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Proteasome inhibitors induce nucleolar aggregation of proteasome target proteins and polyadenylated RNA by altering ubiquitin availability


The ubiquitin–proteasome pathway is essential for most cellular processes, including protein quality control, cell cycle, transcription, signaling, protein transport, DNA repair and stress responses. Hampered proteasome activity leads to the accumulation of polyubiquitylated proteins, endoplastic reticulum (ER) stress and even cell death. The ability of chemical proteasome inhibitors (PIs) to induce apoptosis is utilized in cancer therapy. During PI treatment, misfolded proteins accrue to cytoplasmic aggresomes. The formation of aggresome-like structures in the nucleus has remained obscure. We identify here a nucleolus-associated RNA-protein aggregate (NoA) formed by the inhibition of proteasome activity in mammalian cells. The aggregate forms within the nucleolus and is dependent on nucleolar integrity, yet is a separate structure, lacking nucleolar marker proteins, ribosomal RNA (rRNA) and rRNA synthesis activity. The NoAs contain polyadenylated RNA, conjugated ubiquitin and numerous nucleoplasmic proteasome target proteins. Several of these are key factors in oncogenesis, including transcription factors p53 and retinoblastoma protein (Rb), several cell cycle-regulating cyclins and cyclin-dependent kinases (CDKs), and stress response kinases ataxia-telangiectasia mutated (ATM) and Chk1. The aggregate formation depends on ubiquitin availability, as shown by modulating the levels of ubiquitin and deubiquitinases. Furthermore, inhibition of chromosome region maintenance 1 protein homolog (CRM1) export pathway aggravates the formation of NoAs. Taken together, we identify here a novel nuclear stress body, which forms upon proteasome inactivity within the nucleolus and is detectable in mammalian cell lines and in human tissue. These findings show that the nucleolus controls protein and RNA surveillance and export by the ubiquitin pathway in a previously unidentified manner, and provide mechanistic insight into the cellular effects of PIs.


Ubiquitin is a small conserved protein that is covalently linked to its target proteins by ubiquitylation (Hershko and Ciechanover, 1998; Weissman, 2001; Finley, 2009). Ubiquitin is essential for many if not most cellular processes, including protein quality control, cell cycle and transcriptional control, cellular signaling, protein transport, DNA repair and stress responses. These diverse functions are governed by attachment of mono- and polyubiquitin chains on the targets through ubiquitin lysine residues. Polyubiquitylation generally leads to proteolytic degradation of the target by the proteasome. Polyubiquitylation occurs predominantly through Lys48 residues, although recently ubiquitin chains based on most lysine residues have been implicated in proteasomal targeting (Ikeda and Dikic, 2008; Saeki et al., 2009; Xu et al., 2009). Monoubiquitylation regulates, for example, membrane transport, nucleocytoplasmic protein localization, protein kinase activation, DNA repair and chromatin dynamics (Chen and Sun, 2009). Conjugated ubiquitin is released during proteasomal processing and by deubiquitylating enzymes (Reyes-Turcu et al., 2009), and reused.

The ubiquitin–proteasome pathway is a major proteolytic system of all eukaryotic cells (Hershko and Ciechanover, 1998; Weissman, 2001; Finley, 2009). Inhibition of proteasome activity leads to the accumulation of polyubiquitylated and misfolded proteins, ER stress and eventually apoptosis (Kopito, 2000; Navon and Ciechanover, 2009). Proteasome activity can be inhibited by chemical inhibitors, which fall into several classes based on their chemical structure, mechanism of inhibition and specificity (Kisselev and Goldberg, 2001). Several in vitro studies utilize peptide aldehydes (such as MG132, ALLN), which are substrate analogs, or a non-peptide inhibitor lactacystin. The ability of proteasome inhibitors (PIs) to induce cell death is exploited in clinical cancer treatment with bortezomib (Velcade; PS-341; MG-341), which is a Food and Drug Administration-approved PI in clinical use in mantle cell lymphoma and multiple myeloma (Navon and Ciechanover, 2009). The potential use of PIs against solid tumors is under active investigation.

Upon PI treatment, polyubiquitylated proteins accumulate at sites of proteasomes, forming organized aggregates. Most prominently, this occurs in the pericentrosomal area in the cytoplasm, in structures called aggresomes (Wójcik et al., 1996; Johnston et al., 1998). In the nucleus, chemical inhibition of the proteasome induces transient nuclear stress granules containing stress response proteins such as heat-shock factors (Holmberg et al., 2000), but no aggresome formation in the nucleus has been reported. We and others have shown that proteasome inhibitor MG132 causes translocation of certain stress response-related nuclear proteins (p53, MDM2, PML, Hsp70) to the nucleolus (Klibanov et al., 2001; Mattsson et al., 2001; Latonen et al., 2003; Kurki et al., 2004; Karni-Schmidt et al., 2007). As these proteins are degraded by the proteasome, the question has arisen as to whether the nucleolus has a role in the degradation of these proteins.

The nucleolus is membrane-less nuclear organelle, which is responsible for the assembly of ribosomal subunits (Leary and Huang, 2001; Fatica and Tollervey, 2002). The nucleoli are composed of fibrillar centers, dense fibrillar component and granular component (Olson et al., 2000; Hernandez-Verdun, 2006; Boisvert et al., 2007), in which ribosomal RNA (rRNA) transcription, maturation of pre-RNA transcripts, assembly of pre-ribosomal particles and late RNA processing occur (Lafontaine and Tollervey, 2001; Leary and Huang, 2001). Proteomic analyses have revealed that besides proteins involved in ribosome synthesis, the nucleolus contains a large number of proteins associated with cell cycle and mitotic division regulation, DNA repair and control of tumor suppressor proteins and oncogenes (Andersen et al., 2005; Boisvert et al., 2007).

Here, we investigated the possible role of the nucleolus in the ubiquitin–proteasome pathway. We show that nuclear proteasome targets accumulate to, and are immobilized in, aggregates forming in the nucleoli. These aggregates accumulate polyadenylated RNA (polyA(+)), but not rRNA, possibly reflecting defective RNAs targeted for degradation (Houseley and Tollervey, 2009). Formation of these structures depends on the nucleolar rRNA synthetic activity. We show that an increase in ubiquitin pool overcomes the formation of the aggregate, and conversely, that inhibition of nuclear export aggravates it. We propose that the formation of the aggregates is a consequence of the decreased availability of ubiquitin, which leads to the accumulation and immobilization of multiple proteins and polyA(+) RNA to the aggregate. These findings define a novel nucleolus-dependent structure that forms following severe overload of ubiquitylated, and likely misfolded, proteins.


