Poly(A)-binding proteins are required for diverse biological processes in metazoans

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PABPs [poly(A)-binding proteins] bind to the poly(A) tail of eukaryotic mRNAs and are conserved in species ranging from yeast to human. The prototypical cytoplasmic member, PABP1, is a multifunctional RNA-binding protein with roles in global and mRNA-specific translation and stability, consistent with a function as a central regulator of mRNA fate in the cytoplasm. More limited insight into the molecular functions of other family members is available. However, the consequences of disrupting PABP function in whole organisms is less clear, particularly in vertebrates, and even more so in mammals. In the present review, we discuss current and emerging knowledge with respect to the functions of PABP family members in whole animal studies which, although incomplete, already underlines their biological importance and highlights the need for further intensive research in this area.

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Genome-Wide RNAi of C. elegans Using the Hypersensitive rrf-3 Strain Reveals Novel Gene Functions

Femke Simmer, Celine Moorman, Alexander M. van der Linden … (2003)

Introduction RNA interference (RNAi) is targeted gene silencing via double-stranded RNA (dsRNA); a gene is inactivated by specific breakdown of the mRNA (Fire et al. 1998; Montgomery et al. 1998). It is an ideal method for rapid identification of in vivo gene function. Initial studies on RNAi used microinjection to deliver dsRNA (Fire et al. 1998), but it was subsequently shown that dsRNA can be introduced very easily by feeding worms with bacteria that express dsRNA (Timmons and Fire 1998). Using this technique on a global scale, an RNAi feeding library consisting of 16,757 bacterial clones that correspond to 87% of the predicted genes in Caenorhabditis elegans was constructed (Fraser et al. 2000; Kamath et al. 2003). Upon feeding to worms, these clones will give transient loss-of-function phenotypes for many genes by inactivating the target genes via RNAi. By feeding the clones in this library to wild-type Bristol N2 worms, loss-of-function phenotypes were assigned to about 10% of genes. However, RNAi phenotypes were missed for about 30% of essential genes and 60% of genes required for postembryonic development, probably because RNAi is not completely effective (Kamath et al. 2003). Other global RNAi screens have been recently performed in C. elegans using this RNAi library or other techniques (Gönczy et al. 2000; Maeda et al. 2001; Dillin et al. 2002; Piano et al. 2002; Ashrafi et al. 2003; Lee et al. 2003; Pothof et al. 2003). These screens were done using wild-type worms. We have already shown that mutation of rrf-3, a putative RNA-directed RNA polymerase (RdRP), resulted in increased sensitivity to RNAi (Sijen et al. 2001; Simmer et al. 2002). There are four RdRP-like genes in C. elegans. Two of these, ego-1 and rrf-1, are required for efficient RNAi, as apparent from the fact that these mutants are resistant to RNAi against germline or somatically expressed genes, respectively (Smardon et al. 2000; Sijen et al. 2001). A third gene, rrf-2, appears to have no role in RNAi. The rrf-3 strain, mutated in the fourth RdRP homolog, shows an opposite response to dsRNA; this mutant has increased sensitivity to RNAi (Sijen et al. 2001). A more detailed study of RNAi sensitivity of rrf-3 mutants using a set of 80 genes showed that rrf-3 is generally more sensitive to RNAi than wild-type worms (Simmer et al. 2002). RNAi phenotypes in rrf-3 animals are often stronger, and they more closely approximate a null phenotype, when compared to wild-type. In addition, loss-of-function RNAi phenotypes were detected for a number of genes using rrf-3 that were missed in a wild-type background. For example, known phenotypes were detected for many more neuronally expressed genes in the rrf-3 background. These features suggest that the rrf-3 strain could be used to improve and extend functional information associated with C. elegans genes. We have conducted a genome-wide RNAi screen using the rrf-3 strain. In total, we found reproducible RNAi phenotypes for 423 clones that previously did not induce a phenotype (corresponding to 393 additional genes). To explore the variability of global RNAi screens, we performed the rrf-3 screen twice for Chromosome I and carried out a Chromosome I screen with wild-type. These were cross-compared and also compared to the results of the wild-type screen of Fraser et al. (2000). From this, we find that rrf-3 consistently allowed detection of more phenotypes than wild-type. In addition, we found that there is a significant screen-to-screenvariability (10%–30%). Results Comparative Analysis of RNAi for Chromosome I with Wild-Type and rrf-3 We first conducted a pilot screen of Chromosome I using rrf-3 and found RNAi phenotypes for 456 bacterial clones. We compared these data to those obtained by Fraser et al. (2000) for a screen in the wild-type Bristol N2 strain. For 153 of these 456 clones, no phenotypes were reported by Fraser et al. (2000) and phenotypes were observed for 303 clones in both screens. The N2 screen done by Fraser et al. (2000) resulted in RNAi phenotypes for 40 clones for which no phenotypes were found using rrf-3 (Figure 1A). These results indicate that rrf-3 can be used in a global screen to identify loss-of-function phenotypes for additional genes. However, some phenotypes were missed in the rrf-3 screen. To explore the reproducibility and variability of RNAi screens, we next screened the clones of Chromosome I using N2 and rrf-3 side by side. We detected phenotypes for 447 clones: 140 were found only in rrf-3, 11 only in N2, and 296 in both strains (Figure 1B). These data confirm that rrf-3 is more sensitive to RNAi and, in addition, these data indicate that global RNAi screens with rrf-3 will result in more clones with a detectable phenotype. Variability of the RNAi Effect When we compared the RNAi results that we obtained using N2 with the Fraser et al. (2000) data, we were surprised to find significant differences: we only detected phenotypes for 75% of the clones that gave a phenotype in Fraser et al. (2000), and these researchers reported phenotypes for 84% of clones for which we found a phenotype (Figure 1C). The differences do not appear to be due to false positives. For example, Fraser et al. (2000) detected the predicted phenotype for goa-1 and unc-73, whereas we did not detect a mutant phenotype. Similarly, we detected the known mutant phenotype for egl-30 and cdc-25.1, which were not detected by Fraser et al. (2000). In addition, we found that the false-positive rate is negligible (see below). It is possible that different laboratories or investigators have slightly different results. However, when we compare the results that we obtained with two independent screens of Chromosome I using rrf-3 in our laboratory, we also see differences. For 394 clones we detected a phenotype in both experiments, 54 are specific for the first experiment, and 34 for the second (Figure 1D). Among the clones that only gave an RNAi phenotype in one of the experiments are again clones that induced the predicted phenotype based on the phenotypes of genetic mutants (unc-40, gpc-2, and sur-2). These data show that large-scale RNAi screens done within the same laboratory and by the same investigators also give variable results. A few examples of variable RNAi results are shown in Table 1. In conclusion, we find that RNAi results from different laboratories and from experiments done in the same laboratory vary from 10% to 30%. This appears to be due to a high frequency of false negatives in each RNAi screen, even when the same method is used in the same laboratory. The Genome-Wide RNAi Screen Based on the positive results of the Chromosome I screen using the rrf-3 strain, we next screened the complete RNAi library with rrf-3 mutant animals. We obtained results for 16,401 clones and detected phenotypes for 2,079 (12.7%). Of these, we identified phenotypes for 625 clones for which no phenotype was reported in the Fraser et al. (2000) or Kamath et al. (2003) screens using N2, with the remaining 1,454generating phenotypes in both screens (Table S1). In addition, there are 287 clones for which only Fraser et al. (2000) or Kamath et al. (2003) found phenotypes (23 of these were not done in our screen). The clones for which we only detected an RNAi phenotype once and that were specific for the rrf-3 screen were retested. Subsequently, the phenotypes of the clones corresponding to Chromosomes II to X that were not confirmed by this repetition were tested once more. In this way, the clones specific for the rrf-3 screen had two chances to be confirmed. Of the 625 clones for which no phenotype was found in the Fraser et al. (2000) and Kamath et al. (2003) N2 screens, the phenotypes of 423 clones were confirmed and 202 remained unconfirmed (Table 2; see Table S1). Combining the N2 screens and these 423 clones, the percentage of clones with a phenotype increases from 10.3% to 12.8%. Some of the RNAi phenotypes only found with rrf-3 that remained unconfirmed could be confirmed by RNAi phenotypes detected with other clones of the RNAi library corresponding to the same gene or by other laboratories using different RNAi methods. For example, for the clones corresponding to the predicted genes F56D1.1 (a member of the zinc finger C2H2-type protein family) and F27C8.6 (a member of the esterase-like protein family), we detected sterile progeny (Stp) and embryonic lethality (Emb), respectively; these were also found by Piano et al. (2002). In addition, some unconfirmed RNAi phenotypes are confirmed by comparing to phenotypes of genetic mutants such as gpc-2, hlh-8, and unc-84. This suggests that many of the unconfirmed phenotypes reflect true gene functions. Analysis of the rrf-3 Results To validate the results obtained using rrf-3, we first assayed the rate of false positives in the total dataset (all RNAi results obtained with rrf-3 for the 16,401 clones tested). In the assay used by Kamath et al. (2003), a set of genes for which it is known that genetic mutants display no lethality was selected. A false positive in the RNAi data is then defined as detecting a lethal RNAi phenotype for any of these genes. In the N2 screen, the false-positive rate was 0.4%. We find that the false-positive rate in the rrf-3 data is similarly low (0 of 152 genes). To further determine the effectiveness of the screen, we compared the RNAi phenotypes with loss-of-function phenotypes of genetic mutants. For all chromosomes except for Chromosome I, the rrf-3 data were confirmed by refeeding only if there was no phenotype detected in the N2 screens by Fraser et al. (2000) or Kamath et al. (2003). Therefore, to compare the difference in detection of known phenotypes between the rrf-3 and the N2 screens, we used the Chromosome I datasets, where phenotypes were confirmed independently for the two strains. Of 75 genetic loci on Chromosome I, Fraser et al. (2000) detected 48% of published phenotypes, compared to 59% for rrf-3 (Table S2). Using the genome-wide rrf-3 dataset (excluding the 202 unconfirmed phenotypes), we detected the published phenotype for 54% of 397 selected loci, compared to 52% for N2 (Table 3; see Table S2). We next asked whether using the rrf-3 strain improved general phenotype detection or whether certain types of phenotypes were particularly increased compared to the N2 screens by Fraser et al. (2000) and Kamath et al. (2003). To do this, we analysed the detection rate of different types of Chromosome I loci. First, we looked at a set of 23 loci with nonlethal postembryonic mutant phenotypes. Using rrf-3, we reproducibly detected the published phenotype for 11 of these compared to only two for N2. Of 50 loci required for viability (essential genes), we detected 31 using rrf-3, compared to 33 for N2. Thus, detection of essential genes was similar in the two strains, but detection of postembryonic phenotypes was improved with rrf-3. Finally, for the whole genome using rrf-3, we reproducibly detected the published phenotypes for 34 genetic mutants for which no RNAi phenotype was reported in the N2 screens (nine essential genes, 21 with postembryonic mutant phenotypes, and four with a slow-growth mutant phenotype). By comparison, published phenotypes were detected for 23 loci only with N2 (16 essential genes and seven with postembryonic mutant phenotypes) (see Table S2). We conclude that rrf-3 particularly improves detection of genes with postembryonic mutant phenotypes, a class that is poorly detected using wild-type N2. A striking feature of the rrf-3 dataset