To investigate in vivo the functions of the phosphatidylserine receptor Ptdsr, we generated a null allele in the mouse by gene targeting (Figure 1a,1b,1c). In contrast to previously described Ptdsr-knockout mice 3132, we used Bruce4 embryonic stem (ES) cells for gene targeting 33, thus generating a Ptdsr-null allele in a pure, isogenic C57BL/6J genetic background. The newly established knockout mouse line was named Ptdsr^tm1Gbf (hereafter referred to as Ptdsr ^-/-). Heterozygous Ptdsr^+/- mice were viable and fertile and showed no obvious abnormalities. Ptdsr ^+/- mice were intercrossed to generate homozygous Ptdsr-deficient mice. The absence of Ptdsr expression in Ptdsr ^-/- embryos was confirmed by RT-PCR (data not shown), and by northern and western blotting analyses (Figure 1d,e). Interbreeding of heterozygous mice showed that the mutation was lethal, since homozygous mutants were not detected in over 100 analyzed litters at weaning. To determine the stages of embryonic development affected by the Ptdsr^tm1Gbf mutation, timed breedings were followed by PCR genotyping (Figure 1c) of embryos. We recovered fewer than the expected number of homozygous embryos from intercrosses of Ptdsr^+/-mice. From a total of 1,031 embryos analyzed between gestational day (E) 9.5 and E18.5, 198 (19.2%) Ptdsr-deficient homozygous embryos were harvested, indicating that the introduced mutation is associated with a low rate of embryonic lethality in utero.

From E9.5 to E12.5, Ptdsr ^-/- embryos were viable and of normal size. At E13.5 and thereafter, however, most Ptdsr ^-/-embryos showed morphological abnormalities (Table 1). All homozygous embryos harvested were growth-retarded from E13.5 onwards, had a pale appearance, and displayed multiple developmental dysmorphologies. These included various head and craniofacial malformations, such as exencephaly, cleft palate and abnormal head shape (Figure 1f,g). Gross inspection revealed that eye development was severely affected in 14.1% of homozygous embryos. The affected animals displayed a complete unilateral or bilateral absence of the eyes (Table 1) that was never detected in Ptdsr ^+/+ or Ptdsr ^+/- littermates. Furthermore, homozygous embryos harvested between E12.5 and E15.5 had subcutaneous edema (Figure 1f,g). Because we were able to recover Ptdsr ^-/- embryos until E18.5, we investigated whether Ptdsr-knockout mice could be born alive. Careful observation of timed matings allowed us to recover Ptdsr ^-/- neonates, but homozygous pups died during delivery or within minutes after birth. Ptdsr-deficient neonates were also growth-retarded, had a pale appearance and displayed various malformations. These included cleft palate, abnormal head shape, absence of eyes and edematous skin (Figure 1h). Thus, deletion of the Ptdsr gene resulted in perinatal lethality with variable severity and penetrance of phenotypes.

The observed perinatal lethality indicates that Ptdsr plays an important role during development. Analysis by RT-PCR (data not shown) showed that Ptdsr is expressed early in development, because we were able to detect Ptdsr transcripts in ES cells and embryos at all developmental stages. To analyze in more detail the temporal and spatial expression patterns of Ptdsr, and to correlate expression patterns with observed pathological malformations, we made use of a Ptdsr-β-geo gene-trap reporter mouse line generated from a Ptdsr gene-trap ES cell clone. This line has an insertion of β-galactosidase in the 3' region of the gene (Figure 2a).

We first examined Ptdsr expression by X-Gal staining in heterozygous embryos staged from E9.5 to E12.5. These developmental stages were chosen so as to investigate Ptdsr expression in affected organs prior to the onset of pathological malformations in Ptdsr ^-/- embryos. At E9.5 we found Ptdsr expression in the developing neural tube, somites, heart, gut and branchial arches (Figure 2b). At E10.5, Ptdsr expression remained high in the developing nervous system, with most intense staining in the forebrain, hindbrain and neural tube. At this stage of embryogenesis, high levels of Ptdsr expression could also be detected in the developing limb buds and eyes (Figure 2b). Ptdsr expression was altered at E12.5, with most intensive β-galactosidase staining in the eyes, developing condensations of the limb buds, neural tube and brain (Figure 2b). Transverse sections of X-Gal-stained embryos at E12.5 showed an asymmetric expression pattern in the neural tube with intense staining of the central mantle layer but no expression in the dorsal part of the neural tube (for example, the roof plate; Figure 2c). Expression in dorsal root ganglia lateral to the neural tube and in the somites was observed; Ptdsr was expressed throughout the somite structure (myotome, dermatome and sclerotome; Figure 2d). Expression boundaries between somites were evident, with no expression in the segmental interzones, which correspond to the prospective intervertebral discs (Figure 2d). Transverse sections of the developing eye at E12.5 revealed strong Ptdsr expression in the inner layer of the neural cup, which will later develop into the neural retina. Furthermore, Ptdsr expression was detected in the primary lens fiber cells of the developing lens (Figure 2e). We carefully investigated whether Ptdsr is expressed from E10.5 to E12.5 in the developing kidney and lungs, but no expression could be detected indicating that Ptdsr expression is required only at later stages in the development of these organs (see below).

Hybridization of a multiple-tissue northern blot revealed a single transcript of about 1.8 kb in almost every tissue analyzed in adult mice (Figure 2f). The most prominent expression was observed in testis, thymus, kidney, liver and skin, with moderate to low expression in lung, small intestine, spleen, stomach and skeletal muscle. Thus, Ptdsr is ubiquitously expressed throughout embryogenesis and in adult tissues, although at different levels.

