- Email: [email protected]
Cellular strategies for the assembly of molecular machines
Review
Cellular strategies for the assembly of molecular machines Ashwin Chari and Utz Fischer Department of Biochemistry, Theodor-Boveri Institute, University of Wurzburg, Am Hubland, D-97074 Wurzburg, Germany
Molecular machines are supramolecular assemblies of biomolecules (proteins, RNA and/or DNA) that facilitate a diversity of biological tasks in the cells of all organisms. How these complex structures are built within the crowded cellular environment is, therefore, a central question in the biological sciences. Recent studies on spliceosomal uridine-rich small nuclear ribonucleoproteins (snRNPs) have unveiled cellular assembly strategies for RNA–protein complexes. snRNPs form in vivo by the coordinated action of an elaborate assembly line consisting of assembly chaperones, scaffolding proteins and catalysts. These newly discovered strategies exhibit similarities to those employed by protein complexes such as ribulose-1,5-bisphosphate-carboxylase (Rubisco) and allow the elucidation of general rules for how molecular machines are formed in vivo. Assembly lines in our daily life and at the molecular level Life in our technologically advanced society is greatly simplified by a multitude of gadgets which help us perform our daily tasks. To assemble these machines, pre-manufactured components and an industrial manufacturing design (blueprint) are required. This blueprint essentially specifies where each individual part of the machine should be located in three dimensions and at which temporal stage in assembly the parts should be incorporated into the final product. Then, factory assembly lines add all the components together in a sequential manner, as specified in the blueprint. Macromolecular complexes perform many biochemical reactions crucial for the survival of organisms and are, thus, rightfully also referred to as biological machines [1,2]. Although we understand the underlying mechanisms for many of the biochemical reactions catalyzed by these machines, knowledge about how these machines are assembled in vivo is, in several instances, less complete. An emerging view is that the formation of macromolecular complexes utilizes principles similar to those used in an industrial assembly line. Biological complexes are formed when proteins either bind to each other or to nucleic acids. Prominent examples include the plant enzyme ribulose-1,5-bisphosphatecarboxylase (Rubisco, protein only), the nucleosome (DNA and protein) and the spliceosome (RNA and protein) (Table 1). How are these entities built within cells? Early studies demonstrated that it is possible to reconstitute nucleosomes, or even the 2.6 MDa ribosome, from purified constituents into active assemblies in vitro [3–6]. The structural Corresponding authors: Chari, A. ([email protected]); Fischer, U. ([email protected])
676
information (blueprint) for the formation of macromolecular complexes is therefore encoded within its subunits. Coupled with observations that many molecular machines can be assembled in heterologous expression systems, these findings have given rise to the notion that the machines of life are built spontaneously without the requirement for accessory factors. Despite this general tendency for self-assembly of macromolecular complexes in vitro, it is becoming increasingly clear that assisting factors are required for efficient assembly in vivo. Many of these accessory factors belong to an unrelated group of proteins termed assembly chaperones which act as a cellular counterpart to industrial assembly lines [7,8]. They associate with the individual subunits of a macromolecular complex and promote the formation of the final structure (Figure 1A). In this review, we initially discuss pitfalls encountered during productive macromolecular complex assembly that are introduced by certain properties of the cellular milieu. We then describe strategies employed by cells to overcome these impediments. Rubisco and spliceosomal small nuclear ribonucleoproteins (snRNPs) were chosen for this discussion because both represent well-studied particles exemplifying two distinct classes, namely protein–protein and RNA–protein complexes. Finally, we will delineate common principles that underlie both assembly processes, lending support to the view that the strategies employed for the assembly of these complexes also apply to the assembly of other macromolecular complexes such as proteasomes and ribosomes (Table 1) [9–13]. Principles governing macromolecular complex assembly in vivo From a biophysical point of view, the assembly of macromolecular complexes depends on the diffusion-driven, random collision of subunits [14]. The stability of the resulting assembly is determined by the ratio of the individual dissociation and association rate constants. However, within cells variations to this simplified view appear to be prevalent. First, many subunits of an assembly often complete their individual folding reaction only upon association with their interaction partner(s) [15,16]. Second, the interaction between two or more subunits often involves hydrophobic stretches that are exposed in the individual folded subunit [17]. This property renders isolated proteins of a macromolecular complex particularly prone to irreversible aggregation within cells [18]. Third, an additional pitfall towards the formation of macromolecular entities arises from the fact that, in general, the cellular milieu does not behave as an ideal fluid. Instead, the interior of cells is characterized by a
0968-0004/$ – see front matter ß 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.tibs.2010.07.006 Trends in Biochemical Sciences, December 2010, Vol. 35, No. 12
Review
Trends in Biochemical Sciences
Table 1. Selected list of complexes known to be assembled by assisting factors Macromolecular complex assembly process Bacterial inner membrane proteins Bacterial outer membrane proteins Bacterial pilus Mitochondrial membrane proteins Nucleosomes Eukaryotic ribosomes 20S and 26S proteasomes Form I Rubisco Spliceosomal snRNPs
Refs [73] [74] [75–77] [78] [8,79–81] [9,10] [8,11–13] [28,30,31,33] [55,56,62,71]
high total concentration of many different macromolecules. As a consequence, no single species accumulates in high amounts, but in total, proteins and other macromolecules are so abundant that they occupy 20–30% of the interior of cells. Therefore, a large percentage of the cellular volume is unavailable to other molecules leading to an excluded volume effect that is commonly referred to as macromolecular crowding (Figure 1B) [19,20]. This property of cells lowers the diffusion rate of individual subunits within a cell [21]. Thus, the likelihood of random encounters between the building blocks of a macromolecular entity is expected to be strongly reduced. Simultaneously, specific, as well as nonspecific, association reactions are strongly favored under crowding conditions [20]. Therefore, the propensity for productive interactions between cognate molecules to occur within cells is substantially lowered, resulting in misassembly [22]. [(Figure_1)TD$IG]
Vol.35 No.12
Strategies utilized by cells to encounter pitfalls towards faithful macromolecular complex assembly Cells have evolved several elaborate strategies to encounter the obstacles that can counteract faithful macromolecular assembly. First, the synthesis of the individual constituents of a macromolecular entity is often temporally tightly controlled. This strategy ensures that all components of a macromolecular complex are simultaneously present in sufficient amounts when assembly is initiated. Second, the synthesis of the individual building blocks of a macromolecular machine, their assembly into functional particles and finally their integration into the designated process, by which they help maintain cellular homeostasis, are in many cases spatially segregated. This strategy is particularly (but not exclusively) dominant in the case of RNA–protein complexes (RNPs) and DNA–protein complexes (DNPs), where the nucleic acid moiety is synthesized in the nucleus and the proteins in the cytoplasm. This compartmentalization of synthesis and assembly does not necessarily require organelles, which are surrounded by membranes. In particular, the variety of nuclear subdomains also effectively achieves spatial segregation; a prominent example is the nucleolus in ribosome assembly [9,10]. Spatial separation of assembly and function is also a crucial means by which the interference of biochemical processes of the fully assembled machine by partially assembled complexes is prevented.
Figure 1. (a) The formation of biological machines. The formation of biological machines requires a blueprint, which is intrinsic to the structure of DNA and protein components (histone H3–H4 dimer, from PDB ID: 1TZY). A chaperone then acts as a counterpart for the industrial assembly line within cells (the chaperone Asf1 bound to the histone H3–H4 dimer, PDB ID: 2HUE). It associates with the protein components and orchestrates the formation of the final product; in this case a nucleosome core particle (PDB ID: 1AOI). (b) The cytoplasm of cells is a crowded environment. The illustration depicts an artist’s impression of macromolecular crowding conditions within a cell. A small portion of eukaryotic cytoplasm showing microtubules (light blue), actin filaments (dark blue), ribosomes (yellow and purple), soluble proteins (light blue), kinesin (red), small molecules (white) and RNA (pink) is shown. Illustration by D.S. Goodsell, the Scripps Research Institute.
677
Review Third, cells often utilize the activity of assembly chaperones. These proteins enable the formation of intermediate higher-order structures, thereby ensuring a spatial pre-organization of individual subunits in positions, which they also occupy in the assembled macromolecular complex. Simultaneously, they prevent the premature occurrence of steps that are meant to occur at a later stage in assembly as well as dead-end reactions. This temporal and quality control often leads to the formation of kinetically trapped assembly intermediates, which require the activity of additional factors for assembly to proceed. Fourth, in addition to chaperones, the assembly process of a molecular machine is often assisted by a heterogeneous group of trans-acting factors. These proteins typically enable the resolution of kinetic traps that were introduced by the assembly chaperones. In addition, they stimulate assembly by increasing the local concentration of the individual subunits (or assembly intermediates) of the complex. This activity can sometimes be achieved by subunits of the macromolecular machine itself. In many instances, the fulfillment of one strategy is the prerequisite for the next step, implying a strong synergistic relation between each individual step in biogenesis. Each of these cellular strategies has evolved to ensure the proper and faithful formation of macromolecular complexes in vivo. Assisted assembly of a protein complex: lessons from the Rubisco holoenzyme Rubisco is arguably one of the most important enzymes for life because it catalyzes the primary reaction by which [(Figure_2)TD$IG]
Trends in Biochemical Sciences Vol.35 No.12
inorganic carbon is chemically fixed in the biosphere. This is achieved in the Calvin cycle, where the fixation of CO2 results in the synthesis of usable sugars [23]. Rubisco also catalyses an opposing reaction, termed photorespiration, in which O2 is the substrate. Photorespiration competes with, and determines the efficiency of, carbon fixation [24]. Three forms of quaternary structure for otherwise closely related Rubisco subunits are known [25]. Form I, which is found in plants and cyanobacteria, is a hexadecamer RbcL8S8, containing eight large subunits (RbcL) and eight small subunits (RbcS). Forms II and III Rubisco, by contrast, are much simpler in terms of their biochemical composition and consist of only one (Form II) or five (Form III) RbcL dimers [26]. We will restrict our discussion to the assembly of the form I Rubisco holoenzyme because it has been studied in the greatest detail (Figure 2). The different steps in Rubisco assembly illustrate the four major strategies evolved by cells to encounter the obstacles that can counteract faithful macromolecular assembly. Whereas the eight large subunits (RbcL) of the form I Rubisco holoenzyme are encoded in the chloroplast genome, the eight small subunits (RbcS) are expressed from the nuclear genome. Upon stimulation with light, the syntheses of both chloroplast and nuclear mRNAs are coordinated [27–29], thereby ensuring that both subunits are present simultaneously in sufficient amounts when assembly is initiated. The cytoplasmic synthesis of RbcS makes a subcellular transport event to the chloroplast, the organelle where RbcL is expressed, inevitable. The formation of the form I Rubisco holoenzyme is
Figure 2. Model for the chaperone assisted folding and assembly of Rubisco. Newly synthesized RbcL (green) is sequestered and folded by the chaperonins GroEL (light blue) and GroES (dark blue) (i). The assembly chaperone RbcX2 (yellow) binds to a disordered C-terminal peptide of RbcL (ii). This interaction induces partial folding of the RbcL C-terminus and the formation of RbcL dimers (iii). Stable dimers assemble into RbcL8–(RbcX2)8 complexes (iv). RbcS (gray) binding dissociates RbcX2, which might occur concertedly (v). The catalytically mature Rubisco holoenzyme is then released (vi). Figure modified, with permission, from [33].