is the high number of clones where a slow or arrested growth (Gro/Lva) defect was induced, without associated embryonic lethality or sterility. Overall, 619 clones induced a Gro/Lva defect using rrf-3, compared to 276 for N2, whereas the number of essential genes detected was similar (1,040 versus 1,170, respectively). In addition, in the confirmed set of 423 clones with rrf-3-specific phenotypes, Gro/Lva defects are the largest category (42%), whereas this is only 18% for N2, with the largest category being essential genes (49%). These data suggest that rrf-3 might particularly enhance detection of genes that mutate to a slow-growth phenotype; we cannot easily test this hypothesis, as there are currently few known loci with this mutant phenotype. In some cases, a Gro/Lva phenotype was seen in rrf-3, whereas a different phenotype was seen in N2 (e.g., either lethality or a weak postembryonic phenotype). This suggests that some of the Gro/Lva phenotypes detected are due to incomplete RNAi of an essential gene (where lethality was seen in N2) or by a stronger RNAi effect (where no growth defect was seen in N2). In addition, it is possible that some of the Gro/Lva phenotypes detected are synthetic effects of using the rrf-3 mutant strain. To summarise, using the rrf-3 RNAi supersensitive strain in large-scale screens increases the percentage of clones for which it is possible to detect a phenotype. Detection of postembryonic phenotypes is particularly increased, whereas detection of essential genes is similar in rrf-3 and N2. In addition, using rrf-3, there is a high rate of induction of Gro/Lva defects. Positional Cloning of Genetic Mutants with Visible Phenotypes Despite the advantages of RNAi, genetic mutants remain indispensable for many experiments. In the past decades, forward genetic screens identified a large number of genetic mutants, many of which are not yet linked to the physical map. We used the RNAi phenotypes obtained with the genome-wide screens to test whether we could systematically clone genes that are mutated in existing genetic mutants. First, the genetic map positions of all uncloned genetic mutants with visible phenotypes were checked using WormBase (http://www.wormbase.org, the Internet site for the genetics, genomics, and biology of C. elegans). Second, we searched for clones near the defined map positions that, when fed to N2, rrf-3, or both, gave phenotypes corresponding to the phenotypes of the genetic mutants. For most genetic mutants, more than ten clones with a similar phenotype were found in the interval to which the genetic mutant was mapped. However, for 21 genetic mutants, only one or a few candidate clones were found. The genes corresponding to these clones were subsequently sequenced in the genetic mutant to determine whether a mutation was present. In total, we sequenced 42 predicted genes for the 21 genetic mutants (Table S3). For seven of these—bli-3, bli-5, dpy-4, dpy-6, dpy-9, rol-3, and unc-108—we found a mutation in one of the sequenced genes (Table 4). The mutated gene was confirmed by sequencing the same gene in a second or third allele (or both) of these genetic mutants (Table 4). The identification of mutations in unc-108 encoding the homolog of the small GTPase Rab2 is of particular interest. The RNAi phenotype of this gene gives a clue about the genetic property of the mutations in the mutants of unc-108. With rrf-3, we find that inactivation of Rab2 (F53F10.4) by RNAi causes uncoordinated movement (Table 4). Mutations in unc-108 were isolated in a screen for dominant effects on behaviour; heterozygous unc-108 mutants display dominant movement defects and are indistinguishable from homozygous mutants (Park and Horvitz 1986). RNAi phenocopies a loss-of-function phenotype, suggesting that the dominant movement defects of unc-108 mutants may be due to haplo-insufficiency. In eukaryotes, Rab2 is involved in regulating vesicular trafficking between the endoplasmic reticulum and Golgi. Based on the movement defects of unc-108 mutants, UNC-108 might be involved in vesicle transport in neurons that regulate locomotion. Thus, the RNAi data are a powerful tool to facilitate rapid cloning of the genes identified by genetic mutants and will provide important starting points for further studies of their function. Discussion With this genome-wide RNAi screen using the hypersensitive strain rrf-3, we have significantly increased the functional information on the C. elegans genome, and we confirmed many RNAi phenotypes observed previously. We have assigned RNAi phenotypes for 406 genes (corresponding to the 423 extra clones) using rrf-3. For 13 genes, Kamath et al. (2003) or Fraser et al. (2000) had already found a phenotype using a different clone from the RNAi library that targeted the same gene, and for at least 44 genes a genetic mutant exists (see Table S2). Other investigators have also found RNAi phenotypes for some of the genes using different methods. However, for most genes our result is to our knowledge the first hint about their biological function. Although we have identified new RNAi phenotypes for a substantial number of genes, others will have been missed in our screen for the following reasons. First, besides its increased sensitivity to RNAi, the rrf-3 strain has an increased incidence of males (Him) and displays slightly increased embryonic lethality and a reduced brood size (Simmer et al. 2002). In our rrf-3 experiments, we therefore made some minor adaptations to the original RNAi protocol described by Fraser et al. (2000). We did not score for the Him phenotype and had more stringent criteria for embryonic lethality and sterility. This may have reduced the number of extra clones identified with a phenotype. Moreover, the changes in the protocol can also account for some differences in the detection of RNAi phenotypes between rrf-3 and N2. Second, when an RNAi phenotype is detected with N2 and not with rrf-3, the lack of a detectable phenotype may be the result of variability in the efficiency of RNAi. This is consistent with the fact that we observe differences between experiments done with the same strain. When an RNAi phenotype is detected with rrf-3 and not with N2, this can be due to the increased sensitivity to RNAi of rrf-3. However, besides the higher sensitivity, we may also be observing synthetic effects with rrf-3 (e.g., embryonic lethality, sterility, or developmental delay). In particular, a large number of clones induced a developmental delay phenotype using rrf-3. Synthetic effects cannot be excluded without investigating genetic mutants. Again, variability in the efficiency of RNAi will also contribute to these differences, and a small portion may be false positives. In general, the few false positives that occur in the screen are most likely due to experimental errors, whereas the false negatives are due to reduced efficiency of the RNAi. Finally, differences between rrf-3 and N2 do not only involve the absence and presence of an RNAi phenotype, but also differences in the phenotypes for clones that did induce phenotypes in both screens (e.g., embryonic lethal in one screen and a postembryonic phenotype in the other). For example, we detected for unc-112 a 100% embryonic lethal (Emb) phenotype with rrf-3, whereas Kamath et al. (2003) detected an adult lethal (Adl), uncoordinated (Unc), and paralyzed (Prz) phenotype with N2. Conversely, Kamath et al. (2003) detected for gon-1 a 100% Emb phenotype and other phenotypes with N2, while we did not detect an Emb phenotype with rrf-3. What could be the source of the interexperimental variation of RNAi? Different phenotypes for the same gene can possibly occur owing to slight differences in the developmental stage at which the animals are exposed to dsRNA and owing to changes in temperature during the experiment. However, this probably does not account for the differences we see, as we always used animals of the same larval stage (L3/L4) and used incubators for constant temperature. It was shown previously that the level of induction of dsRNA production by isopropylthio-β-D-galactoside (IPTG) can modify the penetrance of the RNAi phenotype (Kamath et al. 2000). Therefore, differences in the induction of the dsRNA either by changes in the concentration of IPTG, temperature, timing, or the bacteria may be an important source of the variation in the outcome of RNAi. RNAi is starting to be used extensively in other systems experimentally, as well as therapeutically and agriculturally. The relative variability of the RNAi effect is an important fact to take in account also for the use of RNAi in other systems. The RNAi data can be a useful starting point for many new experiments, such as positional cloning of genetic mutants. By sequencing candidate genes based on the RNAi phenotypes, we identified the causal mutation in seven genetic mutants. Identification of these mutated genes gives insight into the biological process in which they are involved. In addition, cloning of these genes increases the resolution of the genetic map of C. elegans, since these mutants have been extensively used as visible markers in linkage studies. The complete set of RNAi phenotypes detected for the 2,079 clones using rrf-3 will be submitted to WormBase, annotated as confirmed or unconfirmed. There the data can be evaluated in the context of information on gene structure, expression profiles, and other RNAi results. Materials and Methods Nematode strains. We used the following C. elegans strains: Bristol N2, NL4256 rrf-3(pk1426), CB767 bli-3(e767), MT1141 bli-3(n259), CB518 bli-5(e518), BC649 bli-5(s277), CB1158 dpy-4(e1158), CB1166 dpy-4(e1166), CB14 dpy-6(e14), CB4452, dpy-6(e2762), F11 dpy-6(f11), CB12 dpy-9(e12), CB1164 dpy-9(e1164), BC119 dpy-24(s71), CB3497 dpy-25(e817), MT1222 egl-6(n592), MT1179 egl-14(n549), MT1067 egl-31(n472), MT151 egl-33(n151), MT171 egl-34(n171), egl-34(e1452), MQ210 mau-4(qm45), CB754 rol-3(e754), BC3134 srl-2(s2507dpy-18(e364); unc-46(e177)rol-3(s1040), CB713 unc-67(e713), CB950 unc-75 (e950), HE177 unc-94(su177), HE33 unc-95(su33), HE151 unc-96(su151), unc-96(r291), HE115 unc-100(su115), MT1093 unc-108(n501), and MT1656 unc-108(n777). RNAi by feeding. RNAi was performed as described elsewhere (Fraser et al. 2000; Kamath et al. 2000) with minor adaptations when the rrf-3 strain was used: after transferring L3- to L4-staged hermaphrodites onto the first plate, we left them for 48 h at 15°C instead of 72 h and then plated single adults onto other plates seeded with the same bacteria. Furthermore, we did not remove the mothers from the second plates. The phenotypes assayed are these: Emb (embryonic lethal), Ste (sterile), Stp (sterile progeny), Brd (low broodsize), Gro (slow postembryonic growth), Lva (larval arrest), Lvl (larval lethality), Adl (adult lethal), Bli (blistering of cuticle), Bmd (body morphological defects), Clr (clear), Dpy (dumpy), Egl (egg-laying defective), Lon (long), Mlt (molt defects), Muv (multivulva), Prz (paralyzed), Pvl (protruding vulva), Rol (roller), Rup (ruptured), Sck (sick), Unc (uncoordinated) Thin and Pale. Emb was defined as greater than 10% dead embryos for N2 and greater than 30% dead embryos for rrf-3. Ste required a brood size of fewer than ten among fed N2 worms and fewer than five among rrf-3. Each postembryonic phenotype was required to be present among at least 10% of the analysed worms. Sequencing of genetic mutants. The coding sequence and the 5′- and 3′-untranslated region (about 500 bp upstream and downstream of the coding sequence) of the predicted genes, as annotated in WormBase, was analysed for mutations by sequencing amplified genomic DNA of the genetic mutants (see Table S3). Nested primers were designed using a modification of the Primer3 program available on our website (http://primers.niob.knaw.nl/). Sequence reactions were done using the ABI PRISM Big Dye terminator sequencing kit (Applied Biosystems, Foster City, California, United States) and were analysed on the ABI 3700 DNA analyser. Sequences were compared to the genomic sequence of C. elegans using the BLAST program (http://www.sanger.ac.uk/Projects/C_elegans/blast_server.shtml) or analysed using the PolyPhred program (available from http://droog.mbt.washington.edu/PolyPhred.html). Supporting Information Table S1 RNAi Phenotypes for Bacterial Clones Using rrf-3 (482 KB PDF). Click here for additional data file. Table S2 Detailed Comparison of RNAi Phenotypes with Those of Known Loci (188 KB PDF). Click here for additional data file. Table S3 Summary of Genes Sequenced in Several Genetic Mutants (25 KB DOC). Click here for additional data file. Accession Numbers RNAi data from this study will be submitted to WormBase (http://www.wormbase.org).

