Bst mapping

Mouse strains were obtained from The Jackson Laboratory (Bar Harbor, ME). An intersubspecific backcross was performed between (C57BLKS Bst/+ × CAST/Ei) F1 and C57BLKS +/+ mice. N2 progeny were scored at weaning for white spotting, white hind feet and tail kinks, and were genotyped using MIT markers (Dietrich et al., 1992) and four new microsatellites:

• D16Ero1

  Forward: 5′-CTTTCTCTAATCATGGCAAAAACA-3′

  Reverse: 5′-TGAGCAAGACAATCACAGAGTATG-3′

• D16Ero2

  Forward: 5′-GAGCAAAATTTACGATGTGATTTG-3′

  Reverse: 5′-ATCTCTATTGCACAAAGATGGACA-3′

• D16Ero4

  Forward: 5′-ACATAGTCAACAGACCGGACCT-3′

  Reverse: 5′-ATGTTCTACTCGGTCTCTCCATCT-3′

• D16Ero7

  Forward: 5′-GCATTTAGTCAACACCATCAGAAT-3′

  Reverse: 5′-CAAGCAAGCATTTCATACAAAAGT-3′.

PCR products were separated using 6% polyacrylamide sequencing or 4% Metaphor agarose gels. To confirm Bst genotypes, critical recombinants and N2 mice with low expressivity were test-crossed to C57BLKS. Opa1 was genotyped using PCR primers 5′-GAGCAGAAGAGCCCTGGAC-3′ and 5′-CAATGGAGGGTGGATGTTTC-3′ and a BlpI polymorphism in the 3′UTR. To type Hes1, a genomic segment corresponding to the terminus of bacterial artificial chromosome (BAC) clone RP24-110I11 was amplified using primers 5′-TTATTTGTTTCTACGACCCAGCTT-3′ and 5′-ACTGGTCTTGAACTCCTAGACTCC-3′ and scored for a single-strand conformational polymorphism (SSCP) by MDE gel electrophoresis (FMC Bioproducts).

Rpl24 genotyping

The Bst allele was typed by PCR, using upstream primer 5′-TTTGCAGCGCACATACGAG-3′ which overlaps the first intron branchpoint deletion and downstream primer 5′-GCTGACTCACATTTTGCATTAAGA-3′ within the third exon. The wild-type allele was amplified using upstream primer 5′-CTCTTTGCAGCGCACATACTAAC-3′, which overlaps the intact first intron branchpoint and downstream primer 5′-GGAAAACCTGCAGTTAACAAATTC-3′ within the second intron. PCR parameters were 3 minutes at 95°C, followed by 40 cycles (30 seconds at 95°C, 1 minute at 56°C, 1 minute at 72°C) and 7 minutes extension at 72°C.

Reverse transcriptase PCR

Total RNA was extracted using Trizol reagent (Invitrogen) from livers of C57BLKS+/+, C57BLKS Bst/+ and SPRET/Ei adult mice; livers of P21 weanlings and embryonic day (E) 18.5 embryos derived from a (C57BLKS Bst/+ × CAST/Ei) F1 × SPRET/Ei cross; and heads of co-isogenic C57BLKS Bst/+ and +/+ E13.5 embryos. First strand cDNA synthesis was primed using random hexamers. To evaluate Bst candidate genes, 40 cycles of RT-PCR were performed on E13.5 head and body RNA using gene-specific primers. To evaluate Bst effects on Rpl24 splicing, three RT-PCRs were performed (designated A, B and C in Fig. 3D,E). These reactions share a common downstream primer (5′-CTGTCTTCTTTGATGCCTGCTTAG-3′), which was ³²P or HEX end-labeled. The upstream primers were 5′-GCAGGCCGACATCTATCAC-3′ (A), 5′-GGATGGCTCCTCTTTGCAG-3′ (B) and 5′-ACATCTATCACCATGAAGGTCGAG-3′ (C). The upstream primer in reaction C spans the exon 1–2 junction and only amplifies from correctly spliced Rpl24 mRNA. PCR parameters were 3 minutes at 95 °C, followed by 35–40 cycles (30 seconds at 95°C, 1 minute at 56°C, 1 minute at 72°C) and 7 minutes at 72°C. To eliminate heteroduplex products in reaction C, a final 10-minute extension step was performed after adding fresh PCR reagents. Samples were then digested overnight with AluI at 37°C. ³²P-labeled products were denatured at 95°C, separated on 6% polyacrylamide sequencing gels and analyzed using a Phosphorimager (Amersham Biosciences). HEX-labeled products were separated on an ABI sequencer.