Proteasome inhibition alters nucleolar morphology

We and others have earlier shown that tumor suppressor p53, a target protein of the proteasome, localizes to nucleoli in PI-treated cells (Klibanov et al., 2001; Latonen et al., 2003; Karni-Schmidt et al., 2007). To study how the nucleoli are affected upon proteasome inhibition, we treated WS1 human skin fibroblasts and HEL-299 human lung fibroblasts with MG132 (10 μM) for 12 h. At this time, cell death is not yet prevailing (Supplementary Figure S1a). We immunostained nucleolar substructures using antibodies against nucleophosmin (NPM; for granular component), fibrillarin (FBL; for dense fibrillar component) and upstream binding factor (UBF; for fibrillar center). Treatment of the cells with MG132 led to drastic changes in the localization of the nucleolar markers (Figure 1a). NPM formed a ring-shaped structure, which partially overlapped by FBL and UBF (Figure 1a). Under phase-contrast illumination, a dense structure became visible that appeared to localize in the center of the reorganized nucleolus. Similar relocalization was observed by using antibodies against nucleolar antigens Ki-67 and ARF (data not shown). To study this effect in more detail, we performed transmission EM (TEM) of mock- and MG132-treated WS1 cells. Although the nucleolar substructures were visible in the mock-treated cells, MG132 treatment caused alterations in the nucleolar morphology (Figure 1b). This was apparent by interruption of the nucleolus by electron-dense material (Figure 1b, arrowhead).

Figure 1

Proteasome inhibition alters nucleolar morphology. (a) WS1 cells treated with MG132 (10 μM) for 12 h or left untreated (ctrl) were stained for NPM (green) and co-stained for FBL and upstream binding factor (red) as indicated and imaged using confocal microscopy. Merged images of cells stained with Hoechst (blue), and phase-contrast images are shown. (b) TEM images of WS1 cells treated with or without MG132 (10 μM) for 12 h. Asterisks show the segregation of the nucleolar structures in MG132-treated cell intervened by an electron-dense structure (arrowhead). (c) In vivo labeling of MG132-treated (10 μM) WS1 cells with fluorouridine (green). Cells were co-stained for FBL (red) and DNA (blue). Differential interference contrast (DIC) image is provided. (d) Cells were treated with proteasome inhibitors MG132 (10 μM), ALLN (50 μM) and lactacystin (10 μM), and lysosomal inhibitors E64 (10 μg/ml) and leupeptin (100 μM) for 12 h. Cells were fixed and stained for p53 (green), FBL (red) and DNA (blue), and imaged using confocal microscopy. (e) Quantification of cells with dense nucleolar aggregates formed by the treatments in (d) (n=3 experiments, error bars denote s.d.). Scale bars, 10 μm. (f) Nucleolar integrity is required for aggregate formation. Cells were pre-treated with Act D (1 μM) for 1 h before the addition of MG132 and incubated for 10 h. Cells were stained for p53 (green) and FBL (red). Arrowheads indicate p53 staining surrounding the nucleolar remnant. Asterisks indicate nucleolar caps. Scale bars, 10 μm.

Previously, Stavreva et al. (2006) have shown that a high dose of MG132 (100 μM) inhibits pre-RNA processing. However, a much smaller, more commonly used dose of MG132 (10 μM), is sufficient to inhibit proteasome activity. Hence, we analyzed the effect of MG132 at this lower concentration, used throughout the study, on nascent rRNA synthesis using fluorouridine (FUrd) incorporation. As shown in Figure 1c, rRNA synthesis was readily detectable in the nucleoli following MG132 treatment for 12 h, and colocalized with FBL, as expected. However, the dense aggregate observed within the nucleolar marker was devoid of any rRNA synthetic activity. To further assess the effect of PI treatment on rRNA synthesis, we performed quantitative PCR using primers for rRNA precursors. Although RNA polymerase I (RNA pol I) inhibitor actinomycin D (Act D) potently suppressed rRNA transcription, MG132 did not (Supplementary Figure S1b). We conclude that while MG132 treatment causes nucleolar reorganization, the nucleoli retain RNA pol I transcriptional activity.

As MG132 may inhibit other enzyme activities in addition to the chymotrypsin-like activity of the proteasome (Kisselev and Goldberg, 2001), we tested the effect of other proteasome and lysosomal inhibitors. We treated WS1 and HEL-299 cells with MG132, ALLN (a peptide aldehyde inhibitor of the proteasome), lactacystin (a highly specific proteasome inhibitor) and two inhibitors of lysosomal proteases, E64 and leupeptin. We assessed nucleolar changes under phase-contrast and FBL staining, and included p53 as marker for a protein undergoing nucleolar translocation. Similarly to MG132 treatment, ALLN caused nucleolar translocation of p53, and this was accompanied by an altered FBL localization in a ring-shaped pattern (Figure 1d). As shown by projection of z-stacks throughout the cell by confocal imaging, FBL was observed surrounding the central p53 aggregate (Supplementary Movie S1). p53 aggregation and FBL reorganization were also detectable following lactacystin treatment (Figure 1d). The formation of nucleolar aggregates was quantified under phase contrast based on the dense structures (Figure 1e). The degree of aggregate formation was concentration-dependent for each PI (data not shown). Inhibition of lysosomal proteases did not lead to alterations in FBL or p53 distribution (Figures 1d and e). In addition, the nucleolar morphology was undisturbed by the lysosomal inhibitors under phase contrast, as compared with the PIs, which all caused formation of dense aggregates surrounded by nucleolar structures (Supplementary Figure S1c). The aggregates were detected irrespective of the fixation method used, as they were visible in both paraformaldehyde and methanol-fixed cells (data not shown). Aggregate formation was detected in several mammalian cell types, regardless of their transformation stage or tissue type of origin (Supplementary Table S1). In addition, analysis of normal and malignant cell lines in their propensity to undergo p53 and MDM2 nucleolar localization following proteasome inhibition showed that the response was irrespective of tissue type or pathological status of the cells (Supplementary Table S1). Furthermore, treatment of ex vivo cultured human prostate tissues (Kiviharju-af Hällström et al., 2007) with MG132 led to the formation of p53 aggregates in epithelial cells of the prostate gland, indicating that the aggregate formation is not only an artifact of cells cultured in monolayer, but is also observed in the context of normal tissue architecture (Supplementary Figure S1d).

As the aggregates form in the nucleoli, we next asked whether nucleolar integrity is essential for the formation of aggregates. To this end, we pretreated the cells for 1 h with Act D, which leads to the inhibition of RNA pol I activity, disintegration of the nucleolar protein content and formation of nucleolar caps (Andersen et al., 2005; Hernandez-Verdun, 2006). Act D treatment abrogated the aggregate formation by MG132, although a little p53 still accrued to the nucleolar remnants (Figure 1f, arrowheads). This finding indicates that the aggregate formation is dependent on nucleolar structures, and possibly nucleolar activity.