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The Central Dogma Decentralized: New Perspectives on RNA Function and Local Translation in Neurons

Christine Elizabeth Holt, Erin M. Schuman (2013)

Main Text Background It is now clear that individual neurons are highly compartmentalized with specific functions and/or signaling that occur in restricted subcellular domains. Extrinsic signals are often spatially localized such that they are “seen” by restricted parts of a neuron, such as synaptic input to a specific dendritic spine or a guidance cue encountered by a growth cone. Twenty-five years ago, when the first issue of Neuron was published, it was well appreciated that the neurons were capable of local information processing, but the potential cellular mechanisms that established and regulated local compartments were not well understood. Dendritic spines had been proposed as biochemical and/or electrical compartments (Harris and Kater, 1994; Koch and Zador, 1993), and polyribosomes had been identified at the base of spines (Steward and Levy, 1982). However, the view that dominated until nearly the end of the twentieth century was that the central dogma (DNA-RNA-protein) was carried out centrally—in the nuclei and somata of neurons. In that context, the localization of mRNA observed in some cells was thought to represent a specialized mechanism that operated in unique biological systems, such as egg cells, where storage of mRNAs is needed for subsequent patterning of the early embryo (see Martin and Ephrussi, 2009 for review). Evidence from a number of studies in the last decade, particularly in neurons, has led to a revolution in our thinking. Although the field is still young, it is becoming clear that RNA-based mechanisms provide a highly adaptable link between extrinsic signals in the environment and the functional responses of a neuron or parts of a neuron. This is accomplished by the localization of both protein-coding and noncoding RNA in neuronal processes and the subsequent regulated local translation of mRNA into protein. Here we discuss some of the key findings that lead us to the view that mRNA localization and RNA-regulated and localized translation underlie many fundamental cellular processes that are regulated by extrinsic signals in neurons, such as memory, dendrite and arbor branching, synapse formation, axon steering, survival, and likely proteostasis. The dynamic regulation of protein synthesis is essential for all cells, including neurons. Over 50 years ago, in vivo experiments (in a variety of species) established a clear functional link between protein synthesis and long-term memory (see Davis and Squire, 1984 for review), indicating that proteome remodeling underlies behavioral plasticity. These observations were paralleled by in vitro studies of synaptic plasticity demonstrating a clear requirement for newly synthesized proteins in the long-term modification of synaptic function (see Sutton and Schuman, 2006 for review; also, Tanaka et al., 2008). This link between protein synthesis and long-term plasticity is most recently reinforced by studies showing that targeted genetic disruption of signaling molecules that regulate protein translation interfere with long-term synaptic or behavioral memories (Costa-Mattioli et al., 2009). The above studies, while indicating a requirement for protein synthesis, do not address the location. We now know dendrites and axons of neurons represent specialized cellular “outposts” that can function with a high degree of autonomy at long distances from the soma, as illustrated by the remarkable ability of growing axons to navigate correctly after soma removal (Harris et al., 1987) or isolated synapses to undergo plasticity (Kang and Schuman, 1996; Vickers et al., 2005). The identification of polyribosomes at the base or in spines (Steward and Levy, 1982) together with metabolic labeling experiments that provided the first evidence of de novo synthesis of specific proteins in axons and dendrites (Feig and Lipton, 1993; Giuditta et al., 1968; Koenig, 1967; Torre and Steward, 1992) indicated the competence of these compartments for translation. Subsequent studies demonstrated that specific subsets of mRNAs localize to synaptic sites (Steward et al., 1998) and directly linked synaptic plasticity with local translation in dendrites (Aakalu et al., 2001; Huber et al., 2000; Kang and Schuman, 1996; Martin et al., 1997; Vickers et al., 2005), providing definitive proof that dendrites are a source of protein during plasticity. In axons, the idea of local protein synthesis has been slower to find acceptance, no doubt hindered by the classical view of axons as information transmitters rather than receivers; so, why would local protein synthesis be required? Although ribosomes were identified in growth cones in early ultrastructural studies (Bunge, 1973; Tennyson, 1970), they were rarely observed in adult axons. It is now thought that at least part of the explanation for their apparent paucity lies in their localization close to the plasma membrane in axons (Sotelo-Silveira et al., 2008) where ribosomal subunits can associate directly with surface receptors (Tcherkezian et al., 2010). In addition, evidence indicates that myelinated axons can tap into an external supply of ribosomes by the translocation of ribosomal proteins from Schwann cells (Court et al., 2011). Growing and navigating axons are clearly information receivers, like dendrites, since their growth cones steer using extrinsic signals. Indeed, the first functional evidence for local protein synthesis in axons came from studies that showed that cue-induced directional steering is abolished by inhibitors of protein synthesis, including rapamycin, in surgically isolated axons (Campbell and Holt, 2001). Subsequent studies confirmed this result in different neurons (Wu et al., 2005; Yao et al., 2006) and revealed that local protein synthesis underlies growth-cone adaptation, gradient sensing, and directional turning in growing axons (Leung et al., 2006; Ming et al., 2002; Piper et al., 2005; Yao et al., 2006). In addition, axonal protein synthesis is elicited in response to injury and plays key roles in axon regeneration and maintenance (Jung et al., 2012; Perry et al., 2012; Verma et al., 2005; Yoon et al., 2012; Zheng et al., 2001). Compartments Neuronal function is highly dependent on spatially precise signaling. Increasing evidence indicates that the complex morphology of neurons has created biological compartments that subdivide the neuron into spatially distinct signaling domains important for neuronal function (Hanus and Schuman, 2013). Dendritic spines represent a specialized (“classical”) cellular compartment in which subsets of specific proteins (e.g. receptors, channels, signaling molecules, and scaffolds) are collected together with a common function for receiving and processing electrical and chemical input. Spines have a distinct structural morphology and, as such, are easy to classify as a compartment. Although spines are small (∼1 μm3), they can still be subdivided into further functional compartments (see Chen and Sabatini, 2012 for review) with multiple microdomains, raising the question of how a compartment is defined. For example, a recent superresolution imaging study demonstrated that, within synapses, AMPA receptors are clustered into small nanodomains (∼70 nm in diameter) that contain on average ∼20 receptors (Nair et al., 2013). These nanodomains are dynamic in both their shape and position and may have a limited lifetime. Anatomically and functionally distinct compartments also exist in axons, such as the growth cone, the axon initial segment, and terminal arbor. Equally, there are examples of compartments that exhibit no obvious “anatomical” specializations. In axons, for example, some membrane proteins are localized to restricted segments of the axon (Fasciclins, Tag1/L1, Robo) (Bastiani et al., 1987; Dodd et al., 1988; Katsuki et al., 2009; Rajagopalan et al., 2000) indicative of plasma-membrane compartmentalization. In addition, second-messenger signaling molecules such as calcium and cyclic nucleotides, once thought to signal extensively throughout a cell, are now known to be highly regulated such that increases in concentration can be confined to a small space, creating a signaling compartment. Selective activation of a single spine on a dendrite, for example, can provide the receiving neuron with information about a specific stimulus (Varga et al., 2011). Compartments may be overlapping or distinct and range in size depending on the biological function. Ultimately, a neuron must integrate the information received from multiple compartments. As such, future experiments aimed at understanding how different compartments emerge and what mechanisms generate such spatially precise intracellular patterning will be very informative. Compartmentalized signaling presents several challenges to the cell, a prime one being the localization of its component parts. Specific molecules must be transported and delivered to the appropriate subcellular destinations. One of the remarkable features of RNA is its ability to be spatially localized and, therefore, potentially contribute to neuronal compartmentalization. Historically, localized mRNAs have been studied during development (see Martin and Ephrussi, 2009). That localized RNA is more often the rule than the exception is spectacularly illustrated by the finding that 71% of the Drosophila embryo transcriptome is localized to specific subcellular compartments (Lécuyer et al., 2007). The proteins encoded by localized mRNAs are also concentrated at the site suggesting that mRNA localization and the ensuing local translation plays an important role in positioning proteins for cellular functions. Asymmetry and Spatial Signaling A general function of mRNA localization is the generation of asymmetry. mRNAs tend to be abundantly localized to the peripheral domains and motile parts of neurons where they are optimally positioned for the arrival of external signals, e.g., in dendrites (synaptic activation) and growth cones. Subcellular asymmetry can lead to highly polarized dynamics and cell morphology that can operate on a remarkably fine scale. Growth-Cone Spatial Signaling To navigate, growth cones must be able to make directional turns, which demands asymmetry. In retinal growth cones, for example, which are only 5 μm in diameter, a polarized external gradient of netrin-1 triggers increases in both the transport and translation of β-actin mRNA on the gradient near side (Leung et al., 2006; Yao et al., 2006). This polarized translation leads to a rapid (5 min) polarized increase in β-actin protein that helps to drive axon turning towards the gradient source. Interestingly, different cues show specificity in their effects on mRNA transport and translation. Different growth factors, for example, trigger the transport of a specific repertoire of mRNAs in axons (Willis et al., 2005, 2007; Zhang et al., 1999), and different guidance cues elicit the translation of specific subsets of mRNAs (Leung et al., 2006; Piper et al., 2006; Shigeoka et al., 2013; Wu et al., 2005; Yao et al., 2006). β-actin mRNA translation is triggered by netrin-1 but not Sema3A, whereas RhoA and cofilin mRNA translation is induced by Sema3A but not netrin-1. This has given rise to the differential translation model suggesting that translation-dependent repulsive and attractive turning in growth cones depends on the differential translation of mRNAs involved in assembly or disassembly of the actin cytoskeleton (Lin and Holt, 2007). Several aspects of this translation-driven cue-induced turning remain to be understood, such as how receptor activation signals mRNA recruitment and, critically, how specific subsets of mRNA are translated. Readout of Spatial Position In Vivo Navigating growth cones encounter a series of patterned molecular cues along the pathway from which they must read out their spatial position. Although there are several examples of stimulus-induced local translation in axons in vitro (Shigeoka et al., 2013), it has only recently become possible to investigate translation in neuronal compartments in vivo. Early studies by Flanagan and colleagues showing compartmentalized expression of EphA2, recapitulated by a translation reporter, in the post-midline crossing segment of commissural spinal cord axons introduced the idea that the growing tip of the axon is stimulated by a regionally expressed cue (e.g., at the midline) that triggers the region-specific translation of proteins needed for pathfinding (Brittis et al., 2002). A recent study provides direct evidence for this type of mechanism in the control of Robo expression and midline guidance (Colak et al., 2013). Two Robo3 receptor isoforms have opposing roles in guiding axons to and away from the midline, and their expression is compartmentalized in pre-crossing (Robo3.1) and postcrossing (Robo3.2) axonal segments (Chen et al., 2008). The switch to Robo3.2 expression at the midline (the transcript of which contains a premature termination codon) is controlled by midline-induced axonal protein synthesis coupled with nonsense-mediated mRNA decay. This provides an elegant mechanism for turning on synthesis time linked to the crossing event (Colak et al., 2013). It was not previously technically possible to inhibit translation of a specific transcript in a compartment-specific manner. Recently, however, new tools have been developed that allow separate manipulation of specific neuronal compartments in vivo such as targeted delivery of siRNAs or antisense morpholinos and conditional targeting of 3′UTRs (Perry et al., 2012; Yoon et al., 2012). These subcellular-directed approaches are beginning to yield information suggesting that local translation is involved in regulating multiple aspects of axonal and dendritic biology. Guidance cues induce