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Human Molecular Genetics Advance Access originally published online on July 11, 2006
Human Molecular Genetics 2006 15(16):2523-2532; doi:10.1093/hmg/ddl173
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© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Ataxin-2 and its Drosophila homolog, ATX2, physically assemble with polyribosomes

Terrence F. Satterfield and Leo J. Pallanck*

Department of Genome Sciences, University of Washington School of Medicine, Seattle, WA 98195-7730, USA

* To whom correspondence should be addressed. Tel: +1 2066165997; Fax: +1 2066857301; Email: pallanck{at}u.washington.edu

Received September 8, 2005; Accepted July 6, 2006


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Mutations resulting in the expansion of a polyglutamine tract in the protein ataxin-2 give rise to the neurodegenerative disorders spinocerebellar ataxia type 2 and Parkinson's disease. The normal cellular function of ataxin-2 and the mechanism by which polyglutamine expansion of ataxin-2 causes neurodegeneration are unknown. Here, we demonstrate that ataxin-2 and its Drosophila homolog, ATX2, assemble with polyribosomes and poly(A)-binding protein (PABP), a key regulator of mRNA translation. The assembly of ATX2 with polyribosomes is mediated independently by two distinct evolutionarily conserved regions of ATX2: an N-terminal Lsm/Lsm-associated domain (LsmAD), found in proteins that function in nuclear RNA processing and mRNA decay, and a PAM2 motif, found in proteins that interact physically with PABP. We further show that the PAM2 motif mediates a physical interaction of ATX2 with PABP in addition to promoting ATX2 assembly with polyribosomes. Our results suggest a model in which ATX2 binds mRNA directly through its Lsm/LsmAD domain and indirectly via binding PABP that is itself directly bound to mRNA. These findings, coupled with work on other ataxin-2 family members, suggest that ATX2 plays a direct role in translational regulation. Our results raise the possibility that polyglutamine expansions within ataxin-2 cause neurodegeneration by interfering with the translational regulation of particular mRNAs.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
CAG repeat expansions within the coding sequences of at least nine genes confer dominantly inherited neurodegenerative diseases by giving rise to proteins bearing expanded polyglutamine tracts (1,2). Increasing evidence suggests that an alteration of the normal cellular functions of the proteins bearing the polyglutamine expansion is at least a partial contributor to polyglutamine pathogenesis. This conclusion is best illustrated by the finding that many of the proteins affected by polyglutamine expansions normally function as transcription factors and that transcriptional dysregulation appears to be a central feature of the neurodegenerative mechanism in the corresponding disorders (3–7). However, the biological functions of many of the proteins affected by polyglutamine expansions are unknown, thus precluding the study of how the normal cellular functions of these proteins may relate to pathogenesis.

Spinocerebellar ataxia type 2 and a dominantly inherited form of Parkinsonism are caused by CAG repeat expansions in ATXN2 (8–15). The biological function of the ATXN2 gene product, ataxin-2, is currently unknown, although previous work on ataxin-2 and several closely related ataxin-2 homologs have implicated this protein family in diverse biological processes, including secretion, cell specification, actin filament formation and apoptotic and receptor-mediated signaling (16–21). Several observations suggest that the influence of the ataxin-2 family on these diverse biological processes results from a conserved biochemical function of these proteins in cytoplasmic RNA metabolism. For example, the N-terminus of ataxin-2 contains Lsm (like-Sm) and Lsm-associated domains (LsmAD), at least one of which, the Lsm domain, is known to function as an RNA-binding domain in proteins that mediate pre-mRNA splicing and mRNA decay (22,23). Moreover, ataxin-2 contains the poly(A)-binding protein (PABP) interacting motif PAM2 (24). PAM2 motifs are known to physically bind to the C-terminal region of the translational regulator PABP, and accordingly, ataxin-2 physically interacts with PABP (25). Finally, ataxin-2 interacts physically with a putative RNA-binding protein called the ataxin-2-binding protein (A2BP) (17).

Genetic analyses of several ATXN2 homologs have begun to suggest that the role of this gene family in RNA metabolism may involve translational regulation. For example, deletion of the yeast ATXN2 homolog, Pbp1, suppresses the lethality caused by deletion of the yeast PABP gene, Pab1 (26). Because Pab1 promotes translation, this genetic interaction suggests that Pbp1 normally serves to oppose the translation-promoting activity of Pab1. Further, genetic evidence that Pbp1 plays a translational role is provided by the observation that overexpression of this gene can phenocopy the effects of treating yeast cells with the translation inhibitor cycloheximide (27). Studies of the Caenorhabditis elegans ATXN2 homolog, ATX-2, indicate that reduced ATX-2 function alters the abundance of particular proteins without altering the abundance of their corresponding transcripts, suggesting that these transcripts may be translationally regulated by ATX-2 (28). Despite the evidence that the ATXN2 family is involved in translational regulation, there is currently no direct evidence that ataxin-2 engages the translational apparatus and is itself a translational regulator.