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Fig. 3

The Bst mutation. (A) Map of the Rpl24 gene showing the informative PCR, the 4 bp deletion within the intron 1 branchpoint, the first codons of the open reading frame, and the premature stop codon in Bst (red). (B) Sequence chromatograms comparing +/+ and Bst/+ PCR products. The exon 2 splice acceptor (arrowheads) and 4 bp Bst deletion (boxed) are indicated. (C) Allele-specific PCR assay used to distinguish Bst and wild-type Rpl24 alleles. The deletion is unique to the C57BLKS Bst/+ strain. (D) Bst causes abnormal Rpl24 splicing with retention of intron 1. RT-PCR was performed using the indicated primers. A common downstream primer (*) was end-labeled with ³²P using T4 DNA kinase. Product A originates from correctly spliced Rpl24 transcripts, whereas A′ and B indicate inclusion of intron 1. The relatively low abundance of product A′ is due to PCR competition and nonsense-mediated decay. The low level of product B in the wild-type lanes most probably reflects amplification from hnRNA. (E) Some Bst transcripts are correctly spliced. RT-PCR was performed using (Bst/+ × SPRET) F1 RNA and the indicated primers. The upstream primer spans the exon 1–2 junction. Product C (388 bp) was end-labeled and digested with AluI to distinguish between musculus (M, 365 bp) and spretus (S, 135 bp) alleles. The molar ratio of M and S products is 1.27 for +/+ mice and 0.26 for Bst/+ littermates, giving a normalized expression level for Bst of 0.21 (±0.02). (F) Similar fluorometric assay performed using a HEX-labeled reverse primer and ABI sequencer following AluI digestion. The Bst expression level is 0.25 (±0.03) relative to wild-type musculus Rpl24. (G) Sequence of correctly spliced Rpl24 cDNAs amplified from F1 mice, showing the informative AluI site in exon 4 (overlined, sense strand). The normalized ratio of Bst to wild-type peak areas is 0.22 (±0.01).

Rpl24 transgenic analysis

BAC clone RP23-10L10 (accession no. AZ117821) contains the 5.2 kb Rpl24 transcription unit flanked by 93 kb 5′ and 83 kb 3′ genomic DNA. Purified BAC DNA was injected into pronuclei of (C57BL/6 × SJL) F2 eggs, generating 17 transgenic founders. The integrity of the BAC ends was assessed in founders by PCR, using primer pairs that span the vector-insert junctions:

• 10L10 Prox-2

  Forward: 5′-TCGAGCTTGACATTGTAGGACTAT-3′

  Reverse: 5′-AGGGTGGTTTCAGTGTGTCG-3′

• 10L10 Dist-1

  Forward: 5′-GATGTTCATGTTCATGTCTCCTTC-3′

  Reverse: 5′-CCCTCAATCTCATTAACCTTCTGT-3′.

To confirm the presence of Rpl24 within each transgene, we crossed founders to SJL mice and compared the ratio of SJL and C57 alleles in offspring using a Sau3A1 RFLP within intron 4 or a microsatellite marker (D16Ero9) located 1 kb upstream of Rpl24. D16Ero9 was typed using primers 5′-CTGGCTAGCATAGCTTTTCTTTTT-3′ and 5′-GGAAAGGTGAGCAGGTTAAGAATA-3′. To evaluate transgene correction, founders were crossed to C57BLKS Bst/+ mice. The resulting progeny were examined for Bst phenotypes and assessed using BAC-specific and Rpl24 allele-specific PCRs.

The human RPL24 cDNA was amplified by PCR from IMAGE clone 5609137 (Open Biosystems) using an upstream primer with an engineered BamHI site (5′-CGTGGATCCGTCGCCATGAAGGTCGAGCTGTG-3′) and a downstream primer with an engineered EcoRI site (5′-TTTAGAATTCTAATCTGCCAGTTTAGCGTTTTCC-3′). Following digestion, the resulting 0.5 kb BamHI-EcoRI fragment was subcloned into BglII- and EcoRI-digested pBROAD3 (Invivogen). The final construct (R26-huL24) contains the entire 471 bp RPL24 coding sequence, is transcribed from the 1.9 kb murine ROSA26 promoter, and terminates in a β-globin polyadenylation site (Fig. 4D). The 3.0 kb transgene was excised with PacI, gel purified and injected into pronuclei of coisogenic C57BLKS Bst/+ and +/+ eggs. Transgenic mice were identified by PCR using primers 5′-ACAGGTGTGAAACAGGAAGAGAAC-3′ and 5′-GGGAACAAAGGAACCTTTAATAGA-3′, which selectively amplify the human cDNA construct.