Proteasome inhibitors cause accumulation of nuclear proteasome targets to the nucleolar aggregate

Previous studies have shown nucleolar translocation of certain proteasome-targeted stress response proteins (Klibanov et al., 2001; Mattsson et al., 2001; Latonen et al., 2003; Kurki et al., 2004; Karni-Schmidt et al., 2007). As 20S proteasome is also localized to the nucleolus upon PI treatment (Mattsson et al., 2001) (Supplementary Figure S2), we asked whether the accumulation of proteins to the nucleoli is restricted to nuclear stress response proteins, or whether this represents a more general phenomenon for proteasome target proteins. We tested several other nuclear and cytoplasmic proteins for their response to PI treatment by immunofluorescent microscopy. All 22 nuclear proteins that produced distinct immunofluorescent signals were found to accumulate to the nucleolar aggregate in the MG132-treated cells (Figure 2 and Table 1). These, in addition to stress response proteins, included several cell cycle proteins, transcription factors and transcriptional regulators. The extent of translocation between the proteins varied, suggesting that the targeting may depend on factors inherent for each protein, possibly protein half-life. Certain proteins, like cyclin D, which is expressed both in the cytoplasm and in the nucleoplasm, was detected both in the nucleolar aggregate and in cytoplasmic aggresomes (Supplementary Figure S2a). To test whether proteasomally degraded cytoplasmic proteins also undergo translocation to nucleolar aggregates, we analyzed IkBα localization following proteasome inhibition. IkBα did not accumulate to nucleolar aggregates (Supplementary Figure S2b). Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and α-smooth muscle actin (SMA), which are proteins degraded by the lysosomal pathway, did not accumulate to the aggregates (Supplementary Figure S2b, data not shown). These results indicate that the accumulation of nuclear proteins to the nucleolar aggregates is not restricted to stress response proteins, and instead represents a common feature of a multitude of nuclear proteasome targets.

Figure 2

Inhibition of proteasome causes nucleolar aggregation of nucleoplasmic proteins. Confocal microsopy of cells mock treated or treated with MG132 for 12 h and stained for the antigens shown (red). Cells were co-stained for p53 (green) and DNA (blue). Scale bars, 10 μm.

Table 1 Proteins detected in nucleolar aggregates

Polyadenylated RNA is entrapped in nucleolar aggregates

The presence of dense nucleolar material in both phase contrast and TEM suggested that the aggregates may, in addition to proteins, contain RNA. We first detected RNA using SYTO12 green, and co-stained for p53. As shown in Figure 3a, SYTO12 positivity was detected in the aggregate and is colocalized with p53. We then used in situ hybridization using highly specific locked nucleic acid probes to detect localization of processed and precursor rRNAs, and used NPM and p53 as co-staining markers for the nucleoli and nucleolar aggregates, respectively. 28S, 18S and 5.8S rRNA and 5′ external transcribed spacer (5′ETS) 47S precursor rRNA largely overlapped with NPM in both mock- and MG132-treated cells (Figure 3b and Supplementary Figure S3). In situ hybridization for 5′ETS and 28S rRNA and co-staining for p53 showed that their staining patterns were distinct (Figures 3c and d). These results indicate that the nucleolar rRNAs do not colocalize with the aggregate formed in the nucleolus by proteasome inhibition, and that they instead colocalize with bona fide nucleolar structures and proteins.

Figure 3

Nucleolar aggregates accumulate polyA(+) RNA. WS1 cells were mock treated or treated with MG132 (10 μM) for 12 h. (a) Cells were stained for p53 (red) and RNA using SYTO 12 (green). Confocal images are shown. Arrowheads indicate nucleolar aggregates. (b) In situ hybridization of 28S rRNA (red) and co-staining for NPM (green). (c) In situ hybridization of 5′ETS rRNA precursor and co-staining of p53. Merged image shows cells co-stained for DNA (blue). (d) In situ hybridization of 28S rRNA and co-staining of p53 (green). (e) In situ hybridization of polyA(+) RNA (red) and co-staining for p53 (green). Arrowheads indicate nucleolar aggregates. (f) Cells were treated with TX-100 and RNase before fixation and in situ hybridization for polyA(+) RNA and p53 immunostaining. DNA is shown in blue. Scale bars, 10 μm.

To resolve the identity of RNA accumulated to the nucleolar aggregates, we tested for the presence of polyA(+) RNA. In situ hybridization using a locked nucleic acid (LNA) probe for polyA(+) showed that proteasome inhibition led to a positive signal within the nucleolar aggregate, and that p53, but not NPM, overlapped with the signal (Figure 3e and Supplementary Figure S3d). Furthermore, the polyA(+) signal colocalized with the prominent aggregate observed under phase contrast (Figure 3e). To test whether the aggregate components are soluble, we exposed MG132-treated cells to treatments with TritonX-100 (TX100) to extract soluble proteins, or to TX100 and RNase. TX100 could not disperse polyA(+) or p53 (Figure 3f). RNase treatment effectively obliterated the polyA(+) signal, whereas p53 staining was still detectable (Figure 3f). Taken together, these results show that polyA(+) RNA is entrapped within the nucleolar aggregate, and that both protein and RNA are in an insoluble state in this structure.

p53 and MDM2 are immobilized within the nucleolar aggregates

If the proteins accumulated to the nucleolar aggregates upon PI treatment are forming inactive precipitates, they should have decreased motility. To test this, we performed fluorescence recovery after photobleaching (FRAP) experiments on representative proteins accumulating to these structures. Fluorescent fusion proteins for p53 and MDM2 were generated using yellow-variant Venus (Nagai et al., 2002) and for NPM using cyan enhanced green fluorescent protein (Sawano and Miyawaki, 2000) (Venus-p53, MDM2-Venus and NPM-ECGPF, respectively). As a test for functionality, we confirmed the ability of Venus-p53 to cause p53-reporter transactivation and the ability of p53 and MDM2-Venus fusion proteins to form complexes (data not shown). We also verified that NPM-ECGPF underwent expected localization changes similar to the endogenous protein in response to RNA pol I inhibition and ultraviolet radiation-induced stress (data not shown). The constructs were expressed in WS1 cells, after which cells were either mock treated or treated with MG132 for 12 h. Proteins localized to different compartments of the cell were photobleached, and the fluorescence recovery in the bleached area was recorded. As expected, Venus-p53 and MDM2-Venus localized to the nucleoplasm in control cells and both proteins were observed in the nucleolus following proteasome inhibition (Figures 4a and b). Nucleoplasmic Venus-p53 and MDM2-Venus in control cells were highly mobile as indicated by fluorescence recovery within 2 min after photobleaching (Figures 4a and b). Inverse FRAP experiments of MG132-treated cells, performed by photobleaching the whole nucleus leaving a single aggregate intact, revealed that Venus-p53 and MDM2-Venus were not able to diffuse from the aggregates, indicating that the mobility of the proteins is significantly decreased in the aggregates (Figures 4a and b). In contrast, based on similar photobleaching experiments of nucleolar NPM-ECGPF, the mobility NPM-ECGPF was only slightly decreased following proteasome inhibition (Figure 4c). The latter finding is consistent with that published earlier (Stavreva et al., 2006). These results show that proteins in nucleolar aggregates, but not bona fide nucleolar proteins, are immobilized upon PI treatment, confirming that the aggregate proteins are in an inactive, immobile state.

Figure 4

Proteins in nucleolar aggregates are immobile. WS1 cells were transfected with fluorescence-tagged proteins (green) and were treated with MG132 or left untreated. FRAP analysis was performed by selecting a region of interest as indicated. Representative images are shown. Normalized intensities are shown to the right (n=3 for each analysis, error bars, s.d.). (a) Venus-p53, (b) MDM2-Venus and (c) NPM-ECGFP. Scale bars, 10 μm. A full colour version of this figure is available at the Oncogene journal online.