In previous work, we demonstrated that the Drosophila melanogaster ATXN2 homolog, ATX2, influences cell specification and actin filament formation (19). To test the hypothesis that the biological processes influenced by ATX2 derive from a role of ATX2 in RNA metabolism, we have now taken a biochemical approach to determine whether ATX2 physically assembles with mRNAs that are engaged with the translational machinery. Here, we demonstrate that ATX2 interacts with the translational machinery by assembling with polyribosomes. The assembly of ATX2 with polyribosomes is mediated independently by the Lsm/LsmAD domain and the PAM2 motif of ATX2. ATX2 also assembles with Drosophila PABP (dPABP) in a PAM2-dependent manner, and this interaction appears to link ATX2 to polyribosomes by allowing ATX2 to interact with polyribosome-bound dPABP. Finally, we show that human ataxin-2 is also able to interact physically with polyribosomes, suggesting that our structure/function studies of Drosophila ATX2 are likely to be relevant to ataxin-2. These findings, together with results from other studies, suggest that ataxin-2 proteins directly influence the translation of particular mRNAs.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Drosophila ATX2 assembles with polyribosomes
The Lsm and PAM2 domains are conserved features of the ataxin-2 family. Given that both of these domains have the potential to promote binding to RNA, we tested whether the Drosophila ataxin-2 homolog, ATX2, is capable of assembling with RNA (23,24). Post-mitochondrial extracts from Drosophila ovaries were subjected to sedimentation through 10–50% (w/v) continuous linear sucrose gradients and the distribution of ATX2 relative to dPABP and the 60S ribosomal protein P0 was analyzed. Ovary extracts were chosen for this analysis because our previous work indicates that ATX2 plays an essential cell-autonomous role in the female germline (19). Many soluble cytoplasmic proteins sediment in the upper, lighter fractions of the sucrose gradient, whereas polyribosomes and their associated proteins sediment into the lower, heavier fractions. ATX2 was detected in all fractions of the gradient, indicating that ATX2 is a component of complexes that are heterogeneous in size (Fig. 1). Also, as would be expected for a protein that assembles with polyribosomes, ATX2 co-sedimented with dPABP and P0 (Fig. 1).


Figure 1731
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  		Figure 1. ATX2 assembles with polyribosomes. Post-mitochondrial extracts from Drosophila ovaries were subjected to sedimentation through 10–50% (w/v) continuous linear sucrose gradients. Western blot analysis was used to test for the presence of ATX2, dPABP and the 60S ribosomal protein P0 (P0) in each of the 12 fractions recovered from each gradient. For EDTA and RNase controls, cell extracts were either prepared in the presence of 25 mM EDTA or treated with 100 µg/ml RNase A prior to sedimentation, respectively.

 
If ATX2 is present in high-density fractions because it physically assembles with polyribosomes, we would expect ATX2 to shift to lower-density fractions when subjected to treatments that dissociate polyribosomes. One such treatment involves the use of the metal chelator EDTA. EDTA treatment reduces the concentration of free Mg2+, a condition that causes ribosomes to disassemble into their 40S and 60S subunits and thus dissociate from mRNAs (29). Upon EDTA treatment, ATX2, dPABP and P0 all shifted to lower-density fractions, suggesting that ATX2, like dPABP and P0, is a component of polyribosomes (Fig. 1). As expected, P0, a component of the 60S subunit, shifted to a distribution that coincides with the 60S subunit upon EDTA treatment. In contrast, ATX2 and dPABP appear to remain in complexes exceeding 80S. This observation indicates that, unlike P0, ATX2 is not a structural component of ribosomes and suggests the possibility that these EDTA-resistant ATX2 complexes represent messenger ribonucleoprotein particles (mRNPs).

To directly test whether the EDTA-resistant ATX2 complexes represent mRNPs, we treated ovary extracts with RNase A prior to centrifugation to disrupt both polyribosomes and mRNPs. In contrast to our results with EDTA treatment, RNase A treatment shifted the distribution of ATX2 and dPABP to low-density gradient fractions containing soluble cytoplasmic proteins and low molecular weight complexes smaller than 60S (Fig. 1). This finding indicates that, like dPABP, ATX2 assembles into mRNP complexes. Neither ATX2 nor dPABP significantly co-sedimented with the 80S ribosomal complexes produced by RNase A treatment, further supporting our conclusion that, although both proteins assemble with polyribosomes, neither is a component of the ribosome itself (Fig. 1).

The Lsm/LsmAD domain and PAM2 motif independently mediate ATX2 assembly with polyribosomes
To identify the sequences in ATX2 that mediate its assembly with polyribosomes, we generated a series of constructs encoding proteins bearing deletions of putative polyribosome interaction regions and tested whether these proteins retained polyribosome assembly (Fig. 2A). We focussed on the Lsm/LsmAD domain and PAM2 motif because these are the only two evolutionarily conserved sequences in the ataxin-2 family and because similar sequences in other proteins have been linked to RNA metabolism (22–25). The Lsm portion of the Lsm/LsmAD domain is a known RNA-binding motif and could thus allow ATX2 to directly bind polyribosomal mRNA, whereas the PAM2 motif is a known PABP-interacting motif that could potentially link ATX2 to polyribosomes indirectly by binding to polyribosome-bound dPABP (23,24). Each of the constructs generated for this analysis encodes a protein bearing an N-terminal FLAG epitope and a deletion of either the Lsm/LsmAD domain or the PAM2 motif. The FLAG–ATX2-{Delta}C construct encodes a protein lacking ATX2 residues 299–1084, and thus encodes little more than the Lsm/LsmAD domain (Fig. 2A). Conversely, the FLAG–ATX2-{Delta}N construct encodes a protein lacking residues 1–298, and thus while the PAM2 motif is intact, the Lsm/LsmAD domain is deleted from this protein (Fig. 2A). These two constructs were transfected into Drosophila S2 cells for expression and subsequent gradient analysis.


