- Open Access
Whole-genome screening indicates a possible burst of formation of processed pseudogenes and Alu repeats by particular L1 subfamilies in ancestral primates
© Ohshima et al.; licensee BioMed Central Ltd. 2003
- Received: 22 July 2003
- Accepted: 25 September 2003
- Published: 28 October 2003
Abundant pseudogenes are a feature of mammalian genomes. Processed pseudogenes (PPs) are reverse transcribed from mRNAs. Recent molecular biological studies show that mammalian long interspersed element 1 (L1)-encoded proteins may have been involved in PP reverse transcription. Here, we present the first comprehensive analysis of human PPs using all known human genes as queries.
The human genome was queried and 3,664 candidate PPs were identified. The most abundant were copies of genes encoding keratin 18, glyceraldehyde-3-phosphate dehydrogenase and ribosomal protein L21. A simple method was developed to estimate the level of nucleotide substitutions (and therefore the age) of PPs. A Poisson-like age distribution was obtained with a mean age close to that of the Alu repeats, the predominant human short interspersed elements. These data suggest a nearly simultaneous burst of PP and Alu formation in the genomes of ancestral primates. The peak period of amplification of these two distinct retrotransposons was estimated to be 40-50 million years ago. Concordant amplification of certain L1 subfamilies with PPs and Alus was observed.
We suggest that a burst of formation of PPs and Alus occurred in the genome of ancestral primates. One possible mechanism is that proteins encoded by members of particular L1 subfamilies acquired an enhanced ability to recognize cytosolic RNAs in trans.
- Additional Data File
- Mismatch Rate
- Neutral Mutation Rate
- Human Genome Draft Sequence
- Respective Subfamily
The abundance of pseudogenes is a remarkable feature of mammalian genomes. Aptly named, pseudogenes are copies of specific genes and are present in every mammalian chromosome [1–5]. In general, pseudogenes are thought to be nonfunctional  as they have accumulated vast numbers of mutations during evolution and have lost the ability to be transcribed. Pseudogenes fall into two distinct categories depending on the mechanism by which they are generated: processed pseudogenes (PPs) are reverse transcribed from mRNAs (and thus do not contain introns) whereas nonprocessed pseudogenes arise from duplications of genomic DNA [2, 4]. Among the abundant PPs, there are a substantial number of 'processed genes' or 'retrogenes' of novel function that also derive from mRNAs of various intron-containing genes [6–8].
In addition to PPs, mammalian genomes contain a large number of retrotransposons (retroposons) that represent a reverse flow of genetic information via RNA [9–13]. In humans, short interspersed elements (SINEs) and long interspersed elements (LINEs) occupy over 30% of the genome . Progress in LINE1 (L1) molecular biology has enabled L1 'retrotransposition' studies in cultured HeLa cells [15, 16]. Recent work [17–21] shows that mammalian L1-encoded proteins may have been involved in the reverse transcription of PP and Alu [22–26]. Furthermore, L1-encoded proteins predominantly mobilize the RNA in which they are encoded [18, 19]. This so-called 'cis preference' explains the fact that, among the overwhelming number of nonfunctional L1 RNAs, recent mutagenic L1 insertions in humans and mice are derived from a progenitor L1 RNA that contained intact open reading frames (ORFs) . In fact, Moran's group estimated that a functional L1 mobilizes nonfunctional L1 RNAs and other cellular mRNAs in trans at frequencies of only 0.2%-0.9% and 0.01%-0.05%, respectively, relative to processes involving cis RNA . This finding also raised the question of how human Alu repeats could have been amplified in trans to their present level of approximately 10% of the human genome, given that L1-encoded proteins preferentially mobilize their own transcripts. Boeke proposed that Alu RNA secondary structure could have positioned this RNA on the ribosome in a manner that promoted effective interactions with L1-encoded proteins [21, 27].
The initial analysis of the human genome draft sequence by the International Human Genome Sequencing Consortium provided the first comprehensive view of retroposons such as LINEs and SINEs, although the description of PPs was largely ignored . The Celera report briefly described a preliminary analysis of PPs . Here, we present the first comprehensive analysis of human PPs using all known human genes as queries. These PPs were derived from 6% of all annotated human genes, and our data suggest a possible burst of PP genesis early in primate evolution.
