ABSTRACT
Although an F-box protein, Mdm30, is found to regulate ubiquitylation of the Sub2 component of TREX (transcription-export) complex for proteasomal degradation in stimulation of mRNA export, it remains unknown whether such ubiquitin-proteasome system (UPS) regulation of Sub2 occurs cotranscriptionally via its interaction with Mdm30. Further, it is unclear whether impaired UPS regulation of Sub2 in the absence of Mdm30 alters mRNA export via splicing defects of export factors and/or mitochondrial dynamics/function, since Sub2 controls mRNA splicing and Mdm30 regulates mitochondrial aggregation. Here, we show that Mdm30 interacts with Sub2, and temporary shutdown of Mdm30 enhances Sub2’s abundance and impairs mRNA export. Likewise, Sub2’s abundance is increased following transcriptional inhibition. These results support Mdm30’s direct role in regulation of Sub2’s cellular abundance in a transcription-dependent manner. Consistently, the chromatin-bound Sub2 level is increased in the absence of Mdm30. Further, we find that Mdm30 does not facilitate splicing of export factors. Moreover, Mdm30 does not have a dramatic effect on mitochondrial respiration/function, and mRNA export occurs in the absence of Fzo1, which is required for mitochondrial dynamics/respiration. Collective results reveal that Mdm30 interacts with Sub2 for proteasomal degradation in a transcription-dependent manner to promote mRNA export independently of splicing or mitochondrial function, thus advancing our understanding of mRNA export.
INTRODUCTION
Nuclear mRNA export, a key regulatory step in the expression of RNA polymerase II genes in eukaryotes, occurs through nuclear pore complex (NPC), which is an assembly of nucleoporins integrated within the nuclear envelope (1–4). Nucleoporins interact with export receptors (which carry mRNAs) to mediate mRNA export (1–4). While a number of export receptors have been identified, an essential receptor, Mex67, in yeast and its human homologue TAP (Tip-associated protein) have been well studied (1–4). These receptors have low affinity toward mRNAs, but adaptor proteins enhance their interactions with mRNAs. Yra1 has been identified as an adaptor protein in yeast. Yra1 and its human homologue, ALY, bridge mRNAs with export receptors Mex67 and TAP, respectively (1–4). Yra1 is recruited to mRNA via Sub2 (an ATPase/RNA helicase) and a tetrameric THO complex (which consists of Hpr1, Thp2, Tho2. and Mft1 and associates with Tex1) (5). Sub2 associates with THO via Hpr1 (6), and THO is cotranscriptionally recruited to the active genes (7–9). Thus, Yra1 is loaded onto nascent mRNAs in a transcription-dependent manner. Following Yra1 recruitment to nascent mRNA, Sub2 is subsequently released to allow the interaction of Yra1 with Mex67 (6) which contacts nucleoporins and transports mRNA across NPC. In the cytoplasmic side of NPC, Mex67 is dissociated from mRNA with the help of Dbp5, InsP6, and Gle1 (3, 10). Subsequently, Mex67 is recycled back to the nucleus for mRNA export. Collectively, these studies support the idea that nuclear mRNA export is coupled to transcription. Yra1, Sub2, and the THO complex are the early players in this process and form a TREX (transcription-export) complex that is conserved from yeast to humans (5, 8, 10, 11).
The TREX components Hpr1 and Yra1 have been previously shown to be regulated by the HECT (homologous to E6-associated protein [E6AP] carboxy terminus) domain-containing ligases Rsp5 and Tom1, respectively (12–17). While ubiquitylated Hpr1 undergoes proteasomal degradation (14), ubiquitylation of Yra1 does not signal its proteasomal degradation (17) but rather promotes its dissociation from completely assembled mRNA-ribonucleoprotein complex prior to mRNA export (17). These studies support the differential roles of UPS (ubiquitin-proteasome system) in regulation of TREX and hence mRNA export. While the roles of HECT domain-containing ligases in regulation of TREX or mRNA export are established, it was not known whether RING (really interesting new gene) or F-box-containing ubiquitin ligase is involved in regulation of TREX and consequently mRNA export. Importantly, we have demonstrated for the first time that an F-box protein, Mdm30, stimulates nuclear mRNA export (18). The F-box protein functions as a component of the SCF (Skp1–cullin–F-box protein) ubiquitin ligase complex which facilitates the interaction between substrate and ubiquitin-conjugating enzyme and results in the covalent transfer of ubiquitin onto the substrate (19–23). While cullin and RING domain proteins form the catalytic core of the SCF ligase complex, substrate specificity is mediated by the F-box protein of this complex (19–25). Like other F-box proteins, Mdm30 copurifies with SCF ligase complex and interacts with the Skp1 component of SCF (26–29). Interestingly, we find that the Sub2 component of TREX is ubiquitylated in the presence of Mdm30 and degraded by the 26S proteasome complex to promote mRNA export (30). These results demonstrate for the first time the role of SCF ubiquitin ligase in mRNA export via Sub2 ubiquitylation and proteasomal degradation (30). Likewise, Sub2’s human homologue, UAP56, is found to be ubiquitylated and degraded by the proteasome (31). Further, increased abundance of UAP56 in Xenopus oocytes has been shown to impair mRNA export (32), consistent with our results (30) and a previous study (6) in yeast. Therefore, Sub2 ubiquitylation and proteasomal degradation in promoting mRNA export appear to be conserved among eukaryotes.