Figure 1732
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  		Figure 2. The Lsm/LsmAD domain and PAM2 motif independently mediate ATX2 assembly with polyribosomes. (A) Schematic depiction of the ATX2 deletion constructs used in sucrose gradient analysis. The cross (X) indicates a deletion of the 15-residue PAM2 motif (ATX2 residues 855–869) in the FLAG–ATX2-{Delta}NC construct. (B) The ATX2 deletion constructs were transfected into Drosophila S2 cells and expressed by adding 500 µM CuSO4 to induce transcription from the metallothionein promoter positioned upstream of the deletion constructs. Forty-eight hours following induction, post-mitochondrial extracts from transfected S2 cells were subjected to sedimentation through 10–50% (w/v) continuous linear sucrose gradients. Western analysis was used to test for the presence of FLAG epitope-tagged proteins and dPABP in each of the 12 fractions recovered from each gradient.

 
To confirm that ATX2 assembles with polyribosomes in S2 cells and that the FLAG epitope does not interfere with polyribosome assembly, we first analyzed the sedimentation profile of a FLAG–ATX2 polypeptide that carries no deletions. Like endogenous ATX2, FLAG–ATX2 co-sediments with dPABP throughout the sucrose gradient (Fig. 2B) and shifts to low-density fractions upon treatment with RNase A (unpublished data), indicating that ATX2 assembles with polyribosomes in S2 cells and that the FLAG sequence does not interfere with this assembly process. We next evaluated the sedimentation properties of the FLAG–ATX2-{Delta}C and FLAG–ATX2-{Delta}N polypeptides. Like FLAG–ATX2, the FLAG–ATX2-{Delta}C and FLAG–ATX2-{Delta}N polypeptides distribute broadly through a gradient indicating that these mutationally altered proteins retain polyribosome assembly (Fig. 2B). Because the FLAG–ATX2-{Delta}C polypeptide consists of little more than the Lsm/LsmAD domain, this result indicates that the Lsm/LsmAD domain is sufficient to bind polyribosomes. However, the FLAG–ATX2-{Delta}N polypeptide, which completely lacks the Lsm/LsmAD domain, also retains polyribosome assembly, indicating that although sufficient, the Lsm/LsmAD domain is not necessary for ATX2 assembly with polyribosomes.

Our findings with the FLAG–ATX2-{Delta}C and FLAG–ATX2-{Delta}N polypeptides indicate that ATX2 bears two or more polyribosome-binding regions, one consisting of the N-terminal Lsm/LsmAD domain and the other consisting of a C-terminal region, possibly the PAM2 motif. To directly test this hypothesis, we created a third ATX2 deletion construct, designated FLAG–ATX2-{Delta}NC which encodes a protein that lacks the Lsm/LsmAD domain, but that additionally bears a small deletion (residues 855–869) spanning only the PAM2 motif (Fig. 2A). The FLAG–ATX2-{Delta}NC construct was transfected into S2 cells for expression and gradient analysis. In contrast to the FLAG–ATX2-{Delta}N and FLAG–ATX2-{Delta}C polypeptides, the FLAG–ATX2-{Delta}NC polypeptide fails to co-sediment with polyribosomes and instead remains in the low-density fractions at the top of the gradient (Fig. 2B). Because the FLAG–ATX2-{Delta}NC polypeptide only differs from the FLAG–ATX2-{Delta}N polypeptide by a deletion of the 15-residue PAM2 motif, this finding indicates that the PAM2 motif is responsible for the polyribosome assembly of FLAG–ATX2-{Delta}N and demonstrates that the Lsm/LsmAD domain and PAM2 motif independently mediate the assembly of ATX2 with polyribosomes.

ATX2 interacts with PABP in a PAM2-dependent manner
Whereas the Lsm/LsmAD domain of ATX2 likely binds RNA directly, the PAM2 motif is predicted to assemble with dPABP. Because dPABP directly assembles with polyribosomes, the presence of the PAM2 motif in ATX2 suggests that this motif facilitates the assembly of ATX2 with polyribosomes by linking ATX2 to polyribosome-bound dPABP. To begin to address this hypothesis, we performed immunoprecipitation experiments to test whether ATX2 physically assembles with dPABP. Using extracts from Drosophila ovaries, we could readily detect dPABP in immunoprecipitates of ATX2 using an ATX2-specific antiserum, but not when using pre-immune serum in place of the ATX2 antiserum (Fig. 3A). Importantly, RNase A treatment of cell extracts prior to immunoprecipitation failed to disrupt the physical interaction of ATX2 with dPABP, indicating that this interaction is not mediated indirectly through RNA (Fig. 3A).