Whole-genome screening for human PPs and their content
Processed pseudogene content of the human genome
Genes that generated PPs
Ribosomal proteins (mitochondrial)
Zinc finger protein‡
Ring finger protein‡
DEAD/H box polypeptide
Total annotated genes
Hypothetical genes §
The most abundant PPs in the human genome
Keratin 18 (KRT18)
Glyceraldehyde-3-phosphate dehydrogenase (GAPD)
Ribosomal protein L21 (RPL21)
Ribosomal protein L31 (RPL31)
Ribosomal protein L34 (RPL34)
Ribosomal protein S15a (RPS15A)
Ribosomal protein L6 (RPL6)
Ribosomal protein L5 (RPL5)
High-mobility group box 1 (HMGB1)
Ribosomal protein S29 (RPS29)
Ribosomal protein S24 (RPS24)
Ribosomal protein S20 (RPS20)
Ribosomal protein L19 (RPL19)
Ribosomal protein S12 (RPS12)
Argininosuccinate synthetase (ASS)
Heterogeneous nuclear ribonucleoprotein C (C1/C2) (HNRPC)
Ribosomal protein S18 (RPS18)
Calponin 2 (CNN2)
Ribosomal protein L27 (RPL27)
Ribosomal protein S6 (RPS6)
Ribosomal protein, large, P1 (RPLP1)
Ribosomal protein L10a (RPL10A)
Tumor protein, translationally-controlled 1 (TPT1)
Heat shock 70 kDa protein 8 (HSPA8)
Ribosomal protein S4, X-linked (RPS4X)
Ribosomal protein S17 (RPS17)
IMP (inosine monophosphate) dehydrogenase 1 (IMPDH1)
Ribosomal protein L24 (RPL24)
Ribosomal protein L3 (RPL3)
Ribosomal protein S5 (RPS5)
Ribosomal protein S13 (RPS13)
Ubiquitin A-52 residue ribosomal protein fusion product 1 (UBA52)
Ribosomal protein S15 (RPS15)
Hypothetical protein MGC4276 similar to CG8198 (MGC4276)
Voltage-dependent anion channel 1 (VDAC1)
Ribosomal protein S25 (RPS25)
Hypothetical protein MGC3062 (MGC3062)
Eukaryotic translation initiation factor 2, subunit 2 beta (EIF2S2)
Glutamic-oxaloacetic transaminase 2, mitochondrial (GOT2)
CGI-35 protein (CGI-35)
NADH dehydrogenase 1 alpha subcomplex, 5, 13 kDa (NDUFA5)
Actin-related protein 2/3 complex, subunit 3, 21 kDa (ARPC3)
TATA box binding protein (TBP)-associated factor, 32 kDa (TAF9)
Lactate dehydrogenase A (LDHA)
Phosphoribosylaminoimidazole carboxylase, (PAICS)
Hypothetical protein FLJ14668 (FLJ14668)
ATP synthase, mitochondrial F0 complex, subunit b (ATP5F1)
Ribosomal protein L22 (RPL22)
Solute carrier family 25, member 5 (SLC25A5)
PHD finger protein 5A (PHF5A)
As shown in Figure 1a and 1b, structural-protein PPs constitute the largest class (34%). The 50 most prolific PP parental genes include 25 ribosomal protein genes (Table 2) which contribute substantially to the high incidence of structural proteins among the total number of PPs presented in Figure 1b.
GC content in PP parental genes
Chromosomal distribution of human PPs
Chromosomal distribution and density of human PPs
Genes that generated PPs
Number of genes (Ensembl 4.28.1)
A simple method for estimating the level of nucleotide substitutions in PPs
To approximate the age of each PP, we developed a method for estimating the level of nucleotide substitutions relative to the parental gene. Initially, this method corrected for the sequence divergence value (a consequence of nucleotide-substitution processes) by removing the contribution of mutations at CpG sites. The C-to-T transition rate in CpG pairs is around 12-fold higher than the rate for other transitions  and causes distortions when comparing different genomic elements of high (for example, Alus) or low (for example, L1s) CpG content. Assuming that CpG frequency (ι) in a genomic element that was generated by duplication of a functional gene of high CpG content decreases over time (t) and reaches a state of equilibrium (ε) (approximately 20% of the frequency [14, 33] expected from the local fraction of cytosines and guanosines ), the time since the duplication (T) was calculated (see Materials and methods) from the given sequence divergence (D) and the neutral mutation rate (μ) of primates:
D = ∫0Tμ(1 + 11((ι - ε)/((0.01ι/(0.99ι-ε))t + 1) + ε))dt
Next, the quantity Σ (=μT) was corrected for multiple substitutions at the same site using the Jukes-Cantor model , giving the average number of substitutions per 100 base-pairs (bp), (K). For PPs, sequence divergences were defined as the mismatch rates of respective PPs relative to the current parental gene sequences. Finally, the levels of substitution that accumulated only in PPs were estimated (see Materials and methods). The estimated levels of substitutions in PPs (K(ψ)) were then calculated as K(ψ) = 0.705 K.
