants is lower than in dmc1Δ cells [10]. Additionally, it was reported that pch2Δ yeast had a meiosis I delay dependent on the RAD17–SAE2 checkpoint that monitors recombination intermediates [12]. However, the exact role of TRIP13 (or Pch2) in recombination is unclear. Because synapsis occurs in TRIP13-deficient spermatocytes and is dependent on DSB formation (activity of SPO11 and MEI1), we suggest that TRIP13 functions after homology recognition and strand exchange, and that recombination events entering the CO repair pathway are either completed or nearly so (because OA treated resulted in bivalent chromosomes). One possibility for TRIP13′s role in recombination is that it is directly involved in a step specific to resolution of NCO recombination intermediates. Another possibility is that TRIP13 is required for disassembly of NCO recombinational repair complexes [51] containing those proteins that persist abnormally on Trip13Gt/Gt pachytene chromosomes. Notably, TRIP13 has two putative ATPase domains, a signature of AAA-ATPase ClpA/B chaperones that perform protein or protein/DNA complex disassembly [52]. These potential recombination roles might not be limited to meiosis, since Trip13 is widely transcribed and the mutant animals exhibited developmental defects. Finally, TRIP13 might play an indirect role, such as providing a “licensing” signal for the resolution of NCO intermediates and completion of meiosis.

Regarding the cause of cell death in Trip13 mutants, our data indicate that this is triggered by defective DSB repair rather than asynapsis. We base this conclusion on two observations: (1) oocyte elimination is dependent upon DSB formation and (2) synapsis is normal in spermatocytes of adult testes. Indeed, this mutant is unique in that recombination defects occur in the absence of asynapsis (e.g., as in Dmc1 knockouts). Thus, the Trip13 mutant provides the first evidence that unrepaired DNA damage alone can trigger the mammalian pachytene checkpoint response. Furthermore, our results allow us to conclude that oocytes and spermatocytes share a similar, if not identical, DNA damage pachytene checkpoint that is decoupled from a synapsis checkpoint.

Interestingly, we found that OA treatment of Trip13Gt/Gt spermatocytes could propel them into MI, despite a report that the same did not occur when wild-type pachytene spermatocytes were treated with the DSB-inducing agents gamma radiation or etoposide [53]. It is possible that the nascent induction of DSBs in pachynema evokes a checkpoint response that cannot be bypassed by OA, whereas the post-strand invasion lesions in TRIP13-deficient spermatocytes do not.

TRIP13 was originally discovered to be an interactor with rat thyroid receptor beta (THRB; [54]), but the relationship between THRB and TRIP13 in meiosis is unknown. Interestingly, we observed that THRB is distributed diffusely throughout wild-type spermatocyte nuclei but is excluded from the XY (sex) body (unpublished observations), a compartmentalized nuclear domain beginning in pachynema, in which the sex chromosomes become heterochromatinized and transcriptionally silenced in the process of MSCI [55]. However, the XY body appeared intact in most mutant spermatocytes upon probing with several markers of XY heterochromatinization (unpublished observations). Considering that THRB knockout mice are viable and fertile [56], the functional relationship between TRIP13 and its receptor THRB in meiosis is unclear.

Given the high similarity of PCH2 orthologs throughout the eukaryotic world, one or more essential functions of this protein must be conserved. Since TRIP13 does not exhibit checkpoint function in mice, we surmise that the TRIP13/PCH2 ancestral protein had a function in recombination that persists to the present. Notably, A. thaliana does not appear to have a meiotic checkpoint activity that eliminates mutant meiocytes in a manner analogous to organisms such as mice, budding yeast, and female Drosophila [11,57], and mammalian TRIP13 is more similar to Arabidopsis PCH2 than the fly or worm proteins (Figures 1A and S1). The unusual relatedness between mammalian and plant PCH2 may therefore be attributable to both the presence of a common conserved function (namely recombination, although the role of PCH2 in plants has yet to be determined), and the absence of checkpoint function. Nevertheless, the evolutionary relationships between animals, fungi, and plants (which are discordant with PCH2 sequence phylogeny) do not allow parsimonious models addressing the points in time that checkpoint functions in PCH2 were gained or lost. It is possible that its checkpoint function evolved independently in worms and budding yeast. The picture will become clearer as the function of PCH2 in other organisms is elucidated.

The nature of the synapsis checkpoint in male mice remains unidentified. One possible candidate is Dot1 (PCH1 in yeast), a histone methyltransferase silencing factor that is required for pachytene arrest of zip1 and dmc1 mutants in yeast [58], and for preventing RAD54-mediated recombinational DSB repair between sister chromatids. However, DOT1 acts upstream of PCH2. Given that TRIP13 doesn't have checkpoint function in mice, a potential role for mammalian DOT1 in the pachytene checkpoint is dubious but awaits investigation. Recently, it was shown that the TRP53 homolog TRP63 is required for DNA damage–induced death of dictyate-stage primordial oocytes, leading to the suggestion that it is involved in monitoring genome integrity [59]. However, this activity occurs subsequent to a pachytene checkpoint. As alluded to earlier, a complicating problem for studying potential meiotic checkpoint genes in mice is that as in yeast, such genes often have mitotic functions (such as RAD24 [7]), and their ablation can cause lethality [42]. Unless mammalian pachytene checkpoint components have orthologs with similar functions in organisms such as yeast, their identities are likely to remain elusive.