Figure 1733
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  		Figure 3. ATX2 physically interacts with dPABP in a PAM2-dependent manner. (A) ATX2 antiserum was used to immunoprecipitate endogenous ATX2 from Drosophila ovary extracts. Western blot analysis was used to test for the presence of dPABP in the immunoprecipitate. To control for specificity, pre-immune serum was used in place of ATX2 antiserum. To rule out an indirect interaction of ATX2 and dPABP mediated by common binding of these factors to RNA, one sample was treated with 100 µg/ml RNase A prior to immunoprecipitation with ATX2 antiserum. The efficiency of RNase A digestion was confirmed by visualizing the RNA from immunoprecipitated cell extracts on an agarose gel. (B) Schematic depiction of the ATX2 deletion constructs used for immunoprecipitations. The cross (X) indicates a deletion of the 15-residue PAM2 motif (ATX2 residues 855–869) in the FLAG–ATX2-{Delta}NC construct. (C) The ATX2 deletion constructs were transfected into Drosophila S2 cells and expressed by adding 500 µM CuSO4 to induce transcription from the metallothionein promoter positioned upstream of the deletion constructs. Forty-eight hours following induction, protein extracts were prepared and subjected to immunoprecipitation with anti-FLAG coated beads. Western analysis was used to test for the presence of dPABP in the immunoprecipitates. Arrowheads designate the locations of mutationally altered ATX2 proteins. The asterisks denote an anti-FLAG cross-reacting species present in S2 cells.

 
If the PAM2 motif is solely responsible for the physical interaction between ATX2 and dPABP, then only ATX2 polypeptides bearing the PAM2 motif should bind dPABP. To test this hypothesis, we performed immunoprecipitation experiments using the ATX2 deletion constructs. Results of this analysis indicate that the FLAG–ATX2 and FLAG–ATX2-{Delta}N polypeptides, which possess the PAM2 motif, are capable of efficiently immunoprecipitating dPABP (Fig. 3B and C). In contrast, FLAG–ATX2-{Delta}C and FLAG–ATX2-{Delta}NC, which lack the PAM2 motif, fail to immunoprecipitate dPABP (Fig. 3B and C). Importantly, although the FLAG–ATX2-{Delta}N and FLAG–ATX2-{Delta}NC polypeptides differ from one another only by a single 15-residue deletion of the PAM2 motif in FLAG–ATX2-{Delta}NC, this small difference is sufficient to abolish the ATX2–dPABP interaction. These results demonstrate that the PAM2 motif is essential for ATX2 to bind dPABP. Together with the results from our gradient analyses, these results support the model that the PAM2 motif links ATX2 to polyribosomes by physically interacting with polyribosome-bound dPABP.

Human ataxin-2 assembles with polyribosomes
To determine the relevance of our findings with Drosophila ATX2 to the function of human ataxin-2, we also tested whether ataxin-2 from a human cell line (HEK293-FT) can assemble with polyribosomes. Like ATX2, ataxin-2 distributes broadly through a gradient, and RNase A and EDTA treatment shifted ataxin-2 from high-density to low-density gradient fractions (Fig. 4). The distribution of ataxin-2 extended into more dense fractions of the gradient than P0 upon EDTA treatment, indicating that ataxin-2 is also a component of large, non-polyribosomal mRNPs. Moreover, though similar to PABP, the distribution of ataxin-2 differed from that of P0 upon either RNase A or EDTA treatment indicating that, like ATX2, ataxin-2 is not a component of the ribosome. Together, these results demonstrate that, like ATX2, ataxin-2 is a component of polyribosomes and non-polyribosomal mRNPs.


Figure 1734
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  		Figure 4. Human ataxin-2 assembles with polyribosomes. Post-mitochondrial extracts from HEK293-FT cells were subjected to sedimentation through 10–50% (w/v) continuous linear sucrose gradients. Western blot analysis was used to test for the presence of ataxin-2, PABP and the 60S ribosomal protein P0 (P0) in each of the 12 fractions recovered from each gradient. For EDTA and RNase controls, cell extracts were either prepared in the presence of 25 mM EDTA or treated with 100 µg/ml RNase A prior to sedimentation, respectively.

 
Because polyglutamine expansions within ataxin-2 are the cause of neurodegeneration in disorders involving this protein, we also examined whether polyglutamine expanded forms of ataxin-2 influence the ability of this protein to assemble with polyribosomes. To address this matter, GFP-tagged versions of ataxin-2 either bearing a normal polyglutamine repeat (Q22) or an expanded (58Q) polyglutamine repeat were expressed in HEK293-FT cells. Protein extracts from these cell lines were subjected to sucrose density gradients to compare the sedimentation properties of the GFP-tagged Q22 and Q58 versions of ataxin-2. Results of this analysis failed to detect obvious differences in the sedimentation properties of these proteins relative to endogeneous ataxin-2, indicating that neither the GFP tag nor the polyglutamine expansion interfere substantially with the ability of ataxin-2 to assemble with polyribosomes (unpublished data).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Ataxin-2 is one of nine proteins known to cause neurodegeneration in response to the expansion of an endogenous polyglutamine tract (1,2). Although the mechanism of neurodegeneration in these disorders is only partially understood, the findings that many polyglutamine disease proteins normally regulate transcription and that transcriptional dysregulation appears to be a central feature of the corresponding disorders suggest that pathogenesis results, at least in part, from an effect of the polyglutamine expansion on the normal cellular functions of these proteins (3–7). These findings raise the possibility that polyglutamine expansions within ataxin-2 might similarly induce pathogenesis by altering the normal biological function of this protein. To provide the necessary foundation to explore this hypothesis, we have been studying the normal cellular function of ATX2, a Drosophila homolog of ataxin-2. In previous work, we showed that altered dosage of ATX2 influences actin filament formation. Our current results demonstrate that ATX2 utilizes multiple independent mechanisms to physically engage the translational machinery, indicating that the phenotypes resulting from altered ATX2 dosage derive from a biochemical defect in mRNA metabolism. Importantly, we also find that human ataxin-2 assembles with polyribosomes. Given that the domains of ATX2 that appear to mediate interactions with polyribosomes are conserved in human ataxin-2, the results of our structure/function analyses of Drosophila ATX2 are likely to be directly relevant to the human protein.