We next examined the role of Ptdsr in organ development. Serial histological sections of Ptdsr ^-/- and control embryos were taken to perform a detailed morphological analysis of all organ systems during development. A significant delay in organ and tissue differentiation was observed at E16.5 in lungs, kidneys and intestine. Lungs of control littermates were properly developed with expanding alveoli (Figure 3a). Terminal bronchi and bronchioles were already well developed, and terminally differentiated epithelial cells with cilia on the luminal cell surface were present. In contrast, almost no alveoli or bronchioles were present in Ptdsr ^-/- lungs, indicating a delay or arrest in lung sacculation and expansion. Instead, we observed an abundance of mesenchyme that appeared highly immature (Figure 3g). A similar delay in tissue differentiation of Ptdsr ^-/- embryos was found in the kidneys (Figure 3h). Kidneys from Ptdsr^+/+ embryos were well developed at E16.5, showing terminally differentiated glomeruli with Bowman's capsule and collecting tubules lined with cuboidal epithelial cells (Figure 3b). In contrast, Ptdsr-deficient kidneys had only primitive glomeruli at E16.5, and collecting tubules were less well-developed. Instead, a large amount of undifferentiated mesenchyme was present in Ptdsr ^-/- kidneys (Figure 3h). A delay in tissue differentiation was also found in the intestine at this stage of development. Ptdsr ^-/- embryos displayed improperly developed villi and an underdeveloped or absent submucosa (Figure 3i). In wild-type embryos (Figure 3c), intestinal cellular differentiation was already highly organized, with intramural ganglion cells between the external and internal muscular layers. Such neuronal cells were absent from the intestine of Ptdsr ^-/- embryos (Figure 3i), however.

Some Ptdsr ^-/- mice (4.5 %) also displayed extensive brain malformations that resulted in externally visible head abnormalities, with occasional ectopic tissue outside the skull or exencephaly (Figure 1f,h). Histological analysis revealed an extensive hyperplasia of brain tissue with herniation of brain tissue either through the skull-cap or through the ventral skull (Figure 3d,j). In the most severe cases, expansion of brain tissue in mutant mice resulted in further perturbations of cortical structures (Figure 3d,j). Of note, a similar brain phenotype was observed in the Ptdsr-deficient mouse line generated by Li and colleagues 31.

In contrast to the study of Li et al. 31, however, we found almost normally developed lungs at birth. Ptdsr ^-/-lungs showed, in comparison to wild-type, only a slight delay in maturation and were fully ventilated in neonates in most cases (Figure 3e,k). This demonstrates that Ptdsr-deficient mice can overcome the delay in embryonic lung differentiation and display normal lung morphology at birth. Thus, it would appear highly unlikely that Ptdsr ^-/- mice die from respiratory failure. Consistent with the observations of Kunisaki and colleagues 32, we found severely blocked erythropoietic differentiation at an early erythroblast stage in the liver (Figure 3f,3l), suggesting an explanation for the grossly anemic appearance that we observed in our Ptdsr ^-/- mice.

By gross morphology we could differentiate two classes of Ptdsr mutants: those that appeared normal with both eyes present (Figure 4) and those that were severely affected and displayed uni- or bilateral anophthalmia (Figure 5).

Analysis of normal or mildly affected embryos revealed no differences between mutant and wild-type embryos in the differentiation of the developing eye until E16.5. In both genotypes, inner and outer layers of the retina displayed a comparable differentiation status, as shown, for example, at E12.5 (Figure 4a,e). At day E16.5, however, retinal layers in Ptdsr ^-/- embryos were much thinner than in wild-type embryos, contained fewer cells and were greatly reduced in size (Figure 4b,f). Comparison of the retinal structures of Ptdsr ^+/+ and Ptdsr ^-/- embryos revealed that all four retinal layers were present in Ptdsr-knockout mice at E16.5 (Figure 4b,f). At E18.5 (Figure 4c,g) and in neonatal animals (postnatal day P0; Figure 4d,h), the differences in retinal differentiation between Ptdsr^+/+ and Ptdsr ^-/- mice were still evident, but the size reduction of the retinal layers was less pronounced in the knockout mice. Ptdsr-deficient animals seem to have compensated for the marked delay in cellular differentiation and expansion of retinal layers. Close examination of retinal structures revealed that the inner granular layer was still less expanded in Ptdsr-deficient animals, however, and that it contained fewer cells and was still severely underdeveloped in comparison with the corresponding retinal layer in control animals (Figure 4c,4g and 4d,4h). Thus, even mildly affected Ptdsr ^-/- mutants had ocular malformations with defects in differentiation of retinal structures.

We next examined Ptdsr ^-/- embryos that displayed unilateral or bilateral absence of eyes (Figure 5a) by serial sectioning of whole embryos. These embryos showed complex malformations of the optical cup, including absence of the lens (Figure 5b). Most surprisingly, we found pigmented epithelial cells in the nasal cavity of all Ptdsr-knockout mice with anophthalmia that were analyzed histopathologically. We could identify black-colored pigmented cells embedded in the epithelium of the maxillary sinus that resembled presumptive retinal-pigmented epithelium (Figure 5b,c). Examination of consecutive serial sections revealed the formation of a primitive eye structure, with induction and subsequent proliferation of ectopic mesenchymal tissue immediately adjacent to the displaced pigmented epithelium (Figure 5d). This structure was clearly induced ectopically, and we failed to identify similar changes in any of the wild-type embryos. In summary, we observed a wide range of ocular malformations in Ptdsr-deficient mice that ranged from differentiation defects in retinal cell layers (for example, the inner granular layer) in mildly affected homozygotes to anophthalmia in severely affected Ptdsr ^-/- mice that was associated with induction of ectopic eye structures in nasal cavities.