Simultaneous burst of processed pseudogenes and Alu repeats in ancestral primates
The above results are reminiscent of the amplification profile of Alu repeats. Alu elements comprise approximately 10% of the human genome  and are restricted to primates [22–26]. It has been proposed that the average age of Alu repeats is around 40 million years and that the majority of Alus were generated around this time [14, 22, 23, 26]. We confirmed these previous results by re-estimating the age distribution of all human Alu repeats (Figure 5b). The Alus also showed a Poisson-like distribution with a sharp peak. Alus are classified into distinct subfamilies that can be identified on the basis of mutations shared among subfamily members [23, 26]. Alu subfamilies were derived from a small number of source or master genes. Accordingly, a consensus sequence constructed from members of each subfamily represents each subfamily's source gene(s) [23, 26]. To evaluate the contribution of each subfamily to the entire distribution of Alus, we estimated the age distribution of respective Alu subfamilies (Figure 5c). The peaks for respective subfamilies are grouped closely, and the subfamily Alu Sx strongly influences the overall distribution of Alus (compare with Figure 5b). Therefore, the Sx subfamily (and thus Alus in general) appears to have been amplified intensively over a relatively short period. To the best of our knowledge, many previous discussions of Alu amplification reflect this viewpoint of Alu evolution [14, 22, 23, 26]. However, our results show that the intensive generation of two distinct elements, PPs and Alus, occurred almost simultaneously suggesting that an unknown change in either the cellular environment or the proliferation mechanism itself enhanced the proliferation of such retroposons in ancestral primates 40-50 million years ago.
Concordant amplification of certain LINE1 subfamilies with PPs and Alus
Recent progress in L1 biology shows that mammalian L1-encoded proteins are likely to have been involved in the reverse transcription of Alus and PPs [17–21]. To elucidate the cause of the elevated retrotransposition of PPs and Alus, we analyzed the age distribution of all human L1s (Figure 5d). Curiously, the rate of amplification (retrotransposition in cells and fixation within a population) of L1s does not peak around 7%, as was the case for PPs and Alus (compare with Figure 5a,b), raising the issue of how the rate of PP/Alu retrotransposition became elevated during a period of moderate change in L1 retrotransposition. To address this problem, L1s were divided into around 80 subfamilies , and age distributions for representative subfamilies are shown in Figure 5e. Although the distributions of respective subfamilies overlap, each subfamily has emerged successively during approximately 150 million years of mammalian evolution. Merging the distribution profiles of all the L1s yields a curve that is rather flat (almost equal to the curve that connects the apices of the respective bars in Figure 5d). Among a large number of L1 subfamilies, certain subfamilies, namely L1PA6, L1PA7 and L1PA8, were amplified intensively around 47 million years ago (the time corresponding to the 7% score). These data suggest that only one or a few L1 subfamilies may have contributed to the increased level of Alu and PP amplification (see Discussion).
Possible mechanisms of a 'retrotranspositional explosion'
A recent extensive survey of the human genome revealed a large number of ribosomal protein pseudogenes derived from the 79 functional ribosomal protein genes . The discussion of the ages of these pseudogenes is problematic, however, in that ages were calculated by simply dividing sequence divergences by mutation rate. As the sequence divergence of a PP relative to its parental gene (K) is dependent on substitutions in both the PP (K(ψ)) and the gene (K(f)) (see Materials and methods), the ages of the ribosomal protein pseudogenes were overestimated. For example, with respect to RPL21 mRNAs (the most predominant source of ribosomal protein pseudogenes in humans), the sequence divergence between human and mouse or rat is approximately 11% (NM_000982, NM_019647, NM_053330, and ). The previous ribosomal protein pseudogene calculations dismissed sequence divergences between the present-day and primordial genes, probably overestimating the ages by around 10 million years (a few percent per 8-10% of divergence). Therefore, it is difficult to compare such values with the ages of Alus/L1s. Our method provides a clear solution to this matter, enabling us to compare the ages of different classes of retroposons. Hence, our method led us to the finding that there was a simultaneous burst of PPs and Alus - a 'retrotranspositional explosion' - in the primate genome.