Our finding that ataxin-2 and its Drosophila counterpart assemble with polyribosomes suggests possible roles for these proteins in mRNA transport, stability and/or translation. Although we cannot at present definitively exclude any of these possibilities, the weight of evidence seems to favor a role for the ataxin-2 protein family in translation. Four ataxin-2 family members are now known to bind the translational regulator PABP, and studies of the yeast ATXN2 homolog, Pbp1, indicate that it interacts genetically with the yeast PABP gene, Pab1 (25,26,28). Although the genetic interaction between Pbp1 and Pab1 could be explained by a role for Pbp1 in mRNA stability, this appears unlikely because overall mRNA abundance is unaltered in Pbp1{Delta} mutants (26). Moreover, overexpression of Pbp1 phenocopies the effect of the translational inhibitor cycloheximide in suppressing the phenotypes resulting from cytoplasmic accumulation of mitochondrial proteins in tim18{Delta} mutants (27), suggesting that Pbp1 plays a translational regulatory role. Finally, studies of the C. elegans ATXN2 homolog, ATX-2, have shown that inactivation of ATX-2 influences the abundance of particular proteins without influencing the abundance of the corresponding mRNAs, suggesting that this factor also plays a translational regulatory role (28).

Although the above evidence suggests that members of the ataxin-2 protein family function as translational regulators, it is unclear whether these proteins play a positive or negative role in translation. The findings in yeast, that deletion of Pbp1 suppresses the effects of a Pab1 deletion and that overexpression of Pbp1 mimics the effects of cycloheximide treatment, suggest that Pbp1 plays an inhibitory role in translation (26,27). However, these findings are difficult to reconcile with the observation that the overall rate of 35S-labeled methionine incorporation into proteins is unaltered in Pbp1{Delta} mutants (26). In C. elegans, loss of ATX-2 increases the abundance of some proteins while reducing the abundance of others, suggesting the possibility that ATX-2 serves both positive and negative regulatory roles in translation (28). Moreover, our previous work with ATX2 mutants indicates that the abundance of most proteins is unaltered in these mutants (19 and unpublished data). Together, these results suggest that only a subset of transcripts are subject to regulation by the ataxin-2 protein family and/or that the ataxin-2 family of proteins play both positive and negative translational regulatory roles. Important future goals will be to identify the specific transcripts bound by the ataxin-2 protein family and to directly explore the effects of the ataxin-2 protein family on the translation of these transcripts.

Given the evidence supporting a role for ataxin-2 in translational regulation, the question arises as to the mechanism by which ataxin-2 imposes this regulation. One possibility is that ataxin-2 directly influences the activity of PABP. PABP promotes translation by facilitating the interaction between the 5' and 3' ends of the mRNA, a process thought to promote the re-initiation of translation of terminating ribosomes (30). PABP accomplishes this task by simultaneously binding to the poly(A) tail and to the PAM2 motif of eIF4G, a component of the 5' cap-binding translation initiation complex (Fig. 5A). Another PAM2 protein, Paip1, mimics the activity of eIF4G by simultaneously binding poly(A)-bound PABP and eIF4A, another component of the 5' cap-binding translation initiation complex (31). In contrast to eIF4G and Paip1, another PAM2 protein, Paip2, inhibits translation. Paip2 accomplishes this task by binding PABP and preventing its assembly onto the poly(A) tail (30,32,33). Our finding that ATX2 is capable of assembling with poly(A)-bound dPABP indicates that, unlike Paip2, ATX2 does not prevent dPABP from assembling with the poly(A) tail. Assuming that ATX2 influences dPABP activity, it appears to do so while dPABP is assembled with the poly(A) tail, possibly by promoting or preventing the interaction between dPABP and the 5' cap-binding translation initiation complex (Fig. 5B). Further work will be required to elucidate the functional significance of the ATX2–dPABP interaction.


Figure 1735
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  		Figure 5. Model depicting how ATX2 might influence translation. (A) In the absence of ATX2, poly(A)-bound dPABP normally promotes successive rounds of translation by physically interacting with eIF4G, a component of the 5' cap-binding translation initiation complex. (B) The PAM2 motif of ATX2 may interfere with the translation promoting activity of dPABP by competitively inhibiting the interaction between dPABP and eIF4G. Alternatively, ATX2 may influence the activity of dPABP in a novel fashion to promote or inhibit translation of particular mRNA targets. The Lsm/LsmAD domain of ATX2 may facilitate interactions between a microRNA and its mRNA target, thereby altering mRNA structure and inhibiting translation.

 
Although previous evidence indicates that ataxin-2 family members interact functionally with PABP, several observations indicate that ataxin-2 does not act solely through PABP. For example, in yeast, Pbp1 deletions suppress the lethality caused by deletion of Pab1, indicating that Pbp1p can perform a functional role in the complete absence of Pab1p (26). Moreover, we have found that an ATX2 transgenic construct that encodes a protein lacking the PAM2 motif significantly extends the lethal phase of ATX2 mutant flies (unpublished data). Although ATX2 null mutants do not develop beyond the second instar larval stage, these mutants can be rescued to the adult stage of development using a wild-type ATX2 transgene (19). Use of ATX2 transgenes bearing a PAM2 deletion can also extend the lethal phase of ATX2 mutants to the pupal stage of development, although none of the partially rescued offspring survives to the adult stage of development. Given that the PAM2 motif is required for ATX2 to interact with dPABP, this observation indicates that ATX2 possesses a biological activity that is independent of its physical interaction with dPABP. Our finding that the Lsm/LsmAD domain of ATX2 is capable of promoting its assembly with polyribosomes independently of the PAM2 motif, together with the observation that the Lsm/LsmAD domain represents the only other evolutionarily conserved sequence in ATX2, suggests that this domain is the source of the dPABP-independent activity of ATX2.