Although Mdm30 is found to be required for Sub2 ubiquitylation and proteasomal degradation (30), it is yet unknown whether Mdm30 is directly involved in such processes via its interaction with Sub2 to promote mRNA export. Further, Sub2 is associated with the coding sequences of active genes, similarly to other TREX components (30, 33–37). Such association of Sub2 promotes transcription as well as mRNA export (5, 7–9, 11, 33–36). However, it is not clear whether transcriptionally engaged Sub2 is targeted for ubiquitylation and proteasomal degradation or whether it is degraded off the gene in a transcription-independent manner. Moreover, Mdm30 is involved in degradation of Fzo1, which is associated with mitochondrial fusion/dynamics and hence respiration/mitochondrial function (26, 38–44). Thus, it is not clear whether the defect in mRNA export in the absence of Mdm30 could result from altered mitochondrial dynamics/function. Further, it is also possible that increased abundance of Sub2 in the absence of Mdm30 impairs splicing of mRNA export factors such as Yra1 and Sus1, leading to the defect in mRNA export, since Sub2 is also involved in regulation of mRNA splicing (45, 46). To address these questions in order to gain more insights into the regulation of mRNA export by an F-box protein, we carried out experiments which reveal that Mdm30 interacts with Sub2 to maintain its abundance in a transcription-dependent manner for efficient mRNA export independently of mitochondrial function and splicing of YRA1 and SUS1, as presented below.
RESULTS
Mdm30 interacts with Sub2.Although Mdm30 is involved in ubiquitylation and proteasomal degradation of Sub2 in stimulation of mRNA export (18, 30), it is unclear whether such a function of Mdm30 occurs directly via its interaction with Sub2. To address this, we analyzed here the interaction between Mdm30 and Sub2 in living cells following formaldehyde-mediated in vivo cross-linking-based coimmunoprecipitation (co-IP) assay, as done previously (47, 48). In this direction, we generated a yeast strain expressing Myc epitope-tagged Mdm30 and hemagglutinin (HA)-tagged Sub2 or Mex67 from their chromosomal loci and then performed an in vivo cross-linking-based co-IP assay as schematically shown in Fig. 1A. We found that an anti-Myc antibody to Myc-tagged Mdm30 pulled down Sub2 (Fig. 1B). As a control, we show that Mdm30 does not pull down Mex67 (Fig. 1C), which is not known to interact with Mdm30. Further, an anti-Myc antibody did not immunoprecipitate Sub2 when Myc epitope tag was not added to Mdm30 (Fig. 1D and E). These results support the idea that Mdm30 interacts with Sub2 in vivo. Next, we analyzed the interaction of Mdm30 with Sub2 using immunopurified Sub2 and Mdm30 by co-IP assay, as done previously (47, 49) and schematically shown in Fig. 1F. Our co-IP analysis reveals that Mdm30 interacts with Sub2 (Fig. 1G). We found that an anti-Myc antibody to Myc-tagged Mdm30 pulled down HA-tagged Sub2 (Fig. 1G). As a control, we show that Mdm30 is not coimmunoprecipitated with Mex67 (Fig. 1G). These results support the idea that Mdm30 interacts with Sub2 in vitro. Such interaction is physiologically relevant as our in vivo cross-linking-based co-IP assay demonstrated the interaction between Mdm30 and Sub2 in living cells (Fig. 1A to E).
Mdm30 interacts with Sub2. (A) Experimental strategy for in vivo formaldehyde cross-linking-based co-IP assay. WCE, whole-cell extract; WB, Western blot; IP, immunoprecipitation; HRP, horseradish peroxidase. (B and C) Mdm30 interacts with Sub2 (B) but not Mex67 (C) in vivo. (D) A control co-IP experiment using the yeast strain that does not bear Myc epitope-tagged Mdm30. (E) Western blot analysis of WCE of yeast strain with or without Myc epitope tag on Mdm30, using an anti-Myc antibody. (F) Experimental strategy for analysis of protein-protein interactions in vitro. (G) Mdm30 interacts with Sub2 but not Mex67 in vitro.