We next tested whether Ptdsr is functionally required for the clearance of apoptotic cells. We started with an investigation of cell death in vivo in the interdigital areas of the developing limbs. Apoptosis of interdigital cells in the distal mesenchyme of limb buds occurs most prominently from developmental stages E12.0 to E13.5 and can be easily examined in situ by whole-mount terminal deoxynucleotide transferase-mediated UTP end-labeling (TUNEL). We compared the pattern of interdigital cell death in fore and hind limb buds from Ptdsr ^-/- (n = 3) and Ptdsr ^+/+ (n = 3) mice at E12.5 and E13.5. No differences in accumulation of TUNEL-positive cell corpses were observed between the two genotypes (Figure 6a). The kinetics of cell death occurrence and regression of the interdigital web was similar in wild-type and mutant littermates, providing no evidence that Ptdsr-deficiency is associated with impaired clearance of apoptotic interdigital cells during limb development.

To investigate further whether removal of apoptotic cells is impaired in Ptdsr ^-/- mice, we stained immunohistochemically for activated caspase 3 (aCasp3) and analyzed additional organs and tissues where apoptosis plays a crucial role in tissue remodeling during development. Starting at E12.5, we analyzed and compared the number and distribution of aCasp3-positive cells in over 140 serial sections of three wild-type and six Ptdsr ^-/- embryos in consecutive and corresponding sections. The sagittal sections were separated by 5 μm, allowing a detailed analysis of apoptosis in several organs and tissues. Tissue restructuring by programmed cell death occurred most notably within the ventral part of the neural tube (Figure 6b,f) and in the developing paravertebral ganglia (Figure 6d,h) with many apoptotic cells being present. In these tissues Ptdsr is highly expressed at E12.5 (Figure 2c) but we observed no difference in the number or distribution of apoptotic cells in Ptdsr^+/+ and Ptdsr ^-/- embryos. The same was true for the developing kidney: apoptotic cells were present in Ptdsr^+/+ and Ptdsr ^-/- embryos, in limited numbers, but we failed to detect any differences in the number of apoptotic cells between the genotypes (Figure 6c,6g). Furthermore, when we continued our analysis of apoptotic cell clearance in vivo at E16.5, E17.5 and E18.5 of embryonic development as well as in neonatal mice, the number and distribution of apoptotic cells was similar in both genotypes. As already observed at E12.5, analysis of aCasp3-stained sections of the developing thymus, heart, diaphragm, genital ridge, eyes and retina convincingly showed that there was no impairment in apoptotic cell removal in Ptdsr ^-/- mice. Moreover, because Li and colleagues 31 reported impaired clearance of dead cells during lung development in Ptdsr-deficient mice, we examined the rate of apoptosis induction and cell clearance in our Ptdsr-knockout mice in the lung. Analysis of aCasp3-stained lung tissue from Ptdsr^+/+ and Ptdsr ^-/- mice at E17.5 and P0 demonstrated that apoptosis was an extremely rare event during lung morphogenesis at this stage. In addition, there were no differences in the number or distribution of apoptotic cells in Ptdsr ^-/- and Ptdsr ^+/+ mice. Furthermore, we were unable to detect any evidence of tissue necrosis in lungs from Ptdsr-deficient mice. In contrast to the report of Li et al. 31, we never observed recruitment of neutrophils or other signs of pulmonary inflammation at any stage of development in our Ptdsr-deficient mice.

To analyze whether macrophages are recruited into areas where apoptosis is prominent during embryogenesis, we stained consecutive serial sections either with the macrophage surface marker F4/80 or with aCasp3. Surprisingly, there was no co-localization of macrophages with apoptotic cells. In virtually all embryonic tissues, apoptotic cells and macrophages were localized in different compartments (Figure 6e,6i; and see also Additional data file 1, Figure S1). This suggests that at this stage of development it is mainly neighboring cells that are involved in removal of apoptotic cells, rather than professional macrophages. In summary, our analysis in vivo did not reveal any impairment in apoptotic cell clearance in Ptdsr-deficient embryos during development and further suggests that phagocytosis of apoptotic cells is mainly mediated by non-professional 'bystander' cells.

To determine whether macrophages from Ptdsr-knockout mice were impaired in the efficacy of apoptotic cell uptake in vitro, we performed phagocytosis assays with fetal-liver-derived macrophages (FLDMs) and quantified their phagocytosis rates. Phagocytosis of apoptotic thymocytes was investigated at 60, 90 and 120 minutes after addition of target cells in the absence of serum. Analysis of phagocytosis rates by flow cytometric analysis (FACS) revealed no differences in the efficacy of apoptotic cell uptake between Ptdsr ^-/- and Ptdsr^+/+ macrophages and demonstrated no differences in apoptotic cell engulfment between selected time points (data not shown). To re-examine and further independently validate the result of normal apoptotic cell uptake by Ptdsr ^-/- macrophages, we performed phagocytosis assays for 60 min and determined the percentage of macrophages that had engulfed apoptotic cells, in a total of at least 300 macrophages counted by fluorescence microscopy. Phagocytosed, 5-carboxytetramethylrhodamine- (TAMRA-) labeled apoptotic cells were identified as being engulfed by inclusion in F4/80-labeled macrophages. Analysis was done independently by three investigators who were not aware of macrophage genotypes (Ptdsr ^-/- or Ptdsr^+/+). Again, no differences were found in the percentage of macrophages that had engulfed apoptotic cells (Figure 7a,c,e) or in the relative number of phagocytosed apoptotic cells per macrophage (phagocytotic index; Figure 7f). Moreover, single Ptdsr ^-/-macrophages could be identified that had engulfed even more apoptotic target cells than had wild-type macrophages (Figure 7b,d). Thus, Ptdsr-deficient macrophages had a normal ability to ingest apoptotic cells and were not impaired in recognition or phagocytosis of cells that had undergone programmed cell death.