Aggregate formation is overcome by an increase in ubiquitin pool

Upon PI treatment, the levels of polyubiquitylated proteins increase (Kisselev and Goldberg, 2001; Navon and Ciechanover, 2009). Ubiquitin has been shown to localize to the nucleoli in cells treated with MG132 (Stavreva et al., 2006). To test whether ubiquitin colocalizes with aggregate proteins, we performed immunofluorescence analysis of ubiquitin and conjugated ubiquitin with p53 (Figure 5) and cyclin D (data not shown). As shown in Figures 5a and b, ubiquitin and conjugated ubiquitin accumulated to cytoplasmic aggresomes and within the nucleolar aggregate, the latter of which overlapped with p53. This suggested that the aggregates contain conjugated ubiquitin. Thus, we hypothesized that ubiquitylation may affect the localization of the proteins to the aggregates. We first confirmed successful extraction of nucleolar preparations in control and PI-treated cells. We treated WS1 cells with MG132 followed by preparation of total cellular extracts or nucleolar extracts and performed western analysis for p53. p53 levels were increased in both total and nucleolar extracts, whereas there was no change in the levels of FBL serving as a nucleolar marker (Figure 6a). We then expressed hemagglutinin (HA)-tagged ubiquitin in HeLa cells and isolated total and nucleolar proteins. Although proteasome inhibition markedly increased the levels of HA-ubiquitin in the cellular extracts, there was no increase of HA-ubiquitin in the nucleolar fraction (Figure 6b, middle panel). The level of nucleostemin, used as a nucleolar loading control, did not change (Figure 6b, bottom panel). However, probing with an antibody that detects conjugated ubiquitin (FK2) showed that its levels increased also in the nucleolar fraction (Figure 6b, upper panel). This suggested that, although nucleolar aggregates contain conjugated ubiquitin, ectopic ubiquitin is, surprisingly, not retained in the aggregates. We then performed immunofluorescence staining on HA-ubiquitin and p53 (Figure 6c). Ectopically expressed HA-ubiquitin was present throughout the cell, but following proteasome inhibition the localization of HA-ubiquitin was primarily cytoplasmic (Figure 6c). Concomitant with the lack of accumulation of ectopic ubiquitin in the nucleoli, we observed that the formation of aggregates was significantly decreased in cells expressing significant levels of HA-ubiquitin (Figures 6c and d). Furthermore, ectopic expression of ubiquitin decreased p53 (Figure 6c) and cyclin D (data not shown) accumulation to the aggregates. We also expressed a mutant ubiquitin with conjugation site mutations in three of seven possible lysines (K29,48,63R), including the canonical polyubiquitylation site for proteasome recognition (K48) (Haglund et al., 2003). Expression of the mutant HA-Ub-K29,48,63R reduced the aggregate formation in MG132-treated cells as effectively as HA-ubiquitin (Figure 6d). These results show that an overall increase in the ubiquitin pool can overcome aggregate formation, and that there may be extensive flexibility of potential ubiquitin conjugation sites involved. This indicates that aggregates form, at least in part, due to the lack of free ubiquitin.

Figure 5

p53 aggregates colocalize with ubiquitin and conjugated ubiquitin. WS1 cells were mock treated or treated with MG132 (10 μM) for 12 h. Cells were stained for (a) ubiquitin (green) and (b) conjugated ubiquitin using FK2 antibody detecting both mono- and polyubiquitin conjugates (green) and p53 (red). Merged images show cells co-stained for DNA (blue). Arrowheads indicate nucleolar aggregates. Scale bars, 50 μm.

Figure 6

Ubiquitin is rate limiting in the formation of nucleolar aggregates. (a) WS1 cells were treated with MG132 (MG) or left untreated (ctrl). Cellular lysates (total) or nucleolar (No) fractions were prepared, boiled in Laemmle sample buffer and equal amounts were resolved by sodium dodecyl sulfate–polyacrylamide gel electrophoresis and blotted for p53 and FBL. (b) HeLa cells were transfected with HA-ubiquitin (Ub) and were treated with MG132 (MG) or left untreated (ctrl). Cellular lysates (total) or nucleolar (No) fractions were prepared and resolved by sodium dodecyl sulfate–polyacrylamide gel electrophoresis and blotted for HA-Ub or antibody detecting conjugated ubiquitin (FK2). Loading, 15:1 nucleolar:total extract. Nucleostemin (GLN3) was used as a loading control. (c) WS1 cells were transfected with HA-Ub and were mock treated or treated with MG132 for 12 h. Cells were fixed and stained for HA-Ub and p53. Arrowheads indicate aggregates. Scale bars, 10 μm. (d) Quantification of nucleolar aggregates of WS1 cells expressing HA-Ub (wt) or HA-Ub-K29,48,63R (Ub-3mut) and treated with MG132 for 12 h (n=3, error bars, s.d.). **P<0.01; ***P<0.001. (e) Quantification of nucleolar aggregates in WS1 cells treated with lactacystin (LC), UBE1-inhibitor (UBE1-i, 10 μM) or both for 12 h (n=3, error bars, s.d.). A full colour version of this figure is available at the Oncogene journal online.

To further address the relevance of ubiquitin conjugation in the aggregate formation, we used an inhibitor against ubiquitin-like modifying activating enzyme E1 ligase (UBE). If the availability of ubiquitin determines the aggregation of the proteins, the E1 ligase inhibitor should further increase the aggregate formation. UBE inhibitor used in combination with MG132 caused such a prominent formation of aggregates that virtually all cells exhibited nucleolar alterations in phase contrast (data not shown). In order to quantify the effect of UBE inhibitor, we used lactacystin treatment of the cells, which causes aggregates in approximately 10% of the cells at the concentrations used (Figure 1e). As shown in Figure 6e, UBE inhibitor enhanced the formation of aggregates by lactacystin by threefold. These results indicate that lack of conjugation-competent ubiquitin can promote the formation of nucleolar aggregates.

To further examine the role of ubiquitin conjugation in the aggregate formation, we tested whether herpesvirus-associated ubiquitin-specific protease (HAUSP) deubiquitinase, which cleaves ubiquitin moieties on p53 and MDM2 (Li et al., 2002), affects p53 and MDM2 accumulation to the aggregates induced by PI treatment. We ectopically expressed HAUSP in HeLa cells and treated the cells with MG132. Quantification of p53-containing aggregates in the control and HAUSP-expressing cells indicated that HAUSP expression significantly decreased the accumulation of p53 to the aggregates (Figure 7a). Conversely, the absence of HAUSP should potentiate the formation of p53 aggregates following proteasome inhibition. We hence used HCT116 cells with knockdown of HAUSP (USP7) expression by homologous recombination (HAUSP−/− cells; Cummins et al., 2004). Treatment of HAUSP−/− cells with MG132 resulted in increased accumulation of p53 to the nucleolar aggregates compared with the parental HCT116 (Figure 7b and Supplementary Table S1). Furthermore, accumulation of MDM2 to the aggregates was markedly enhanced in HAUSP−/− cells as compared with the parental cell line following MG132 treatment (Figure 7c). These results indicate that HAUSP-targeted ubiquitin conjugation regulates p53 and MDM2 accrual to the nucleolar aggregates. Taken together, these results indicate a role for ubiquitin in aggregate formation, and suggest that proteins are retained in the nucleolar aggregates due to lack of sufficient pool of ubiquitin.