678
Review dependent on the molecular chaperones GroEL, GroES and RbcX [30,31]. In the first phase, the chaperonin GroEL, together with its co-chaperonin GroES, ensures proper folding of the RbcL subunit and thus represents a quality control checkpoint for Rubisco formation (Figure 2i) [32]. During the folding reaction, RbcL monomers are transiently released from GroEL [33]. Upon release, the partially disordered C-terminal region of RbcL is recognized by the dimeric assembly chaperone RbcX2 (Figure 2ii). RbcX2 then induces the formation of stable RbcL dimers, which imparts partial folding in the C-terminus of RbcL (Figure 2iii). As a result, the RbcX2-bound RbcL dimers assemble into tetramers forming the RbcL8–(RbcX2)8 complex (Figure 2iv). RbcL8–(RbcX2)8 can be considered a catalytically immature roadblock in Rubisco holoenzyme assembly. Finally, this roadblock is salvaged by the Rubisco small subunit, RbcS (Figure 2v). Its association with RbcL8–(RbcX2)8 triggers a conformational change in RbcL, thereby eliciting RbcX2 dissociation and holoenzyme formation (Figure 2vi). This final assembly step is coincident with the attainment of catalytic maturity of Rubisco. Spliceosomal snRNPs have a segmented biogenesis pathway Although the major principles of protein complex assembly have been established, much less is known about the formation of molecular complexes composed of RNA and proteins (RNPs) [34]. The spliceosomal snRNPs provide an ideal model for the analysis of RNP assembly strategies because their in vivo maturation pathway can be faithfully recapitulated in vitro and dissected biochemically [35]. snRNPs are abundant nuclear complexes and the major components of the nuclear splicing machinery [36]. This entity catalyzes the removal of introns from mRNA precursors (pre-mRNAs) and the precise joining of exons to yield a functional mRNA. Four different snRNPs are part of the major spliceosome, named after their RNA moiety: U1, U2, U4/6 or U5. A second, so-called minor spliceosome, also exists, in which the snRNPs U1, U2, U4/U6 are replaced by U11, U12 and U4atac/U6atac, respectively. The minor spliceosome catalyzes the splicing of a small subset of introns [37]. It is assumed, but not yet experimentally proven, that the principles discussed here for the major snRNPs are identical for the minor snRNPs. Each individual particle contains a set of snRNP-specific proteins (for example U1-70K, U1A & U1C in the case of the U1 snRNP) and assumes a designated role in splicing (Figure 3) [35,36]. In addition, the U1, U2, U4 and U5 snRNPs also contain a set of seven common proteins termed B/B0 , D1, D2, D3, E, F and G; they are often also referred to as the Smith (Sm) antigen because patients suffering from rheumatoid diseases (including patient Smith) frequently develop antibodies against these factors [38]. The seven Sm proteins bind and encircle a singlestranded region (Sm site) of the snRNAs and form a toroidal RNP core which constitutes the structural framework of all snRNPs. Only the U6 snRNP is divergent in this respect. Instead of the seven canonical Sm proteins, it contains a set of seven evolutionarily related proteins ‘like Sm proteins 2–8 (LSm2–8)’ [39].
Trends in Biochemical Sciences
Vol.35 No.12
snRNP biogenesis is an intricate, segmented pathway (Figure 3) [35]. It begins with the RNA polymerase II-dependent transcription of a monomethyl-guanosine (m7G) capped snRNA-precursor, followed by its subsequent nuclear export [40,41]. Within the cytoplasm, the snRNA encounters Sm proteins, which are synthesized and stockpiled in this compartment [42]. Sm proteins then bind to the Sm site and form the Sm core domain. This event, in turn, triggers the hypermethylation of the m7G-cap, thereby generating the trimethylguanosine (TMG) m2,2,73G-cap [43,44]. A bipartite nuclear localization signal (NLS) consisting of the assembled Sm core domain and the TMG cap ensures the translocation of snRNPs to the nucleus [45–47]. Before they reach their site of function in the nucleoplasm, snRNPs are transiently directed to subnuclear domains termed Cajal bodies, where further maturation steps, such as modification of the RNA moiety, complete the biogenesis cycle [48–50]. The biogenesis pathway ends when the snRNP particles leave Cajal bodies and become part of the splicing machinery within the nucleoplasm. It is still largely unknown at which stages in the biogenesis cycle specific proteins join the assembled snRNP. The U6 snRNP does not follow the biogenesis pathway outlined above. Instead, it is thought that assembly of this particle occurs exclusively in the nuclear compartment [35]. snRNP modification, nuclear transport and splicing crucially depend on the assembly of the Sm core domain [51]. Hence, the formation of this common RNP domain constitutes the key event in the cytoplasmic maturation phase of all snRNPs. Studies using purified Sm proteins and RNA revealed some important principles of in vitro Sm core assembly (Figure 4A) [52]. First, Sm proteins do not exist as individual polypeptides; instead they are organized into three preformed hetero-oligomers: B/B0 –D3, D1–D2 and E–F–G. Second, none of the Sm protein hetero-oligomers is individually capable of stable association with U snRNA. Third, the cooperative joining of D1–D2 and E–F–G with snRNA generates the earliest RNP assembly intermediate, the Sm subcore. Fourth, the addition of the B/B0 –D3 heterodimer leads to the conversion of the Sm subcore into the mature Sm core structure. Finally, the structural information required for Sm core formation is present in both the Sm protein and U snRNA counterparts of this domain, rendering the core capable of self-assembly. Although these spontaneous reactions can certainly occur in dilute, aqueous solutions, ample evidence now suggests that in vivo Sm core assembly requires a large number of assisting factors and, hence, does not follow a ‘self-assembly route’ [53]. Assisting factors mediate snRNP assembly in vivo Insights into the in vivo mechanism of snRNP assembly came from an unexpected direction. Mapping experiments identified a gene on human chromosome 5 that was systematically altered in the devastating neurological disorder spinal muscular atrophy (SMA) [54]. Because all SMA patients suffer from the degeneration of motor neurons in the spinal cord, the gene was named survival motor neuron 1 (SMN1). This gene was subsequently shown to encode a ubiquitously expressed protein, which, together with eight other factors (the gem-associated proteins (GEMINs) 2–8 and UNRIP), forms the macromolecular SMN complex 679
(Figure_3)TD$IG][ Review
Trends in Biochemical Sciences Vol.35 No.12
Figure 3. The biogenesis pathway of spliceosomal snRNPs. The newly synthesized snRNAs are transiently exported to the cytoplasm (snRNA Export). In the cytoplasm the seven Sm proteins assemble onto the Sm site to form the Sm core domain (snRNP assembly). Formation of the Sm core domain triggers the conversion of the m7G cap moiety to the m3G trimethylguanosine cap structure common to spliceosomal snRNPs (snRNP hypermethylation). Together, the m3G cap and Sm core domain constitute a bipartite NLS, which retranslocates the assembled snRNP into the nucleus (snRNP Import). The first station of the newly assembled snRNP in the nucleus is the Cajal body, where the RNA is further modified. Eventually, the mature snRNP is incorporated into spliceosomes in the nucleoplasm. At which stage the specific snRNP proteins join the assembled snRNP particle is, in many cases, unknown.