Mdm30 interacts with Sub2 to regulate its abundance in orchestrating mRNA export.Although Mdm30 is known to be involved in Sub2 ubiquitylation and proteasomal degradation (30), the above-identified Mdm30-Sub2 interaction implicates Mdm30’s direct role in UPS regulation of Sub2. Consistently, we found an increased steady-state level of Sub2 in the absence of Mdm30 (30). In such an analysis, Sub2 synthesis and degradation occurred simultaneously. However, Sub2’s stability in the absence of its synthesis has not been analyzed in the Δmdm30 strain in comparison to the wild-type equivalent. To address this, we analyzed Sub2’s stability in the Δmdm30 strain and its isogenic wild-type equivalent following inhibition of its synthesis. For this purpose, we blocked translation or protein synthesis by cycloheximide in the wild-type and Δmdm30 strains and then analyzed Sub2’s level at different time points. Our cycloheximide chase assay (Fig. 2A) revealed increased stability of Sub2 in the Δmdm30 strain in comparison to the wild-type equivalent (Fig. 2B to D). Thus, null mutation of Mdm30 increased the steady-state level of Sub2 or decreased its degradation.
Analysis of Sub2’s stability in the wild-type and Δmdm30 strains following translational inhibition by cycloheximide. YPD, yeast extract-peptone plus 2% dextrose. (A) Schematic diagram for the cycloheximide chase assay. (B) Sub2’s stability is increased in the absence of Mdm30 following cycloheximide treatment. Sub2’s levels in the wild-type and Δmdm30 strains at 0 h were set to 100, and its levels at other time points were normalized with respect to 100. (C and D) Western blot analysis of Sub2 (C) and actin (D) following cycloheximide treatment in wild-type and Δmdm30 strains.
Although the null mutation of Mdm30 had increased the abundance of Sub2 (Fig. 2) (30) and impaired mRNA export (30), it is quite possible that the complete loss of Mdm30 for the long term has indirect/secondary/compounding effects. To address this issue, we shut down the expression of Mdm30 temporarily and then analyzed its effect on Sub2’s abundance and mRNA export. In this direction, we replaced the endogenous promoter of MDM30 by a galactose-inducible GAL1 promoter (which is repressed in dextrose-containing growth medium) (50, 51) and then temporarily shut down the expression of Mdm30 in dextrose-containing growth medium for 2 h (Fig. 3A). Subsequently, we analyzed the abundance of Sub2 and mRNA export. We found that short-term loss of Mdm30 (Fig. 3A) increased the level of Sub2 (Fig. 3A and B), as observed in the Δmdm30 strain (Fig. 2B and C) (30). As a control, we show that the actin level was not similarly increased (Fig. 3A and B). Further, such temporary shutdown of Mdm30 expression decreased the ADH1 mRNA level in the cytoplasm (Fig. 3C and D). However, the total ADH1 mRNA level was not altered (Fig. 3C and D). The 18S rRNA level was monitored as a loading control, and its level was not altered following temporary shutdown of Mdm30 expression (Fig. 3C and D). Further, as expected in the cytoplasmic fractionation, we did not observe nuclear DNA (i.e., GAL1) in the cytoplasmic fraction (Fig. 3E), consistent with our previous studies (30). These results support the idea that temporary loss of Mdm30 increases Sub2’s abundance and impairs mRNA export (Fig. 3A to D), in agreement with our results for defects in export of ADH1 and other mRNAs following the complete loss of Mdm30 (30).
Short-term loss of Mdm30 enhances Sub2’s abundance and decreases mRNA export. (A) Western blot analysis of Mdm30, Sub2, and actin levels following transcriptional shutdown of MDM30 under the control of the GAL1 promoter (PGAL1-MDM30-HA) in dextrose-containing growth medium for 2 h. Although both Mdm30 and Sub2 bear the HA epitope tag, they were identified in the Western blot analysis based on their different molecular weights. (B) Fold increase of Sub2’s stability relative to actin following transcriptional shutdown of MDM30 for 2 h. (C) RT-PCR analysis of total and cytoplasmic ADH1 mRNA and 18S rRNA levels following 2-h transcriptional shutdown of MDM30 in dextrose-containing growth medium. (D) Results from panel C are plotted in the form of a histogram. RNA level at 0 h was set to 100. RNA level at the 2-h time point was normalized with respect to 100. (E) Analysis of quality control of cytoplasmic fractionation by determining the absence of the nuclear GAL1 gene in the cytoplasmic fraction. A primer pair targeted to the GAL1 core promoter was used in the PCR analysis. Set1, Set2, and Set3 represent different biological sets of cytoplasmic fractionations.