Regarding the cause of the retrotranspositional explosion, it is worth considering the effect of a 'bottleneck' [34, 41] during primate evolution. Only individuals that experienced extensive genomic retrotransposition might have propagated to become a majority within a population of ancestral primates, via a mechanism involving a rapid reduction in the general population. Studies on the molecular phylogeny and demographic history of humans show, however, that the primate lineage leading to humans never experienced an extensive bottleneck, at least since its divergence from the prosimian lineage . Therefore, the effect of a bottleneck can be largely ignored.
The retrotranspositional explosion could be due to a change in the cellular environment of ancestral primates 40-50 million years ago, such as a higher transcriptional potential of parental (master) genes of PPs and Alus. A specific environment of the genome during the period of the retrotranspositional explosion, such as more available target sites of PPs and Alus, might have facilitated this event. Alternatively, a change in the proliferation mechanism of PPs and Alus, such as an increased amount of reverse transcriptase or an enhanced activity of enzymes for retrotransposition, might have promoted the explosion.
Recent studies on the L1 retrotransposons show that mammalian L1-encoded proteins may have been involved in the reverse transcription of Alus and PPs [17–21]. Here, we have shown that the intensive amplification of distinct genetic elements, namely PPs and Alus, seems to have occurred almost simultaneously around 40-50 million years ago, and suggests that only one or a few L1 subfamilies may have contributed to the observed high levels of Alu/PP retrotransposition.
How could a specific L1 subfamily (or subfamilies) have generated Alus and PPs at such an accelerated rate? We propose that L1s within specific subfamilies mobilized RNAs in trans at accelerated rates in ancestral primate genomes. Thus, a specific L1 subfamily may have mediated the Alu/PP retrotranspositional explosion. The age distributions estimated in this study allow the prediction of the most probable L1 subfamilies responsible for the explosion (care must be exercised when comparing ages between distinct genetic elements; see Materials and methods). The most probable candidate subfamilies are L1PA6, L1PA7 and L1PA8 (Figure 5e). As mentioned above, although the youngest L1 subfamily mobilizes cellular RNAs in trans at very low frequencies (0.01-0.05%) in HeLa cells, the frequency is not necessarily intrinsic to L1s. In fact, in cultured feline cells the frequency of L1-mediated PP formation in trans is 5% relative to that of L1 retrotransposition in cis . Moreover, an eel LINE family exhibits a high level of trans retrotransposition (up to 30% ), and the frequency of L1-mediated Alu retrotransposition in HeLa cells is 100-1,000 times higher than control mRNAs . Although L1 subfamilies such as L1PA6, L1PA7 and L1PA8 appear to have been extinguished by cumulative mutations, the possibility that an ancient L1 subfamily exhibited an enhanced ability to mobilize RNAs in trans could be verified experimentally in HeLa cells using reconstructed L1 subfamilies  as sources for reverse transcription of trans RNAs.
The impact of the retrotranspositional explosion on the ancestral primate genome
Alu insertions mediate many genomic rearrangements, such as unequal crossing over, induction of alternative splicing, and the introduction of new promoters, poly(A) signals and even new exons . Inactivation of CMP-N-acetylneuraminic acid hydroxylase (around 2.8 million years ago) before brain expansion during human evolution occurred by an Alu-mediated inactivating mutation , representing yet another example of the impact of the Alu expansion. The current frequency of human endogenous insertional mutations caused by Alu retrotransposition is estimated at around 1 in every 16-200 individuals [26, 46]. The frequency of Alu insertion at the time of the retrotranspositional explosion is estimated to have been 30-200 times higher than the frequency over the last 10 million years ( and data not shown). This implies that at least one in seven individuals at the time carried new Alu insertions in their genomes (a maximum of 12 insertions per individual). This high Alu insertion rate may have had a much greater impact on ancestral primate genomes compared with the impact of present-day mutations.