[55,56]. This unit is essential for the formation of the Sm core domain of all splicing RNPs in cells: it provides a binding platform for Sm proteins and facilitates their targeted delivery onto snRNA [53,57,58]. The SMN complex cooperates in snRNP assembly with three trans-acting factors termed WD45 (also called MEP50 and WDR77), pICln and the protein arginine methyltransferase 5 (PRMT5) [57,59,60]. These trans-acting factors unite in the PRMT5 complex and sequester Sm proteins in the early phase of assembly (Figure 4Bi). PRMT5 (and possibly other methyltransferases) then methylates a subset of bound Sm proteins (B/B0 , D1 and D3) on designated arginine residues [57,59–61], thereby leading to the formation of two distinct higher-order Sm protein subcomplexes [62]. The first is ring-shaped and composed of pICln bound to the Sm proteins D1–D2 and E– F–G (Figure 4Biia) [62]. This unit dissociates from the PRMT5 complex after its formation, and is characterized by a sedimentation coefficient of 6S. It is most probably identical to the assembly precursor of snRNPs that was described more than 25 years ago, but whose biochemical composition remained elusive [63]. A second complex forms between pICln and B–D3, i.e. the other two Sm proteins 680
required for snRNP assembly (Figure 4Biib). However, in contrast to the 6S complex, pICln–B–D3 appears to stay stably associated with the PRMT5 complex [62]. The association of pICln with distinct sets of Sm proteins therefore dominates the early phase of assembly. Curiously, rather than promoting assembly, this interaction efficiently prevents Sm protein binding onto snRNA [60,62,64]. The basis for this inactivation became clear when the 6S complex was analyzed by electron microscopy: the Sm proteins form a toroidal structure, where pICln occupies the position of the Sm B–D3 heterodimer in the mature Sm core. Hence, the inhibition of RNP formation is the result of a steric occlusion of the RNA to gain access to its binding site on the inner face of the Sm protein ring [15,65]. Thus, a kinetic trap is imposed on Sm proteins by bound pICln. Owing to its capacity to induce the formation of higher-order Sm protein structures and prevent their premature association with RNA, pICln is an assembly chaperone that specifically directs snRNP formation in vivo. How can the pre-assembled Sm proteins be released from the pICln-induced kinetic trap? The biochemical dissection of the assembly pathway in vivo suggested that the
(Figure_4)TD$IG][ Review
Trends in Biochemical Sciences
Vol.35 No.12
Figure 4. Formation of the Sm core domain follows an ordered pathway. (a) Assembly of the Sm core domain in the absence of assisting factors. Sm protein hetero-oligomers are neither capable of binding to RNA, nor able to associate with each other in the absence of RNA. Sm core domain formation is induced by the initial, cooperative binding of D1–D2 and E–F–G to snRNA, giving rise to the snRNP subcore domain. This intermediate is then matured into the snRNP core domain upon the addition of D3–B. (b) Mechanism of the assisted assembly of SnRNPs. The chaperone pICln initially binds and recruits D1–D2 and D3–B to the PRMT5 complex (i). E–F–G is then recruited to pICln–D1–D2 to form the 6S complex (iia). The pICln–D3–B complex might transiently dissociate from the PRMT5–WD45 heterodimer (iib). Transfer of pICln-bound Sm proteins onto the SMN complex coincides with displacement of pICln (iiia and b) and forms the loaded SMN complex. The SMN complex then allows RNA binding (iv), ring closure and RNP release (v).
SMN complex is the crucial factor for this reaction. In support of this notion, the SMN complex contacts the 6S complex on its outer surface [62]. Notably, the prior methylation of Sm proteins stimulates this reaction [66,67]. Binding results in the dissociation of pICln and opening of the 6S ring (Figure 4Biiia). In a similar reaction, the B– D3 heterodimer is delivered to the SMN complex (Figure 4Biiib). As a consequence, the snRNA is capable of gaining access to the inner site of the open Sm ring, which is bound by the SMN complex (Figure 4Biv). The SMN complex then mediates ring closure around the snRNA and Sm core particle release (Figure 4Bv). The fact that Sm proteins are reactivated by the SMN complex suggests that the latter lowers the activation energy bar-
rier for snRNP assembly and hence acts as a catalyst for Sm core formation from 6S complexes. snRNP assembly factors: what are they good for? Why is the highly ordered interplay of PRMT5- and SMNcontaining complexes necessary for the in vivo assembly of an RNP that can form spontaneously in vitro? If we assume that similar principles apply for snRNP formation as for the assembly of protein complexes, one can hypothesize that RNP assembly factors primarily fulfill two functions: first, they increase assembly efficiency by assisting the formation of higher order building blocks, and second, they prevent aggregation of Sm proteins and/or assembly of snRNP cores which contain ‘incorrect’ RNA (i.e. RNAs that do not contain 681
Review an Sm site). Although studies on the kinetics of in vivo assembly are needed, several reports have addressed the proposed ‘proofreading’ function of the assembly machinery [58,68,69]. These studies clearly demonstrate that Sm proteins can only bind specifically to target snRNAs (under physiological conditions), when they are pre-bound to the SMN complex. By contrast, Sm proteins on their own exhibit no specificity for a particular RNA species under the same conditions. Thus, it is clear that the SMN complex functions as an RNP chaperone that prevents binding of Sm proteins onto non-cognate RNA. However, it remains unknown how this is achieved at the molecular level. Several reports showed that the SMN complex subunit GEMIN5 can specifically identify the Sm-site of snRNA, and hence ensure faithful assembly of the Sm core domain [70–72]. By contrast, the Drosophila melanogaster and Trypansome SMN complexes, which lack Gemin5, mediate faithful assembly, thus suggesting that other mechanisms might also exist [68,69]. One alternative model is that the catalytic activity of the SMN complex confers assembly specificity [62]. Assuming that a high activation energy barrier exists between free Sm proteins and snRNA on one side, and the assembled snRNP on the other, the SMN complex might favor formation of RNPs that are very stable, such as Sm core domains. In this scenario, the specificity for cognate RNP assembly would arise from a kinetic proofreading activity of the SMN complex. Concluding remarks and future perspectives Recent findings have provided detailed insight into the cellular mechanisms that ensure the efficient and specific assembly of various classes of macromolecular complexes. As we have outlined here, similar strategies are employed by such structurally diverse molecular machines as Rubisco and snRNPs. An emerging concept is that assembly chaperones, i.e. proteins that induce the formation of higher order building blocks, act in an early phase of assembly (pICln and RbcX2 in RNP and Rubisco assembly, respectively). In fact, the past few years revealed a large number of proteins with this activity for the assembly of complexes as diverse as ribosomes and proteasomes [9–13]. To exert their functions, these proteins interact with their respective target in a stable manner and with high avidity, employing several low-affinity interaction surfaces. Kinetic traps often result as a consequence, thus pointing to the need for additional factors that can drive assembly to completion. In general, these factors obstruct the association of the chaperone with the building blocks and instead promote formation of the final complex. To release the building blocks from the kinetic traps, many of these late factors act catalytically (e.g. the SMN complex) and consume energy to drive the required conformational changes (either thermal or derived from the hydrolysis of ATP). An additional function of these late assembly factors is the survey and control of proper complex formation and hence a quality control. Assuming that the assembled molecular machine is the thermodynamically most favored state of the individual subunits, proofreading of correct macromolecular complex assembly is anticipated to be kinetic. Despite being assembled by very similar principles in vivo, it is remarkable that each molecular machine employs 682
Trends in Biochemical Sciences Vol.35 No.12
a unique set of assembly chaperones. For example, there is so far no evidence that pICln has any role in the formation of complexes other than the Sm-class of snRNPs. The same most probably holds true for RbcX or assembly chaperones acting in the formation of other complexes. This might reflect the fact that unlike folding chaperones, assembly chaperones are tailored to induce higher-order structures only within distinct sets of proteins. One might therefore predict that the biochemical dissection of the assembly pathways of molecular machines, such as the ribosome or the spliceosome, will uncover factors and mechanisms similar to those exemplified here for snRNPs and Rubisco. Acknowledgments We thank David Goodsell, Manajit Hayer-Hartl and Franz Ulrich Hartl for kindly providing Figures 1B and 2, respectively. We apologize to our colleagues whose work could not be cited owing to size constraints. Work in the authors laboratory is supported by a grant of the German Research Foundation (DFG) to U.F.
References 1 Hartwell, L.H. et al. (1999) From molecular to modular cell biology. Nature 402, C47–C52 2 Alberts, B. (1998) The cell as a collection of protein machines: preparing the next generation of molecular biologists. Cell 92, 291–294 3 Kornberg, R.D. (1977) Structure of chromatin. Annu. Rev. Biochem. 46, 931–954 4 Nierhaus, K.H. and Dohme, F. (1974) Total reconstitution of functionally active 50S ribosomal subunits from Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 71, 4713–4717 5 Nomura, M. et al. (1968) Hybrid 30S ribosomal particles reconstituted from components of different bacterial origins. Nature 219, 793–799 6 Kushner, D.J. (1969) Self-assembly of biological structures. Bacteriol. Rev. 33, 302–345 7 Frydman, J. (2001) Folding of newly translated proteins in vivo: the role of molecular chaperones. Annu. Rev. Biochem. 70, 603–647 8 Ellis, R.J. (2006) Molecular chaperones: assisting assembly in addition to folding. Trends Biochem. Sci. 31, 395–401 9 Fromont-Racine, M. et al. (2003) Ribosome assembly in eukaryotes. Gene 313, 17–42 10 Zemp, I. and Kutay, U. (2007) Nuclear export and cytoplasmic maturation of ribosomal subunits. FEBS Lett. 581, 2783–2793 11 Gallastegui, N. and Groll, M. (2010) The 26S proteasome: assembly and function of a destructive machine. Trends Biochem. Sci. DOI: 10.1016/ j.tibs.2010.05.005 12 Park, S. et al. (2010) Assembly manual for the proteasome regulatory particle: the first draft. Biochem. Soc. Trans. 38, 6–13 13 Besche, H.C. et al. (2009) Getting to first base in proteasome assembly. Cell 138, 25–28 14 Berg, O.G. and von Hippel, P.H. (1985) Diffusion-controlled macromolecular interactions. Annu. Rev. Biophys. Biophys. Chem. 14, 131–160 15 Pomeranz Krummel, D.A. et al. (2009) Crystal structure of human spliceosomal U1 snRNP at 5.5 A resolution. Nature 458, 475–480 16 Luger, K. et al. (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389, 251–260 17 Nooren, I.M. and Thornton, J.M. (2003) Diversity of protein-protein interactions. EMBO J. 22, 3486–3492 18 Ellis, R.J. and Minton, A.P. (2006) Protein aggregation in crowded environments. Biol. Chem. 387, 485–497 19 Ellis, R.J. (2001) Macromolecular crowding: obvious but underappreciated. Trends Biochem. Sci. 26, 597–604 20 Zimmerman, S.B. and Minton, A.P. (1993) Macromolecular crowding: biochemical, biophysical, and physiological consequences. Annu. Rev. Biophys. Biomol. Struct. 22, 27–65 21 Zhou, H.X. (2004) Protein folding and binding in confined spaces and in crowded solutions. J. Mol. Recognit. 17, 368–375 22 Wenner, J.R. and Bloomfield, V.A. (1999) Crowding effects on EcoRV kinetics and binding. Biophys. J. 77, 3234–3241 23 Gutteridge, S. and Gatenby, A.A. (1995) Rubisco synthesis, assembly, mechanism, and regulation. Plant Cell 7, 809–819