immediate steering responses in growth cones via classical signaling pathways that involve receptor activation and phosphorylation of downstream signaling molecules (Bashaw and Klein, 2010). Some of these “immediate” steering responses also involve local translation, as discussed above. Thus, local translation can provide new proteins on demand at subcellular sites for “immediate” use. Interestingly, local translation in response to extrinsic cues has recently been shown to provide proteins for “delayed” use in axon growth and regeneration. Examples of this are the de novo synthesis of proteins that shuttle back to the nucleus where they influence transcriptional output (Cox et al., 2008; Perry et al., 2012), and another is the de novo synthesis of surface receptor proteins that are employed later in a growth cone’s journey (Leung et al., 2013). Spatial Signaling in Dendrites Recent advances in experimental procedures, allowing the stimulation of individual synapses, have shown that synapses can be independently regulated by synaptic activity (Matsuzaki et al., 2004). On the other hand, other studies emphasize the consideration of the dendritic branch as a computational unit (Govindarajan et al., 2011). Taken together, it seems reasonable to consider a range of spatial domains over which signaling can occur, which would span the scale from subdomains in spines to dendritic branches to the entire neuron. These data can be compared to what we know about the quantitative localization of the protein-synthesis machinery. Indeed, it is clear that many synapses possess a polyribosome nearby (Ostroff et al., 2002). Moreover, recent high-resolution in situ hybridization data suggest that mRNA molecules are distributed in local domains (Cajigas et al., 2012), but not necessarily specific to individual synapses. Preliminary estimates of mRNA numbers indicate that there may not be sufficient copies of individual mRNA species for each synapse to have an exclusive and dedicated molecular toolbox. These data imply that there is local sharing of cell biological machineries, including the machinery for protein synthesis and degradation. It remains unclear, however, over what spatial scale local translation can be regulated and stimulated in dendrites. For example, is stimulation of a single spine sufficient to regulate local translation, and, if so, over what spatial domain do the newly synthesized proteins function? RNA in Neurons The past view that RNA acts primarily as an inert intermediate between genes and proteins has undergone a revolution in recent years with discoveries of both new classes of RNAs (e.g., noncoding RNAs, (see Ulitsky and Bartel, 2013 for review) and new RNA-based mechanisms of gene regulation (e.g., microRNA and RNAi silencing) (see McNeill and Van Vactor, 2012 for review). Indeed, given the relatively constrained diversity of proteomes across cells and organisms, RNA-based mechanisms (diverse RNA species and RNA functions) represent a unique platform to diversify and specialize cells, especially neurons. Numerous new roles for RNA have been found in recent years, expanding the role of RNA to controlling many and diverse cellular processes, including stimulus-induced local translation that underlie adaptive responses in neurons (e.g., memory, axon guidance, and maintenance). In addition, RNA’s role may not be limited to the cells where it is synthesized, as new studies indicate it can be transferred between cells (via exosomes) (Sharma et al., 2013) and even between organisms (Sarkies and Miska, 2013), bringing a whole new era of RNA function in cellular communication into focus. mRNA The demonstrations that local protein translation functions during synaptic development and plasticity led to the hunt for specific mRNAs that could be translated in these local compartments. For many years, in situ hybridization was the method of choice, and several individual mRNAs were visualized in dendrites, including the mRNA for the Ca2+-calmodulin-dependent protein kinase alpha subunit, CaMKIIα (Burgin et al., 1990; Mayford et al., 1996), MAP2 (Garner et al., 1988), Shank (Böckers et al., 2004), and β-actin (Tiruchinapalli et al., 2003). In growth cones and axons, in situ hybridization provided evidence for several different mRNAs, including β-actin (Bassell et al., 1998; Kaplan et al., 1992; Wu et al., 2005). Recent microarray approaches and deep RNA sequencing have dramatically expanded the local transcriptome in both dendrites and axons (Poon et al., 2006; Zhong et al., 2006). One of the most surprising findings to come out of these studies is the vast number of mRNAs that are present in these neuronal compartments. Growing axons have 1000–4500 mRNAs (Zivraj et al., 2010), while dendrites have >2500 mRNAs (Cajigas et al., 2012). The mRNAs resident in these compartments span many different functional classes of molecules: metabolism, translation, degradation, receptors/channels, cytoskeleton, etc. Many functional categories are shared between the two compartments, although there are numerous distinct compartment-specific subsets of mRNAs, e.g., GAP43 mRNA in axons and neurotransmitter receptor subunits in dendrites. The localization of mRNA to cellular compartments involves recognition of information that is contained in the 3′ and/or 5′ untranslated (UTR) sequences. The use of mRNA localization to achieve protein localization may arise from the fact that, at least theoretically, unlimited address information can be built into the 3′ and/or 5′ UTRs of mRNA without altering its gene-coding function, whereas there is a tight limit to how much additional coding sequence can be added to a protein without ramifications for function. The family of proteins that bind, transport, localize, and regulate the translation of mRNAs are known as RNA-binding proteins (RBPs) (see Darnell, 2013 for review). RBPs bind to cis-elements in the 3′ and 5′ UTRs of mRNAs. RNA-binding proteins complexed with mRNA, other RNA species, and accessory proteins are thought to be assembled in the cell body and form RNA granules (Kiebler and Bassell, 2006). During transport on microtubules and microfilaments to its destination (e.g., Hirokawa, 2006 and Czaplinski and Singer, 2006), the mRNA cargo is thought to be “silenced” by translational repressors (Krichevsky and Kosik, 2001). Once transported, it is unclear how or whether mRNAs are anchored near translational sites—or if they show continued dynamics. Both stationary and anchored particles have been observed in dynamic mRNA imaging experiments (Lionnet et al., 2011). RNA-binding proteins are an important class of regulatory molecule that recognizes specific nucleotide sequences in RNA (Ray et al., 2013). IP-Seq analysis has revealed, unexpectedly, that some RBPs can bind hundreds of different mRNAs (see Darnell, 2013 for review). Some RBPs, however, appear to be cell-type specific, such as Hermes (RPBMS2) that is expressed exclusively in retinal ganglion cells in the CNS and its knockdown causes severe defects in axon terminal branching (Hörnberg et al., 2013). The number of mRNA-binding proteins identified by known RNA-binding domains is relatively small (around 270) given the increasingly large number of transcripts found in axons and dendrites. Recent work using interactome capture in embryonic stem cells has significantly expanded the number of RBPs, adding a further ∼280 proteins to the repertoire, including, remarkably, many enzymes such as E3 ubiquitin ligases with previously unknown RNA-binding function (Kwon et al., 2013). Several RBPs have been implicated in neurological disorders, such as FMRP in Fragile X syndrome and survival of motor neuron protein (SMN) in spinal muscular atrophy (Bear et al., 2008; Liu-Yesucevitz et al., 2011), and translation dysregulation has recently been implicated as a major factor in autism (Gkogkas et al., 2013; Santini et al., 2013). Noncoding RNAs In recent years the discovery of noncoding RNAs, including miRNAs (which use sequence complementarity to recognize target mRNA), has revealed unanticipated and enormous potential for the regulation of mRNA stability and translation, as well as other functions. Given the huge and unanticipated number of mRNAs detected in axons and dendrites, it is perhaps not surprising that these noncoding RNAs also exist—and are even enriched—in neuronal compartments. One might even argue the complex morphology and functional specialization of neurons provides a hotbed for mRNA regulation that can potentially be mediated by noncoding RNAs. Indeed, an analysis of 100 different miRNAs discovered the differential distribution of some miRNAs in dendrites versus somata and copy numbers in individual neurons as high as 10,000—equivalent to the number of synapses a typical pyramidal neuron possesses (Kye et al., 2007). Recently, the differential distribution of miRNAs has been also reported in axons versus soma (Natera-Naranjo et al., 2010; Sasaki et al., 2013) and recently emerged as regulators of axon growth and branching (Kaplan et al., 2013). Moreover, the enrichment of miRNAs in synaptosomes isolated from specific brain regions has also been reported (Pichardo-Casas et al., 2012). miRNAs have now been shown to regulate many synaptic functions (see Schratt, 2009 for review). In addition, miRNAs themselves are regulated by behavioral experience (Krol et al., 2010) as well as synaptic plasticity (Park and Tang, 2009). More recently, the appreciation of other types of noncoding RNAs have come into focus, though very little is known about their function in neurons. This includes small-nucleolar RNA-derived and transfer RNA-derived small RNAs, firstly identified as degradation products, and long noncoding RNA known as regulators of gene transcription, that may regulate gene expression posttranscriptionally. A recent study demonstrated, for example, that a long noncoding RNA that is anti-sense to a K+ channel subunit (Kcna2) is upregulated following peripheral nerve injury, leading to a downregulation of the K+ channel and a resulting increase in the excitability of DRG neurons, increasing neuropathic pain (Zhao et al., 2013). Technical Hurdles and Advances Isolating Compartments In the early years, the study of local translation was hampered by the technical difficulty of obtaining pure and sufficient quantities of dendrites and axons for analysis. Pioneering studies used metabolic labeling to demonstrate the synthesis of specific proteins such as tubulin in axons (Giuditta et al., 1968; Koenig, 2009), but the possibility that the signal arose from cell-body contamination could not be eliminated due to these technical limitations. Localized translation was convincingly demonstrated by surgically severing the soma from its processes (Aakalu et al., 2001; Campbell and Holt, 2001; Kang and Schuman, 1996) and, more recently, by the use of chambers in which the processes (dendrites or axons) are fluidically isolated from cell bodies (Eng et al., 1999; Taylor et al., 2010). Other methods for isolating neuronal processes include substrates with limited pore size that allow axons to penetrate but not cell bodies (Torre and Steward, 1992; Zheng et al., 2001) and laser capture microdissection (Zivraj et al., 2010). These methods combined with the rapid increase in the sensitivity of profiling techniques have enabled genome-wide transcriptome analyses to be performed on axons and dendrites in a variety of neurons (see below). Tagging Newly Synthesized Proteins The visualization and identification of newly synthesized proteins has also been a hurdle due to issues of sensitivity (detecting low levels of newly synthesized proteins) as well as difficulties in distinguishing between the movement of existing proteins and the synthesis of new proteins. Puromycin, a tRNA analog, can be used together with fluorescent tags (Smith et al., 2005) or antibodies (Schmidt et al., 2009) to label sites of protein synthesis. Fluorescent reporters, such as photo-switchable Kaede, fused to the 3′UTR regulatory region of mRNAs of interest have enabled de novo protein synthesis to be monitored live in neuronal processes (Aakalu et al., 2001; Brittis et al., 2002; Leung et al., 2006). In addition, new methods have been developed to selectively label the pool of newly synthesized proteins, to ascertain a given cell type or cellular compartment as the site of synthesis, and to visualize the newly synthesized proteins. These methods make use of noncanonical amino acids that cross cell membranes and get charged onto tRNAs by the cell’s own tRNA synthetases and then incorporated into new protein during protein synthesis. These techniques, bio-orthogonal noncanonical amino acid tagging (BONCAT) and fluorescent noncanonical amino acid tagging (FUNCAT) can be used to selectively identify (Dieterich et al., 2006) or visualize (Dieterich et al., 2010) newly synthesized proteins. A modification of the NCAT method, which in principle enables one to label newly synthesized proteins in specific cell types, has also recently been developed (Ngo et al., 2012), and NCAT can be used in combination with 2D difference gel electrophoresis (DIGE-NCAT) to compare the proteomes of specific subcellular (e.g. axonal) compartments (Yoon et al., 2012). Looking Ahead There are many questions for the future, as noted below. 1. How Should We Think about Subcellular Compartments? We know that some compartments (like spines) have plasma membrane as a boundary that can serve to compartmentalize chemical and electrical signals. Other compartments could be determined by the spatial arrangement of molecules, cytoskeleton, or limited diffusion. Are compartments “static” when bounded by anatomy (e.g., a spine) but dynamic when determined by signaling molecule volumes? What defines a subcellular compartment such that mRNAs contain specific addresses to target them there? 