In addition to its suggested importance for phagocytosis of apoptotic cells, it has been proposed that Ptdsr fulfils a second crucial role in regulating and maintaining a non-inflammatory environment upon the recognition of apoptotic cells by macrophages 26. We therefore tested whether Ptdsr ^-/- macrophages were able to release anti-inflammatory cytokines after ingestion of apoptotic cells. We examined levels of TGF-β1 and interleukin-10 (IL-10) after stimulation of FLDMs with lipopolysaccharide (LPS), with and without co-culture of apoptotic cells. Quantification of TGF-β1 and IL-10 levels after 22 hours of culture demonstrated that Ptdsr ^-/- macrophages were able to secrete these anti-inflammatory cytokines upon ingestion of apoptotic cells, although at a slightly lower level than wild-type (Figure 8a,b). This indicates that ablation of Ptdsr function does not compromise in general the ability of macrophages to release immune-suppressive cytokines after recognition and engulfment of apoptotic cells.

To analyze whether pro-inflammatory signaling is affected in Ptdsr ^-/- macrophages, we stimulated FLDMs from Ptdsr^+/+ and Ptdsr ^-/- mice with LPS and measured levels of tumor necrosis factor-α (TNF-α) at different time points after stimulation (Figure 8c). Ptdsr ^-/- macrophages produced significantly less TNF-α than did wild-type macrophages. The difference in TNF-α secretion was first visible after 3 h of LPS stimulation and became more prominent during the course of the experiment (for example, after 9 h and 12 h of LPS stimulation; Figure 8c). To analyze whether TNF-α release by Ptdsr ^-/- macrophages can be affected by engulfment of apoptotic cells, we stimulated FLDMs with LPS, apoptotic cells or both. Quantification of TNF-α levels by ELISA after 22 h showed that Ptdsr-deficient macrophages release less TNF-α after stimulation with LPS alone, and also after double stimulation of macrophages with LPS and apoptotic cells (Figure 8d). Moreover, the double stimulation demonstrated that the LPS-induced TNF-α release by Ptdsr ^-/- macrophages could be inhibited by co-administration of apoptotic cells to an extent comparable to that seen in wild-type macrophages. Similar results were obtained when other pro-inflammatory cytokines, such as interleukin-6 and monocyte chemoattractant protein-1, were analyzed (data not shown). These results indicate that Ptdsr is not required in macrophages for the inhibition of pro-inflammatory signaling after recognition and engulfment of apoptotic cells. Ptdsr-deficiency does, however, affect the overall release of pro- and anti-inflammatory cytokines after stimulation with LPS and after double treatment with LPS and apoptotic cells, indicating that Ptdsr-deficient macrophages have a reduced capacity to produce or secrete pro- and anti-inflammatory cytokines.

In this study, we have generated a null mutation in the phosphatidylserine receptor (Ptdsr) gene in C57BL/6J mice. We show that ablation of Ptdsr results in profound differentiation defects in multiple organs and tissues during embryogenesis, although with variable penetrance. While this work was in progress, two other groups reported the generation of Ptdsr-deficient mice 3132. In all three knockout mouse lines, the first two exons (31 and this study) or exons one to three 32 were deleted by replacement with a neomycin-selection cassette. The Ptdsr-knockout mouse lines differ in the genetic background in which the mutation was generated and maintained, however. In our case, the Ptdsr-null allele was generated in an isogenic C57BL/6J background, whereas Li et al. 31 and Kunisaki et al. 32 investigated the phenotype of their Ptdsr-knockout mice in a mixed 129 × C57BL/6 background. The ablation of Ptdsr function results in perinatal lethality in all cases, but there are interesting differences in severity or expressivities of phenotypes among the different Ptdsr-deficient mouse lines. This might be due either to differences in genetic background or because the phenotypes that have been investigated in this study have not been analyzed in such detail before.

In the Ptdsr-knockout mouse line reported here, growth retardation started from E12.5 onwards and was associated with delayed differentiation in several organs in which Ptdsr is expressed either during embryogenesis or later in adulthood. At E16.5 almost no branching morphogenesis of the lung epithelium was observed in Ptdsr ^-/- lungs. Similarly, epithelial structures were only partially developed in mutant kidneys, without terminal differentiation of Bowman's capsule and with a severe reduction in the number of differentiated collecting tubules. Likewise, the differentiation of the intestine was also severely delayed at this developmental stage. When compared with wild-type controls, intestinal tissues of Ptdsr knockout mice appeared unstructured, with an absence of enteric ganglia and of differentiated smooth muscle tissue. Interestingly, defects in kidney and intestine differentiation were not described in the Ptdsr-knockouts generated by Li et al. 31 and Kunisaki et al. 32. Surprisingly, when we examined Ptdsr^-/- embryos shortly before birth (E18.5) or neonatally, we found only mild differentiation delays in organs that appeared severely affected at mid-gestation. This 'recovery' was most visible in Ptdsr ^-/- lungs: at P0 we found expanded lungs in the knockout mice that showed normal branching patterns, with differentiated alveoli and bronchioles.