Figure 7

p53 and MDM2 accumulations are modulated by HAUSP deubiquitinase. (a) Quantification of nucleolar p53 in HeLa cells transfected with control vector (ctrl) or Flag-HAUSP (HAUSP), and treated with MG132 for 12 h (n=2, error bars, s.d.). *P<0.05. (b, c) HCT116 (wt) and HCT116 HAUSP−/− (HAUSP−/−) cells were treated with MG132 (10 μM) for 12 h or left untreated (control), and were co-stained for p53 (b), MDM2 (c) and DNA. Arrowheads indicate nucleolar aggregates. Scale bars, 10 μm. A full colour version of this figure is available at the Oncogene journal online.

Formation of nucleolar aggregates by proteasome inhibition is enhanced by inhibition of nuclear export

Conjugated ubiquitin, especially in the form of monoubiquitin, has been previously linked to cellular trafficking (Li et al., 2003; Chen and Sun, 2009). Given that many proteasomally targeted nuclear proteins are exported to the cytoplasm for degradation via CRM1/exportin-1-mediated nuclear export pathway, and that the CRM1 pathway is involved in the export of ribosomes and certain mRNAs (Ho et al., 2000; Gadal et al., 2001; Thomas and Kutay, 2003; Hutten and Kehlenbach, 2007), we tested whether CRM1-mediated nuclear export is involved in the aggregate formation consequent to proteasome inhibition. First, we studied the effect of proteasome inhibition on CRM1. We treated HeLa cells with MG132 either alone or in combination with CRM1 inhibitor leptomycin B (LMB, 5 ng/ml), and stained the cells for CRM1. In control cells, CRM1 was localized to the nucleoplasm and nucleoli, but the number of cells with nucleolar CRM1 expression decreased following either MG132 or LMB treatment, and their combination (Figures 8a and b). A similar effect was observed following lactacystin and combinatory lactacystin/LMB treatment of the cells (Figure 8b). These results indicate that inhibition of either proteasome or CRM1 hampers the normal nucleolar localization of CRM1.

Figure 8

Inhibition of nuclear export enhances accumulation of p53 and polyA(+) RNA to aggregates. (a) HeLa cells were treated with MG132 (10 μM), LMB (5 ng/ml) or both, or left untreated. CRM1 was detected by immunostaining. Scale bars, 10 μm. (b) HeLa cells were treated with lactacystin (LC, 10 μM) or MG132 (MG, 10 μM) either alone or in combination with LMB (5 ng/ml). Cells with the expression of nucleolar CRM1 were quantified (n=2–3, error bars, s.d.). *P<0.05; **P<0.01; ***P<0.001. (c) WS1 cells were treated with lactacystin (LC, 10 μM), LMB (5 ng/ml) or both. Cells with aggregates were quantified (n=4, error bars, s.d.). (d) WS1 cells treated as in (a) were stained for p53. Scale bars, 20 μm. (e) In situ hybridization for polyA(+) RNA (red) of WS1 cells treated as in (a). Cells were co-stained for NPM (green) and DNA (blue). Merged images are shown. Scale bars, 10 μm. (d, e) Arrowheads indicate nucleolar aggregates.

To assess whether inhibition of nuclear export could affect the formation of aggregates, we treated WS1 cells with lactacystin either alone or in combination with LMB, and assessed aggregate formation under phase contrast. Treatment of cells with LMB alone did not induce nucleolar aggregates (Figures 8c–e), indicating that inhibition of nuclear export is insufficient to induce nucleolar aggregates under conditions where nuclear proteasome activity is maintained. However, LMB significantly enhanced the formation of the aggregates by PI treatment. As nearly all cells treated with MG132 and LMB exhibited nucleolar aggregates (data not shown), the potency of LMB to increase nucleolar aggregates in combination with lactacystin was quantified by phase-contrast imaging and was 2.5-fold (Figure 8c). We further analyzed whether LMB, in combination with PI treatment, affected the accumulation of p53 and polyA(+) RNA to the aggregates. The number of cells containing p53 and polyA(+) aggregates by MG132 was substantially enhanced by LMB (Figures 8d and e). Similar results were obtained using antibody against cyclin D (data not shown). Thus, impaired nuclear export has an additive effect on the formation of aggregates by proteasome inhibition.


Proteasome inhibition leads to an increase in polyubiquitylated and misfolded proteins, induces cell death and is considered an effective strategy in cancer therapy. In this work, we studied the molecular events taking place in the nucleus of PI-treated cells. We identify here a nuclear aggregate, NoA, which forms in the nucleolus. We show that the nucleolus undergoes major reorganization following inhibition of the proteasome, as the nucleolar structures and activity surround a dense aggregate that contains proteins and polyA(+) RNA. Strikingly, we observe aggregation of over 20 proteasomally degraded nucleoplasmic proteins in this structure, several of which are key cellular regulators and relevant for cancer. We provide evidence, using p53 and MDM2 as model proteins, that the aggregate represents immobilized and insoluble proteins. We further show that these aggregates form owing to the lack of availability of conjugation-competent ubiquitin under proteasome inhibition. Free ubiquitin is required to prevent formation of nucleolar aggregates, likely through promoting nucleocytoplasmic export. These results provide mechanistic insight into the events taking place during proteasome inhibition, and identify a common mechanism for the inactivation of several cancer-related proteins during PI treatment.

Several stress-associated proteins (Hsp70, PML, p53, MDM2) have previously been described to undergo nucleolar localization following proteasome stress (Klibanov et al., 2001; Mattsson et al., 2001; Latonen et al., 2003; Kurki et al., 2004; Karni-Schmidt et al., 2007). Here we significantly extend the previous findings, and expand the number and type of proteins to include cyclins, CDKs, transcription factors, stress-inducible kinases and signaling molecules. These represent central, highly regulated proteins functioning in stress responses, cell cycle and transcription. Their common denominator is that all are nucleoplasmic proteins that can be targeted to proteasomal degradation, and most of them have been shown to be ubiquitylated. The NoAs resemble cytoplasmic aggresomes, as both contain aggregated proteasome target proteins. However, whereas the cytoplasmic aggresomes accumulate cytoplasmic proteins (Johnston et al., 1998), the NoAs seem specific for nuclear proteins. Thus, it seems that the NoAs are a nuclear counterpart of cytoplasmic aggresomes, and that both bodies can coexist in the same cells. The most striking difference between these stress-induced bodies is that RNA, detected in NoAs, is not present in cytoplasmic aggresomes. As the proteins in NoA are virtually immobile and insoluble, proteasome inhibitor treatment may result in the loss of function of many cellular activities through irrevocable retention of key proteins in the NoA.