2. How Do mRNAs Reach Neuronal Compartments? Some mRNAs are targeted specifically to axons and dendrites and even to the growth cone—how is this targeting achieved? While we have in hand several “zip codes,” there are certainly many messages for which a clear consensus sequence in the UTR has not emerged. In addition, in some cases the signal for recognition by an RNA-binding protein may reside in the secondary structure of the mRNA, rather than the nucleotide sequence. The fact that current secondary structure prediction techniques are limited to small stretches of nucleotides (∼100) complicates our ability to identify binding motifs in 3′UTRs. Adding to the complexity is the recent observation that low-complexity regions of RNA-binding proteins are sufficient to create reversible RNA granule-like structures (Kato et al., 2012). The expanded identification of RBPs as well as the ability to define the binding sites with methods like HITS-CLIP (Licatalosi et al., 2008) should dramatically enhance our knowledge of the binding sites. Future studies should focus on the dynamics of the RNA-protein interactions in cellular contexts. In addition, the possibility that RNA might be delivered from extracellular sources (e.g., via exosomes from neighboring neurons or glia) is a recently suggested exciting idea. 3. How Is the Repertoire of Localized mRNAs Regulated? Unbiased genome-wide analyses have shown that the mRNA repertoire is dynamically regulated with the mRNA repertoire changing over time (Gumy et al., 2011; Zivraj et al., 2010). In addition, it is clear that synaptic activity can lead to the regulated trafficking of mRNA to the distal processes (e.g., Steward et al., 1998). Is this regulated at the level of transcription, or is there some “gating” mechanism that regulates the trafficking of specific transcripts into dendrites/axons? Evidence with ephrinB in RGCs indicates that although the transcripts are present in somas early in development, they do not move into axons until later, suggesting that some kind of specific gating mechanism may exist. 4. How Many Molecules of mRNA Are in a Compartment? Currently, little is known about the quantitative aspects of mRNA localization and translation in neurons. For example, how many RNA molecules are needed to provide a functionally significant amount of protein? How many proteins are synthesized from a single mRNA? One might speculate that some classes of proteins, such as cytoskeletal, would be translated much more than others—such as receptors or channels—and transcript abundance could reflect this difference. In theory, just a few new channel or receptor proteins could be sufficient to alter signaling characteristics within a neuronal microdomain. In addition, a low abundant transcript could be stable and translated with high efficiency. Thus, low-abundance transcripts could exert a significant physiological effect and should not be overlooked in profiling analyses. This also raises the intriguing question of whether translation from monosomes, rather than polysomes, may be more common in distal neuronal compartments where there could be demand for a few highly localized proteins. New high-resolution single molecule detection methods (Cajigas et al., 2012; Park et al., 2012) and live-imaging methods for translation (Chao et al., 2012) will be valuable when answering these sorts of questions. 5. What mRNAs Are Translated in Subcellular Compartments In Vivo? With the advent of TRAP (translating affinity purification) technology (Heiman et al., 2008) it will be possible in the future to answer this question in specific neuronal compartments of specific subsets of neurons. For example, cell-type specific Cre-driver lines can be crossed with the RiboTag mouse (Sanz et al., 2009), which expresses HA-tagged endogenous ribosomal protein (Rrl22), thereby generating mice with specific neurons expressing HA-tagged ribosomes. These can be isolated from mouse brains by immunoprecipitation at different ages and under different conditions (and diseased), and RNA-Seq analysis can identify the ribosome-protected, and therefore, actively translating transcripts. This will be of huge importance in characterizing and understanding the translatome of neuronal compartments. Thus, current technology now offers the exciting possibility of being able to discover differences in the dendritic or axonal translatome of diseased (e.g., autosomal models) individuals. 6. How Is Translation Regulated in Space? How does the spatial morphology of the dendrite, axon, or spine contribute to or constrain protein synthesis? It was recently shown that spines enhance the cooperative interaction among multiple inputs (Harnett et al., 2012). These observations suggest that the amplifying and coordinating properties of dendritic spines have an effect on neuronal input processing and may influence information storage by promoting the induction of clustered forms of synaptic and dendritic plasticity among coactive spines. This could allow spines to enhance the ability of neurons to detect, uniquely respond to, and store distinct synaptic input patterns (Harnett et al., 2012). Different patterns of synapse activation can lead to protein synthesis-dependent or -independent plasticity (Govindarajan et al., 2011). However, the importance and mechanism of specific protein translation remains to be examined in this cooperativity. Since there are mRNAs that are differentially distributed in the length of the dendrites, it is tempting to speculate that there is a role for protein synthesis in regulating the functional compartment in dendrites and spines. Thus, while it is clear that protein synthesis occurs in the dendrite and that it is regulated by neuronal activity, the extent to which the activity of single synapses or synaptic regions stimulates protein synthesis, or alters protein localization, remains unknown. Moreover, the importance and impact of synapse location along the dendrite or axon for protein synthesis is unknown. 7. How Is Translation Coordinated with Degradation? In the small cytoplasmic volume of a dendritic spine or growth cone, there is a limit to the amount of protein that can fit into the space before molecular crowding becomes a problem. While it is clear that changes in synaptic transmission involve extensive regulation of the synaptic proteome via the regulated synthesis and degradation of proteins (Fonseca et al., 2006; Wang et al., 2009), it is not well understood how these two processes are coordinately regulated to achieve the desired level of individual proteins at synapses. Indeed, this is another level of homeostatic control that must exist in order for synapses to maintain the desired level of receptors, scaffolds, and signaling molecules. Changes in the steady-state level of a protein have to be particularly fast and fine-tuned in neurons, due to the fast nature of synaptic transmission and the rapid induction of plasticity. 8. How Specific Is mRNA Translation, and How Is It Regulated? How are specific mRNAs translated and not others? Studies using either global activity manipulations (TTX/APV) (Sutton et al., 2004) or application of an D1/D5 agonist (Hodas et al., 2012) have suggested large-scale (at least ∼100 distinct proteins synthesized) changes in the dendritic proteome. Similarly, global cue stimulation of axons elicits the de novo translation of hundreds of new proteins (Yoon et al., 2012). In these studies, however, the stimulation was applied to the entire network (dish of cultured neurons or brain slice). Under physiological conditions the spatial and temporal profile of synaptic and cue stimulation is on a much finer scale and the translational readout is likely limited. Indeed, we know that different cues can trigger translation of specific subsets of mRNAs in the growth cone (Lin and Holt, 2007). The mechanisms by which specific patterns of synaptic signals (e.g., different frequencies of stimulation, different concentrations or gradients of agonists) and receptor activation lead to activation of the translation machinery are not well understood. Mechanistically, it is clear that elements contained in the 5′ and 3′UTR of mRNAs can regulate their translation initiation. In addition, it is probably the case that the spatial proximity of an mRNA to an active translation site plays a role. The use of high-resolution imaging techniques and focal stimulation should provide answers to these questions. 9. What Roles Do MicroRNA and Other Noncoding RNAs Play in Regulating Local Translation and Neuronal Function? In neurons, the miRNA function has been explored both individually and on a population level, but a broad conceptual understanding is still lacking. Moreover, if miRNAs regulate mRNA translation and expression in different neuronal compartments, what regulates the expression of miRNA themselves? The accessibility of deep sequencing has enabled the detection of other noncoding RNA species in neurons. These additional RNA classes can directly regulate translation, regulate miRNA function, or serve as scaffolds for other molecules, making the levels of regulation and interactions potentially extremely complicated. In addition, the recent appreciation of the abundance and regulatory potential of other noncoding RNAs, mostly in nonneuronal cell types, adds another level of complexity, including the recent demonstration of regulation by circular RNAs that may serve as either shuttles, assembly factories, or sponges for miRNAs and/or RBPs (Hentze and Preiss, 2013). Based on this, it is likely that a real understanding of the complexity of RNA function in neurons will require not only investigation of individual molecules but also a systems biology perspective where the entire network of RNA molecules and their targets can be considered together (see Peláez and Carthew, 2012). 10. Do Specialized Ribosomes Exist, and Can They Tune Translation? While ribosomes are readily visible in dendrites spines (Ostroff et al., 2002) and growth cones (Bassell et al., 1998; Bunge, 1973) how they are transported and whether they are sequestered or anchored is not well understood. A mechanism that could provide specificity or docking would be the specialization of ribosomes by accessory proteins or subunits. One of the most intriguing questions raised by recent work is whether ribosomes are tuned to translating specific mRNAs. This possibility is suggested by recent studies showing that haplo-insufficiency of several different ribosomal proteins give rise to specific phenotypes rather than affecting all cells ubiquitously (Kondrashov et al., 2011; Uechi et al., 2006; Xue and Barna, 2012). This has given rise to the notion of a “ribocode” that suggests heterogeneity in the composition of ribosomes, enabling ribosomes to be tuned to translate specific mRNAs via specific ribosomal proteins (Xue and Barna, 2012). In addition, a striking and curious feature of many recent sequencing studies is the detection of many ribosomal subunits in dendritic or axonal fractions. Indeed, the single most abundant class of mRNAs encode ribosomal proteins in axons (Andreassi et al., 2010; Gumy et al., 2011; Taylor et al., 2009; Zivraj et al., 2010). Thus, an additional intriguing possibility suggested by the abundance of ribosomal protein mRNA in axons and dendrites is that ribosomal proteins may be synthesized de novo. This could provide proteins for in situ repair of ribosomes, or even more interestingly could provide onsite “tuning” of translation (Lee et al., 2013). 11. Does Dysregulated Protein Synthesis Underlie a Wide Range of Neurological Disorders? One of the most exciting clinically relevant findings to emerge from recent work is the link between dysregulated synaptic protein synthesis and neurological disorders (Bear et al., 2008; Darnell and Klann, 2013; Liu-Yesucevitz et al., 2011). Mouse models of neurodevelopmental disorders such as autism spectrum disorder (ASD) show significant improvement on treatment with reagents that target the protein-synthesis pathway (Bear et al., 2008; Darnell and Klann, 2013; Gkogkas et al., 2013; Santini et al., 2013), opening up new possibilities in terms of potential therapeutics. Much of the focus has been on the postsynaptic side of the synapse, the predominant site of plasticity and learning. Recent evidence indicates that regulated protein synthesis in the presynaptic compartment is also important for synapse formation (Taylor et al., 2013) and axon arborization (Hörnberg and Holt, 2013; Hörnberg et al., 2013; Kalous et al., 2013), raising the question of whether defects in axonal protein synthesis contribute to the miswiring aspects of neurodevelopmental disorders. Dysregulated protein synthesis may also underlie a broad range of neurodegenerative disorders (Fallini et al., 2012; Liu-Yesucevitz et al., 2011) consistent with axonal protein synthesis being required for axon maintenance (Hillefors et al., 2007; Yoon et al., 2012). Indeed, the first “effective” oral drug treatment that prevents neurodegeneration in a prion disease/Alzheimer’s mouse model targets a kinase (PERK) that shuts down protein synthesis as part of the unfolded protein response (Moreno et al., 2013). Summary Recent years have witnessed a transformation in our appreciation of RNA function in dendrites/axons on the one hand and of neuronal compartments as spatially distinct signaling/processing units on the other. Here we have highlighted the convergence of these two areas and have sought to define some of the many interesting questions and challenges that lie ahead. As technical approaches become increasingly sensitive for unbiased profiling there is the promise of improved “understanding” of the qualitative concepts that govern the various active RNA species and formation and function of compartments as well as quantitative details on the stoichiometries of all of the players positioned within the morphological framework of the neuron and its remarkable dendritic and axonal arbor.