Assuming that the Lsm/LsmAD domain is responsible for the dPABP-independent activity of ATX2 and that this activity serves a translational regulatory role, the question arises as to the precise mechanism by which this domain regulates translation. Although eukaryotic Lsm and the related Sm proteins are not currently known to regulate translation, one well-studied bacterial Sm protein, Hfq, does appear to regulate translation. Hfq functions as an RNA chaperone and regulates translation by stabilizing basepairing interactions between small non-coding RNAs (sRNAs) and their mRNA targets (34). These sRNA–mRNA interactions influence translation by altering the physical structure of the mRNA target. Although only limited sequence homology exists between Hfq and other Sm and Lsm proteins, the crystal structures of Staphylococcus aureus Hfq and eukaryotic Sm proteins are nearly identical, indicating that these proteins may function in a similar fashion (35). Furthermore, studies of eukaryotic Sm and Lsm proteins suggest that these proteins also function by mediating RNA–RNA interactions (36,37). The structural similarity of Sm and Lsm proteins raises the possibility that the Lsm/LsmAD domain of ATX2 might function, like Hfq, to regulate translation by mediating RNA–RNA interactions. An attractive potential target of ataxin-2 regulation in eukaryotes is the group of sRNAs known as microRNAs. MicroRNAs are known to play translational regulatory roles by basepairing with particular target mRNAs on polyribosomes (Fig. 5B) (38–41). Studies are currently underway in the laboratory to investigate this possible mode of ATX2 regulation.

Previous work on several other polyglutamine disorders indicates that pathogenesis results from a reduction in transcriptional efficiency (3–7). Although the cytoplasmic localization of ataxin-2 indicates that this protein does not directly influence transcription, our finding that ATX2 physically assembles with polyribosomes, coupled with other work on ataxin-2 homologs, raises the possibility of a conserved mechanism of polyglutamine pathogenesis involving dysfunctional gene expression. In contrast to the transcriptional alterations associated with other polyglutamine diseases, polyglutamine expansions within ataxin-2 may adversely affect gene expression by impairing translation. Although polyglutamine expansion of human ataxin-2 does not detectably influence the binding of ataxin-2 to polyribosomes, it remains conceivable that polyglutamine expansions within ataxin-2 influence a function of ataxin-2 in translational regulation that lies downstream of polyribosome binding. The finding that polyglutamine expansions within ataxin-2 also cause a heritable form of Parkinsonism (9–15) further suggests that altered translational regulation of particular targets might trigger the degeneration of dopaminergic neurons in the substantia nigra. As increasing evidence indicates that an overabundance of the protein alpha-synuclein plays an important role in the pathogenesis of Parkinson's disease (42), our findings raise the interesting possibility that polyglutamine expansions within ataxin-2 lead to increased translation of alpha-synuclein. Future studies aimed at a better understanding of ataxin-2 function and the effects of polyglutamine expansions on that function will be required to directly address the hypothesis that translational dysregulation underlies ataxin-2-mediated neurodegeneration. Our current work provides a foundation for these studies.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Molecular constructs
The FLAG–ATX2 construct was generated using the ATX2 cDNA clone GH27029 as a template in a PCR reaction involving the primers FLAG–ATX2-FORWARD (AAAGATCTGAGCTCATGGACTATAAAGACGACGACGACAAAAACAACAATAGCA) and DSCA-CK-1 (TGCACCAAGTGCCGCAAA). The FLAG–ATX2-FORWARD primer incorporates the restriction sites BglII and SacI for re-insertion in the ATX2 cDNA clone GH27029 situated in the pOT2a vector and subsequent subcloning into the cell expression vector pRmHa3-NTAP (a gift from T. Furuyama), an ATG start codon, and the FLAG epitope coding sequence. DSCA-CK-1 lies downstream of a unique KpnI restriction site in the ATX2 cDNA. The PCR product was digested with BglII and KpnI and inserted between the unique BglII and KpnI sites of the ATX2 cDNA clone GH27029 situated in the pOT2a vector. This modification yielded pOT2a::FLAG–ATX2.

To create the FLAG–ATX2-{Delta}C construct, the ATX2 cDNA clone GH27029 was used as a template in a PCR reaction involving the primers FLAG–ATX2-FORWARD and ATX2-NTRUNC (AAACTCGAGTCAGCGATCACGATCACGGTC). FLAG–ATX2-FORWARD lies upstream of a unique XmnI site in the ATX2 cDNA. ATX2-NTRUNC incorporates a premature TGA termination codon and an XhoI site for insertion in pOT2a::FLAG–ATX2 and subsequent subcloning into pRmHa3-NTAP. The PCR product was digested with XmnI and XhoI and inserted between the unique XmnI and XhoI sites of the pOT2a::FLAG–ATX2 construct. This modification yielded pOT2a::FLAG–ATX2-{Delta}C.