The ubiquitin–proteasome system has several previously identified links to nucleoli and ribosome biosynthesis. Ubiquitin is present in the nucleoli (Stavreva et al., 2006; this work), and several ribosomal (r) proteins are ubiquitylated or translated as fusion proteins with ubiquitin (Finley et al., 1989; Spence et al., 2000; Matsumoto et al., 2005). Defects in ubiquitin conjugation or reduced deubiquitylation alter the nucleolar structure (Sudha et al., 1995; Endo et al., 2009). Ubiquitin conjugation is also an essential step in the quality control of rRNA (Fujii et al., 2009). Inhibition of the proteasome activity results in the accumulation of ribosomal proteins in the nucleus, suggesting that ribosomal proteins that have failed to assemble into ribosome subunits undergo proteasomal degradation (Lam et al., 2007). We find here that ubiquitin serves as a highly central modifier of NoA formation and protein mobility. NoAs contained conjugated ubiquitin as determined both by immunofluorescence and biochemical analysis of the nucleoli, suggesting the accumulation of ubiquitin-tagged proteins either as mono- or polyubiquitin conjugates or both. However, it is possible that some of the NoA proteins are not modified by ubiquitylation, but directed to these structures via protein-protein interactions. By the use of HAUSP deubiquitinase, we further showed that HAUSP-driven deubiquitylation of p53 decreases its localization to NoAs. Thus, at least in the case of p53, the accumulation to nucleolar aggregates is likely mediated by ubiquitylation.

Ubiquitin, however, seems to have a dual role in the formation of nucleolar aggregates. Ectopic expression of ubiquitin effectively rescued NoA formation, an effect that we linked to enhanced cytoplasmic export of ubiquitin. Interestingly, p53 targeted to degradation has previously been suggested to be exported via the nucleolus (Sherr and Weber, 2000; Rubbi and Milner, 2003), and the stress-induced nuclear accumulation of p53 could depend on the loss of nucleolar integrity (Rubbi and Milner, 2003). In addition, p53 trafficking is known to rely on ubiquitylation; monoubiquitylation promotes nuclear export of p53, whereas high levels of MDM2 activity promote polyubiquitylation and nuclear degradation of p53 (Li et al., 2003). In this work, ectopically expressed ubiquitin caused dissolution of p53 and cyclin D from the aggregates. These results indicate that as proteins may need conjugated ubiquitin to be targeted to the nucleolus, more ubiquitin is needed for proteins to exit. This suggests that complex, sequential ubiquitylation events are required for export of proteasome targets, at least under conditions where nuclear proteasomes are inactive. We cannot currently identify the type or types of ubiquitylation patterns involved, and whether mono- or polyubiquitylation, or both, are required. Expression of a triple-mutant ubiquitin shows that the ‘classical’ proteasome-targeting K48 link is not essential for ubiquitin to mitigate the aggregate formation, nor are the K63 and K29 links. However, four additional lysines on ubiquitin are competent of ubiquitylation, in addition to the C-terminal glycine which mediates monoubiquitylation. Numerous possibilities thus exist as to how ubiquitin could serve to relieve nucleolar aggregate formation. In addition, recent literature describes increasing complexity of possible ubiquitylation chains, and even heterologous chains involving ubiquitin and ubiquitin-like modifiers, such as small ubiquitin-related modifier (SUMO) (Ikeda and Dikic, 2008). We and Mattsson et al. (2001) have detected SUMO accumulating to the nucleoli upon proteasome inhibition, and a recent proteomic study provides evidence of SUMO conjugates accumulated in the nucleoli upon PI treatment (Matafora et al., 2009). Furthermore, SUMO-1 modification can promote cytoplasmic export of p53 in concert with ubiquitylation (Carter et al., 2007; Carter and Vousden, 2008). In the future, it will be interesting to determine whether a specific ubiquitin and/or ubiquitin-like modification signature exists targeting proteasome targets from the nucleus to the cytoplasm via the nucleolus.

Inhibition of CRM1-dependent nuclear export potentiated NoA formation. Moreover, proteasome inhibition caused exclusion of CRM1 from the nucleoli. CRM1 nucleolar localization has been noted in previous studies (Fornerod et al., 1997; Daelemans et al., 2005; Ernoult-Lange et al., 2009). CRM1 pathway is involved in the export of ribosomes and certain mRNAs (Ho et al., 2000; Gadal et al., 2001; Thomas and Kutay, 2003; Hutten and Kehlenbach, 2007), and for transport of small nucleolar RNA from Cajal bodies to the nucleolus (Boulon et al., 2004). Inhibition of CRM1 leads to 40S and 60S ribosome subunit retention in the nucleoli (Ho et al., 2000; Gadal et al., 2001; Thomas and Kutay, 2003). Based on these results, the nucleoli may serve as a platform promoting CRM1 associations with certain cargo, including ribosomal particles. However, inhibition of CRM1 alone did not induce the formation of NoAs, indicating that the inability to transport ribosome particles alone is insufficient for NoA formation. When CRM1 is inhibited, but the nuclear proteasomes remain functional, protein degradation and ubiquitin recycling can take place in the nucleus despite defective nucleocytoplasmic export. When nuclear proteasomes are inhibited, ubiquitylated proteins may be directed to the export pathway; however, exhaustion of ubiquitin prevents the later stages of export and induces formation of nucleolar aggregates. Concomitant inhibition of nuclear export with proteasome inhibition could thus promote aggregate formation by accelerating the accumulation of degradation targets.

NoA contained polyA(+) RNA, but not mature or precursor rRNA, and lacked nascent rRNA synthesis. Further studies are required to resolve the identity of the polyA(+) RNA accumulating in the nucleolar aggregates, as polyA(+) RNA may represent either nascent mRNA exported via the CRM1 route or RNA destined for degradation by polyA(+) tag. However, the finding that ubiquitin is rate limiting for the aggregate formation suggests that ubiquitin-mediated licensing is required for RNA surveillance and export also in mammalian cells, as previously described in yeast (Thomsen et al., 2008; Houseley and Tollervey, 2009).

Impairment of the proteasome function and aggresomes are linked with aging-related neurodegenerative diseases, and aggresome-like structures are formed in the nucleoplasm in relation to Huntington's disease (Klement et al., 1998; Saudou et al., 1998). Certain neurodegenerative diseases, such as polyglutamine (polyQ) repeat diseases, are characterized by the aggregation of misfolded proteins to nuclear aggregates in DNA-sparse areas resembling the nucleoli (Bennett et al., 2005). PolyQ variant of ataxin-1 forms nuclear aggregates that co-stain with ubiquitin (Bennett et al., 2005). Ataxin-1 binds RNA depending on the polyQ tract length and has been proposed to participate in RNA metabolism (Yue et al., 2001). Moreover, in Drosophila, RNA increases toxicity of the ataxin-3 polyQ repeat (Li et al., 2008). Lastly, HAUSP has been shown to bind ataxin-1 and to lose this interaction with the polyQ variant (Hong et al., 2002). Although these are presently circumstantial observations, we raise the possibility that these neuronal aggregates may represent the nucleolus-associated aggregates described here. Taken together, these findings may indicate that the underlying causes in protein aggregation diseases may be exacerbated by defects in RNA processing in addition to ubiquitin turnover.