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Fig. 4

Transgene correction of the Bst phenotype. (A) BAC clone RP23-10L10 (blue) contains Rpl24 and five other genes, and spans 181 kb (Ensembl mouse genome database, v. 12.3.1). (B) Founder Tg321 carries an intact RP23-10L10 transgene, which suppresses external Bst phenotypes in all doubly heterozygous offspring (compare mice 4514 and 4516 with 4515). Genotypes were determined using BAC end (red) and allele-specific Rpl24 PCRs. (C) Correction data summary. Four BAC transgenic founders were mated to C57BLKS Bst/+ mice. Tg321 and Tg326, which are intact, correct Bst phenotypes but Tg898 or Tg901, which lack Rpl24, do not. (D) Human RPL24 transgene (R26-huL24). The cDNA is ubiquitously expressed using the murine ROSA26 promoter.

Pulse-labeled rRNA analysis

Sex-matched adult littermates were fasted for 48 hours and re-fed for 2 hours. This fasting protocol decreases liver mass twofold (Hutson and Mortimore, 1982; Volarevic et al., 2000). Immediately before re-feeding, each mouse was given an intraperitoneal injection of ³²P-orthophosphate (30 μCi per gram body weight). Livers were snap frozen after re-feeding. Total liver RNA (1–10 μg per lane) was separated by formaldehyde agarose gel electrophoresis, transferred to a nitrocellulose membrane and exposed to X-ray film (Biomax, Kodak). Density tracings were analyzed using NIH Image software (Scion). The membrane was stained with Methylene Blue to confirm equal RNA loading.

Polysome profiles

Livers from sex-matched adult littermates were homogenized in ice-cold hypotonic lysis buffer (1.5 mM KCl, 2.5 mM MgCl₂, 5 mM Tris-HCl pH 7.4, 1% Na deoxycholate, 1% Triton X-100) using 1 ml buffer per 10 mg liver. After centrifugation for 15 minutes at 2500 g, heparin (1 mg/ml) and cycloheximide (100 μg/ml) were added, and 700 μl of the supernatants were applied to 13 ml 7–47% (w/v) sucrose gradients. Samples were centrifuged in a Beckman SW-41 rotor at 288,000 g for 2 hours at 4°C. Gradients were collected using a Biocomp fractionator with UV photometer.

Murine embryonic fibroblasts

Murine embryonic fibroblast (MEF) cultures were generated at E13.5 from co-isogenic C57BLKS Bst/+ and +/+ embryos and from (CAST/Ei × C57BLKS Bst/+) F2 embryos as described (Robertson, 1987) and tested at equivalent early passages (<P8). Experiments were performed in triplicate unless otherwise noted.

To evaluate protein synthesis, MEFs were plated at a density of 5×10⁴ cells per 35 mm dish. After 24 hours, cells were incubated for 2 hours in 2 ml fresh media containing 10 μCi [³H]leucine, washed with ice-cold phosphate buffered saline (PBS), and treated with 5% trichloroacetic acid (TCA) for at least 2 hours at 4°C. [³H]leucine incorporation was measured after lysis in 0.5 N NaOH, 0.5% sodium dodecyl sulphate (SDS) and was normalized to mitochondrial dehydrogenase activity in parallel cultures, measured using a water soluble tetrazolium (WST) assay (Roche).

To measure cell-doubling time, parallel cultures were plated in complete media at a density of 5×10⁴ cells per 35 mm dish. Starting 36 hours after plating, adherent cells were harvested at 12-hour intervals and counted using a Coulter Z2 counter. The growth rate was determined between 36 and 60 hours after plating.

Bivariate cell cycle analysis was performed on asynchronous cultures using a BrdU Flow Kit (BD Pharmingen). Cultures were labeled with 10 μM BrdU for 75 minutes, stained with an FITC-conjugated anti-BrdU antibody (new DNA synthesis) and 7-AAD (total DNA content), and sorted using a Coulter Elite ESP Fluorescence Cell Sorter. The fraction of cells in G1, S and G2/M is directly related to the duration of each phase in the cell cycle (Gray et al., 1990). Flow cytometry data were analyzed using WinMDI v.2.8 software.

To evaluate S-phase entry after serum starvation, parallel MEF cultures were plated in complete media at 5×10⁴ cells per 35 mm dish. After plating, cells were washed twice with serum-free media and re-fed Dulbecco’s modified Eagle’s medium (DMEM) containing 0.5% fetal bovine serum (FBS) for 48 hours. Cultures were released from serum starvation (Herrera et al., 1996) by the addition of complete DMEM containing 15% FBS. DNA synthesis was measured periodically by adding 0.8 μCi [³H]thymidine per dish for 1 hour. These cultures were washed twice with ice-cold PBS and treated with 5% trichloroacetic acid (TCA) for at least 3 hours at 4°C. The cells were washed, lysed in 0.5 N NaOH, 0.5% SDS and counted for [³H]thymidine incorporation. This experiment was repeated using MEFs derived from three separate litters.