We investigated the occurrence of programmed cell death during lung development in wild-type and Ptdsr-knockout mice throughout embryogenesis (E16.5 to P0). Comparative immunohistochemistry for aCasp3 revealed that apoptosis is a rare event during lung morphogenesis. Furthermore, we failed to detect any differences in the number of apoptotic cells in Ptdsr-knockout and wild-type animals in the rare cases where we could detect apoptotic cells within lung tissues. These findings are contrary to the results reported by Li et al. 31, who suggested that impaired clearance of apoptotic mesenchymal and epithelial cells causes a failure in lung morphogenesis in Ptdsr-deficient mice. In contrast, our findings are in line with the current view on lung development during embryogenesis. Accordingly, formation of the epithelial lung via branching morphogenesis can be subdivided into a series of sequential steps that involve: first, formation of the organ anlage in the form of a placode; second, primary bud formation by placode invagination; third, branch initiation and branch outgrowth; fourth, further reiteration of the branching process; and fifth, terminal differentiation of organ-specific proximal and distal structures 3435. In contrast to other invagination processes during embryogenesis, such as mammary gland formation, the lumen of the lungs is expanded by successive branching events, branch outgrowth and elongation, rather than by apoptosis 3436. Finally, because the lungs of Ptdsr ^-/-neonates were almost fully expanded and appeared normal in structure in comparison to wild-type littermates, it is highly unlikely that Ptdsr mutants die of respiratory lung failure. In addition, Li and colleagues 31 demonstrated that surfactant expression is normal in Ptdsr-deficient animals, supporting the idea of normal maturation of surfactant-producing type II alveolar epithelial cells and lung function. Other defects must therefore be responsible for the death of Ptdsr-mutant mice. The frequently observed subcutaneous edema of various extents in Ptdsr-deficient homozygotes gave us a hint that Ptdsr-deficiency and lethality might be associated with cardiovascular problems. Indeed, very recently we have obtained strong evidence that Ptdsr-knockout mice die as a result of defects in heart development that are associated with specific cardiopulmonary malformations; (J.E. Schneider, J.B., S.D. Bamfort, A.D.G., C. Broadbent, K. Clarke, S. Neubauer, A.L. and S. Battacharya, unpublished observations).

In addition, we demonstrate that eye development requires a functional Ptdsr gene. Ptdsr-deficient embryos can be roughly divided into two categories. The first, severely affected group develops anophthalmia that correlates with formation of ectopic retinal-pigmented epithelium and induction of proliferation of underlying mesenchyme in the nasal cavity. This phenotype represents a completely novel lesion that to our knowledge has not been described before in any other mouse mutant. The second group shows normal external eye structures, although in this case retinal development is temporally delayed during mid-gestation, with persistent, abnormal morphogenesis of the inner granular retinal layer at later stages of embryogenesis. A possible explanation for these two phenotypes can be found in the expression pattern of the Ptdsr gene. Initially, Ptdsr is expressed throughout the whole developing nervous system, with exceptionally high levels in the anterior part of the forebrain. Later expression becomes more restricted to the developing retina and lens. Thus, Ptdsr might play an important role in early events of ocular morphogenesis, such as establishment and bisection of eye fields and formation of optic cups. These early eye-formation steps are closely interconnected with development of the forebrain 3738 and the nose 394041.

Interestingly, we occasionally observed serious malformations of forebrain and nasal structures in Ptdsr-knockout embryos that were associated with bilateral anophthalmia (see for example the mutant embryo in Figure 1g). This suggests that Ptdsr is involved in the regulation of differentiation processes within forebrain regions, and that ablation of Ptdsr function might secondarily affect early eye formation. Li et al. 31 found smaller lenses in Ptdsr-knockout mice and described the formation of retinal protrusions, although anophthalmia and specific differentiation defects of retinal cell layers were not reported in their study. Li et al. proposed 31 that the eye phenotype they observed could be explained by failed removal of apoptotic cells during eye development, but we think that the observed defects are unrelated to a failure of apoptotic cell clearance. A recent comprehensive kinetic analysis of apoptosis induction during mouse retinal development described four major peaks of apoptotic cell death 42. This study demonstrated that there is an initial phase of cell death during the invagination of the optic cup (E10.5), followed by subsequent waves of apoptosis induction immediately before and after birth (E18.5 to postnatal day P2), and from postnatal days P9 to P10 and P14 to P16 42. Thus, besides the formation of the inner and outer layers of the optic cup in early eye development, other major phases of retinal cell apoptosis take place only postnatally and correspond to important periods in the establishment of neuronal connections. Furthermore, cell death during normal retinal development occurs in retinal layers distinct from the inner granular layer where we observed the most pronounced differentiation defects in the Ptdsr ^-/- mutants described here. Other studies that connect the postnatal elimination of apoptotic photoreceptor cells to Ptdsr-mediated macrophage engulfment 43 should be interpreted with extreme caution as these studies were based on the monoclonal anti-Ptdsr antibody mAb 217G8E9 2643 (see below).

Consistent with the results of Li et al. 31, we found particular brain malformations in our Ptdsr ^-/- mice. Exencephaly and hyperplastic brain phenotypes were observed at a low penetrance in Ptdsr-mutant mice (less then 4.5% of homozygotes), but these do not resemble to any extent the brain-overgrowth phenotypes of caspase- or Apaf1-knockout mice (44, and references therein) in that we failed to identify any differences in the number or distribution of apoptotic cells or pyknotic cell clusters in the neuroepithelium of Ptdsr ^-/- and Ptdsr^+/+ mice. Thus, reduced cell death or diminished clearance of apoptotic neural progenitor cells is unlikely to be the cause of the brain hyperplasia.

In summary, our studies demonstrate that Ptdsr is required for normal tissue differentiation, especially during the mid-gestation period when we observed the most severe differentiation delays in several organs of Ptdsr-knockout mice. The multiple defects in tissue differentiation cannot be explained by failure of apoptotic cell clearance, as this process is normal in our Ptdsr-knockout line. This result therefore indicates that Ptdsr has a novel, hitherto unexpected, role in promoting tissue maturation and terminal differentiation. Additional studies with conditionally targeted Ptdsr-deficient mice are required to investigate the role of spatial and temporal Ptdsr expression and function during tissue differentiation.