Sub2’s abundance is regulated in a transcription-dependent manner.Although Sub2 is targeted for ubiquitylation and proteasomal degradation by Mdm30 (30), it is not clear whether Sub2’s degradation occurs cotranscriptionally at the gene or off the gene in a transcription-independent manner. Sub2 is cotranscriptionally associated with the coding sequences as are other TREX components (30, 33–37) and promotes transcription and mRNA splicing and export (5, 7, 9, 11, 33–36, 45, 46). We find that increased abundance of Sub2 in the absence of Mdm30 does not impair transcription or splicing but rather decreases mRNA export (30). Further, the Sub2 level at the coding sequence of the active gene is increased in the absence of Mdm30 (30). These results suggest that transcriptionally engaged Sub2 might be targeted for degradation. If so, Sub2’s abundance would be increased following inhibition of transcriptional elongation, as Sub2 is associated with the coding sequence in a transcription-dependent manner or an elongating RNA polymerase II-dependent manner (30, 33–37). To test this, we analyzed the level of Sub2 following pharmacological inhibition of transcriptional elongation by α-amanitin. α-Amanitin binds strongly with RNA polymerase II, blocks its translocation, and hence inhibits transcriptional elongation (52–55). We find that the abundance of Sub2 is increased following α-amanitin treatment (Fig. 4A and B). As a control, we show that actin level is not similarly increased (Fig. 4A and B). These results support the idea that Sub2’s abundance is regulated by active transcription. However, we did not observe a manyfold increase in Sub2’s level following α-amanitin treatment (Fig. 4A and B), unlike the mdm30 null mutant (30). This could be due to incomplete transcriptional blockage by α-amanitin. To test this, we analyzed transcriptional defects following α-amanitin treatment. In this direction, we analyzed the association of RNA polymerase II with the coding sequence of ADH1 and its mRNA level following α-amanitin treatment. The largest subunit of RNA polymerase II, Rpb1, is essential to maintain the structural and functional integrities of RNA polymerase II and thus served as a representative component for our chromatin immunoprecipitation (ChIP) analysis of RNA polymerase II association with ADH1. Our ChIP analysis revealed that the association of RNA polymerase II with the ADH1 coding sequence was decreased by ∼2-fold following α-amanitin treatment (Fig. 4C and D). Consistently, the ADH1 mRNA level was also decreased by ∼2-fold (Fig. 4E). Thus, α-amanitin treatment blocks transcription by ∼2-fold, and such transcriptional blockage enhances Sub2 level relative to actin by ∼2-fold. Therefore, Sub2’s abundance is regulated in a transcription-dependent manner.
Sub2’s abundance is increased following transcriptional inhibition by α-amanitin. (A) Western blot analysis of Sub2 and actin levels following transcriptional inhibition by α-amanitin. (B) Results from panel A are plotted in the form of a histogram. Sub2 and actin levels without α-amanitin treatment were set to 100, and their levels following α-amanitin treatment were normalized with respect to 100. Statistical analysis was performed using four sets of biologically independent experiments. (C) ChIP analysis for the association of the largest subunit (Rpb1) of RNA polymerase II with the ADH1 ORF following α-amanitin treatment. The ratio of immunoprecipitate over the input is indicated below each band. IP, immunoprecipitate. (D) Quantitation of the results from panel C. (E) RT-PCR analysis of ADH1 mRNA and 18S rRNA following α-amanitin treatment.
To further support the above conclusion genetically, we analyzed Sub2’s level following deletion of SAN1. San1 promotes transcriptional elongation (49), and thus, deletion of SAN1 would increase the abundance of Sub2, if Sub2’s abundance is regulated in a transcription-dependent manner (as observed above following pharmacological inhibition of transcription) (Fig. 4). We found increased abundance of Sub2 in the absence of San1 (Fig. 5A). However, the actin level (loading control) was not similarly increased in the absence of San1 (Fig. 5A). Thus, inhibition of transcriptional elongation in the absence of San1 (49) enhances Sub2’s abundance. However, it is possible that the impairment of RNA polymerase II association with the active coding sequence (and hence transcriptional elongation) in the null mutation of SAN1 in our recent studies (49) is due to indirect/secondary/compounding effects, leading to increased abundance of Sub2. To address this issue, we shut down the expression of SAN1 temporarily and then analyzed its effect on RNA polymerase II association with active coding sequence and transcription. In this direction, we replaced the endogenous promoter of SAN1 with the GAL1 promoter and then temporarily shut down the expression of San1 in dextrose-containing growth medium (Fig. 5B). Subsequently, we analyzed the association of RNA polymerase II with the coding sequence of ADH1 and its transcription. We found that short-term loss of San1 (Fig. 5B) decreased the association of RNA polymerase II with the ADH1 coding sequence (Fig. 5C and D). Consistently, transcription of ADH1 is also impaired following temporary shutdown of San1 (Fig. 5E). Thus, like the results in Δsan1 (49), temporary shutdown of San1 expression impairs transcription and RNA polymerase II association with active gene. Further, we find that temporary loss of San1 increases the abundance of the Spt16 subunit (Fig. 5B) of FACT (facilitates chromatin transcription that regulates transcription by facilitating chromatin disassembly at the promoter and reassembly at the coding sequence) (56–62) and its association with the coding sequence (Fig. 5F and G), consistent with the results in the null mutation of SAN1 (49). Such upregulation of Spt16 occurs via the loss of San1’s function in causing Spt16 ubiquitylation and proteasomal degradation and is associated with transcriptional impairment (49). Thus, both short-term and complete loss of San1 impair transcription, which increases the abundance of Sub2. These results were further corroborated by the fact that temporary shutdown of the Paf1 (RNA polymerase II-associated factor 1) component of Paf1C (Paf1 Complex, which promotes transcription elongation) enhances Sub2’s abundance (Fig. 5H to J). Together, our results demonstrate that pharmacological and genetic inhibitions of transcriptional elongation (which promotes the association of Sub2 with the coding sequence) (30, 33–37) enhances Sub2’s abundance. These results indicate that transcriptionally engaged Sub2 is targeted for proteasomal degradation, and hence Sub2’s abundance is increased following transcriptional inhibition.