Molecular identification of Bst

Bst arose spontaneously in the inbred C57BLKS strain (Southard and Eicher, 1977) and was mapped to a 10 cM interval on mouse chromosome 16 (Rice et al., 1995). To further localize Bst, we performed an intersubspecific backcross between (C57BLKS Bst/+ × CAST/Ei) F1 and C57BLKS+/+ mice. N2 progeny were scored for Bst pigmentation and tail phenotypes and genotyped with informative microsatellite DNA markers. Among 828 N2 progeny, 386 were Bst/+. This segregation ratio (386:442) is less than 1:1 (P=0.092), consistent with decreased viability of Bst/+ mice. Bst maps within a 0.5 cM interval, delimited by D16Mit199 and D16Ero1 (Fig. 2A). Our data formally exclude Opa1, the mouse homolog of the human optic nerve atrophy gene (MIM605290), and the Hes1 basic helix-loop-helix transcription factor, which has roles in retinal and somite development (Saga and Takeda, 2001; Tomita et al., 1996). Both genes lie more than 5cM centromeric to Bst (Fig. 2A).

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Fig. 2

Positional cloning of Bst. (A) Recombination data localizing Bst to a 0.5 cM interval on mouse chromosome 16 (shaded in red) and excluding Opa1 and Hes1. Bst and microsatellite alleles transmitted by F1 parents are indicated together with the number of N2 progeny in each class. The 135 N2 mice typed with the extended marker set (left) are included in the total (right). □, CAST/Ei; ■, C57BLKS. (B) Physical map of critical Bst region derived from Ensembl mouse genome database (3.35 Mb, v. 4.1.1). The map shows informative microsatellite markers, candidate transcription units and relevant BAC clones (blue). The nonrecombinant interval maps within cytogenetic band B4, spans 1.5 Mb (green arrows) and includes Rpl24 (red).

The non-recombinant interval spans 1.5 Mb (Fig. 2B) and contains at least 18 known or predicted genes (Ensembl mouse and UC Santa Cruz human databases, April 2002). Eleven are expressed in E13.5 head and E15.5 eye (data not shown) and were evaluated as Bst candidates by comparing genomic DNA sequences from C57BLKS Bst/+ and +/+ mice. These studies revealed only a single change, within Rpl24 (Fig. 3A,B).

Rpl24 encodes a protein component of the large (60S) ribosomal subunit. L24 is a highly basic protein of 157 amino acids and is conserved among eukaryotes and archaebacteria. Its position near the polypeptide exit and elongation factor binding sites (Ban et al., 2000), its contacts with the 40S subunit (Spahn et al., 2001) and the properties of L24 mutant yeast (Dresios et al., 2000) suggest that L24 helps catalyze the peptidyl transferase reaction. Rpl24 spans six exons, extends over 5.2 kb and contains a characteristic 5′ terminal oligopyrimidine (TOP) initiator element (Hariharan et al., 1989). More than 20 processed Rpl24 pseudogenes are interspersed throughout the genome (data not shown), similar to other riboprotein genes (Zhang et al., 2002). However, only one of these loci, within the Bst critical interval, contains introns and is therefore the bona fide Rpl24 gene.

Rpl24 branchpoint mutation alters mRNA splicing

Bst destroys the first intron splice branchpoint of Rpl24, which matches the consensus sequence (UACUAAC) exactly. The deletion removes four nucleotides, including the critical adenosine (underlined) that reacts to form the lariat intermediate (Fig. 3A,B). No other obvious branchpoints are present within this 108 bp intron. Using an allele-specific PCR assay, we determined that the deletion in Rpl24 is unique to the Bst mice (Fig. 3C). This branchpoint is intact in the parental C57BLKS strain, highly related C57 strains, different Mus species (M. spretus, M. castaneus and M. molossinus) and rat Rpl24 (accession no. X78443).

To determine how Bst affects Rpl24 splicing, we performed RT-PCR experiments (Fig. 3D). Using primers in exons 1 and 5, we amplified a novel 500 bp product from Bst/+ RNA, which reflects inclusion of intron 1. No other products were detected, including a 320 bp product that would result if exon 2 were skipped. This result was confirmed by direct PCR sequencing and a second RT-PCR using primers in intron 1 and exon 5 (Fig. 3D). Rpl24 translation begins in the first exon (Fig. 3A). Retention of intron 1 would cause premature termination of the L24 polypeptide after the first two codons. The first intron contains two other stop codons and the next AUG codon (M91) is in the fourth exon.