Review 24 Andersson, I. (2008) Catalysis and regulation in Rubisco. J. Exp. Bot. 59, 1555–1568 25 Watson, G.M. and Tabita, F.R. (1997) Microbial ribulose 1,5bisphosphate carboxylase/oxygenase: a molecule for phylogenetic and enzymological investigation. FEMS Microbiol. Lett. 146, 13–22 26 Andersson, I. and Taylor, T.C. (2003) Structural framework for catalysis and regulation in ribulose-1,5-bisphosphate carboxylase/ oxygenase. Arch. Biochem. Biophys. 414, 130–140 27 Dean, C. et al. (1989) Structure, evolution, and regulation of RbcS genes in higher plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 40, 415–439 28 Gatenby, A.A. and Ellis, R.J. (1990) Chaperone function: the assembly of ribulose bisphosphate carboxylase-oxygenase. Annu. Rev. Cell Biol. 6, 125–149 29 Gruissem, W. (1989) Chloroplast gene expression: how plants turn their plastids on. Cell 56, 161–170 30 Hemmingsen, S.M. et al. (1988) Homologous plant and bacterial proteins chaperone oligomeric protein assembly. Nature 333, 330–334 31 Saschenbrecker, S. et al. (2007) Structure and function of RbcX, an assembly chaperone for hexadecameric Rubisco. Cell 129, 1189–1200 32 Roy, H. (1989) Rubisco assembly: a model system for studying the mechanism of chaperonin action. Plant Cell 1, 1035–1042 33 Liu, C. et al. (2010) Coupled chaperone action in folding and assembly of hexadecameric Rubisco. Nature 463, 197–202 34 Moore, M.J. (2005) From birth to death: the complex lives of eukaryotic mRNAs. Science 309, 1514–1518 35 Will, C.L. and Luhrmann, R. (2001) Spliceosomal UsnRNP biogenesis, structure and function. Curr. Opin. Cell Biol. 13, 290–301 36 Wahl, M.C. et al. (2009) The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 37 Tarn, W.Y. and Steitz, J.A. (1997) Pre-mRNA splicing: the discovery of a new spliceosome doubles the challenge. Trends Biochem. Sci. 22, 132– 137 38 Lerner, M.R. and Steitz, J.A. (1979) Antibodies to small nuclear RNAs complexed with proteins are produced by patients with systemic lupus erythematosus. Proc. Natl. Acad. Sci. U. S. A. 76, 5495–5499 39 Achsel, T. et al. (1999) A doughnut-shaped heteromer of human Sm-like proteins binds to the 30 - end of U6 snRNA, thereby facilitating U4/U6 duplex formation in vitro. EMBO J. 18, 5789–5802 40 Hamm, J. and Mattaj, I.W. (1990) Monomethylated cap structures facilitate RNA export from the nucleus. Cell 63, 109–118 41 Ohno, M. et al. (2000) PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell 101, 187–198 42 Zeller, R. et al. (1983) Nucleocytoplasmic distribution of snRNPs and stockpiled snRNA-binding proteins during oogenesis and early development in Xenopus laevis. Cell 32, 425–434 43 Mattaj, I.W. (1986) Cap trimethylation of U snRNA is cytoplasmic and dependent on U snRNP protein binding. Cell 46, 905–911 44 Plessel, G. et al. (1994) m3G cap hypermethylation of U1 small nuclear ribonucleoprotein (snRNP) in vitro: evidence that the U1 small nuclear RNA-(guanosine-N2)- methyltransferase is a non-snRNP cytoplasmic protein that requires a binding site on the Sm core domain. Mol. Cell Biol. 14, 4160–4172 45 Fischer, U. et al. (1994) Nuclear transport of U1 snRNP in somatic cells: differences in signal requirement compared with Xenopus laevis oocytes. J. Cell Biol. 125, 971–980 46 Fischer, U. and Luhrmann, R. (1990) An essential signaling role for the m3G cap in the transport of U1 snRNP to the nucleus. Science 249, 786– 790 47 Hamm, J. et al. (1990) The trimethylguanosine cap structure of U1 snRNA is a component of a bipartite nuclear targeting signal. Cell 62, 569–577 48 Stanek, D. and Neugebauer, K.M. (2004) Detection of snRNP assembly intermediates in Cajal bodies by fluorescence resonance energy transfer. J. Cell Biol. 166, 1015–1025 49 Stanek, D. et al. (2008) Spliceosomal small nuclear ribonucleoprotein particles repeatedly cycle through Cajal bodies. Mol. Biol. Cell 19, 2534–2543 50 Kiss, T. (2004) Biogenesis of small nuclear RNPs. J. Cell Sci. 117, 5949– 5951 51 Zhang, D. et al. (2001) A biochemical function for the Sm complex. Mol. Cell 7, 319–329
Trends in Biochemical Sciences
Vol.35 No.12
52 Raker, V.A. et al. (1996) The snRNP core assembly pathway: identification of stable core protein heteromeric complexes and an snRNP subcore particle in vitro. EMBO J. 15, 2256–2269 53 Meister, G. et al. (2001) A multiprotein complex mediates the ATPdependent assembly of spliceosomal U snRNPs. Nat. Cell Biol. 3, 945– 949 54 Lefebvre, S. et al. (1995) Identification and characterization of a spinal muscular atrophy-determining gene. Cell 80, 155–165 55 Neuenkirchen, N. et al. (2008) Deciphering the assembly pathway of Sm-class U snRNPs. FEBS Lett. 582, 1997–2003 56 Gubitz, A.K. et al. (2004) The SMN complex. Exp. Cell Res. 296, 51–56 57 Meister, G. and Fischer, U. (2002) Assisted RNP assembly: SMN and PRMT5 complexes cooperate in the formation of spliceosomal UsnRNPs. EMBO J. 21, 5853–5863 58 Pellizzoni, L. et al. (2002) Essential role for the SMN complex in the specificity of snRNP assembly. Science 298, 1775–1779 59 Friesen, W.J. et al. (2001) The methylosome, a 20S complex containing JBP1 and pICln, produces dimethylarginine-modified Sm proteins. Mol. Cell. Biol. 21, 8289–8300 60 Meister, G. et al. (2001) Methylation of Sm proteins by a complex containing PRMT5 and the putative U snRNP assembly factor pICln. Curr. Biol. 11, 1990–1994 61 Gonsalvez, G.B. et al. (2007) Two distinct arginine methyltransferases are required for biogenesis of Sm-class ribonucleoproteins. J. Cell Biol. 178, 733–740 62 Chari, A. et al. (2008) An assembly chaperone collaborates with the SMN complex to generate spliceosomal SnRNPs. Cell 135, 497–509 63 Fisher, D.E. et al. (1985) Small nuclear ribonucleoprotein particle assembly in vivo: demonstration of a 6S RNA-free core precursor and posttranslational modification. Cell 42, 751–758 64 Pu, W.T. et al. (1999) pICln inhibits snRNP biogenesis by binding core spliceosomal proteins. Mol. Cell. Biol. 19, 4113–4120 65 Urlaub, H. et al. (2001) Sm protein-Sm site RNA interactions within the inner ring of the spliceosomal snRNP core structure. EMBO J. 20, 187–196 66 Brahms, H. et al. (2001) Symmetrical dimethylation of arginine residues in spliceosomal Sm protein B/B0 and the Sm-like protein LSm4, and their interaction with the SMN protein. RNA 7, 1531–1542 67 Friesen, W.J. et al. (2001) SMN, the product of the spinal muscular atrophy gene, binds preferentially to dimethylarginine-containing protein targets. Mol. Cell 7, 1111–1117 68 Kroiss, M. et al. (2008) Evolution of an RNP assembly system: a minimal SMN complex facilitates formation of UsnRNPs in Drosophila melanogaster. Proc. Natl. Acad. Sci. U. S. A. 105, 10045–10050 69 Palfi, Z. et al. (2009) SMN-assisted assembly of snRNP-specific Sm cores in trypanosomes. Genes Dev. 23, 1650–1664 70 Battle, D.J. et al. (2006) The Gemin5 protein of the SMN complex identifies snRNAs. Mol. Cell 23, 273–279 71 Yong, J. et al. (2010) Gemin5 delivers snRNA precursors to the SMN complex for snRNP biogenesis. Mol. Cell 38, 551–562 72 Lau, C.K. et al. (2009) Gemin5-snRNA interaction reveals an RNA binding function for WD repeat domains. Nat. Struct. Mol. Biol. 16, 486–491 73 Facey, S.J. and Kuhn, A. (2010) Biogenesis of bacterial innermembrane proteins. Cell. Mol. Life Sci. 67, 2343–2362 74 Hagan, C.L. et al. (2010) Reconstitution of outer membrane protein assembly from purified components. Science 328, 890–892 75 Nishiyama, M. et al. (2008) Reconstitution of pilus assembly reveals a bacterial outer membrane catalyst. Science 320, 376–379 76 Vetsch, M. et al. (2006) Mechanism of fibre assembly through the chaperone-usher pathway. EMBO Rep. 7, 734–738 77 Waksman, G. and Hultgren, S.J. (2009) Structural biology of the chaperone-usher pathway of pilus biogenesis. Nat. Rev. Microbiol. 7, 765–774 78 Becker, T. et al. (2009) Biogenesis of mitochondrial membrane proteins. Curr. Opin. Cell Biol. 21, 484–493 79 Park, Y.J. and Luger, K. (2008) Histone chaperones in nucleosome eviction and histone exchange. Curr. Opin. Struct. Biol. 18, 282–289 80 Rocha, W. and Verreault, A. (2008) Clothing up DNA for all seasons: histone chaperones and nucleosome assembly pathways. FEBS Lett. 582, 1938–1949 81 Segal, E. and Widom, J. (2009) What controls nucleosome positions? Trends Genet. 25, 335–343
683
Cellular strategies for the assembly of molecular machines Ashwin Chari and Utz Fischer Department of Biochemistry, Theodor-Boveri Institute, University of Wurzburg, Am Hubland, D-97074 Wurzburg, Germany
Molecular machines are supramolecular assemblies of biomolecules (proteins, RNA and/or DNA) that facilitate a diversity of biological tasks in the cells of all organisms. How these complex structures are built within the crowded cellular environment is, therefore, a central question in the biological sciences. Recent studies on spliceosomal uridine-rich small nuclear ribonucleoproteins (snRNPs) have unveiled cellular assembly strategies for RNA–protein complexes. snRNPs form in vivo by the coordinated action of an elaborate assembly line consisting of assembly chaperones, scaffolding proteins and catalysts. These newly discovered strategies exhibit similarities to those employed by protein complexes such as ribulose-1,5-bisphosphate-carboxylase (Rubisco) and allow the elucidation of general rules for how molecular machines are formed in vivo. Assembly lines in our daily life and at the molecular level Life in our technologically advanced society is greatly simplified by a multitude of gadgets which help us perform our daily tasks. To assemble these machines, pre-manufactured components and an industrial manufacturing design (blueprint) are required. This blueprint essentially specifies where each individual part of the machine should be located in three dimensions and at which temporal stage in assembly the parts should be incorporated into the final product. Then, factory assembly lines add all the components together in a sequential manner, as specified in the blueprint. Macromolecular complexes perform many biochemical reactions crucial for the survival of organisms and are, thus, rightfully also referred to as biological machines [1,2]. Although we understand the underlying mechanisms for many of the biochemical reactions catalyzed by these machines, knowledge about how these machines are assembled in vivo is, in several instances, less complete. An emerging view is that the formation of macromolecular complexes utilizes principles similar to those used in an industrial assembly line. Biological complexes are formed when proteins either bind to each other or to nucleic acids. Prominent examples include the plant enzyme ribulose-1,5-bisphosphatecarboxylase (Rubisco, protein only), the nucleosome (DNA and protein) and the spliceosome (RNA and protein) (Table 1). How are these entities built within cells? Early studies demonstrated that it is possible to reconstitute nucleosomes, or even the 2.6 MDa ribosome, from purified constituents into active assemblies in vitro [3–6]. The structural Corresponding authors: Chari, A. ([email protected]); Fischer, U. ([email protected])