Sub2’s abundance is increased following genetic inhibition of transcription. (A) Western blot analysis of Sub2 and actin levels in the wild-type and Δsan1 strains. (B) Western blot analysis of San1, Spt16, and actin levels following transcriptional shutdown of SAN1 under the control of the GAL1 promoter (PGAL1-SAN1-HA) in dextrose-containing growth medium. (C) ChIP analysis of RNA polymerase II association with the ADH1 ORF or coding sequence. The ratio of immunoprecipitate over the input is indicated below each band. (D) Quantitation of the results from panel C. (E) RT-PCR analysis of ADH1 mRNA and 18S rRNA levels following transcriptional shutdown of SAN1 in dextrose-containing growth medium. (F) ChIP analysis of Spt16 association with the ADH1 ORF following transcriptional shutdown of SAN1. The ratio of immunoprecipitate over the input is indicated below each band. (G) Quantitation of the results from panel F. (H and I) Western blot analysis of Sub2 and actin levels following transcriptional shutdown of PAF1 under the control of the GAL1 promoter (PSUB2-SUB2-HA, PGAL1-PAF1-HA) in dextrose-containing growth medium for 2 and 4 h. Sub2-HA is expressed under the control of its own promoter (PSUB2-SUB2-HA). Although both Paf1 and Sub2 bear the HA epitope tag and their molecular weights are quite close, the Paf1 level decreases dramatically in dextrose as presented on the right of panel H, and the Sub2 level under the control of its own promoter (PSUB2-SUB2-HA) does not dramatically increase in dextrose (I). Thus, the increase in the Western blot signal of Sub2 under the control of its own promoter (PSUB2-SUB2-HA) following temporary shutdown of Paf1 expression under the control of the GAL1 promoter (PGAL1-PAF1-HA) at 4 h in dextrose-containing growth medium is primarily due to the increased level of Sub2 in the absence of Paf1 (left of panel H). Further, since both Sub2 and Paf1 are present at 0 h (see Paf1 and Sub2 levels at 0 h in panel I and the right of panel H) and only Sub2 is present at 4 h in the left panel (also see Paf1 signal at 4 h in the right of panel H), the absolute increase in Sub2 level at 4 h following underexpression of Paf1 (or in the absence of Paf1) would be more than what is seen in this panel. (J) Quantitation of the results from panel H.
To further complement the above observations, we analyzed the level of Sub2 in the chromatin-bound fraction in the wild-type and Δmdm30 strains, following the experimental strategy as schematically shown in Fig. 6A. We find a relatively smaller amount of Sub2 in the chromatin-bound fraction (Fig. 6B), similar to the Rpb1 subunit of RNA polymerase II and Mdm30 (Fig. 6C). The quality of such fractionation was monitored by analyzing histone H3 and tubulin. As expected, histone H3 and tubulin were found in the chromatin-bound and unbound fractions, respectively (Fig. 6B). The Sub2 level in the chromatin-bound fraction was significantly increased in the absence of Mdm30 (Fig. 6D to F). Histone H3 levels in the chromatin-bound fractions of the wild-type and Δmdm30 strains served as loading controls, and thus, the increased level of Sub2 in the chromatin-bound fraction in the Δmdm30 strain in comparison to the wild-type equivalent was normalized with respect to histone H3 (Fig. 6F). Further, we analyzed chromatin-bound Sub2 level in the wild-type and Δmdm30 strains following formaldehyde-based in vivo cross-linking. We found predominantly chromatin-bound Sub2 following cross-linking (Fig. 6G), similar to the Rpb1 subunit of RNA polymerase II and Mdm30 (Fig. 6H), and its abundance was increased in the absence of Mdm30 (Fig. 6I and J). These results indicate that chromatin-bound Sub2 is targeted for degradation by Mdm30, and hence, the Sub2 level in the chromatin-bound fraction is increased in the absence of Mdm30 (Fig. 6D to F, I, and J). Consistently, Sub2 level is increased in response to pharmacological and genetic inhibitions of transcription (Fig. 4 and 5). Further, ChIP analysis revealed an increased level of Sub2 at the active coding sequence in the absence of Mdm30 (30). Collectively, our results support the idea that transcriptionally engaged Sub2 is degraded by Mdm30.