Activation of a cryptic branchpoint could restore proper splicing (Ruskin et al., 1985). Because Bst homozygotes are not available, we evaluated Rpl24 splicing in interspecies hybrid mice, derived from a (C57BLKS Bst/+ × CAST/Ei) F1 × SPRET/Ei cross, by a PCR strategy, using a polymorphic AluI restriction site in exon 4 and an upstream primer spanning the exon 1–2 junction. This allowed us to resolve and compare the amount of correctly spliced Rpl24 transcripts from wild-type and Bst alleles (Fig. 3E–G). These results show that 20–25% of Bst transcripts are correctly spliced. Bst is therefore hypomorphic. Inbred C57BLKS Bst heterozygotes and homozygotes thus have approximately 60% and 20% of normal Rpl24 transcript levels, respectively.

Rpl24 BAC and cDNA transgenes correct Bst

Like the Drosophila Minutes, Bst has general effects on somatic growth and specific effects on different tissues (Fig. 1). To confirm that the Rpl24 mutation is solely responsible for the pleiotropic Bst phenotypes, we created transgenic mouse lines expressing wild-type Rpl24 from a 181 kb mouse BAC (RP23-10L10) or a human cDNA expression cassette (Fig. 4A,D). Four BAC transgenic lines were bred to C57BLKS Bst/+ mice (Fig. 4C). Founders Tg321 and Tg326 have intact transgenes that completely correct the Bst phenotypes (Fig. 4B,C). Founders Tg898 and Tg901 have partial BAC transgenes lacking Rpl24 and these fail to correct the Bst phenotype. We generated cDNA transgenics using human RPL24 and the ubiquitously active murine ROSA26 promoter (Zambrowicz et al., 1997). Human and mouse L24 proteins are identical. We injected co-isogenic C57BLKS Bst/+ and +/+ eggs with the R26-huL24 construct (Fig. 4D) and recovered seven transgenic founders. Three of the four Bst/+ founders were indistinguishable from wild-type littermates (data not shown), proving that RPL24 alone is sufficient to correct the Bst phenotype and that Rpl24 is Bst. Taken together, our data show Bst acts via a partial loss-of-function mechanism and should be considered a mouse Minute.

Bst impairs ribosome biogenesis

Ribosome biogenesis requires highly coordinated synthesis of ribosomal proteins and rRNAs (Warner, 1999). These are assembled in a tightly ordered process involving more than 200 accessory proteins and small nucleolar ribonucleoproteins (Fatica and Tollervey, 2002). Decreased availability of individual components can impair these events (Saveanu et al., 2001; Volarevic et al., 2000; Zhao et al., 2003).

Although large and small subunits assemble independently, transcription and processing of the 45S rRNA precursor into 18S and 28S rRNAs couples formation of both subunits (Fatica and Tollervey, 2002). As such, rRNA processing is a sensitive indicator of ribosome biogenesis. We therefore examined rRNA synthesis in a pulse labeling experiment. Sex-matched co-isogenic C57BLKS Bst/+ and wild-type littermates were fasted for 48 hours, injected with ³²P-orthophosphate and re-fed for 2 hours. Profiles of unlabeled (steady state) rRNA were similar after re-feeding (Fig. 5A, Methylene Blue), but the amount of newly processed rRNA in the Bst/+ mouse was dramatically decreased (Fig. 5A, autoradiogram). The 28S rRNA was reduced to a greater extent than the 18S rRNA, consistent with a defect in large subunit assembly (Fig. 5A, densitometry). In yeast, mutations specific to the 60S subunit can affect 18S rRNA processing indirectly, most probably through the exosome (Zanchin et al., 1997). This nuclease complex degrades improperly spliced mRNAs, excess rRNA precursors and unused rRNA intermediates, and is required for rRNA processing (Allmang et al., 2000; Butler, 2002; Mitrovich and Anderson, 2000). Likewise, ribosome subunits are stable once assembled and most unincorporated riboproteins are rapidly eliminated (Warner, 1977). Bst is therefore unlikely to affect the abundance of L24 at steady state. The labeling of the 34S and 45S pre-rRNAs was similar in both mice (Fig. 5A, autoradiogram), suggesting that the decrease in 28S rRNA is due to increased turnover rather than a specific bottleneck in rRNA processing. This cellular response is consistent with a dynamic shortage of L24 polypeptide.