676
information (blueprint) for the formation of macromolecular complexes is therefore encoded within its subunits. Coupled with observations that many molecular machines can be assembled in heterologous expression systems, these findings have given rise to the notion that the machines of life are built spontaneously without the requirement for accessory factors. Despite this general tendency for self-assembly of macromolecular complexes in vitro, it is becoming increasingly clear that assisting factors are required for efficient assembly in vivo. Many of these accessory factors belong to an unrelated group of proteins termed assembly chaperones which act as a cellular counterpart to industrial assembly lines [7,8]. They associate with the individual subunits of a macromolecular complex and promote the formation of the final structure (Figure 1A). In this review, we initially discuss pitfalls encountered during productive macromolecular complex assembly that are introduced by certain properties of the cellular milieu. We then describe strategies employed by cells to overcome these impediments. Rubisco and spliceosomal small nuclear ribonucleoproteins (snRNPs) were chosen for this discussion because both represent well-studied particles exemplifying two distinct classes, namely protein–protein and RNA–protein complexes. Finally, we will delineate common principles that underlie both assembly processes, lending support to the view that the strategies employed for the assembly of these complexes also apply to the assembly of other macromolecular complexes such as proteasomes and ribosomes (Table 1) [9–13]. Principles governing macromolecular complex assembly in vivo From a biophysical point of view, the assembly of macromolecular complexes depends on the diffusion-driven, random collision of subunits [14]. The stability of the resulting assembly is determined by the ratio of the individual dissociation and association rate constants. However, within cells variations to this simplified view appear to be prevalent. First, many subunits of an assembly often complete their individual folding reaction only upon association with their interaction partner(s) [15,16]. Second, the interaction between two or more subunits often involves hydrophobic stretches that are exposed in the individual folded subunit [17]. This property renders isolated proteins of a macromolecular complex particularly prone to irreversible aggregation within cells [18]. Third, an additional pitfall towards the formation of macromolecular entities arises from the fact that, in general, the cellular milieu does not behave as an ideal fluid. Instead, the interior of cells is characterized by a
0968-0004/$ – see front matter ß 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.tibs.2010.07.006 Trends in Biochemical Sciences, December 2010, Vol. 35, No. 12
Review
Trends in Biochemical Sciences
Table 1. Selected list of complexes known to be assembled by assisting factors Macromolecular complex assembly process Bacterial inner membrane proteins Bacterial outer membrane proteins Bacterial pilus Mitochondrial membrane proteins Nucleosomes Eukaryotic ribosomes 20S and 26S proteasomes Form I Rubisco Spliceosomal snRNPs
Refs [73] [74] [75–77] [78] [8,79–81] [9,10] [8,11–13] [28,30,31,33] [55,56,62,71]
high total concentration of many different macromolecules. As a consequence, no single species accumulates in high amounts, but in total, proteins and other macromolecules are so abundant that they occupy 20–30% of the interior of cells. Therefore, a large percentage of the cellular volume is unavailable to other molecules leading to an excluded volume effect that is commonly referred to as macromolecular crowding (Figure 1B) [19,20]. This property of cells lowers the diffusion rate of individual subunits within a cell [21]. Thus, the likelihood of random encounters between the building blocks of a macromolecular entity is expected to be strongly reduced. Simultaneously, specific, as well as nonspecific, association reactions are strongly favored under crowding conditions [20]. Therefore, the propensity for productive interactions between cognate molecules to occur within cells is substantially lowered, resulting in misassembly [22]. [(Figure_1)TD$IG]
Vol.35 No.12
Strategies utilized by cells to encounter pitfalls towards faithful macromolecular complex assembly Cells have evolved several elaborate strategies to encounter the obstacles that can counteract faithful macromolecular assembly. First, the synthesis of the individual constituents of a macromolecular entity is often temporally tightly controlled. This strategy ensures that all components of a macromolecular complex are simultaneously present in sufficient amounts when assembly is initiated. Second, the synthesis of the individual building blocks of a macromolecular machine, their assembly into functional particles and finally their integration into the designated process, by which they help maintain cellular homeostasis, are in many cases spatially segregated. This strategy is particularly (but not exclusively) dominant in the case of RNA–protein complexes (RNPs) and DNA–protein complexes (DNPs), where the nucleic acid moiety is synthesized in the nucleus and the proteins in the cytoplasm. This compartmentalization of synthesis and assembly does not necessarily require organelles, which are surrounded by membranes. In particular, the variety of nuclear subdomains also effectively achieves spatial segregation; a prominent example is the nucleolus in ribosome assembly [9,10]. Spatial separation of assembly and function is also a crucial means by which the interference of biochemical processes of the fully assembled machine by partially assembled complexes is prevented.
Figure 1. (a) The formation of biological machines. The formation of biological machines requires a blueprint, which is intrinsic to the structure of DNA and protein components (histone H3–H4 dimer, from PDB ID: 1TZY). A chaperone then acts as a counterpart for the industrial assembly line within cells (the chaperone Asf1 bound to the histone H3–H4 dimer, PDB ID: 2HUE). It associates with the protein components and orchestrates the formation of the final product; in this case a nucleosome core particle (PDB ID: 1AOI). (b) The cytoplasm of cells is a crowded environment. The illustration depicts an artist’s impression of macromolecular crowding conditions within a cell. A small portion of eukaryotic cytoplasm showing microtubules (light blue), actin filaments (dark blue), ribosomes (yellow and purple), soluble proteins (light blue), kinesin (red), small molecules (white) and RNA (pink) is shown. Illustration by D.S. Goodsell, the Scripps Research Institute.
677
Review Third, cells often utilize the activity of assembly chaperones. These proteins enable the formation of intermediate higher-order structures, thereby ensuring a spatial pre-organization of individual subunits in positions, which they also occupy in the assembled macromolecular complex. Simultaneously, they prevent the premature occurrence of steps that are meant to occur at a later stage in assembly as well as dead-end reactions. This temporal and quality control often leads to the formation of kinetically trapped assembly intermediates, which require the activity of additional factors for assembly to proceed. Fourth, in addition to chaperones, the assembly process of a molecular machine is often assisted by a heterogeneous group of trans-acting factors. These proteins typically enable the resolution of kinetic traps that were introduced by the assembly chaperones. In addition, they stimulate assembly by increasing the local concentration of the individual subunits (or assembly intermediates) of the complex. This activity can sometimes be achieved by subunits of the macromolecular machine itself. In many instances, the fulfillment of one strategy is the prerequisite for the next step, implying a strong synergistic relation between each individual step in biogenesis. Each of these cellular strategies has evolved to ensure the proper and faithful formation of macromolecular complexes in vivo. Assisted assembly of a protein complex: lessons from the Rubisco holoenzyme Rubisco is arguably one of the most important enzymes for life because it catalyzes the primary reaction by which [(Figure_2)TD$IG]
Trends in Biochemical Sciences Vol.35 No.12
inorganic carbon is chemically fixed in the biosphere. This is achieved in the Calvin cycle, where the fixation of CO2 results in the synthesis of usable sugars [23]. Rubisco also catalyses an opposing reaction, termed photorespiration, in which O2 is the substrate. Photorespiration competes with, and determines the efficiency of, carbon fixation [24]. Three forms of quaternary structure for otherwise closely related Rubisco subunits are known [25]. Form I, which is found in plants and cyanobacteria, is a hexadecamer RbcL8S8, containing eight large subunits (RbcL) and eight small subunits (RbcS). Forms II and III Rubisco, by contrast, are much simpler in terms of their biochemical composition and consist of only one (Form II) or five (Form III) RbcL dimers [26]. We will restrict our discussion to the assembly of the form I Rubisco holoenzyme because it has been studied in the greatest detail (Figure 2). The different steps in Rubisco assembly illustrate the four major strategies evolved by cells to encounter the obstacles that can counteract faithful macromolecular assembly. Whereas the eight large subunits (RbcL) of the form I Rubisco holoenzyme are encoded in the chloroplast genome, the eight small subunits (RbcS) are expressed from the nuclear genome. Upon stimulation with light, the syntheses of both chloroplast and nuclear mRNAs are coordinated [27–29], thereby ensuring that both subunits are present simultaneously in sufficient amounts when assembly is initiated. The cytoplasmic synthesis of RbcS makes a subcellular transport event to the chloroplast, the organelle where RbcL is expressed, inevitable. The formation of the form I Rubisco holoenzyme is
Figure 2. Model for the chaperone assisted folding and assembly of Rubisco. Newly synthesized RbcL (green) is sequestered and folded by the chaperonins GroEL (light blue) and GroES (dark blue) (i). The assembly chaperone RbcX2 (yellow) binds to a disordered C-terminal peptide of RbcL (ii). This interaction induces partial folding of the RbcL C-terminus and the formation of RbcL dimers (iii). Stable dimers assemble into RbcL8–(RbcX2)8 complexes (iv). RbcS (gray) binding dissociates RbcX2, which might occur concertedly (v). The catalytically mature Rubisco holoenzyme is then released (vi). Figure modified, with permission, from [33].