To create the FLAG–ATX2-{Delta}N construct, the ATX2 cDNA clone GH27029 was used as a template in a PCR reaction involving the primers ATX2-DELTAN2 (AAAAGATCTGAGCTCATGGACTATAAAGACGACGACGACAAAGGTAACAAGCCCCGTGGC) and DSCA-CK-1. ATX2-DELTAN2 incorporates the restriction sites BglII and SacI for re-insertion in the ATX2 cDNA clone GH27029 situated in the pOT2a vector and subsequent subcloning into the cell expression vector pRmHa3-NTAP, an ATG start codon, and the FLAG epitope coding sequence. DSCA-CK-1 lies downstream of a unique SphI site in the ATX2 cDNA. The PCR product was digested with BglII and SphI and inserted between the unique BglII and SphI sites of the GH27029 cDNA clone situated in the pOT2a vector to yield pOT2a:: FLAG–ATX2-{Delta}N.

To create the FLAG–ATX2-{Delta}NC construct, the ATX2 cDNA clone GH27029 was used as a template in a PCR reaction involving the primers DELTA-C-FINAL (TAAGGTGATGCGCAACAATG) and ATX2-DELTA-C (AAGGGCCCAACCACCGGAGTGGTTCC). DELTA-C-FINAL lies upstream of a unique SphI site in the ATX2 cDNA, and ATX-DELTA-C incorporates an ApaI site for subcloning. The PCR product was digested with SphI and ApaI and inserted between the unique SphI and ApaI sites of pOT2a::FLAG–ATX2-{Delta}N to yield pOT2a::FLAG–ATX2-{Delta}NC.

For expression in cultured Drosophila S2 cells, all FLAG–ATX2 inserts were removed from pOT2a as SacI–XhoI fragments and then ligated between the SacI and SalI sites of the Drosophila expression vector pRmHa3-NTAP.

To create an ATX2 transgenic construct lacking the 15-residue PAM2 motif (residues 855–869), we replaced the SphI–ApaI fragment of the ATX2 clone GH27029 residing in the pOT2a vector (pOT2a::ATX2) with the SphI–ApaI fragment from pOT2a::FLAG–ATX2-{Delta}NC, a modification that yields pOT2a::ATX2-{Delta}C1. The modified cDNA insert was then removed from the pOT2a vector and inserted into the Drosophila transformation vector pUAST as a BglII–XhoI fragment to yield pUAST::ATX2-{Delta}C1. Except for the small deletion of the PAM2 motif, this construct is identical to the previously described ATX2 transgenic construct that was used to generate the UAS-Datx2.1B, UAS-Datx2.3 and UAS-Datx2.4 transgenic lines that are able to rescue ATX2 mutants to adulthood (19).

Maintenance and transfection of cell lines
Drosophila S2 cells were maintained at 25°C in Schneider's media (Sigma) supplemented with 10% fetal bovine serum (Invitrogen) and 100 U of penicillin and 100 µg of streptomycin/ml (Invitrogen). For each transfection, 6 ml of log-phase cells were washed twice with serum-free Schneider's media and then pelleted and resuspended in 780 µl of serum-free media in a 0.4 cm gap electroporation cuvette (BioRad). Twenty micrograms (1 µg/µl) of expression plasmid DNA was added, and cells were electroporated (400 V, 960 µF) using a Gene Pulser II (BioRad). Following transfection, samples were resuspended in 6 ml of complete Schneider's media. For each construct, two separate transfections were plated in a total of 12 ml into a 75 cm2 tissue culture flask. Twenty-four hours following the transfection, protein expression was induced by adding CuSO4 to 500 µM. Cells were harvested for gradient analysis 48 h after induction.

HEK293-FT cells were cultured in Dulbecco's modified Eagle's medium (Invitrogen) supplemented with 10% fetal bovine serum (Invitrogen). For experiments involving constructs encoding GFP-tagged ataxin-2 proteins, the plasmids GFP-ataxin-2[Q22] and GFP-ataxin-2[Q58] (18) were transfected into HEK293-FT cells using the Lipofectamine 2000 reagent (Invitrogen) according to manufacturer's instructions.

Sucrose density gradient fractionation
To prepare extracts for sucrose gradient analysis, groups of 50 ovaries were dissected in cold PBS containing 100 µg/ml cycloheximide (Sigma), transferred to 0.7 ml cold modified Drosophila extract buffer (125 mM sucrose, 25 mM HEPES, pH 6.9, 100 mM KCl, 5 mM MgCl2, 1 mM DTT) supplemented with Complete® protease inhibitor cocktail (Roche) and subjected to 30 strokes in a pre-chilled 2 ml dounce homogenizer. Drosophila S2 cells in log phase were incubated with 100 µg/ml cycloheximide for 10 min, washed twice with cold PBS containing 100 µg/ml cycloheximide, resuspended in 1 ml polyribosome lysis buffer (25 mM HEPES, pH 6.9, 100 mM KCl, 5 mM MgCl2, 1 mM DTT, 0.5% NP-40) supplemented with Complete® protease inhibitor cocktail and incubated on ice for 10 min. Post-mitochondrial supernatants from ovaries and S2 cell extracts were obtained by centrifugation of protein extracts at 10 000g for 10 min at 4°C. Post-mitochondrial extracts (15–20 OD A260 units) were layered onto 11 ml 10–50% (w/v) continuous linear sucrose gradients in polyribosome buffer (25 mM HEPES, pH 6.9, 100 mM KCl, 5 mM MgCl2, 1 mM DTT) supplemented with Complete® protease inhibitor cocktail and 100 µg/ml cycloheximide. For EDTA disruption of polyribosomes, 25 mM EDTA was used in place of MgCl2 in the lysis and gradient buffers. For RNase disruption of polyribosomes, 100 µg/ml RNase A (Sigma) was added to post-mitochondrial protein extracts and incubated for 10 min at room temperature prior to sucrose density gradient centrifugation.