Retrotransposition of PPs causes not only insertional mutations but also the propagation of new genes. These 'retrogenes' comprise PPs that inserted themselves next to resident promoter/enhancer elements and thereby escaped transcriptional silencing and PPs that were initially inactive but were reactivated at a later time when flanking regulatory elements became activated by mutation . Retrogenes are often observed in primate genomes , one example being the testis-specific human gene CDY (on the Y chromosome), which arose during primate evolution by retrotransposition of the ubiquitous mRNA of the gene CDYL located on chromosome 13 . From the observed distribution of CDY homologs in primates, this event appears to have occurred in the simian lineage after its divergence from prosimians but before the split between Old and New World monkeys  during the period of the retrotranspositional explosion. We predict that further studies will demonstrate that many human retrogenes were generated during this period, and postulate that such retrogenes were involved in generating new characteristics that are specific to simian primates [8, 47].
Determining a set of processed pseudogenes
PPs were searched for in an assembled human genome sequence (Human Genome Project Working Draft, April 1 2001)  using BLAT . The BLAT setting was as follows: Assembly: April 1, 2001; Query type: DNA; Sort output: query, score; Output type: hyperlink. 'Confirmed cDNAs' (23,929 entries) in Ensembl DB (v1.1.0)  were used as queries. The subject with the highest score was regarded as the gene encoding the transcript. Multiple hits were subjected to analysis.
Subjects that contained over 90% of the query length were used. The number of aligning blocks, which usually corresponds to the number of exons, was compared between a gene and other subjects. If the number of aligning blocks was smaller than that of the gene, the subject was further analyzed, thus eliminating pseudogenes generated by DNA duplications. Subjects that were identified by intronless genes (single exon genes) were not included in the analysis to avoid confusing PPs and pseudogenes generated by DNA duplications. To avoid confusing phylogenetic relationships, loci (subjects) that were identified by multiple query hits were not included in the analysis. A series of Perl scripts were designed to analyze the BLAT search results.
Evaluating PP annotation
To evaluate our annotation of PPs, our results for chromosomes 21 and 22 were compared with those from other studies. For chromosome 21, the PP total in this study was 34 whereas previous studies reported 41 [52, 53] and 57 . The number of annotations common to two studies totaled 18 (this study and ), 14 (this study and ) and 21 [4, 52]. Annotations common to all studies totaled 10. For chromosome 22, the PP total in this study was 62, whereas previous studies reported 91 [54, 55] and 73 . The number of common annotations totaled 37 (this study and ), 28 (this study and ) and 52 [4, 54]. Annotations common to all studies totaled 27. Differences between the numbers appear to derive mainly from differences in the gene sets used for the analyses .
Identification of Alus, L1s, and their subfamilies
For each Alu and L1 repeat, the genomic location and sequence divergence was obtained from the output file of the RepeatMasker program applied to the human genome draft sequence (22 December 2001 ). Sequence divergences were defined as the mismatch rates of respective repeats relative to the consensus sequence of respective subfamilies.
Analysis of sequence divergence
The level of substitutions that accumulated in a PP (K(ψ)) was estimated using the following method.