678
Review dependent on the molecular chaperones GroEL, GroES and RbcX [30,31]. In the first phase, the chaperonin GroEL, together with its co-chaperonin GroES, ensures proper folding of the RbcL subunit and thus represents a quality control checkpoint for Rubisco formation (Figure 2i) [32]. During the folding reaction, RbcL monomers are transiently released from GroEL [33]. Upon release, the partially disordered C-terminal region of RbcL is recognized by the dimeric assembly chaperone RbcX2 (Figure 2ii). RbcX2 then induces the formation of stable RbcL dimers, which imparts partial folding in the C-terminus of RbcL (Figure 2iii). As a result, the RbcX2-bound RbcL dimers assemble into tetramers forming the RbcL8–(RbcX2)8 complex (Figure 2iv). RbcL8–(RbcX2)8 can be considered a catalytically immature roadblock in Rubisco holoenzyme assembly. Finally, this roadblock is salvaged by the Rubisco small subunit, RbcS (Figure 2v). Its association with RbcL8–(RbcX2)8 triggers a conformational change in RbcL, thereby eliciting RbcX2 dissociation and holoenzyme formation (Figure 2vi). This final assembly step is coincident with the attainment of catalytic maturity of Rubisco. Spliceosomal snRNPs have a segmented biogenesis pathway Although the major principles of protein complex assembly have been established, much less is known about the formation of molecular complexes composed of RNA and proteins (RNPs) [34]. The spliceosomal snRNPs provide an ideal model for the analysis of RNP assembly strategies because their in vivo maturation pathway can be faithfully recapitulated in vitro and dissected biochemically [35]. snRNPs are abundant nuclear complexes and the major components of the nuclear splicing machinery [36]. This entity catalyzes the removal of introns from mRNA precursors (pre-mRNAs) and the precise joining of exons to yield a functional mRNA. Four different snRNPs are part of the major spliceosome, named after their RNA moiety: U1, U2, U4/6 or U5. A second, so-called minor spliceosome, also exists, in which the snRNPs U1, U2, U4/U6 are replaced by U11, U12 and U4atac/U6atac, respectively. The minor spliceosome catalyzes the splicing of a small subset of introns [37]. It is assumed, but not yet experimentally proven, that the principles discussed here for the major snRNPs are identical for the minor snRNPs. Each individual particle contains a set of snRNP-specific proteins (for example U1-70K, U1A & U1C in the case of the U1 snRNP) and assumes a designated role in splicing (Figure 3) [35,36]. In addition, the U1, U2, U4 and U5 snRNPs also contain a set of seven common proteins termed B/B0 , D1, D2, D3, E, F and G; they are often also referred to as the Smith (Sm) antigen because patients suffering from rheumatoid diseases (including patient Smith) frequently develop antibodies against these factors [38]. The seven Sm proteins bind and encircle a singlestranded region (Sm site) of the snRNAs and form a toroidal RNP core which constitutes the structural framework of all snRNPs. Only the U6 snRNP is divergent in this respect. Instead of the seven canonical Sm proteins, it contains a set of seven evolutionarily related proteins ‘like Sm proteins 2–8 (LSm2–8)’ [39].
Trends in Biochemical Sciences
Vol.35 No.12
snRNP biogenesis is an intricate, segmented pathway (Figure 3) [35]. It begins with the RNA polymerase II-dependent transcription of a monomethyl-guanosine (m7G) capped snRNA-precursor, followed by its subsequent nuclear export [40,41]. Within the cytoplasm, the snRNA encounters Sm proteins, which are synthesized and stockpiled in this compartment [42]. Sm proteins then bind to the Sm site and form the Sm core domain. This event, in turn, triggers the hypermethylation of the m7G-cap, thereby generating the trimethylguanosine (TMG) m2,2,73G-cap [43,44]. A bipartite nuclear localization signal (NLS) consisting of the assembled Sm core domain and the TMG cap ensures the translocation of snRNPs to the nucleus [45–47]. Before they reach their site of function in the nucleoplasm, snRNPs are transiently directed to subnuclear domains termed Cajal bodies, where further maturation steps, such as modification of the RNA moiety, complete the biogenesis cycle [48–50]. The biogenesis pathway ends when the snRNP particles leave Cajal bodies and become part of the splicing machinery within the nucleoplasm. It is still largely unknown at which stages in the biogenesis cycle specific proteins join the assembled snRNP. The U6 snRNP does not follow the biogenesis pathway outlined above. Instead, it is thought that assembly of this particle occurs exclusively in the nuclear compartment [35]. snRNP modification, nuclear transport and splicing crucially depend on the assembly of the Sm core domain [51]. Hence, the formation of this common RNP domain constitutes the key event in the cytoplasmic maturation phase of all snRNPs. Studies using purified Sm proteins and RNA revealed some important principles of in vitro Sm core assembly (Figure 4A) [52]. First, Sm proteins do not exist as individual polypeptides; instead they are organized into three preformed hetero-oligomers: B/B0 –D3, D1–D2 and E–F–G. Second, none of the Sm protein hetero-oligomers is individually capable of stable association with U snRNA. Third, the cooperative joining of D1–D2 and E–F–G with snRNA generates the earliest RNP assembly intermediate, the Sm subcore. Fourth, the addition of the B/B0 –D3 heterodimer leads to the conversion of the Sm subcore into the mature Sm core structure. Finally, the structural information required for Sm core formation is present in both the Sm protein and U snRNA counterparts of this domain, rendering the core capable of self-assembly. Although these spontaneous reactions can certainly occur in dilute, aqueous solutions, ample evidence now suggests that in vivo Sm core assembly requires a large number of assisting factors and, hence, does not follow a ‘self-assembly route’ [53]. Assisting factors mediate snRNP assembly in vivo Insights into the in vivo mechanism of snRNP assembly came from an unexpected direction. Mapping experiments identified a gene on human chromosome 5 that was systematically altered in the devastating neurological disorder spinal muscular atrophy (SMA) [54]. Because all SMA patients suffer from the degeneration of motor neurons in the spinal cord, the gene was named survival motor neuron 1 (SMN1). This gene was subsequently shown to encode a ubiquitously expressed protein, which, together with eight other factors (the gem-associated proteins (GEMINs) 2–8 and UNRIP), forms the macromolecular SMN complex 679
(Figure_3)TD$IG][ Review
Trends in Biochemical Sciences Vol.35 No.12
Figure 3. The biogenesis pathway of spliceosomal snRNPs. The newly synthesized snRNAs are transiently exported to the cytoplasm (snRNA Export). In the cytoplasm the seven Sm proteins assemble onto the Sm site to form the Sm core domain (snRNP assembly). Formation of the Sm core domain triggers the conversion of the m7G cap moiety to the m3G trimethylguanosine cap structure common to spliceosomal snRNPs (snRNP hypermethylation). Together, the m3G cap and Sm core domain constitute a bipartite NLS, which retranslocates the assembled snRNP into the nucleus (snRNP Import). The first station of the newly assembled snRNP in the nucleus is the Cajal body, where the RNA is further modified. Eventually, the mature snRNP is incorporated into spliceosomes in the nucleoplasm. At which stage the specific snRNP proteins join the assembled snRNP particle is, in many cases, unknown.