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Poly(A)-binding proteins: multifunctional scaffolds for the post-transcriptional control of gene expression

David A Mangus, Matthew C. Evans, Allan Jacobson (2003)

Gene organization and evolutionary history RNA-binding proteins are often purified and classified on the basis of the RNA sequences with which they interact [1]. One class of these factors comprises proteins recognizing the homopolymeric polyadenylate tracts that are added to the 3' end of most mRNAs. Poly(A)-binding proteins have been identified in many eukaryotes, but appear to be absent from prokaryotes. PABP genes have been cloned from a number of organisms, and their sequences are available in several databases; a current list with database links is available as an additional data file and on our website [2]. Typically, only one gene encoding cytoplasmic PABP (PABPC) is present in the single-cell eukaryotes, whereas multiple PAPBC genes are present in metazoans and plants (Table 1, Figure 1). A single gene encoding a nuclear PABP (PABPN) has also been identified in cow, frog, human, mouse, fly, worm, and yeasts (Figure 1). A phylogenetic analysis comparing all known PABP protein sequences groups PABPs by organism type (such as metazoans, yeast, and plants) and also identifies similarities among the PABP family members (Figure 1). To date, genes encoding a single nuclear PABP and four cytoplasmic PABPs, as well as four pseudogenes, have been identified in human cells, and their chromosomal locations have been mapped (Table 2). In humans, three lineages of PABP proteins are observed: cytoplasmic PABPs (PABPC1, PABPC3, and iPABP); nuclear PABP (PABPN1); and X-linked PABP (PABPC5). Within the PABPC group, PABPC1 and PABPC3 are most closely related. Interestingly, the mouse gene encoding the alternate PABP, mPABPC2, seems to be a retroposon, as it has no introns and its promoter is distinct from that of mPABPC1 [3]; mPABPC2 is most closely related to hPABPC3, which also lacks introns [4]. Similarly, all the characterized PABPC5 genes lack introns [5], suggesting that they too may be derived from retrotransposition events. A comparable evolutionary analysis was reported for the eight PAB genes identified in the plant Arabidopsis thaliana [6]. Phylogenetic comparisons coupled with expression analyses identified four classes of PABP proteins. In class I (PAB3 and PAB5), expression is limited to reproductive tissue; class II members (PAB2, PAB4 and PAB8) are highly and broadly expressed; class III PABPs (PAB6 and PAB7) have a restricted, weak expression pattern; and the sole member of class IV (PAB1) has low, tissue-specific expression. Comparison of the Arabidopsis PABPs with those from rice indicates that the duplication events which gave rise to classes I-III in flowering plants occurred prior to the divergence of monocots and dicots, more than 200 million years ago [6]. By analyzing the conservation and loss of introns within the PABP gene family, an evolutionary model has been derived in which an ancestral PABP independently gave rise to classes II, III and IV, with class I subsequently derived from class II [6]. Although all eight of the Arabidopsis PABPs are more closely related to the set of nuclear PABPs than to the PABPs of most other eukaryotes (Figure 1), none of these proteins appears to be an authentic PABPN1 species. One interesting characteristic conserved among the PABPC1 genes is an adenylate-rich region in the 5' untranslated region (UTR). Several studies have suggested that PABP regulates its own expression by binding to these sequences [7-9]. Characteristic structural features The association of PABPs with poly(A) requires a minimal binding site of 12 adenosines, and multiple PABP molecules can bind to the same poly(A) tract, forming a repeating unit covering approximately 27 nucleotides [10-13]. In vitro binding affinities of PABP for poly(A) are of the order of 2-7 nM [13-15]. PABPs interact with poly(A) via RNA-recognition motifs (RRMs; Figure 2). The RRM is the most prevalent domain used in the recognition of RNA, as shown by its presence in hundreds of different proteins [16]. RRMs, which are typically 90-100 amino acids in length, appear to be present in proteins in all types of organisms, suggesting that this is an ancestral motif with important functions in RNA biology. Solution nuclear magnetic resonance (NMR) and X-ray crystallographic studies have determined that the RRM is a globular domain composed of a four-stranded anti-parallel β sheet backed by two α helices (Figure 3a) [17]. The central two β strands of each RRM include two highly conserved sequence motifs, octameric RNP1 ((K/R)-G-(F/Y)-(G/A)-F-V-X-(F/Y), where X is any amino acid) and hexameric RNP2 ((L/I)-(F/Y)-(V/I)-(G/K)-(N/G)-(L/M)) (Figure 3a). The electron density map of the human PABPC1-oligo(A) complex identifies eight adenylate residues extending through a trough lined by the β-sheets of the RNPs (Figure 3b) [17]. Specificity for recognition of poly(A) is primarily mediated via van der Waals contacts, hydrogen bonds, and stacking interactions with conserved residues within the RNP motifs [17]. Cytoplasmic PABPs The overall structure of the cytoplasmic PABPs is highly conserved and consists of four RRMs connected to a carboxy-terminal helical domain by an unstructured linker region rich in proline and methionine residues [12,18]. Phylogenetic analysis suggests that the four RRMs arose from successive duplications before the divergence of yeast and mammals [19]. The first two RRMs make up one functional unit and the latter two make up a second. This conclusion is derived partly from the observation that residues participating in RNA recognition within RRM1 are most similar to those in RRM3, while those of RRM2 are most like those of RRM4 [17]. Although each RRM is capable of binding RNA, they are not functionally equivalent, as they have differing affinities for poly(A) [15]. The carboxy-terminal helical domain is highly conserved. In humans it is composed of five helices (Figure 3c), while the yeast protein has only four, lacking an ortholog of the first helix [20,21]. The carboxy-terminal domain is not required for RNA recognition, is dispensable for cell viability in yeast [13,15], and is missing from PABPC5 proteins [5]. This domain is shared with HECT domain proteins in the hyperplastic disc (HYD) family of ubiquitin-protein ligases [22], but there is no evidence that PABPs play any role in protein degradation. The carboxy-terminal domain is, however, the site of interaction with factors regulating polyadenylation, deadenylation, translation initiation, and translation termination (see below). Nuclear PABPs The structure of the nuclear PABPs is not as well understood as that of the cytoplasmic PABPs, largely because crystal and NMR structures have yet to be determined, but it is known that they typically have an acidic amino terminus followed by a single RRM and an arginine-rich carboxy-terminal domain. Recognition of poly(A) requires both the RRM and the arginine-rich domain [23]. A run of alanines in PABN1 is expanded in the recessive disease oculopharyngeal muscular dystrophy (see Figure 2) [24,25]. In yeast, the nuclear PABP is essential for viability and is encoded by the NAB2 gene [26]. Unlike other poly(A)-binding proteins, Nab2p uses an Arg-Gly-Gly (RGG) domain for binding. This protein also contains a Cys-Cys-Cys-His zinc-binding motif, similar to one in RNA polymerase subunits, and a glutamine-rich region that contains a variable number of Gln-Gln-Gln-Pro segments, the number of which is strain-dependent. Localization and function PABPs have crucial roles in the pathways of gene expression. They bind the poly(A) tails of newly synthesized or mature mRNAs and appear to act as cis-acting effectors of specific steps in the polyadenylation, export, translation, and turnover of the transcripts to which they are bound. Lacking any evident catalytic activity, PABPs provide a scaffold for the binding of factors that mediate these steps and also apparently act as antagonists to the binding of factors that enable the terminal steps of mRNA degradation. Polyadenylation Messenger RNAs synthesized in the nucleus generally contain a 3' poly(A) tail; the rare exceptions to this rule are principally the transcripts of replication-dependent histone genes. Newly synthesized poly(A) tails of different mRNAs are relatively homogeneous in length and approximately 200-250 residues in mammals and 70-90 residues in yeast [27]. These poly(A) tracts are not encoded within genes but are added to nascent pre-mRNAs in a two-step processing reaction that involves site-specific cleavage and subsequent polyadenylation of the upstream cleavage product [23,28-30]. Throughout eukaryotes, pre-mRNA cleavage and polyadenylation take place in a large complex (500-1,000 kDa) that includes poly(A) polymerase (PAP) and many additional factors. In general, the factors regulating PAP stimulate both the specificity and processivity of an otherwise marginally active and indiscriminate enzyme. In so doing, they not only regulate the process of polyadenylation but also determine the ultimate size of the poly(A) tail. In mammalian cells, PABPN1 binds nascent tracts of 11-14 adenylate residues [31] and, along with cleavage and polyadenylation specificity factor (CPSF), stimulates PAP to switch from distributive synthesis (dropping off after synthesis of a few nucleotides) to processive (continuous, highspeed) synthesis [32,33]. PABPN1 monomers continue to bind available, nascent adenylates until the full-length poly(A) tail has been synthesized and the polymerase then reverts back to its distributive mode [34]. This sequential binding is accompanied by the formation of linear filaments and 21 nm spherical particles: the latter are thought to serve as 'molecular rulers' that dictate the final length of the poly(A) tail [34]. In this model, the particle is postulated to encompass a stable polyadenylation complex and to tolerate PABPN1-poly(A) oligomers until the tail reaches 200-300 nucleotides. Beyond that point, increased poly(A) length is believed to be compromised by disruption of critical interactions between PAP and CPSF [34]. PABPs also play a role in the polyadenylation of yeast pre-mRNAs. Recent studies indicate that Nab2p is the most likely candidate for the yeast equivalent of PABPN1 function, at least for a subset of mRNAs. Mutations in NAB2 promote hyperpolyadenylation of mRNA that cannot be reversed by overexpression of Pab1p [35]. The failure to detect this activity of Nab2p in earlier studies may be attributable to inhibitory interactions between Nab2p and its nuclear import receptor Kap104p, and/or to the preponderance of Pab1p in whole-cell extracts used for in vitro polyadenylation and the consequent obstruction of Nab2p activity by Pab1p bound to nascent poly(A) [35]. Interestingly, mutations in the yeast gene encoding cytoplasmic PABP, PAB1, cause a significant increase in mRNA poly(A) tail lengths in vivo and in vitro [36-38], and this effect, too, is partly attributable to a switch of PAP (Pap1p) between processive and distributive activities. Unlike the process in mammalian cells, the yeast switch appears to be directly regulated by Fip1p and Yth1p, two factors unrelated to nuclear or cytoplasmic PABPs, and only indirectly regulated by Pab1p [39-41]. Pab1p interactions underlying this indirect effect may include its binding to the nascent mRNA [28] or a direct interaction with the RNA-processing factor Rna15p [37]. Evidence for a direct role for Pab1p in yeast poly(A) length control comes from experiments analyzing the Pab1p-mediated regulation of poly(A) nuclease (PAN). This exonuclease, comprising the Pan2p and Pan3p proteins, appears to trim up to 20 residues from excessively long newly synthesized poly(A) tails in an mRNA-specific manner [42-44]. Pan2p, the subunit with apparent exonuclease activity, is positively and negatively regulated by interactions with Pan3p and Pbp1p, respectively; both of the latter interact with Pab1p (D.M. and A.J., unpublished observations; [43-45]). Nuclear export A second role for PABPs in the nuclear maturation of mRNA can be inferred from experiments in which impaired 3' processing interferes with export of mRNAs to the cytoplasm. In both mammalian cells and yeast, mRNAs are generally retained in the nucleus when they lack a functional polyadenylation signal or when polyadenylation is inhibited by the absence or inactivity of specific catalytic factors [46-49]. Since the failure to polyadenylate an mRNA would deprive it of bound PABPs, nuclear retention of mRNA could be attributable to an essential role for PABPs in mRNA export. As noted above, PABPs coat the nascent poly(A) tail and play a role in determining its ultimate length. How, then, might this poly(A)-PABP complex facilitate the exit of mRNAs and their associated proteins (mRNPs) from the nucleus? Consistent with the propensity of PABPs to form interactions critical to specific functions, yeast Pab1p has been shown to interact with specific nucleoporins [50] and the nuclear export signal export receptor, Xpo1p [49], and Nab2p has been shown to interact with Gfd1p, a nuclear-pore-associated protein [51]. The presence of bound Pab1p or Nab2p could serve as a determinant of an mRNP's export competence, in a manner analogous to the function of the RNA export factor Yra1p [52]. This view is consistent with the observed nucleocytoplasmic shuttling of yeast and mammalian PABPs [53-56] and with the inhibitory effects on mRNA export caused by interactions between the influenza virus NS1A protein and PABPN1 [57]. The notion of a direct role for PABPs in mRNA export may, however, be too simplistic. It does not accommodate examples of mRNAs that enter the cytoplasm without conventional 3' processing [58,59], viable mutants devoid of PABP [36], or functional interactions between the 3' processing apparatus and the factors that promote mRNA export [49,60]. The latter reflect a quality control mechanism that leads to retention of an mRNA in the nucleus (often at its transcription site) in the event of processing problems [49,61,62]. This apparent checkpoint illustrates the interdependence of many steps in gene expression and the manner in which such regulatory mechanisms can make indirect effects appear to be direct. Translation initiation After an mRNA enters the cytoplasm, the association of PABP with its poly(A) tail promotes 5'-3' interactions that stimulate initiation of its translation [27,63]. Formation of this 'closed loop' [27] was shown by Sachs and colleagues [64-66] to promote the recruitment of 40S ribosomal subunits and to be dependent, at a minimum, on interactions between initiation factor eIF4G and PABP and concurrent interactions between eIF4G and the cap-binding protein eIF4E (Figure 4). The existence of a translational regulatory network involving PABP, eIF4G, and eIF4E is consistent with the impaired-translation phenotypes of yeast strains lacking functional Pab1p [36] and provides a mechanistic basis for the synergistic effects on translation known to occur when mRNAs are both capped and polyadenylated [65,67,68]. The combined cooperative interactions enhance the affinity of eIF4E for the 5' cap of the mRNA by lowering its dissociation rate [69-72], stimulate the RNA-binding activity of PABP [73], and increase the ATPase and RNA helicase activities of eIF4A, eIF4B, and eIF(iso)4F [74]. The combination of these effects also provides an effective means for the protein synthesis apparatus to ensure preferential translation of mRNAs containing both a cap and a poly(A) tail [74] and may create an opportunity for ribosomes to recycle from the 3' to the 5' end of the same mRNA [27,75]. Studies in yeast and mammalian cells have shown that the Pab1p-eIF4G interaction requires RRM1 and RRM2 of Pab1p (the same RRMs required for poly(A) recognition) and an amino-terminal domain of eIF4G [65,76-78]. Several