Our studies demonstrate that Ptdsr is not a primary receptor for the uptake of apoptotic cells. Investigation of apoptotic cell clearance in vivo in Ptdsr ^-/- embryos conclusively showed that removal of apoptotic cells is not compromised by ablation of Ptdsr function. Comparative analysis of ten different tissues and organs in Ptdsr^+/+ and Ptdsr ^-/- animals at several stages of embryonic development and in neonates failed to identify impaired uptake of apoptotic cells at any time during development. Furthermore, phagocytosis assays in vitro demonstrated a completely normal uptake of apoptotic cells by Ptdsr ^-/- macrophages, with some knockout macrophages showing loads even higher than wild-type of engulfed dead cells. These results are contrary to the expected role of Ptdsr in apoptotic cell clearance and to the reported findings of Li et al. 31 and Kunisaki et al. 32, as well as to a study done with a phosphatidylserine receptor null allele in C. elegans 45. In previous studies in the mouse, the distribution and amount of apoptotic cells in Ptdsr-knockout and control animals were investigated in only a few tissues and at one 31 or two 32 developmental stages. Li et al. 31 examined lung, midbrain and retina at day E17.5 of gestation and identified apoptotic cells by TUNEL staining. Their findings must be interpreted with caution because remodeling of cellular structures by apoptosis in specific retina layers is known to occur mainly postnatally 42, and apoptosis plays an important physiological role in the maintenance and homeostasis of lung epithelium after birth or in pathological conditions involving pulmonary inflammation and not during lung development 46. This postnatal role for apoptosis is in accordance with our data, as we rarely observed apoptotic cells in retina or lung tissue throughout embryogenesis in Ptdsr^+/+ and Ptdsr ^-/- mice. Kunisaki et al. 32 analyzed TUNEL-stained sections of liver and thymus at days E13.5 and E16.5 of development in Ptdsr^+/- and Ptdsr ^-/- embryos and found reduced rather than increased numbers of TUNEL-positive cells in Ptdsr-deficient embryos. Using co-localization of TUNEL-positive cells with F4/80-positive macrophages they suggested that Ptdsr ^-/- embryos exhibited a three-fold increase in the frequency of unphagocytosed TUNEL-positive cells together with a severely reduced number of F4/80-positive cells. These results must be interpreted very carefully, however, as it is technically difficult to unambiguously identify engulfed target cells in individual macrophages in solid tissues by fluorescence microscopy.

In addition, our data suggest that during embryogenesis, macrophage-mediated clearance of apoptotic cells is not the only - or even the primary - mechanism for the removal of apoptotic cells. In many tissues where programmed cell death occurs as a prominent event during embryogenesis, such as remodeling of the genital ridge during gonad morphogenesis and differentiation of the neural tube, we found almost no co-localization of apoptotic cells and macrophages. This indicates that in these cases clearance of apoptotic cells is directly mediated by neighboring 'bystander' cells rather than by macrophages that have been recruited into areas where apoptosis occurs. Obviously these in vivo clearance mechanisms are not compromised by Ptdsr-deficiency in our knockout mutant. This finding is in line with studies in macrophageless Sfpi1-knockout embryos that are deficient for the hematopoietic-lineage-specific transcription factor PU.1. Here, the phagocytosis of apoptotic cells during embryogenesis is taken over by 'stand-in' mesenchymal neighbors 47. As recognition of phosphatidylserine is thought to be a universal engulfment mechanism for all cells that are able to phagocytose apoptotic cells, it is very striking that apoptotic cell clearance mediated by non-professional bystander cells is also not compromised by Ptdsr-deficiency.

In contrast to Li et al. 31, we did not observe any impairment in the uptake of apoptotic cells by Ptdsr ^-/- macrophages in vitro. We performed phagocytosis assays in vitro with fetal-liver-derived macrophages, while in their assays, Li and colleagues used thioglycollate-elicited peritoneal macrophages after adoptive transfer of Ptdsr ^-/- hematopoietic stem cells. The different results obtained in the two studies are puzzling; they might be due to the use of different macrophage or cell populations. We and Kunisaki et al. 32 found that Ptdsr-deficiency is to some extent associated with defects in hematopoiesis. Thus, it seems possible that recruitment and activation/differentiation of macrophages after adoptive transfer and thioglycollate elicitation are affected by Ptdsr-deficiency. We do not think that the different results observed in Ptdsr-knockout mice in a mixed C57BL/6 × 129 background and in a pure C57BL/6J background can be attributed to genetic background effects: comparison of apoptotic cell engulfment efficacies of thioglycollate-elicited macrophages from 129P2/OlaHsd and C57BL/6J mice did not show any differences in apoptotic cell uptake (J.B. and A.L., unpublished observations). Moreover, in contrast to our studies, neither Li et al. 31 nor Kunisaki et al. 32 determined phagocytotic engulfment indexes for Ptdsr-deficient macrophages.

Interestingly, we observed differences between Ptdsr^+/+ and Ptdsr ^-/- macrophages in the secretion of pro- and anti-inflammatory cytokines after stimulation with LPS and apoptotic cells. This provides evidence that cellular activation and effector mechanisms are impaired in Ptdsr-deleted macrophages. It remains to be determined which classical pathways of macrophage activation and function involve Ptdsr. This is especially important in light of recent findings that demonstrated nuclear localization of the Ptdsr protein 29.