Materials and Methods

PCR analysis of Trip13 cDNA.

Trip13 was amplified from samples of Clontech's Mouse Multiple Tissue cDNA Panel I (http://www.clontech.com), using the following primers: 5′-GCACCATTGCACTTCACATC-3′ (TRP3-6F) and 5′-TGACCATCAGACTGTCGAGC-3′ (TRP3-6R). These primers correspond to exons 3 and 6, respectively, and amplify a 330-bp cDNA product. The cDNAs in this panel are equalized to allow quantitative analysis by RT-PCR.

Generation of Trip13-deficient mice.

The mouse embryonic stem cell line RRB047 (strain 129/Ola) containing a gene trap insertion in Trip13 was obtained from BayGenomics (http://www.baygenomics.ucsf.edu/). The gene-trapping vector used to create this line, pGT1lxf, was designed to create an in-frame fusion between the 5′ exons of the trapped gene and a reporter, βgeo (a fusion of β-galactosidase and neomycin phosphotransferase II). The gene-trapped locus creates a fusion transcript containing exons 1–3 of Trip13 and βgeo. To identify the exact insertion site within intron 3, PCR was performed using one primer within the gene trap vector, and the other primer at various positions in intron 3 pointing towards the 3′ end of the gene. Product from a productive reaction was sequenced, revealing that the insertion site was 445 bp into intron 3.

Genotyping of mice.

Three primers were used to distinguish wild-type and mutant alleles of Trip13: primer 1, 5′-CGTCGCTCCATTGCTTTGTGC-3′; primer 2, 5′-AGTAGTGGTACACTGTATTTTTGCTTTCATTGA-3′; and primer 3, 5′-GTAGATCCCGGCGCTCTTACCAA-3′. Primers 1 and 2 are located upstream and downstream, respectively, of the gene trap insertion within the intron 3. Primer 3 corresponds to pGTlxf sequence. Primers 1 and 2 amplify a 700-bp band from a wild-type allele; primers 1 and 3 amplify a 540-bp fragment from a mutant allele. Separate reactions were used to assay the presence or absence of each amplicon from a DNA sample. The cycling conditions were: 94 °C 2 min; 35 cycles of 94 °C 30 s, 57 °C 45 s, and 72°C 50 s; and 72 °C 2 min.

RT-PCR.

Total RNA was isolated from adult testes with the RNeasy Mini Kit (Qiagen, http://www.qiagen.com), and 4.0 μg was oligo dT–primed and reverse-transcribed with Superscript II (Stratagene, http://www.stratagene.com). The entire Trip13 protein-coding sequence was amplified with primers 5′-ATGGACGAGGCGGTG-3′ and 5′-TCAAACATAAGCTGAAAGTT-3′. The cycling conditions were: 94 °C 2 min; 94 °C 30s, 55 °C 45 s, and 72 °C 80 s for 35 cycles; and 72 °C 2 min. The primers for amplifying the Med31 coding sequence as control were : 5′-ATGGCCGCGGCCGTCGCTATGG-3′ and 5′-TCATTTCCCTGCTGTGTTATTCTGCTGCTGCTGC-3′. The cycling conditions were: 94 °C 2 min; 94 °C 30 s, 55 °C 30 s, and 72 °C 35 s for 35 cycles; and 72 °C 2 min.

Development and purification of chicken antibodies.

A peptide corresponding to amino acids 25–40 of TRIP13, VLQRSGSTAKKEDIK, was conjugated to KLH and used to immunize chickens (done by Sigma Genosys, http://www.sigmaaldrich.com). Polyclonal IgY was isolated from eggs with the Eggcellent Chicken IgY Purification kit (Pierce, http://www.piercenet.com). IgY antibodies were then affinity purified using the immunizing synthetic peptide.

Western blotting.

50 μg of testis extract in RIPA buffer was separated by 8% SDS-PAGE and electrotransferred onto a Pure Nitrocellulose membrane (Bio-Rad, http://www.biorad.com). The membrane was incubated with a polyclonal rabbit anti-human TRIP13 antibody (18-003-42687; Genway, http://www.genwaybio.com). According to the manufacturer, the immunogen was a synthetic peptide embedded in sequence we deduced to correspond to exon 3. Binding was detected by chemiluminescence ECL kit (Pierce) using a rabbit anti-chicken IgG horseradish peroxidase conjugate (Pierce).

Histological analyses.

Testes or ovaries were fixed in Bouin's, embedded in paraffin, sectioned at 6 μm, and stained by hematoxylin and eosin. Antigen retrieval for immunohistochemistry of testis sections was as described [60]. Oocyte and follicle numbers were counted as described [61]. Only follicles containing an oocyte with a clearly visible nucleus were scored.

Immunocytochemistry.