Chromatin-bound Sub2 level is increased in the absence of Mdm30. (A) Schematic diagram outlining chromatin fractionation. (B) Western blot analysis of Sub2, tubulin, and histone H3 in the chromatin-bound and unbound fractions. (C) Western blot analysis of Rpb1 and Mdm30 in the chromatin-bound and unbound fractions. (D) Western blot analysis of chromatin-bound Sub2 and histone H3 in the wild-type and Δmdm30 strains. (E) Quantitation of the results from panel D. (F) Quantitation of alteration of Sub2 levels relative to histone H3 of panel D. (G) Western blot analysis of Sub2 and histone H3 in the chromatin-bound and unbound fractions following formaldehyde-based in vivo cross-linking. (H) Western blot analysis of Rpb1 and Mdm30 in the chromatin-bound and unbound fractions following cross-linking. (I) Western blot analysis of chromatin-bound Sub2 and histone H3 in the wild-type and Δmdm30 strains following formaldehyde-based in vivo cross-linking. (J) Quantitation of the results from panel I.
Mdm30 regulates mRNA export independently of mitochondrial respiration/function.Although our results reveal that Mdm30 interacts with Sub2 to regulate its abundance in a transcription-dependent manner to control mRNA export, it is possible that Mdm30 can also indirectly contribute to the regulation of mRNA export by altering mitochondrial function, as Mdm30 is involved in degradation of Fzo1, which is required for mitochondrial fusion/function (26, 38, 44). To address this, we analyzed mitochondrial function in the wild-type and Δmdm30 strains. The Δfzo1 strain was used as a control. If mitochondrial function is impaired, yeast cells will be unable to metabolize via the tricarboxylic acid cycle and oxidative phosphorylation, leading to respiration incompetency. Therefore, we analyzed the growth of the yeast cells in medium containing dextrose or acetate as a sole carbon source. Yeast cells growing in acetate-containing medium would require an active tricarboxylic acid cycle and oxidative phosphorylation (or respiration) for their survival. If mutant cells do not grow in acetate-containing medium, the corresponding gene(s) would be attributed to promote mitochondrial respiration/function. We found that both the wild-type and Δmdm30 strains grew in acetate-containing medium with modest/moderate defects in the Δmdm30 strain (Fig. 7A). These results support the idea that Mdm30 modestly/moderately promotes mitochondrial respiration/function. However, the Δfzo1 strain did not grow in acetate-containing medium (Fig. 7B). Thus, the null mutation of FZO1 impairs respiration, consistent with the fact that Fzo1 is essential for mitochondrial fusion (26, 38, 44) that is required for maintenance of mitochondrial DNA, formation of intracellular mitochondrial networks to promote dissipation of energy in the cell, and complementation of mitochondrial gene products to defend against cellular aging (63–66). Even though Mdm30 regulates the steady-state level of Fzo1 (Fig. 7C) (38, 42, 43), it does not greatly control mitochondrial respiration/function. This could be due to the fact that mitochondria may not be dramatically fragmented in the Δmdm30 strain but rather in the absence of Fzo1. Indeed, previous studies found that mitochondria are mostly fragmented in the Δfzo1 strain but not in the Δmdm30 strain (38). Thus, a dramatic respiration defect was observed not in the Δmdm30 strain (Fig. 7A) but in the Δfzo1 strain (Fig. 7B). Similarly to the growth defect in acetate-containing medium, the fzo1 null mutant did not grow in nonfermentable glycerol, while a modest/moderate growth defect was observed in the absence of Mdm30 (Fig. 7A and B). Likewise, previous studies also found that the Δfzo1 strain, but not the Δmdm30 strain, did not grow in glycerol-containing medium (38, 44). Further, mRNA export occurs in dextrose-containing growth medium in the null mutation of FZO1 (18). Thus, the modest/moderate defect in mitochondrial respiration/function in the absence of Mdm30 does not appear to primarily cause significant decrease of mRNA export.
Growth and Western blot analysis. (A and B) Growth analysis of the wild-type, Δmdm30 (A), and Δfzo1 (B) strains in the medium containing 2% dextrose, 2% glycerol, or 2% acetate as a sole carbon source. YP, yeast extract and peptone. (C) Western blot analysis of Fzo1 and actin in the wild-type and Δmdm30 strains.