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Fig. 5

Bst impairs ribosome biogenesis. (A) Bst/+ causes a decrease in processed rRNAs. C57BLKS Bst/+ and +/+ littermates were fasted for 48 hours and re-fed for 2 hours. Tissue RNAs were radiolabeled by an intraperitoneal ³²P injection at the start of the re-feeding period. (left) Autoradiogram of pulse-labeled liver RNA (5 μg/lane). Mature 18S and 28S rRNAs and 34S and 45S precursors are indicated. (center) Methylene Blue stain of the same filter, showing total RNA. (right) Density tracing of autoradiogram showing that Bst has a greater effect on 28S rRNA. The Bst/+ profile is scaled twofold to equalize 18S rRNA levels. The relative peak areas measured for 45S, 34S, 28S and 18S rRNAs are 0.9, 1.1, 5.9 and 4.2 (+/+) and 0.9, 1.1, 1.5 and 1.8 (Bst/+), respectively. (B) Sucrose gradients of liver homogenates from normally fed +/+ and Bst/+ adult littermates. UV absorbance peaks corresponding to ribosomal subunits and polysomes are labeled. A prominent shoulder is present between the 60S and 80S subunits (arrowhead).

There were no changes in the ratio of small to large subunits or in the polyribosome fraction of Bst/+ liver homogenates (Fig. 5C). However, a prominent shoulder between the 60S subunit and 80S monosome peaks was consistently observed in sucrose gradients from Bst/+ mice, compatible with an alteration in 60S subunit structure. This change was observed under fed, fasted and re-fed conditions (data not shown).

Bst/+ fibroblasts have altered properties

In spite of its apparent tissue specificity, L24 deficiency should affect every cell in the Bst/+ mouse, in the same way that cell growth is universally slower in Minute flies (Simpson and Morata, 1981). To test this hypothesis, we examined cell size, protein synthesis, proliferation rates and cycle kinetics in exponentially growing mouse embryonic fibroblast (MEF) cultures. We found no difference in size between Bst/+ and wild-type MEFs by flow cytometry (forward light scatter, data not shown). This is consistent with recent studies in Drosophila showing no size difference between Minute and wild-type imaginal disc cells (Montagne et al., 1999). We examined overall protein synthesis in MEFs by [³H]leucine incorporation, normalized to cell viability (Fig. 6A). Bst/+ cultures showed a 31.2±5.8% reduction in the rate of protein synthesis, similar to the 30% reduction described for M(3)i⁵⁵/+ Drosophila Minute embryos (Boring et al., 1989).

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Fig. 6

Cellular effects of the Bst mutation. (A) Protein synthesis ([³H]leu incorporation) is decreased in Bst/+ MEFs (P<0.001, two-tailed unpaired t-test). (B) Growth rates of subconfluent +/+ and Bst/+ MEF cultures (cell counts), showing decreased doubling time of Bst/+ cells (P<0.001, two-tailed unpaired t-test). (C) Bivariate cell cycle analysis. Cells were sorted by total DNA content (7-AAD fluorescence) and BrdU incorporation (FITC-conjugated antibody). Density plots represent combined data from two experiments (20,000 cells each). The histogram shows the proportion of cells in G1, S and G2/M sectors (red boxes); bars indicate the range for two experiments. The distribution of cells in G1, S and G2/M differ significantly between Bst/+ and +/+ (P<0.001, χ²=233 for 2×3 contingency table). (D) S-phase entry of Bst/+ MEFs is delayed following 48 hour serum starvation. [³H]thymidine incorporation was normalized for each genotype to the highest level measured during the experiment. The panel shows one of three independent experiments performed. Each point shows the average of three cultures; bars indicate standard deviation. (E) Model for increased cell cycle length, based on bivariate FACS analysis and doubling times. Assuming the length of S phase is constant, Bst/+ MEFs remain significantly longer in G1 before reaching the restriction point (R).

Bst/+ MEFs proliferate significantly more slowly than wild-type cultures. The population doubling time (T_d) is 25.8±0.4 hours for Bst/+ MEFs compared with 21.6±0.2 hours for wild-type MEFs (Fig. 6B). To determine what part of the cell cycle is elongated, we performed bivariate flow cytometry on asynchronous MEF cultures using BrdU incorporation and DNA content as sorting parameters. The ratio of G1 to S-phase cells was 1.45 (±0.05) for Bst/+ MEFs and 1.04 (±0.01) for control MEFs (Fig. 6C). This suggests that Bst causes actively growing cells to remain longer in G1 phase. A small increase in the G2/M fraction was also observed. If the absolute length of S phase is constant (9.1 hours) and the cell cycle is increased by 20% overall (Fig. 6B), then G1 phase is increased by 40% in Bst/+ MEFs (Fig. 6E).