In conclusion, we define here a novel nucleolus-dependent structure that arises consequent to severe overload of nuclear proteasome targets. We propose that these nucleolar aggregates accumulate polyA(+) RNA destined to degradation and ubiquitylated nuclear proteins targeted to nucleocytoplasmic export. The data presented here identify a remarkable number of cancer-related proteins accumulating in these PI treatment-induced stress bodies. This set of factors most likely represent only a fraction of proteins targeted to nucleolar aggregates upon proteasome inhibition, and suggest that the event is common among nuclear proteasome targets. Our results identify a close link between nuclear export, nucleoli and the proteasome, and provide mechanistic insight into how PI treatment affects several of the key regulators in cancer.

Materials and methods

Cell culture, chemicals and treatments

WS1 skin fibroblasts (CRL-1502, ATCC, Manassas, VA, USA) and HEL-299 lung fibroblasts (CCL-137, ATCC) were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum and non-essential amino acids, HeLa cells in high glucose Dulbecco's modified Eagle's medium and 10% fetal calf serum, and HCT116 cells (wild type and HAUSP−/−) in Dulbecco's modified Eagle's medium and 10% fetal calf serum. HAUSP−/− cells were a kind gift from Dr B Vogelstein (Johns Hopkins University, Baltimore, MD, USA). Cells were kept at +37 °C in a humidified atmosphere containing 5% CO2. All cell culture reagents were obtained from Gibco-BRL (Rockville, MD, USA). Chemicals used in the treatments were MG132 (Z-Leu-Leu-Leu-CHO; Biomol, Enzo Life Sciences Inc., Farmingdale, NY, USA), ALLN (MG101/calpain inhibitor I/N-Ac-Leu-Leu-norleucinal, Calbiochem, San Diego, CA, USA), lactacystin (clasto-lactacystin-b-lactone, Calbiochem), E64 (Sigma, Sigma-Aldrich, St Louis, MO, USA), leupeptin (Sigma), Act D (Sigma), LMB (Calbiochem) and UBEI-41 (Biogenova, Rockville, MD, USA). For combination treatments, cells were pre-incubated with Act D, UBEI or LMB of 1 h, after which PIs were added.

Immunofluorescence analysis

Cells grown on glass coverslips were fixed with 3.5% paraformaldehyde and permeabilized with 0.5% NP-40 in phosphate-buffered saline (PBS) at room temperature or, alternatively, were fixed with methanol and permeabilized with acetone at –20 °C. For in vivo labeling of nascent RNA synthesis, cells were treated with 1 mM 5-fluorouridine (Sigma) for 30 min, fixed with 1% paraformaldehyde for 5 min, permeabilized with 0.5% Triton-X100 for 10 min and detected using anti-bromodeoxyuridine antibody (B2531, Sigma). Primary antibodies were detected using fluorescence-conjugated secondary antibodies (Molecular Probes, Invitrogen, Carlsbad, CA, USA). DNA was stained with Hoechst 33258 (Sigma). RNA was detected with SYTO 12 (Molecular Probes). The antibodies were obtained from the following sources, Abcam (Cambridge, UK; ARF, 14PO2; FBL); Affiniti Research Products and Biomol (now Enzo Life Sciences; FK2, S20); BD Transduction Laboratories (BD Transduction Laboratories, Franklin Lakes, NJ, USA; CRM1); Bethyl Laboratories (Montgomery, TX, USA; HDMX), Epitomics (Burlingame, CA, USA; SMA, E184); Europa Bioproducts Ltd (Cambridge, UK; GAPDH, 9.B.88), DAKO (Glostrup, Denmark; Ki-67, MIB-1); Neomarkers (Thermo Scientific, Fremont, CA, USA; cyclin B1, Ab-3; cyclin D1, Ab-4; p73, Ab5); Pharmingen (Franklin Lakes, NJ, USA; Ku80, M040337; retinoblastoma, 14001, 14006); Santa Cruz Biotechnology Inc. (Santa Cruz, CA, USA; ATM, 2C1; CDK2, M2; CDK4, C-22; Chk1, G-4; cyclin A, C-19; cyclin E, C-19; IkB, C-21; MDM2, 2A10, SMP-14; p21, M-19, C-19; p27, C-19; p300, N-15; p53, DO-1, FL-393; PML, H-238; SMAD1/2/3, H-2; Sp1, 1C6; Sp3, D-20; ubiquitin, P4D1; Wee1A, H-300; YY1, H-414); and Zymed (San Francisco, CA, USA; SUMO-1, 33–2400; SUMO-3, 51–9100). For quantification, a minimum of 300 non-transfected or 100 transfected cells were analyzed per sample. The nucleolar aggregates were quantified by counting the percentage of cells with dense nucleolar aggregates visible under phase-contrast imaging. All analyses were performed from two to four independent experiments. Results were scored according to their statistical significance using Student's one-tailed t-test (*P<0.05; **P<0.01; ***P<0.001).

The fluorochromes were visualized either with Zeiss Axioplan 2 Imaging MOT (Zeiss, Jena, Germany) equipped with × 20/0.5 NA or × 400.75 NA Plan-Neofluar objectives and Chroma 31000v2, Chroma 41001, and Chroma 41004 filters (Chroma, Bellows Falls, VT, USA), or Zeiss AxioImager MOT Z1 (Jena), equipped with EC-PlanNeofluar × 40/0.75 objectives, and Zeiss filters sets 38, 45 and 49. Images were captured with Zeiss AxioCam HRm 14-bit grayscale CCD camera and AxioVision program version 4.6.

Confocal imaging was performed with Zeiss LSM510 META microscope equipped × 63/1.25 NA Plan-Neofluar objective, diode, argon and HeNe lasers. Emissions were detected with the following filter settings: BP 420–480 for Hoechst, BP 505–530 for Alexa 488 and LP 585 for Alexa 594. HFT405/488/543 was used as dichroic beam splitter and NFT545 as emission splitter.


Cells were harvested by pelleting and fixed by 2.5% glutaraldehyde. Samples were post-fixed by 1% osmium tetroxide for 1 h at room temperature, dehydrated in graded ethanol and embedded in epoxy resin LX-112. Serial sections were cut at 60 nm and stained with uranyl acetate and lead citrate using Leica EM STAIN automatic stainer (Leica Microsystems GmbH, Wetzlar, Germany). The sections were examined with a JEM-1400 (Jeol) TEM at 80 kV at × 5000 magnification. Images were captured using Olympus-SIS Morada digital camera (Olympus, Münster, Germany).

Transient transfections

Transfections to WS1 cells were performed using electroporation essentially as described earlier (Latonen et al., 2003). Cells were trypsinized, suspended to Opti-MEM (Gibco BRL), DNA was added and the mixture was electropulsed (300V, 800 μF) with Gene Pulser II (BioRad, Hercules, CA, USA). HeLa cells were transfected using JetPei transfection reagent (Polyplus-transfection Inc., New York, NY, USA) according to the manufacturer's protocol.