additional experiments have indicated, however, that the network of 5'-3' interactions regulating translation initiation goes well beyond the communication of a single domain in PABP with another in eIF4G. This was initially suggested by the existence of viable yeast pab1 mutants in which the Pab1p-eIF4G interaction could not occur [65] and others that had defects in poly(A)-dependent translation but no defects in eIF4G binding [76]. The potential complexity of PABP's translation-promoting interactions is illustrated by interactions of PABPs in wheat germ with the initiation factor eIF4B [73] and in mammals with the PABP-interacting proteins Paip1 and Paip2 [79-82]. Paip1 is homologous to the central segment of mammalian eIF4G and binds with high affinity and 1:1 stoichiometry to two sites in PABP, one in RRMs 1 and 2 and the other in the carboxy-terminal domain [79,80]. The region of eIF4G to which Paip1 is homologous encompasses one of two binding sites for the RNA helicase eIF4A. Not surprisingly, Paip1 also interacts with eIF4A, and is capable of stimulating the translation of a reporter mRNA when overexpressed in cultured cells [79]. Paip2, a low-molecular-weight acidic protein, binds PABP at two sites, one in RRMs 2 and 3 and one in the carboxyl terminus [81,82]. Binding of Paip2 to the RRM2-3 region competes effectively for binding of Paip1 to PABP, reduces PABP binding to poly(A), and inhibits the translation of polyadenylated mRNA [81,82]. Tethered-function assays in yeast and Xenopus that exploit PABP fusions to the bacteriophage MS2 coat protein also underscore the intricate nature of PABP's stimulatory effects on translation [83]. PABP tethered at specific MS2 coat binding sites stimulates translation of a reporter mRNA in cis, but not in trans, and can do so without its poly(A)-binding activity and in the absence of a poly(A) tail [83]. With the exception of the yeast requirement that Pab1p be bound to poly(A) in order to interact with eIF4G [64], this implies that, at least with respect to translational stimulation, poly(A) simply provides a binding site for PABP. The failure of yeast Pab1p to function in the absence of bound poly(A) may reflect the selective inability of yeast eIF4G to stabilize the packing of poly(A)-associated RRMs 1 and 2 in a manner comparable to that achieved by the eIF4Gs of other species [84]. Tethered function assays also reveal that RRMs 1 and 2, or RRMs 3 and 4, of Xenopus PABP are as capable of translational stimulation as the full-length protein, despite the fact that RRMs 3 and 4 lack the ability to interact with eIF4G or Paip1 [83]. Like the Pab1p-eIF4G interaction mutants in yeast [65,76], the novel PABP interactors in mammals and plants [73,79-82], and the unique domain requirements for transactivation of translation by Pab1p [78], this observation implies that interaction with eIF4G is not likely to be the only mechanism by which PABP stimulates translation. One alternative model for PABP function, supported by genetic analyses in yeast [85] and the biochemical properties of poly(A)-deficient mRNAs in vitro [67], suggests that PABP is also a regulator of the joining of the 60S subunit to the 40S preinitiation complex. The studies in yeast indicate that PABP controls 60S joining by regulating the activities of two RNA heli-cases, Ski2p and Slh1p [85]. Additional insight into the translational networks affected by the presence of PABP is derived from studies of the tactics that viruses and cells use to modulate PABP structure and/or activity. For example, rotaviruses reroute translation for their own purposes by synthesizing a protein, NSP3, which serves as a PABP analog. NSP3 binds to specific 3' sequences on viral mRNAs and effectively circularizes those transcripts, and mimics PABP, by also binding to eIF4G [86]. Enteroviruses, on the other hand, choose to eliminate the activity of PABP, rather than replace it. As part of a general assault on host cap-dependent translation, these viruses express two proteases, 2A and 3C, that not only remove the PABP-interacting domain of eIF4G but also cleave PABP into several fragments [87,88]. PABP interactions and activity, at least in plants, are also altered by changes in its phosphorylation status [89] and may be affected by arginine methylation within the domain separating RRM4 from the carboxy-terminal helices [90]. In addition to their global effects on translation initiation, PABPs can also selectively affect the translation of individual mRNAs. PABPs can bind oligoadenylate tracts in the 5' UTRs of their own mRNAs, thereby repressing their own translation (and possibly their stability [17]) [9,91]. This autoregulation can be mimicked both in vitro and in vivo, can be abolished by deletion of the adenylate-rich region and can be conferred on other mRNAs by insertion of the adenylate-rich tract within their 5' UTRs. In each case, the presence of PABP is required to mediate the observed effects. The inhibition of translation has been ascribed to an inability of the 60S ribosomal subunit to join the pre-initiation complex [92]. PABP can also facilitate the binding of translational repressors specific for other mRNAs, such as that encoding the iron-oxidizing protein ceruloplasmin [93], and can activate the translation of a large number of mRNAs whose polyadenylation is developmentally controlled [94], as well as functioning as an mRNA-specific activator. Cytoplasmic PABP in Chlamydomonas reinhardtii, normally a 69 kDa polypeptide, is imported into chloroplasts where it is processed to a 47 kDa form that binds the 5' UTR of the psbA mRNA and activates its translation [95]. The latter role of PABP is particularly intriguing in light of the generally prokaryotic nature of chloroplast translation systems. Translation termination The eukaryotic translation termination factor eRF1, which is responsible for catalyzing polypeptide hydrolysis in response to recognition of any of the three nonsense codons by the ribosome, appears to be activated by the GTPase eRF3 [96]. The amino-terminal region of eRF3 does not participate in this interaction with eRF1, but does interact directly with the carboxy-terminal domain of cytoplasmic PABPs [21,97,98]. The eRF3-PABP interaction appears to enhance the efficiency of termination in cells with mutated or aggregated eRF3 [98] and to promote ribosome recycling for multiple rounds of translation on the same mRNA [99]. It also seems to minimize the multimerization of PABP monomers on poly(A), possibly expediting access of poly(A) shortening enzymes to their substrate and linking translational termination to normal mRNA decay [97]. Additional insights into the role of PABPs in translation termination come from analyses of instances in which termination occurs abnormally, such as at premature nonsense codons. In this case, termination is thought to be aberrant because of the creation of a 'faux' UTR, an untranslated region lacking at least one of the factors required for efficient polypeptide hydrolysis and ribosome release that are normally positioned 3' to a termination codon by interaction with poly(A)-associated PABP [100]. Decay of mRNA The process of mRNA decay can be initiated by three distinct events: endonucleolytic cleavage, removal of the 5' cap, and poly(A) shortening [101]. In yeast, in which the process of mRNA decay has been extensively analyzed, most wild-type mRNAs decay by a mechanism in which the initial nucleolytic event is the shortening of the poly(A) tail to an oligo(A) length of 10-15 nucleotides. After poly(A) shortening, transcripts are decapped by the Dcp1p-Dcp2p complex. Decapped and deadenylated mRNAs are then digested exonucleolytically by the 5'-to-3' exoribonuclease, Xrn1p, and/or the 3'-to-5' multi-subunit exosome [102] (Figure 4). All three decay-initiating events eliminate the closed-loop state of the mRNP by removing or separating the binding sites for the respective 5' and 3' interacting proteins [27], and these events also render the remaining mRNA fragments substrates for further degradation. At a minimum, then, mRNA decay generally occurs concurrently with the conversion of an mRNP from a translatable to an untranslatable (or poorly translatable) form [27,101], that is, in parallel with the termination of PABP's role in the enhancement of translation initiation. Although the onset of mRNA decapping does coincide with the loss of PABP's binding site, and efficient translation initiation does, indeed, antagonize mRNA decay [103,104], PABP's role in the maintenance of mRNA stability is more complicated than that of a mere translation enhancer. Several observations suggest that loss of PABP's binding site, and presumptive disruption of the closed loop state, may not always trigger immediate degradation of the remainder of the mRNA. These observations include, first, that poly(A) shortening or removal is the rate-determining event in the decay of some mRNAs whereas for others, it may be an obligate event in their degradation but not the rate-determining step [105,106]; second, that yeast pab1 mutations that unlink mRNA decapping from poly(A) shortening do not necessarily accelerate the rate of mRNA decay [107,108]; and third, that the domains of tethered Pab1p that provide yeast mRNA stabilization and translation functions are different [83,109]. Additional roles for PABP in the regulation of mRNA stability range from being an antagonist or promoter of poly(A) shortening to a facilitator of the binding of additional factors that promote or retard rapid mRNA decay. In vitro, excess poly(A) is an effective competitor of PABP binding to mRNA [63,110-112]. Such competition accelerates the rate of poly(A) shortening, indicating that the presence of PABP on the poly(A) tail provides a protective effect [110-112]. This effect is, in part, attributable to physical hindrance of the deadenylase, because poly(A) tails are often shortened in discrete lengths equivalent in size to a PABP 'footprint' [113]. It is also known, however, that the principal yeast deadenylase, the Ccr4p-Pop2p-Notp complex (Figure 4), and a major mammalian deadenylase, PARN, are both inhibited in the presence of PABP [114]. In contrast, the yeast Pan2p-Pan3p deadenylase, an enzyme responsible for the initial trimming of the poly(A) tail (see above), requires Pab1p for its activity [42,44]. Much like its role in translational initiation, PABP also influences mRNA decay by interacting with key regulatory proteins, either influencing their activity or being influenced by them. Two proteins that bind the 3' UTR of the α-globin mRNA and enhance its stability, αCP1 and αCP2, interact with human PABP [113]. PABP appears to stimulate the ability of the αCP proteins to bind to their target sequence in the 3' UTR, thereby precluding access of an endonuclease (ErEN) to its recognition site [115]. Interestingly, the binding of PABP to the poly(A) tail is also enhanced by the αCP proteins, implying that α-globin mRNA stabilization is mediated by multiple interdependent events [115]. Stability of the mRNA encoding the transcription factor c-Fos is regulated by sequence elements in its 3' UTR and coding region [116]. The coding region stability element, also known as the major protein-coding-region determinant (mCRD), interacts with a complex of RNA-binding proteins that includes PABP, Paip1, hnRNPD, NSAP1, and Unr [117]. Translation through the mCRD destabilizes c-fos mRNA by a mechanism that is thought to disrupt interactions with this complex and, in turn, promote poly(A) shortening [117]. As noted above, PABP also interacts with the termination factor eRF3 [21,97,98], a consequence of which is a decrease in the number of PABP multimers associated with the poly(A) tail. This observation links translation termination to poly(A) shortening and suggests one mechanism for orchestrating a standardized 'clock' that limits the lifetime of a poly(A) tail and, in turn, the mRNA to which it is appended [97]. Additional roles for PABP in mRNA decay are illustrated by events that occur after the poly(A) tail has been removed. As shown in Figure 4, mRNA deadenylation is accompanied by an mRNP rearrangement that allows binding of a decapping activator complex containing the proteins Lsm1p-Lsm7p and Pat1p [118]. This complex appears to promote interaction of the mRNP with the Dcp1p-Dcp2p decapping complex, thereby creating a substrate for terminal 5'-to-3', and/or 3'-to-s' exonucleolytic degradation [119] (Figure 4). Recent studies indicate that all steps subsequent to association of the Lsm1-7p-Pat1p complex occur at a limited number of subcellular sites called P bodies [119]. In principle, therefore, both the terminal steps of mRNA decay (from decapping onwards), and their localization to a specific subcellular site, are prevented from occurring by the presence of bound Pab1p. Pab1p may simply maintain the mRNP in its translation-favorable mode, but the formal possibility that it directly inhibits mRNA association of the Lsm1-7p-Pat1p complex has not been excluded. In the latter hypothesis, PABP's exit from the mRNP would complete a cycle in which its initial association with mRNA assists in mRNP formation, then leads to efficient mRNP utilization, and culminates in destruction of the mRNA. Frontiers Considering the number of functions associated with the PABPs, and their simultaneous interactions with both RNA and other proteins, the number of questions for which we have no answers far exceeds the number of those for which we do. Does the presence or absence of PABP determine an mRNP's competence for export, or does it play a more active role? How is nuclear PABP exchanged for cytoplasmic PABP and where does that exchange take place? How does interaction with PABP actually affect eIF4G and eRF3, and vice versa; in other words, do these proteins influence each other's conformations and interactions with other factors? Does autoregulatory PABP simply 'block' the 5' UTR or does it promote interactions with other factors that are the ultimate regulators? Why do plants have so many PABP genes? Have they separated PABP's functions into distinct polypep-tides? A key question is which of PABP's many functions are essential. Of the functions enumerated in this review, most appear to be dispensable. For example, PABP mutants lacking the ability to interact with factors governing polyadenylation, translation initiation, and translation termination are all viable. Cross-species complementation experiments assessing the essential nature of PABP demonstrate that Arabidopsis Pab3p can restore PABP's role in mRNA biogenesis but fails to complement defects in mRNA decay and translation initiation [120]. What does appear to be required is the ability of PABP to recognize RNA. The possibility remains that the essential nature of PABP lies not with a single function but with a combination of functions. That, of course, raises the final question: have all of PABP's functions been enumerated? That seems unlikely. A hint of PABP's untapped versatility is apparent from its role in the replication of zucchini yellow mosaic potyvirus, a plant virus whose RNA-dependent RNA polymerase appears to exploit PABP for viral replication [121]. Who knows - maybe PABP will find its way into splicing and transcription, completing its act as the one-man band of gene expression. Additional data files A list of the currently known PABP genes with accession numbers and links to their entries in the nucleotide and protein sequence databases (Additional data file 1) and the sequences of these proteins in FASTA format (Additional data file 2) are available with the online version of this article. Supplementary Material Additional data file 1 A list of the currently known PABP genes with accession numbers and links to their entries in the nucleotide and protein sequence databases Click here for additional data file Additional data file 2 The sequences of the currently known PABP proteins in FASTA format Click here for additional data file