Most strikingly, the recently published data regarding the genetic ablation or perturbation of phosphatidylserine receptor function in C. elegans are also contradictory. Wang et al. 45 reported that psr-1, the C. elegans homolog of Ptdsr, is important for cell-corpse engulfment, whereas psr-1 RNAi studies performed by Arur et al. 25 yielded, in this respect, no phenotype. Moreover, Wang and colleagues hypothesized on the basis of their data that psr-1 might act to transduce an engulfment signal upstream of Ced-2 (Crk II), Ced-5 (Dock 180), Ced-10 (Rac 1) and Ced-12 (Elmo) in one of the two cell-corpse engulfment pathways in the worm 45. But the loss-of-function phenotype of psr-1 mutants and the complementation phenotypes in overexpressing transgenic worms shown by Wang et al. 45 are rather weak as compared to the classical C. elegans engulfment mutants 8.

Many previous functional studies that reported a requirement for Ptdsr for the phagocytosis of apoptotic cells used the monoclonal anti-Ptdsr antibody mAb 217G8E9 26. This antibody was used in Ptdsr binding and blocking experiments, as well as in subcellular localization studies, which led to the conclusion that Ptdsr is a transmembrane receptor critical for signal transduction at the engulfment interface. More recently it was used in binding assays to show that the human and worm Ptdsr molecules can recognize phosphatidylserine 45. In the course of the study presented here, we stained immunohistochemically for Ptdsr with mAb 217G8E9 on wild-type and Ptdsr-deficient macrophages and fibroblasts (see Additional data file 1, Figure S2 and data not shown). To our surprise, we observed similar staining patterns with cells of both genotypes. Furthermore, using a Ptdsr-peptide array we found that mAb 217G8E9 can bind weakly to a Ptdsr peptide, explaining the original isolation of Ptdsr cDNA clones by phage display 26; but the antibody mainly recognizes additional, as-yet unknown, membrane-associated protein(s) (see Additional data file 1, Figure S2). Experiments that have used this antibody should therefore be interpreted with great caution as they might come to be viewed in a different light.

The genotypes of embryos or animals were determined by PCR analysis and confirmed by Southern blot. Genomic DNA for PCR was prepared from extraembryonic membranes or tail clips using a non-organic tail-DNA extraction protocol 49. High molecular weight genomic DNA for Southern blotting was prepared according to standard protocols. For PCR analysis the wild-type Ptdsr allele was detected using forward primer 1 (5'-GACACTGTCCATGGCAAACAC-3') and reverse primer 2 (5'-TAAAGTCGCCTTCCAGAAGATT-3'). The primer 1 site is located 5' to the deletion and the primer 2 site within the deletion. This primer pair amplified a fragment of approximately 300 bp from wild-type and Ptdsr^+/- mice but not from Ptdsr ^-/-mutants. To detect the mutant Ptdsr allele, genomic DNA was also amplified using primer 1 and reverse primer 3 (5'-CCACACGCGTCACCTTAATA-3'), which corresponds to a sequence in the neo cassette. In this case, a 500 bp fragment was detected in mice heterozygous or homozygous for the mutant allele, while no signal was detected in wild-type mice. For Southern blot analysis, genomic DNA (30 μg) was digested overnight with BamHI (30 U; Roche Diagnostics GmbH, Mannheim, Germany) and ScaI (30 U; Roche), fractionated on a 0.8 % agarose gel, transferred to a nylon membrane (Hybond N; Amersham Biosciences Europe GmbH, Freiburg, Germany) and hybridized with 5' and 3' flanking probes. The BamHI digest was hybridized with a Ptdsr-specific 5' flanking probe, and Southern blot gave a single 17.2 kb band for wild-type (^+/+), an 11.6 kb band for homozygous (^-/-) and both bands for heterozygous (^+/-) mice. The ScaI digest was hybridized using a 3' flanking probe, and Southern blot gave a single 12.4 kb band for wild-type, a 17.2 kb band for homozygous and both bands for heterozygous mice.

Total RNA was isolated from homogenized embryos using TRIZOL reagent (Invitrogen GmbH, Karlsruhe, Germany). For northern blots, either total RNA (30 μg) was extracted from embryos, electrophoresed and transferred to a nylon membrane (Hybond N; Amersham) or a polyA⁺ RNA northern blot (OriGene Technologies Inc., Rockville, USA) was hybridized using as the probe a Ptdsr fragment amplified from wild-type cDNA using forward primer 5'-GTTCCAGCTCGTCAGACTCG-3' and reverse primer 5'-TGCCCCTAAGACATGACCAC-3'. In all experiments the same membrane was re-hybridized with a β-actin probe (OriGene) to confirm that equivalent RNA levels were present in each lane. Northern blotting indicated that homozygous mutant embryos did not express Ptdsr mRNA and heterozygous mutant embryos expressed only reduced amounts of Ptdsr mRNA.

Embryos (E13.5) for protein isolation were homogenized in lysis buffer containing 1 × PBS, 1% Nonidet P-40, 0.5% sodium deoxycholate, 0.1% SDS and protease inhibitor cocktail (CompleteMini; Roche). Equal amounts (25 μg) of protein lysate were separated by SDS-polyacrylamide gel electrophoresis and transferred onto a PVDF membrane (Millipore, Billerica, USA) according to standard protocols. Western blots were done using a specific antibody to Ptdsr (PSR N-20, sc-11632; Santa Cruz Biotechnology Inc., Santa Cruz, USA) and β-actin (ab-6276; Abcam, Cambridge, UK) as described by the supplier. Secondary antibodies conjugated to horseradish peroxidase were from Santa Cruz and Abcam, used as described by the supplier, and detection was performed with an enhanced chemiluminescence system (ECLPlus; Amersham).