NPM-ECGFP fusion protein was generated by excision of NPM1 cDNA from B23-GFP (a kind gift from Dr M Olson, University of Mississippi Medical Center, MS, USA; Dundr et al., 2000) and ligation to ECGFP-pRSETb (a kind gift from Dr A Miyawaki, Brain Science Institute, RIKEN, Saitama, Japan; Sawano and Miyawaki, 2000). The construct was further subcloned to pCDNA3.1+ (Invitrogen) to yield NPM-ECGFP. Venus-p53 was generated by ligating p53 cDNA to C-terminus of Venus/pCS2 (a kind gift from Dr A Miyawaki; Nagai et al., 2002). MDM2-Venus was generated by ligating MDM2 insert to N-terminus of Venus/pCS2 to generate MDM2-Venus in-frame fusion. Expression vectors for ubiquitin (HA-Ub-wt/pcDNA3 and HA-Ub-K29, 48, 63R/pcDNA3) were a kind gift from Dr I Dikic (Goethe University, Frankfurt, Germany; Haglund et al., 2003), and pCIneo-HAUSP-Flag (USP7) vector was kindly provided by Dr B Vogelstein (Johns Hopkins University, Baltimore, MD, USA; Cummins et al., 2004).


Before imaging, normal growth medium was replaced with Dulbecco's modified Eagle's medium without phenol red (Gibco-BRL). In MG132-treated cells, MG132 was present throughout imaging. FRAP was performed using LSM 510 META confocal laser scanning microscope (Zeiss) equipped with a heating stage. Argon laser line (488 nm) was set at 100% during the bleaching and at 2% during imaging with 50% output using Plan-Neofluar × 63/1.25 NA oil-immersion objective. The region of interest was bleached after three pre-bleach scans with 30 iterations, and 97 post-bleach images were captured. For each fusion protein, at least three independent experiments were performed. Fluorescence intensities were measured using the LSM 510 Physiology Software. Data from three representative cells for each fusion protein and from each condition were used for the FRAP analysis. Recovery curves from untreated, nucleoplasmic protein intensities were corrected to total cell intensity and to scan controls.

In situ hybridization

Cells grown on coverslips were fixed in 4% paraformaldehyde with 10% acetic acid in diethylpyrocarbonate-treated water for 20 min. The cells were washed three times in PBS and immersed in cold 70% ethanol overnight. Cells were then rehydrated in PBS for 10 min and pre-hybridized in 40% formamide (Sigma) in 2 × SS (sodium chloride–sodium phosphate–ethylenediaminetetraacetic acid buffer) for 20 min. Locked nucleic acid probes were diluted in hybridization buffer (50% formamide, 5 × SSC, 250 μg/ml Escherichia coli tRNA (Roche, Basel, Switzerland), 500 μg/ml salmon sperm DNA (Invitrogen), 2% Roche blocking reagent (Roche), 0.02% Tween-20, 0.05% CHAPS (Sigma) in diethylpyrocarbonate-treated water) and incubated at 37 °C for 5 h. Coverslips were washed with 5 × SSC for 15 min at 37 °C, twice for 35 min each at 37 °C in 0.2 × SSC and then once in PBS for 15 min atroom temperature. Coverslips were blocked in 4% sheep serum and 3% bovine serum albumin in PBS for 1 h and incubated overnight in anti-digoxigenin-AP (Roche) solution at 4 °C followed by washing seven times in PBST (0.01% Tween in PBS) for 5 min and once in PBS. For detection of AP, Fast Red tablets (Roche) were dissolved in 0.1 M Tris–HCl (pH 8.2) and the color reaction was carried out in dark for up to 5 h. Finally, the coverslips were stained for DNA using Hoechst 33258 and mounted in VECTASHIELD mounting medium (Vector Laboratories, Burlingame, CA, USA). The following LNA probe sequences were used: 28S, IndexTermCCTTAGAGCCAATCCTTATCCC; 18S, IndexTermCTGATCGTCTTCGAACCTCCGA; 5′ETS, IndexTermGACGTCACCACATCGATCGAAG; 5.8S, IndexTermTTCTTCATCGACGCACGAGCCG; and polyT(25)Vn, IndexTermTTTTTTTTTTTTTTTTTTTTTTTTTVN. All probes were from Exiqon (Vedbaek, Denmark). In indicated experiments, the cells were incubated with 0.05% TX-100 and 0.05% Tween-20 in PBS for 10 min, followed by RNase treatment (1 mg/ml, Roche) for 30 min at room temperature before fixation and hybridization as above.


Cells were lysed in RIPA buffer (50 mM Tris–HCl (pH 7.5), 150 mM NaCl, 1% NP-40, 0.1% sodium dodecyl sulfate, 1% sodium deoxycholate containing 1 mM Na3VO4, 1 mM phenylmethanesulfonyl fluoride, 1 mM dithiothreitol, 20 mM N-ethylmaleimide and 10 mg/ml of each E64, leupeptin and SBTI) to obtain total cellular lysates. Nucleolar fractionation was performed as described previously (Andersen et al., 2005). N-ethylmaleimide (20 mM) was added to extraction buffers in experiments assessing ubiquitin conjugation. Proteins were separated by 5% sodium dodecyl sulfate–polyacrylamide gel electrophoresis, and immunoblotting was performed as described previously (Latonen et al., 2003). The following antibodies were used: nucleostemin (H-270; Santa Cruz Biotechnology) and HA.11 (16B12; Covance, Princeton, NJ, USA). Horse radish peroxidase-conjugated secondary antibodies were from DAKO.


Act D:

actinomycin D


ataxia telangiectasia mutated


cyclin-dependent kinase


chromosome region maintenance 1 protein homolog


endoplastic reticulum


external transcribed spacer








herpes virus-associated ubiquitin-specific protease


leptomycin B


locked nucleic acid


nucleolar aggregate




proteasome inhibitor



RNA pol I:

RNA polymerase I


polyadenylated RNA






Triton X-100


upstream binding factor


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We thank Drs M Olson, A Miyawaki, I Dikic, B Vogelstein and A Salminen for providing reagents and cell lines. M Salo and H Liu are thanked for excellent technical assistance. Members of Laiho lab at Helsinki and Hopkins, O Matilainen and C Holmberg are thanked for helpful discussions. University of Helsinki Molecular Imaging Unit is thanked for assistance in image acquisition. University of Helsinki Advanced Imaging Unit is thanked for assistance in TEM imaging and sample preparation. This work was supported by Academy of Finland (ML Grant No. 129699, LL Grant No. 108828), Biocentrum Helsinki and Helsinki Biomedical Graduate School (HMM).

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Correspondence to M Laiho.

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Latonen, L., Moore, H., Bai, B. et al. Proteasome inhibitors induce nucleolar aggregation of proteasome target proteins and polyadenylated RNA by altering ubiquitin availability. Oncogene 30, 790–805 (2011).

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  • proteasome inhibitor
  • proteasome
  • ubiquitin
  • polyadenylated RNA
  • nuclear export
  • aggresome

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