First, the sequence divergence value (D) was corrected by removing the contribution of mutations at CpG sites. Sequence divergence (δ) of a sequence (at a given time point) of length (N) including the number of CpG dinucleotides (n) is given as a function of the mutation rate at non-CpG dinucleotides (α) and CpG dinucleotides (β) as follows:
δ = α(1/2 - n/N) + βn/N (1)
From the result of Sved and Bird , the ratio of β to α is ∼ 6.5. Therefore, designating α/2 = μ and n/N = ν in Equation 1 gives the following:
δ = μ(1 + 11ν) (2)
Assuming that CpG frequency (ι) in a genomic element that was generated by duplication of a functional gene of high CpG content decreases over time (t) and reaches an equilibrium state (ε) (approximately 20% of the frequency [14, 33] expected from the local fraction of cytosines and guanosines ), the CpG frequency (ν) at time (t) was calculated as follows:
ν = 1/(At + 1/(ι - ε)) + ε (3)
If we accept the value of 1.5 × 10-9 per nucleotide per year [34, 35] as the neutral mutation rate  and equate this to the mutation rate at non-CpG dinucleotides (μ) and use a time unit of 1 million years, then the mutation rate at CpG dinucleotides, β/2, will be around 1 per 100 nucleotides per million years (that is, ν will be reduced by 1% every million years). Therefore, ν(t = 1)/ν(t = 0) in Equation 3 gives:
(1/(A + 1/(ι - ε)) + ε)/ι ≈ 0.99
Solving for A gives:
A = 0.01ι/((0.99ι - ε)(ι - ε)) (4)
The sequence divergence value (D) is given as an integral of the sequence divergence (δ) from the present (t = 0) to the time of the duplication (t = T): D = ∫0Tδdt. From Equations 2, 3 and 4,
D = ∫0Tμ(1 + 11((ι - ε)/((0.01ι/(0.99ι - ε))t + 1) + ε))dt (5)
Alu (0.077, 0.020); L1 (0.012, 0.008); PPs (0.015, 0.010)
The substitution level (Σ) at sites other than CpG is given from the time since the duplication (T) and the neutral mutation rate (μ) of primates : Σ = μT. The quantity Σ was corrected for multiple substitutions at the same site using the Jukes-Cantor model , giving the average number of substitutions per 100 bp (K): K = - (3/4)ln(1 - (4/3)Σ).
For PPs, sequence divergences were defined as the mismatch rates of respective PPs relative to the current sequences of their parental genes. The mismatch rate of a PP relative to its parental gene (K) consists of the level of substitutions that accumulated only in the PP (K(ψ)) and the level of substitutions that accumulated only in the gene (K(f)): K = K(f) + K(ψ). K(f) and K(ψ) can be further subdivided into the number of synonymous (Ks) and nonsynonymous (Ka) substitutions [58–60]: K(f) = Ks(f) + Ka(f), K(ψ) = Ks(ψ) + Ka(ψ). Kuma and Miyata evaluated the average nucleotide substitution rates of 31 pairs of human PPs and their parental genes using homologs of other species as outgroups (K. Kuma and T. Miyata, personal communication). They used the following genes: ADP-ribosylation factor 1, aldolase A, aldose reductase, alpha-E-catenin, alpha-L-fucosidase, alpha-enolase, arylamine N-acetyltransferase, beta-tubulin, c-Raf protooncogene, cAMP-dependent protein kinase regulatory subunit, calmodulin, ceruloplasmin, creatine kinase, cyclophilin, cytochrome b5, cytochrome c, ferrochelatase, gamma-actin, glucocerebrosidase, glutamine synthetase, glyceraldehyde-3-phosphate dehydrogenase, histone H3.3, hsc70, hsp27, hsp60, lactate dehydrogenase-A, neurotrophin-4, phosphoglycerate kinase, prothymosine alpha, topoisomerase-I, triose phosphate isomerase. They calculated the following ratios: Rs(ψ), the synonymous substitutions in PPs to synonymous substitutions in their parental genes; Ra(f), the ratio of nonsynonymous substitutions in genes to synonymous substitutions in genes; and Ra(ψ), the ratio of nonsynonymous substitutions in PPs to synonymous substitutions in genes. The mean values of Rs(ψ), Ra(ψ) and Ra(f) were:
Rs(ψ) = Ks(ψ)/Ks(f) = 1.40 (6.1)
Ra(ψ) = Ka(ψ)/Ks(f) = 1.13 (6.2)
Ra(f) = Ka(f)/Ks(f) = 0.06 (6.3)
From Equations 6.1-6.3, and Equations K = K(f) + K(ψ), K(f) = Ks(f) + Ka(f), and K(ψ) = Ks(ψ) + Ka(ψ), the estimated level of substitutions in PPs (K(ψ)) is given by:
K(ψ) = 0.705K (7)
We thank Katsuhiko Murakami (RIKEN-GSC) for helpful discussions and Kei-ichi Kuma and Takashi Miyata (Kyoto University) for providing the data on the average nucleotide substitution rates of 31 pairs of human PPs. This work was partially supported by the Ministry of Education, Culture, Sports, Science and Technology of Japan, Grant-in-Aid for Scientific Research. This work was also supported by a grant from BIRD of Japan Science and Technology Corporation (JST) for K.O.
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