Wild-type C57BL/6J and 129P2/OlaHsd mice were obtained from Jackson Laboratories (Bar Harbor, USA) and Harlan UK (Bicester, UK), respectively. All mice were housed in individually ventilated cages in a specific pathogen-free environment with a 12 h light-dark cycle and were fed a regular unrestricted diet. The GBF's routine surveillance program screened for selected pathogens. The Ptdsr^tm1Gbf mutant was crossed to C57BL/6J mice to establish the co-isogenic C57BL/6J-Ptdsr^tm1Gbf mouse line. All studies were approved by the appropriate authorities.

Heterozygous male and female mice were intercrossed in order to obtain Ptdsr-deficient progeny. Females were daily monitored for vaginal plugs, and noon of the day of plug detection was defined as E0.5. Embryos at indicated time points were dissected in sterile PBS, washed in ice-cold PBS and transferred to cold fixative. Extra-embryonic membranes were kept and used for genotyping. Ptdsr ^-/- embryos and their wild-type littermates were used for experiments.

Embryos for histology and immunohistochemistry were harvested and fixed in 10% neutral-buffered formalin, dehydrated through a graded series of alcohol, embedded in paraffin, sagittally sectioned at 5 μm intervals, and every fifth section was processed for hematoxylin and eosin (H&E) staining according to standard protocols. Remaining sections of wild-type and Ptdsr ^-/- specimens were used for immunohistochemistry. For detection of apoptotic cells and macrophages, anti-aCasp3 (an antibody specific for activated caspase 3; R&D Systems, Minneapolis, USA) and anti-F4/80 (Serotec GmBH, Düsseldorf, Germany; #MCA 1957) antibodies were used as described by the supplier. Detection was performed using indirect streptavidin with biotinylated secondary antibodies and cobalt-enhanced diaminobenzidine (brown) or fast-red (red) as chromogens. Sections were counterstained with hematoxylin. For whole-mount terminal deoxynucleotidyl transferase-mediated UTP end labeling (TUNEL), limb buds were dissected from E12.5 and E13.5 embryos, fixed in 4% paraformaldehyde and processed for analysis as previously described 50.

Fetal livers were excised from embryos at E12.5 and E13.5, respectively, washed in PBS and dissociated enzymatically for 60 min at 37°C. The digestion buffer (150 μl per liver) comprised 0.6 U/ml dispase I (Roche), 0.1% collagenase D (Roche), 10 U DNase (Roche), and 20% FCS in PBS. X-Vivo 15 medium (Cambrex, East Rutherford, USA) was added to the resulting cell suspension, and after centrifugation (200 × g; 3 min) cells were resuspended in X-Vivo 15 medium supplemented with 50 ng/ml macrophage colony-stimulating factor (M-CSF; Sigma-Aldrich, St. Louis, USA) and cultured on non-treated tissue-culture dishes at 37°C with 5% CO₂. Every second or third day the medium was changed by centrifugation. Following withdrawal of M-CSF on day 6 after excision, adherent cells were cultured for an additional 24-48 h in X-Vivo 15 medium.

For preparation of monolayer cultures of macrophages, FLDMs were plated on glass coverslips in 24 well plates (2 × 10⁵ cells per well) in X-Vivo 15 medium. For preparation of apoptotic target cells, primary thymocytes were harvested from the thymus of 4- to 8-week-old C57BL/6J mice, stained with TAMRA for 15 min, and apoptosis was induced either by treating cells with 5 μM staurosporine in medium for 4 h at 37°C or by culturing cells in medium overnight. The efficacy of apoptosis induction was compared in thymic target cells and controls by FACS analysis. On average, 60% of the cells of the resulting population were apoptotic, with exposed PS on their surface, and less than 5% of the cells were necrotic, as confirmed by FITC-annexin V and propidium iodide staining. The apoptotic thymocytes obtained were washed with PBS and added to the prepared FLDM cultures (ratio 10:1). Phagocytosis was then allowed to proceed at 37°C and 5% CO₂. After the indicated time periods, the uptake of apoptotic cells by FLDMs was stopped by intensive washing of co-cultures with cold PBS to remove unphagocytosed cells. To measure phagocytosis of apoptotic thymocytes, macrophages were further processed for immunofluorescence analysis. Cells were fixed in 4% paraformaldehyde, blocked in 0.5% BSA/PBS and stained with an anti-F4/80 antibody (Serotec) followed by a secondary antibody coupled to Alexa 488 (Molecular Probes Inc., Eugene, USA). Coverslips were mounted on slides and engulfed thymocytes were enumerated by fluorescence microscopy. The percentage of phagocytosis was calculated by counting at least 300 macrophages and determining the number of macrophages that had engulfed apoptotic thymocytes. The phagocytotic index was calculated according to the following formula: phagocytotic index = (total number of engulfed cells/total number of counted macrophages) × (number of macrophages containing engulfed cells/total number of counted macrophages) × 100. The experiments were performed at least three times, each time in triplicate, and the counting was done by three different investigators.

Monolayer cultures of FLDMs and apoptotic thymocytes were prepared as described above. FLDMs were incubated with medium, LPS (10 ng/ml), apoptotic cells (ratio 1:10) or both for the determination of IL-10, TGF-β1 or TNF-α levels after co-culture for 22 h. For TNF-α quantification at various time points, FLDMs were cultured with a high concentration of LPS (100 ng/ml). Culture supernatants were harvested and TNF-α (Mouse TNF-α OptEIA set; BD Biosciences, Heidelberg, Germany) and TGF-β1 (Quantikine, TGF-β1 immunoassay; R&D Systems) were measured by ELISA as described by the supplier. IL-10 in culture supernatants was determined by a cytometric bead assay (Mouse inflammation CBA; BD Biosciences) as indicated in the manual. Data are presented as mean ± SEM from at least three independent experiments, each carried out in triplicate. Analysis of the results used the Wilcoxon-signed rank test; p values below 0.05 were considered significant.