[55,56]. This unit is essential for the formation of the Sm core domain of all splicing RNPs in cells: it provides a binding platform for Sm proteins and facilitates their targeted delivery onto snRNA [53,57,58]. The SMN complex cooperates in snRNP assembly with three trans-acting factors termed WD45 (also called MEP50 and WDR77), pICln and the protein arginine methyltransferase 5 (PRMT5) [57,59,60]. These trans-acting factors unite in the PRMT5 complex and sequester Sm proteins in the early phase of assembly (Figure 4Bi). PRMT5 (and possibly other methyltransferases) then methylates a subset of bound Sm proteins (B/B0 , D1 and D3) on designated arginine residues [57,59–61], thereby leading to the formation of two distinct higher-order Sm protein subcomplexes [62]. The first is ring-shaped and composed of pICln bound to the Sm proteins D1–D2 and E– F–G (Figure 4Biia) [62]. This unit dissociates from the PRMT5 complex after its formation, and is characterized by a sedimentation coefficient of 6S. It is most probably identical to the assembly precursor of snRNPs that was described more than 25 years ago, but whose biochemical composition remained elusive [63]. A second complex forms between pICln and B–D3, i.e. the other two Sm proteins 680
required for snRNP assembly (Figure 4Biib). However, in contrast to the 6S complex, pICln–B–D3 appears to stay stably associated with the PRMT5 complex [62]. The association of pICln with distinct sets of Sm proteins therefore dominates the early phase of assembly. Curiously, rather than promoting assembly, this interaction efficiently prevents Sm protein binding onto snRNA [60,62,64]. The basis for this inactivation became clear when the 6S complex was analyzed by electron microscopy: the Sm proteins form a toroidal structure, where pICln occupies the position of the Sm B–D3 heterodimer in the mature Sm core. Hence, the inhibition of RNP formation is the result of a steric occlusion of the RNA to gain access to its binding site on the inner face of the Sm protein ring [15,65]. Thus, a kinetic trap is imposed on Sm proteins by bound pICln. Owing to its capacity to induce the formation of higher-order Sm protein structures and prevent their premature association with RNA, pICln is an assembly chaperone that specifically directs snRNP formation in vivo. How can the pre-assembled Sm proteins be released from the pICln-induced kinetic trap? The biochemical dissection of the assembly pathway in vivo suggested that the
(Figure_4)TD$IG][ Review
Trends in Biochemical Sciences
Vol.35 No.12
Figure 4. Formation of the Sm core domain follows an ordered pathway. (a) Assembly of the Sm core domain in the absence of assisting factors. Sm protein hetero-oligomers are neither capable of binding to RNA, nor able to associate with each other in the absence of RNA. Sm core domain formation is induced by the initial, cooperative binding of D1–D2 and E–F–G to snRNA, giving rise to the snRNP subcore domain. This intermediate is then matured into the snRNP core domain upon the addition of D3–B. (b) Mechanism of the assisted assembly of SnRNPs. The chaperone pICln initially binds and recruits D1–D2 and D3–B to the PRMT5 complex (i). E–F–G is then recruited to pICln–D1–D2 to form the 6S complex (iia). The pICln–D3–B complex might transiently dissociate from the PRMT5–WD45 heterodimer (iib). Transfer of pICln-bound Sm proteins onto the SMN complex coincides with displacement of pICln (iiia and b) and forms the loaded SMN complex. The SMN complex then allows RNA binding (iv), ring closure and RNP release (v).
SMN complex is the crucial factor for this reaction. In support of this notion, the SMN complex contacts the 6S complex on its outer surface [62]. Notably, the prior methylation of Sm proteins stimulates this reaction [66,67]. Binding results in the dissociation of pICln and opening of the 6S ring (Figure 4Biiia). In a similar reaction, the B– D3 heterodimer is delivered to the SMN complex (Figure 4Biiib). As a consequence, the snRNA is capable of gaining access to the inner site of the open Sm ring, which is bound by the SMN complex (Figure 4Biv). The SMN complex then mediates ring closure around the snRNA and Sm core particle release (Figure 4Bv). The fact that Sm proteins are reactivated by the SMN complex suggests that the latter lowers the activation energy bar-
rier for snRNP assembly and hence acts as a catalyst for Sm core formation from 6S complexes. snRNP assembly factors: what are they good for? Why is the highly ordered interplay of PRMT5- and SMNcontaining complexes necessary for the in vivo assembly of an RNP that can form spontaneously in vitro? If we assume that similar principles apply for snRNP formation as for the assembly of protein complexes, one can hypothesize that RNP assembly factors primarily fulfill two functions: first, they increase assembly efficiency by assisting the formation of higher order building blocks, and second, they prevent aggregation of Sm proteins and/or assembly of snRNP cores which contain ‘incorrect’ RNA (i.e. RNAs that do not contain 681
Review an Sm site). Although studies on the kinetics of in vivo assembly are needed, several reports have addressed the proposed ‘proofreading’ function of the assembly machinery [58,68,69]. These studies clearly demonstrate that Sm proteins can only bind specifically to target snRNAs (under physiological conditions), when they are pre-bound to the SMN complex. By contrast, Sm proteins on their own exhibit no specificity for a particular RNA species under the same conditions. Thus, it is clear that the SMN complex functions as an RNP chaperone that prevents binding of Sm proteins onto non-cognate RNA. However, it remains unknown how this is achieved at the molecular level. Several reports showed that the SMN complex subunit GEMIN5 can specifically identify the Sm-site of snRNA, and hence ensure faithful assembly of the Sm core domain [70–72]. By contrast, the Drosophila melanogaster and Trypansome SMN complexes, which lack Gemin5, mediate faithful assembly, thus suggesting that other mechanisms might also exist [68,69]. One alternative model is that the catalytic activity of the SMN complex confers assembly specificity [62]. Assuming that a high activation energy barrier exists between free Sm proteins and snRNA on one side, and the assembled snRNP on the other, the SMN complex might favor formation of RNPs that are very stable, such as Sm core domains. In this scenario, the specificity for cognate RNP assembly would arise from a kinetic proofreading activity of the SMN complex. Concluding remarks and future perspectives Recent findings have provided detailed insight into the cellular mechanisms that ensure the efficient and specific assembly of various classes of macromolecular complexes. As we have outlined here, similar strategies are employed by such structurally diverse molecular machines as Rubisco and snRNPs. An emerging concept is that assembly chaperones, i.e. proteins that induce the formation of higher order building blocks, act in an early phase of assembly (pICln and RbcX2 in RNP and Rubisco assembly, respectively). In fact, the past few years revealed a large number of proteins with this activity for the assembly of complexes as diverse as ribosomes and proteasomes [9–13]. To exert their functions, these proteins interact with their respective target in a stable manner and with high avidity, employing several low-affinity interaction surfaces. Kinetic traps often result as a consequence, thus pointing to the need for additional factors that can drive assembly to completion. In general, these factors obstruct the association of the chaperone with the building blocks and instead promote formation of the final complex. To release the building blocks from the kinetic traps, many of these late factors act catalytically (e.g. the SMN complex) and consume energy to drive the required conformational changes (either thermal or derived from the hydrolysis of ATP). An additional function of these late assembly factors is the survey and control of proper complex formation and hence a quality control. Assuming that the assembled molecular machine is the thermodynamically most favored state of the individual subunits, proofreading of correct macromolecular complex assembly is anticipated to be kinetic. Despite being assembled by very similar principles in vivo, it is remarkable that each molecular machine employs 682
Trends in Biochemical Sciences Vol.35 No.12
a unique set of assembly chaperones. For example, there is so far no evidence that pICln has any role in the formation of complexes other than the Sm-class of snRNPs. The same most probably holds true for RbcX or assembly chaperones acting in the formation of other complexes. This might reflect the fact that unlike folding chaperones, assembly chaperones are tailored to induce higher-order structures only within distinct sets of proteins. One might therefore predict that the biochemical dissection of the assembly pathways of molecular machines, such as the ribosome or the spliceosome, will uncover factors and mechanisms similar to those exemplified here for snRNPs and Rubisco. Acknowledgments We thank David Goodsell, Manajit Hayer-Hartl and Franz Ulrich Hartl for kindly providing Figures 1B and 2, respectively. We apologize to our colleagues whose work could not be cited owing to size constraints. Work in the authors laboratory is supported by a grant of the German Research Foundation (DFG) to U.F.