Post-mitochondrial extracts were subjected to centrifugation in a Beckman SW41-Ti rotor at 36 000 r.p.m. for 1 h and 50 min at 4°C followed by collection of 12–1 ml fractions. Proteins in each fraction were precipitated with 10% trichloroacetic acid (Sigma), and following two washes with cold 95% ethanol, protein pellets were resuspended by incubation at 37°C for 10 min in 20 µl 100 mM Tris, pH 8.0 containing 1% SDS. Twenty microliters of 2x SDS–PAGE sample buffer was added, and proteins were denatured by boiling for 10 min. Twenty microliters of each sample was used for analysis by SDS–PAGE as described subsequently.

The preparation of HEK293-FT cellular lysates and the sucrose gradient centrifugation of these lysates were carried out using the same methods that were used to perform these experiments with Drosophila S2 cell extracts, except that 25 mM Tris, pH 7.8, was used in place of 25 mM HEPES in all buffers.

Immunoprecipitations
To generate ovary protein extracts for immunoprecipitation, 50 ovaries were dissected in cold PBS, transferred to 100 µl of cell lysis buffer (50 mM Tris, pH 6.9, 150 mM NaCl, 0.5% NP-40, 1 mM DTT) supplemented with Complete® protease inhibitor cocktail and manually homogenized with a small pestle in a 1.5 ml Eppendorf tube. To prepare S2 cell protein extracts, 2x107 transfected cells were pelleted, washed twice with cold PBS and lysed in 100 µl cell lysis buffer supplemented with Complete® protease inhibitor cocktail. Post-mitochondrial supernatants from ovaries and S2 cells were obtained as described above. Protein concentrations of post-mitochondrial supernatants were determined using a commercial protein assay (BioRad), adjusted to 2 µg/µl with PBS, and 500 µl aliquots were pre-cleared with 10 µl of protein A beads (Amersham) for 30 min at 4°C. For RNase A controls, cell extracts were treated with 100 µg/ml RNase A before adjusting the protein concentration. To generate antibody-coated beads, 10 µl of undiluted rabbit anti-ATX2 or pre-immune serum was incubated overnight at 4°C with 100 µl of a 50% slurry of protein A beads in PBS, followed by three washes with PBS. Ten microliters of either anti-ATX2 or pre-immune serum coated beads were then incubated with the pre-cleared post-mitochondrial protein extracts overnight at 4°C. The beads were collected and washed four times with PBS the following day. Following the washes, beads were boiled in 20 µl 2x SDS–PAGE sample buffer for 10 min, and the supernatants were analyzed by SDS–PAGE as described subsequently.

Western blot analysis
Proteins were separated by SDS–PAGE on 8% acrylamide gels and electrophoretically transferred onto nitrocellulose membranes (Amersham) as previously described (43). Immunodetection of FLAG-modified proteins, ATX2, dPABP, ribosomal P0, PABP and ataxin-2 was performed using anti-FLAG M2 monoclonal antibody (1:10 000; Sigma), rabbit anti-ATX2 (1:15 000) (19), rabbit anti-dPABP (1:5000) (33), human anti-P0 (1:3000; Immunovision), rabbit anti-PABP (1:1000) (44) and mouse anti-ataxin-2 (1:250; BD Transduction Labs), respectively.

Transgenic rescue of ATX2 mutants
All fly crosses were performed on standard cornmeal/molasses food at 25°C. Using standard procedures, the pUAST::ATX2-{Delta}C1 construct was used to generate two independent transgenic lines (UAS-ATX2-{Delta}C1a and UAS-ATX2-{Delta}C1b) that were subsequently crossed into a homozygous ATX2X1 null background. ATX2X1 homozygotes bearing a single copy of the transgene were monitored until lethality occurred. As a positive control for the transgenic rescue, simultaneous crosses were performed using the previously described UAS-Datx2.4 transgenic line (19). A minimum of 300 ATX2X1 heterozygotes were scored for each rescue cross performed.


   ACKNOWLEDGEMENTS
 
The authors would like to thank D. Morris, V. Mackay and C. Connolly for advice and assistance with the sucrose gradient experiments, T. Furuyama for providing the pRmHa3-NTAP vector and for advice on cell transfections, N. Sonenberg and G. Pavlakis for the dPABP and PABP antisera, respectively, S. Pulst for GFP-ataxin-2 plasmids, R. Ciosk for advice on immunoprecipitations, N. Smith and L. Andrews for technical support, A. LaSpada and S. Gartler for the use of laboratory equipment and members of the Pallanck laboratory for scientific advice and critical reading of this manuscript. This work was supported by the Stroum Endowed Minority Fellowship (through the University of Washington Graduate Opportunities and Minority Achievement Program to T.F.S.) and a training grant from the National Institutes of Health (GM07735-23).

Conflict of Interest statement. None declared.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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