Mdm30 promotes mRNA export independently of splicing of YRA1 and SUS1 mRNAs.Our results reveal that Mdm30 interacts with Sub2 for proteasomal degradation in a transcription-dependent manner to promote mRNA export independently of mitochondrial function. Such stimulation of mRNA export by Mdm30 is mediated via enhanced recruitment of Yra1 (30). However, it is possible that reduced mRNA export via decreased recruitment of Yra1 in the Δmdm30 mutant can result from impaired splicing of YRA1 mRNA in the absence of Mdm30, as YRA1 has one intron and Sub2 regulates mRNA splicing (45, 46). We rule out this possibility based on our recent results (30) demonstrating that increased abundance of Sub2 in the Δmdm30 mutant does not impair splicing of the intron-containing genes such as RPL7B and RPS17B. However, Sub2 can have a gene-specific role in regulating mRNA splicing (45, 46, 67–69). Thus, it is possible that increased abundance of Sub2 in the absence of Mdm30 impairs Yra1 mRNA splicing and hence mRNA export. To test this, we analyzed the splicing of YRA1 mRNA in the wild type and Δmdm30 strains, using oligo(dT)-based reverse transcriptase PCR (RT-PCR) assay as done previously (30). For this purpose, we prepared cDNAs from total RNAs of the wild-type and Δmdm30 strains and then amplified cDNAs using primer pairs targeted to the exon and intron regions of YRA1. The PCR signals using exon-specific primers in the wild-type and Δmdm30 strains are equal (Fig. 8A), supporting the idea that Mdm30 does not alter transcription of YRA1, consistent with previous studies (30). The absence of PCR signals using intron-specific primers in the wild-type and Δmdm30 strains (Fig. 8A) indicates that the splicing of YRA1 mRNA is not impaired in the Δmdm30 mutant. Otherwise, the PCR signal using the intron-specific primer pair would have been observed in the Δmdm30 strain, as observed in the sub2-ts mutant (30). However, the lack of the PCR signals could be due to the fact that intron-specific primers did not work. To test this possibility, we performed PCR analysis of yeast genomic DNA using YRA1 intron-specific primers and observed the PCR signal (Fig. 8B), supporting the idea that intron-specific primers worked. Therefore, our results demonstrate that Mdm30 promotes mRNA export independently of the splicing of YRA1 mRNA. These results were further corroborated by radioactive PCR (Fig. 8C).
Mdm30 does not promote the splicing of YRA1 and SUS1 mRNAs. (A) Oligo(dT)-based RT-PCR analysis reveals that null mutation of MDM30 does not impair splicing of SUS1 and YRA1 mRNAs. PCR signals were analyzed by agarose gel electrophoresis with ethidium bromide staining. (B) Amplification of genomic DNA using PCR primer pairs targeted to the intron regions of SUS1 and YRA1. (C) RT-PCR analysis by radioactive PCR for YRA1 mRNA. (D) Radioactive RT-PCR analysis for SUS1 mRNA. The arrows (at the bottom) refer to the primers used in the PCR analysis. Amplified SUS1 fragments are shown on the left side of the autoradiogram.
Like Yra1, another mRNA export factor, namely, Sus1, has introns and participates in mRNA export via the TREX-2 (transcription-export 2) complex. TREX-2 consists of Sus1, Sac3, Thp1, Cdc31, and Sem1 and interacts with NPC and mRNA export receptor to mediate nuclear export of mRNA. The Sus1 component of TREX-2 is the product of two-intron-containing SUS1 mRNAs and is conserved among eukaryotes. To test whether Mdm30 facilitates mRNA export by promoting splicing of SUS1 mRNA, we analyzed SUS1 mRNA splicing in the wild-type and Δmdm30 strains. For this purpose, we prepared SUS1 cDNAs from the wild-type and Δmdm30 strains and then amplified cDNAs using primer pairs targeted to the exon and first intron regions of SUS1. Using exon-specific primers in the PCR analysis, we found that the SUS1 mRNA level was not altered in the Δmdm30 strain (Fig. 8A). However, we did not observe PCR signals using the intron-specific primers in the wild-type and Δmdm30 strains (Fig. 8A). Such absence of PCR signals was not due to the fact that intron-specific primers did not work, as these primers generated PCR signal when yeast genomic DNA was used as a template (Fig. 8B). These results support the idea that Mdm30 does not promote SUS1 mRNA splicing. These observations were further corroborated by radioactive PCR (Fig. 8D). Thus, our results support the idea that Mdm30 does not facilitate SUS1 mRNA splicing. Taken together, our results reveal that Mdm30 promotes mRNA export independently of splicing of YRA1 and SUS1 mRNAs.