To further investigate the prolongation of G1 by Bst, we compared the time required for Bst/+ MEF cultures to enter S phase after serum stimulation. Parallel cultures were arrested in G1 by serum starvation, released by the addition of serum-rich media, and assayed periodically for DNA synthesis by [³H]thymidine incorporation. We observed a 2.4±0.5 hour increase in the time required to reach half-maximal incorporation in Bst/+ cultures compared with wild type (Fig. 6D). These results confirm the increased length of G1 in Bst/+ MEFs and are consistent with classical studies in which the protein synthesis inhibitor cycloheximide was shown to specifically lengthen G1 phase following serum stimulation (Brooks, 1977).

Paucity of mammalian riboprotein mutations

Riboprotein mutations appear to be rare in mammals, in contrast to bacteria, yeast and Drosophila. The paucity of mutations can be explained if heterozygous phenotypes are too mild, too severe, heterogeneous or otherwise poorly ascertained. The fact that Bst is hypomorphic suggests that a threshold level of gene expression might be required for survival. Null mutations may be lethal or subvital. Likewise, ribosomes may tolerate mutations in few of their components. L24 does not exist in eubacteria and is one of seven dispensable riboproteins in yeast (Baronas-Lowell and Warner, 1990). Yeast lacking L24 have an increased doubling time and a reduced peptidyl transferase rate (Baronas-Lowell and Warner, 1990; Dresios et al., 2000). These observations suggest that L24 enhances translation efficiency but does not have a unique structural role. Indeed, L24 is added to the 60S subunit relatively late, most probably during export through the nuclear pore or in perinuclear cytoplasm (Harnpicharnchai et al., 2001; Saveanu et al., 2001). A special case of redundancy might also exist for L24. An evolutionarily conserved paralog, Rlp24 (ribosomal-like protein 24), associates with pre-60S complexes (Saveanu et al., 2001). Mouse Rlp24 and L24 are 31% identical and equally similar to the archaean L24 homolog. It has been proposed that Rlp24 plays an essential role in ribosome biogenesis and that Rlp24 and L24 sequentially occupy the same docking site on the 60S subunit, in the nucleus and cytoplasm, respectively (Saveanu et al., 2001). If Rlp24 can reach the cytoplasm, it might in principle partially substitute for L24 in the mature ribosome. Alternatively, a large number of unrecognized riboprotein mutations may exist in mice as they do in Drosophila, with semidominant spotting, tail defects and small size as core phenotypes, and homozygous lethality (Hrabe de Angelis and Balling, 1998; Morgan, 1950; Tease and Fisher, 1993).

Bst phenotypes

Since Rpl24 is essential for protein translation, Bst homozygotes are likely to die at or before the blastocyst stage, when the pool of maternal ribosomes is exhausted (Copp, 1995), or slightly later, owing to low-level Rpl24 expression from the Bst allele (Fig. 4E-G). In a normal mouse blastocyst, ≤20% of the 2.5×10⁸ ribosomes originate from the oocyte (Piko and Clegg, 1982) and ribosomal proteins account for 8% of total protein synthesis (LaMarca and Wassarman, 1979). By contrast, 0-nu Xenopus mutants lacking rRNA genes can survive to the tadpole stage using maternal ribosomes only (Brown and Gurdon, 1964).

The specific defects in Bst/+ mice (Fig. 1) are difficult to reconcile with a global reduction in protein synthesis. Developing retinas, somites and melanocytes may be hypersensitive because mRNAs critical for these tissues are translated at or near a required threshold. Indeed, different mRNAs are translated with widely varying rates of initiation and elongation (Dever, 2002; Lodish, 1976). However, Bst phenotypes may be more easily explained by tissue differences in (1) the extent of coupling between cell division and organogenesis, (2) levels of riboprotein gene expression or (3) extraribosomal functions.

The cell cycle alterations in Bst/+ mice (Fig. 6) provide the basis for the first model. In Drosophila, mutations in the promitotic dmyc gene are phenotypically similar to Minutes (Johnston et al., 1999). Mice with graded c-myc alleles are likewise small, have short tails and prolonged cell cycles (Trumpp et al., 2001). Bst phenotypes could arise in tissues in which the timecourse of organogenesis is relatively uncoupled from cell proliferation. During the life cycle of Minute flies, delayed eclosion allows for complete development. In mice, however, gestation cannot be prolonged; Bst/+ pups are born at the same time as wild-type littermates. Organogenesis must proceed within the time allotted to ensure viability and must be relatively uncoupled from cell proliferation. In principle, defects could result from situations in which development outpaces cell proliferation. Such a dyschronic mechanism may explain the phenotypes in the retina and vertebral column, where timing of developmental events is particularly critical.