References 1 Hartwell, L.H. et al. (1999) From molecular to modular cell biology. Nature 402, C47–C52 2 Alberts, B. (1998) The cell as a collection of protein machines: preparing the next generation of molecular biologists. Cell 92, 291–294 3 Kornberg, R.D. (1977) Structure of chromatin. Annu. Rev. Biochem. 46, 931–954 4 Nierhaus, K.H. and Dohme, F. (1974) Total reconstitution of functionally active 50S ribosomal subunits from Escherichia coli. Proc. Natl. Acad. Sci. U. S. A. 71, 4713–4717 5 Nomura, M. et al. (1968) Hybrid 30S ribosomal particles reconstituted from components of different bacterial origins. Nature 219, 793–799 6 Kushner, D.J. (1969) Self-assembly of biological structures. Bacteriol. Rev. 33, 302–345 7 Frydman, J. (2001) Folding of newly translated proteins in vivo: the role of molecular chaperones. Annu. Rev. Biochem. 70, 603–647 8 Ellis, R.J. (2006) Molecular chaperones: assisting assembly in addition to folding. Trends Biochem. Sci. 31, 395–401 9 Fromont-Racine, M. et al. (2003) Ribosome assembly in eukaryotes. Gene 313, 17–42 10 Zemp, I. and Kutay, U. (2007) Nuclear export and cytoplasmic maturation of ribosomal subunits. FEBS Lett. 581, 2783–2793 11 Gallastegui, N. and Groll, M. (2010) The 26S proteasome: assembly and function of a destructive machine. Trends Biochem. Sci. DOI: 10.1016/ j.tibs.2010.05.005 12 Park, S. et al. (2010) Assembly manual for the proteasome regulatory particle: the first draft. Biochem. Soc. Trans. 38, 6–13 13 Besche, H.C. et al. (2009) Getting to first base in proteasome assembly. Cell 138, 25–28 14 Berg, O.G. and von Hippel, P.H. (1985) Diffusion-controlled macromolecular interactions. Annu. Rev. Biophys. Biophys. Chem. 14, 131–160 15 Pomeranz Krummel, D.A. et al. (2009) Crystal structure of human spliceosomal U1 snRNP at 5.5 A resolution. Nature 458, 475–480 16 Luger, K. et al. (1997) Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389, 251–260 17 Nooren, I.M. and Thornton, J.M. (2003) Diversity of protein-protein interactions. EMBO J. 22, 3486–3492 18 Ellis, R.J. and Minton, A.P. (2006) Protein aggregation in crowded environments. Biol. Chem. 387, 485–497 19 Ellis, R.J. (2001) Macromolecular crowding: obvious but underappreciated. Trends Biochem. Sci. 26, 597–604 20 Zimmerman, S.B. and Minton, A.P. (1993) Macromolecular crowding: biochemical, biophysical, and physiological consequences. Annu. Rev. Biophys. Biomol. Struct. 22, 27–65 21 Zhou, H.X. (2004) Protein folding and binding in confined spaces and in crowded solutions. J. Mol. Recognit. 17, 368–375 22 Wenner, J.R. and Bloomfield, V.A. (1999) Crowding effects on EcoRV kinetics and binding. Biophys. J. 77, 3234–3241 23 Gutteridge, S. and Gatenby, A.A. (1995) Rubisco synthesis, assembly, mechanism, and regulation. Plant Cell 7, 809–819
Review 24 Andersson, I. (2008) Catalysis and regulation in Rubisco. J. Exp. Bot. 59, 1555–1568 25 Watson, G.M. and Tabita, F.R. (1997) Microbial ribulose 1,5bisphosphate carboxylase/oxygenase: a molecule for phylogenetic and enzymological investigation. FEMS Microbiol. Lett. 146, 13–22 26 Andersson, I. and Taylor, T.C. (2003) Structural framework for catalysis and regulation in ribulose-1,5-bisphosphate carboxylase/ oxygenase. Arch. Biochem. Biophys. 414, 130–140 27 Dean, C. et al. (1989) Structure, evolution, and regulation of RbcS genes in higher plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 40, 415–439 28 Gatenby, A.A. and Ellis, R.J. (1990) Chaperone function: the assembly of ribulose bisphosphate carboxylase-oxygenase. Annu. Rev. Cell Biol. 6, 125–149 29 Gruissem, W. (1989) Chloroplast gene expression: how plants turn their plastids on. Cell 56, 161–170 30 Hemmingsen, S.M. et al. (1988) Homologous plant and bacterial proteins chaperone oligomeric protein assembly. Nature 333, 330–334 31 Saschenbrecker, S. et al. (2007) Structure and function of RbcX, an assembly chaperone for hexadecameric Rubisco. Cell 129, 1189–1200 32 Roy, H. (1989) Rubisco assembly: a model system for studying the mechanism of chaperonin action. Plant Cell 1, 1035–1042 33 Liu, C. et al. (2010) Coupled chaperone action in folding and assembly of hexadecameric Rubisco. Nature 463, 197–202 34 Moore, M.J. (2005) From birth to death: the complex lives of eukaryotic mRNAs. Science 309, 1514–1518 35 Will, C.L. and Luhrmann, R. (2001) Spliceosomal UsnRNP biogenesis, structure and function. Curr. Opin. Cell Biol. 13, 290–301 36 Wahl, M.C. et al. (2009) The spliceosome: design principles of a dynamic RNP machine. Cell 136, 701–718 37 Tarn, W.Y. and Steitz, J.A. (1997) Pre-mRNA splicing: the discovery of a new spliceosome doubles the challenge. Trends Biochem. Sci. 22, 132– 137 38 Lerner, M.R. and Steitz, J.A. (1979) Antibodies to small nuclear RNAs complexed with proteins are produced by patients with systemic lupus erythematosus. Proc. Natl. Acad. Sci. U. S. A. 76, 5495–5499 39 Achsel, T. et al. (1999) A doughnut-shaped heteromer of human Sm-like proteins binds to the 30 - end of U6 snRNA, thereby facilitating U4/U6 duplex formation in vitro. EMBO J. 18, 5789–5802 40 Hamm, J. and Mattaj, I.W. (1990) Monomethylated cap structures facilitate RNA export from the nucleus. Cell 63, 109–118 41 Ohno, M. et al. (2000) PHAX, a mediator of U snRNA nuclear export whose activity is regulated by phosphorylation. Cell 101, 187–198 42 Zeller, R. et al. (1983) Nucleocytoplasmic distribution of snRNPs and stockpiled snRNA-binding proteins during oogenesis and early development in Xenopus laevis. Cell 32, 425–434 43 Mattaj, I.W. (1986) Cap trimethylation of U snRNA is cytoplasmic and dependent on U snRNP protein binding. Cell 46, 905–911 44 Plessel, G. et al. (1994) m3G cap hypermethylation of U1 small nuclear ribonucleoprotein (snRNP) in vitro: evidence that the U1 small nuclear RNA-(guanosine-N2)- methyltransferase is a non-snRNP cytoplasmic protein that requires a binding site on the Sm core domain. Mol. Cell Biol. 14, 4160–4172 45 Fischer, U. et al. (1994) Nuclear transport of U1 snRNP in somatic cells: differences in signal requirement compared with Xenopus laevis oocytes. J. Cell Biol. 125, 971–980 46 Fischer, U. and Luhrmann, R. (1990) An essential signaling role for the m3G cap in the transport of U1 snRNP to the nucleus. Science 249, 786– 790 47 Hamm, J. et al. (1990) The trimethylguanosine cap structure of U1 snRNA is a component of a bipartite nuclear targeting signal. Cell 62, 569–577 48 Stanek, D. and Neugebauer, K.M. (2004) Detection of snRNP assembly intermediates in Cajal bodies by fluorescence resonance energy transfer. J. Cell Biol. 166, 1015–1025 49 Stanek, D. et al. (2008) Spliceosomal small nuclear ribonucleoprotein particles repeatedly cycle through Cajal bodies. Mol. Biol. Cell 19, 2534–2543 50 Kiss, T. (2004) Biogenesis of small nuclear RNPs. J. Cell Sci. 117, 5949– 5951 51 Zhang, D. et al. (2001) A biochemical function for the Sm complex. Mol. Cell 7, 319–329
Trends in Biochemical Sciences
Vol.35 No.12
52 Raker, V.A. et al. (1996) The snRNP core assembly pathway: identification of stable core protein heteromeric complexes and an snRNP subcore particle in vitro. EMBO J. 15, 2256–2269 53 Meister, G. et al. (2001) A multiprotein complex mediates the ATPdependent assembly of spliceosomal U snRNPs. Nat. Cell Biol. 3, 945– 949 54 Lefebvre, S. et al. (1995) Identification and characterization of a spinal muscular atrophy-determining gene. Cell 80, 155–165 55 Neuenkirchen, N. et al. (2008) Deciphering the assembly pathway of Sm-class U snRNPs. FEBS Lett. 582, 1997–2003 56 Gubitz, A.K. et al. (2004) The SMN complex. Exp. Cell Res. 296, 51–56 57 Meister, G. and Fischer, U. (2002) Assisted RNP assembly: SMN and PRMT5 complexes cooperate in the formation of spliceosomal UsnRNPs. EMBO J. 21, 5853–5863 58 Pellizzoni, L. et al. (2002) Essential role for the SMN complex in the specificity of snRNP assembly. Science 298, 1775–1779 59 Friesen, W.J. et al. (2001) The methylosome, a 20S complex containing JBP1 and pICln, produces dimethylarginine-modified Sm proteins. Mol. Cell. Biol. 21, 8289–8300 60 Meister, G. et al. (2001) Methylation of Sm proteins by a complex containing PRMT5 and the putative U snRNP assembly factor pICln. Curr. Biol. 11, 1990–1994 61 Gonsalvez, G.B. et al. (2007) Two distinct arginine methyltransferases are required for biogenesis of Sm-class ribonucleoproteins. J. Cell Biol. 178, 733–740 62 Chari, A. et al. (2008) An assembly chaperone collaborates with the SMN complex to generate spliceosomal SnRNPs. Cell 135, 497–509 63 Fisher, D.E. et al. (1985) Small nuclear ribonucleoprotein particle assembly in vivo: demonstration of a 6S RNA-free core precursor and posttranslational modification. Cell 42, 751–758 64 Pu, W.T. et al. (1999) pICln inhibits snRNP biogenesis by binding core spliceosomal proteins. Mol. Cell. Biol. 19, 4113–4120 65 Urlaub, H. et al. (2001) Sm protein-Sm site RNA interactions within the inner ring of the spliceosomal snRNP core structure. EMBO J. 20, 187–196 66 Brahms, H. et al. (2001) Symmetrical dimethylation of arginine residues in spliceosomal Sm protein B/B0 and the Sm-like protein LSm4, and their interaction with the SMN protein. RNA 7, 1531–1542 67 Friesen, W.J. et al. (2001) SMN, the product of the spinal muscular atrophy gene, binds preferentially to dimethylarginine-containing protein targets. Mol. Cell 7, 1111–1117 68 Kroiss, M. et al. (2008) Evolution of an RNP assembly system: a minimal SMN complex facilitates formation of UsnRNPs in Drosophila melanogaster. Proc. Natl. Acad. Sci. U. S. A. 105, 10045–10050 69 Palfi, Z. et al. (2009) SMN-assisted assembly of snRNP-specific Sm cores in trypanosomes. Genes Dev. 23, 1650–1664 70 Battle, D.J. et al. (2006) The Gemin5 protein of the SMN complex identifies snRNAs. Mol. Cell 23, 273–279 71 Yong, J. et al. (2010) Gemin5 delivers snRNA precursors to the SMN complex for snRNP biogenesis. Mol. Cell 38, 551–562 72 Lau, C.K. et al. (2009) Gemin5-snRNA interaction reveals an RNA binding function for WD repeat domains. Nat. Struct. Mol. Biol. 16, 486–491 73 Facey, S.J. and Kuhn, A. (2010) Biogenesis of bacterial innermembrane proteins. Cell. Mol. Life Sci. 67, 2343–2362 74 Hagan, C.L. et al. (2010) Reconstitution of outer membrane protein assembly from purified components. Science 328, 890–892 75 Nishiyama, M. et al. (2008) Reconstitution of pilus assembly reveals a bacterial outer membrane catalyst. Science 320, 376–379 76 Vetsch, M. et al. (2006) Mechanism of fibre assembly through the chaperone-usher pathway. EMBO Rep. 7, 734–738 77 Waksman, G. and Hultgren, S.J. (2009) Structural biology of the chaperone-usher pathway of pilus biogenesis. Nat. Rev. Microbiol. 7, 765–774 78 Becker, T. et al. (2009) Biogenesis of mitochondrial membrane proteins. Curr. Opin. Cell Biol. 21, 484–493 79 Park, Y.J. and Luger, K. (2008) Histone chaperones in nucleosome eviction and histone exchange. Curr. Opin. Struct. Biol. 18, 282–289 80 Rocha, W. and Verreault, A. (2008) Clothing up DNA for all seasons: histone chaperones and nucleosome assembly pathways. FEBS Lett. 582, 1938–1949 81 Segal, E. and Widom, J. (2009) What controls nucleosome positions? Trends Genet. 25, 335–343
683