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Journal ID (nlm-ta): Biochem Soc Trans

Journal ID (iso-abbrev): Biochem. Soc. Trans

Journal ID (pmc): bst

Journal ID (publisher-id): BST

Title: Biochemical Society Transactions

Publisher: Portland Press Ltd.

ISSN (Print): 0300-5127

ISSN (Electronic): 1470-8752

Publication date (Electronic): 11 August 2014

Publication date (Print): 1 August 2014

Volume: 42

Issue: Pt 4

Pages: 1229-1237

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*MRC Centre for Reproductive Health, Queen's Medical Research Institute, University of Edinburgh, 47 Little France Crescent, Edinburgh EH16 4TJ, Scotland, U.K.

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¹Correspondence may be addressed to either of these authors (email r.smith@ 123456ed.ac.uk or nicola.gray@ 123456ed.ac.uk ).

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Publisher ID: BST20140111

DOI: 10.1042/BST20140111

PMC ID: 4128646

PubMed ID: 25110030

SO-VID: 0f70eeca-ecdb-4073-b498-a7d98eb2efd6

Copyright © © 2014 The author(s) This is an Open Access article distributed under the terms of the Creative Commons Attribution Licence (CC-BY)(http://creativecommons.org/licenses/by/3.0/) which permits unrestricted use, distribution and reproduction in any medium, provided the original work is properly cited.

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