In the case of the developing retina, a single progenitor population gives rise to seven major cell types in a characteristic order (Turner et al., 1990). The progenitors are thought to proceed through a sequence of competency states, each of which favors the differentiation of one or a few particular cell types (Livesey and Cepko, 2001). RGCs are the first-born retinal neurons in all vertebrate species (Altshuler et al., 1991). Although histogenesis can occur when cell division is blocked (Harris and Hartenstein, 1991), the timing of cell cycle exit greatly influences retinal development and progenitor cell fate (Dyer and Cepko, 2001). Furthermore, cell cycle length is significantly modulated during the normal course of retinal histogenesis (Alexiades and Cepko, 1996; Li et al., 2000). In the Bst/+ retina, there is no change in the fraction of cells undergoing proliferation or apoptosis, but the total number of neuroblasts is reduced by roughly 60% (E10.5) to 20% (E13.5), and the onset of terminal mitoses is delayed from E10.5 to E12.5 (Tang et al., 1999). These findings can now be considered in light of the general increase in Bst/+ cell-cycle length (Fig. 6). Assuming our MEF results apply in vivo, Bst is likely to have a disproportionately greater effect during the early stages of retinal histogenesis, when the cycle time is shorter (Alexiades and Cepko, 1996), protein synthetic demands are greatest, and RGCs are born (Livesey and Cepko, 2001). The subretinal neovascularization (Smith et al., 2000) is probably a secondary consequence of RGC deficiency, similar to that observed in Math5 mutant mice and humans with optic nerve aplasia (Brown et al., 2001; Lee et al., 1996; Brzezinski et al., 2003).

Timing is similarly critical during somite development. Somites are added iteratively to the caudal end of the neural tube, in response to periodic waves of gene expression (Saga and Takeda, 2001). This process is controlled by an intrinsic clock that is sensitive to cycloheximide (Hirata et al., 2002; Palmeirim et al., 1997). The vertebral defects in Bst/+ mice are more severe posteriorly, consistent with a progressive deterioration in periodicity and phase coherence of the intrinsic somite clock (Saga and Takeda, 2001). This pattern could occur if high levels of the protein synthesis were needed to propagate successive cycles of somitogenesis.

In the second model, pleiotropic Bst phenotypes may reflect tissue-specific differences in Rpl24 expression. To achieve unimolar stoichiometry, dispersed riboprotein genes are coordinately expressed via 5′TOP elements, which define transcriptional start sites and control translation of riboprotein mRNAs (Hariharan et al., 1989; Jefferies et al., 1997). In spite of this convergently evolved feature, 5′TOP sequences may vary in potency across tissues. Differential regulation of riboprotein mRNAs has been described in pluripotent embryonal carcinoma cells, various cancer cell lines and in RNA profiling studies (Blackshaw et al., 2001; Panda et al., 2002). If the developing retina, somites and melanoblasts express relatively low levels of Rpl24 mRNA, they may be more sensitive to quantitative reduction caused by the Bst mutation.

In the third model, Bst phenotypes may occur because L24 has unique extraribosomal functions, apart from protein translation (Wool, 1996). For example, the small subunit riboprotein S3 functions as an endonuclease during DNA repair, while S0 acts on the cell surface, as the primary laminin receptor (Ardini et al., 1998). This explanation would be supported if mice with mutations in other riboprotein genes have phenotypes unlike Bst. The high incidence of RPS19 mutations among DBA patients (Draptchinskaia et al., 1999) suggests that S19 has extraribosomal functions in erythropoiesis. Recent studies show that L24 from a variety of species interacts directly with the transactivator (TAV) protein of cauliflower mosaic virus to promote translation re-initiation on polycistronic mRNAs (Park et al., 2001). In principle, the unique Bst phenotypes could reflect tissue specificity of cellular mRNAs that utilize internal ribosome entry (Hinnebusch, 1997; Johannes et al., 1999; Morris and Geballe, 2000).

We thank Margaret Van Keuren, Elizabeth Hughes, Donalyn Scheuner, Mark Adams, Deanna Kulpa, Stephanie Schmoll, Bob Lyons, Dave Siemieniak, Sean Morrison, Ömer Yilmaz, Phil Andrews, John Strahler, Sinisa Volarevic, Joe Brzezinski and Sara Schulz for technical advice; Elizabeth Robertson and Philippe Soriano for providing ES cells; and David Ginsburg, Sally Camper, Dave Turner, John Moran, Miriam Meisler, Randy Kaufman and Steve Weiss for helpful discussions and critically reading the manuscript. E.R.O. was supported by MSTP and UM Organogenesis NIH training grants (T32 GM07863 and HD07505). This work was supported by NIH grants to T.G. (EY14259, EY12994 and GM067840) and the UM Diabetes Research and Training Center (DK20572).