Immunolabeling of surface-spread spermatocytes and oocytes was performed as described [39,62]. To reach conclusions on the pattern of staining for various proteins, 30 (unless otherwise indicated) well-spread nuclei of particular meiotic stages were first identified under the fluorescent microscope on the basis of SYCP3 or STAG3 staining, then imaged at both appropriate wavelengths to determine the pattern of second proteins with focal patterns such as RAD51 or RPA. Unless otherwise indicated, the panels shown in the figures were the exclusive or predominant patterns seen. The exception for this approach was in the case of staining for MLH1 or MLH3 plus RAD51 (in which case SYCP3 or STAG3 was not available to find chromosome cores). Nuclei in this situation were identified first by MLH1/3 foci clustering, then imaged for both fluorescent wavelengths.

Primary antibodies used in this study were as follows: mouse anti-SCP3 (1:500; Abcam, http://www.abcam.com); rabbit anti-SYCP1 (1:1,000; a gift from C. Heyting) [63]; rabbit anti-REC8 (1:100; a gift from C. Heyting); rabbit anti-RAD51 (1:250, this polyclonal antibody recognizes both RAD51 and DMC1; Oncogene Research Products, http://www.merckbiosciences.co.uk); rabbit anti-γH2AX (1:500; Upstate Biotechnology, http://www.upstate.com/); rabbit anti-STAG3 (1:1,000; a gift from R. Jessberger); rabbit anti-MLH3 (1:400; a gift from P. Cohen); mouse-anti-human MLH1 (1:50; BD Biosciences, http://www.bdbiosciences.com); rabbit-anti-TopBP1 (1:100; a gift from J. Chen) [22]; mouse-anti-ubiquityl-histone H2A (1:200; Upstate Biotechnology); rabbit-anti-TRF2 (1:500; a gift from T. de Lange); and rabbit-anti-BLM (1:50; a gift from R. Freire). All secondary antibodies conjugated with either Alexa Fluor 488 or 594 (Molecular Probes, http://probes.invitrogen.com/) were used at a dilution of 1:1,000. All images were taken with a 100× objective lens under immersion oil.

Metaphase I spermatocyte spreads and OA treatment.

Metaphase fixed spermatocytes from 8-mo-old Trip13RRB047 homozygotes, using 23-d-old wild-type mice as control, were prepared and stained with Giemsa as described [64].

For OA treatment, cells were exposed to 5 μM OA (Calbiochem, http://www.emdbiosciences.com) for 6 h at 32 °C in a humidified environment of 5% CO2 before spreading [65]. These preparations were stained with DAPI to visualize metaphase nuclei and chromosomes.

Phylogenetic analyses.

TRIP13 orthologs were identified by BLASTP searches of Genbank and other sources providing gene models such as Ensembl. The selected orthologs can be found in Table S1. Amino acid alignments were done with Clustal W, using the default settings with and without removing the regions outside of the AAA-ATPase central domain. The trees were constructed by using the neighbor-joining method with Poisson correction. The reliability of internal branches was assessed by using 500 bootstrap replicates, and sites with gaps were ignored in this analysis. Neighbor-joining searches were conducted by using the computer program MEGA3 [66].

Supporting Information

Figure S1

Depiction of Conserved Regions of Mouse TRIP13 and Its PCH2 Orthologs

(75 KB PDF)

Click here for additional data file.

Figure S2

Surface-Spread Chromosomes Immunolabeled with the Indicated Antibodies and Fluorophores

(947 KB PDF)

Click here for additional data file.

Figure S3

Trip13 Mutant Spermatocytes That Progress Beyond Pachynema Have Repaired DSBs and Form Bivalents at Metaphase I

(1.5 MB PDF)

Click here for additional data file.

Table S1

Sources of TRIP13 Amino Acid Sequences Used to Construct the Phylogenetic Tree in Figure 1

(22 KB PDF)

Click here for additional data file.

Acknowledgements

We thank P. Burgoyne, P. Cohen, E. Alani, and P. Moens for helpful discussions, R. Viswanatha for helpful comments on the manuscript, W. Pawlowski and T. Pawlowska for input on the evolutionary tree of PCH2, R. Freire and J. Chen for antibodies, M. Jasin for Spo11− embryonic stem cells, and R. Munroe for generating chimeric mice. We also thank an anonymous reviewer for pointing out that Pch2p might be sensing synaptic polymerization in yeast, not initiation.

Abbreviations

CO - crossover

DSB - double strand break

MSCI - meiotic sex chromosome inactivation

MSUC - meiotic silencing of unpaired chromatin

NCO - noncrossover

OA - okadaic acid

RT-PCR - reverse-transcriptase PCR

SC - synaptonemal complex

Footnotes

A previous version of this article appeared as an Early Online Release on June 21, 2007 (doi:10.1371/journal.pgen.0030130.eor).

Author contributions. XL and JCS conceived and designed the experiments and analyzed the data. XL performed the experiments. JCS wrote the paper.

Funding. This work was supported by NIH grant GM45415 to JS.

Competing interests. The authors have declared that no competing interests exist.