DISCUSSION
Here, we show that Mdm30 interacts with Sub2 and regulates Sub2’s abundance/stability in a transcription-dependent manner to orchestrate mRNA export. Consistently, Mdm30 is found to be localized in the nucleus (39). In addition, Mdm30 also regulates the steady-state level of Fzo1 (38, 42, 43) but does not greatly alter mitochondrial respiration/function. However, the loss of Fzo1 dramatically impairs mitochondrial fusion or respiration but not mRNA export. Thus, the decrease in mRNA export in the absence of Mdm30 does not appear to be mediated via the modest/moderate defect in mitochondrial respiration/function in the Δmdm30 strain. Further, Mdm30 does not promote mRNA splicing of export factors. Collectively, Mdm30 interacts with Sub2 to regulate its abundance in a transcription-dependent manner in orchestrating mRNA export independently of mRNA splicing or mitochondrial function (Fig. 9), thus advancing our understanding of mRNA export.
Schematic diagram showing the Mdm30 regulation of Sub2 in a transcription-dependent manner in stimulation of mRNA export independently of splicing or mitochondrial respiration/function. The double-headed arrow indicates interaction, and the single-headed arrows represent stimulation. Sub2 associates with the gene in a transcription-dependent manner and becomes ubiquitylated via direct interaction with the F-box protein Mdm30 for proteasomal degradation. Such degradation of Sub2 promotes recruitment of mRNA export adaptor Yra1 to enhance mRNA export (30). Further, splicing of mRNA export factors Sus1 and Yra1 is not promoted by Mdm30, and thus, Mdm30 facilitates mRNA export independently of YRA1 and SUS1 mRNA splicing. Moreover, Mdm30 has a moderate effect on mitochondrial respiration/function and regulates the stability of Fzo1, which is required for mitochondrial fusion/dynamics and function. However, mRNA export occurred in the fzo1 null mutant in dextrose-containing growth medium (18).
Increased abundance of Sub2 is found to impair mRNA export in yeast (6, 30) as well as in Xenopus oocytes (32). However, it was unknown until recently how the cellular level of Sub2 is maintained for optimal mRNA export. Recently, we have demonstrated that Sub2 undergoes ubiquitylation and proteasomal degradation in orchestrating mRNA export (30). Such UPS regulation of Sub2 is mediated by the F-box protein, Mdm30 (30), which functions as a component of SCF ligase (26–29). However, it is not clear whether Mdm30 directly regulates Sub2 ubiquitylation and proteasomal degradation. If so, it would interact with Sub2. We find here that Mdm30 interacts with Sub2 in vitro (Fig. 1F and G) as well as in vivo (Fig. 1A to E). Further, temporary or long-term loss of Mdm30 increases Sub2’s abundance via impaired ubiquitylation and decreases mRNA export (Fig. 3) (30). Cycloheximide chase experiments also revealed increased stability of Sub2 in the absence of Mdm30 (Fig. 2), and increased abundance of Sub2 impairs mRNA export (30). These results support the idea that Mdm30 directly regulates Sub2’s abundance via UPS in controlling mRNA export.
In addition to promoting mRNA export, Sub2 is also involved in splicing (45, 46, 67). Thus, increased abundance of Sub2 in the absence of Mdm30 might alter the splicing of intron-containing genes associated with mRNA export. We find that the loss of Mdm30 does not impair splicing of intron-containing genes, including mRNA export factors such as Yra1 and Sus1 (Fig. 8) (30). Thus, Mdm30 promotes mRNA export by regulating Sub2’s abundance, but independently of mRNA splicing. However, increased abundance of Sub2 in the absence of Mdm30 might alter transcription and hence cotranscriptional mRNA export, as Sub2 is recruited to the active coding sequence and promotes transcription (30, 70). We rule out this possibility, since increased abundance of Sub2 or loss of Mdm30 does not alter transcription (Fig. 3C and D) (30). In addition to regulating Sub2’s abundance, Mdm30 is also involved in degrading Fzo1 (38, 42, 43) (Fig. 7C), which is associated with mitochondrial fusion/function. Thus, the decrease in mRNA export in the absence of Mdm30 could be mediated via the defect in mitochondrial fusion/function. However, we do not observe a dramatic defect in mitochondrial respiration/function in the absence of Mdm30 (Fig. 7A), while the loss of Fzo1 leads to severe impairment of mitochondrial respiration/function (Fig. 7B). Such a modest/moderate defect in mitochondrial respiration or function in the absence of Mdm30 does not appear to contribute to the significant defect in mRNA export, as mRNA export occurred in the null mutant of Fzo1 (which impairs mitochondrial respiration/function) (18). Taken together, our results reveal that Mdm30 regulates mRNA export by controlling the abundance of Sub2, but independently of transcription, splicing, or mitochondrial function. Although Mdm30 is found to promote mRNA export primarily via UPS regulation of Sub2 independently of mitochondrial function in dextrose-containing growth medium, mitochondrial dynamics/function would be generally required for mRNA export and other cellular processes under the growth conditions that would require mitochondrial respiration/function.
Sub2 is cotranscriptionally recruited to the coding sequence by the THO complex and subsequently targets Yra1, which interacts with mRNA export receptor Mex67 for mRNA export through NPC. Interestingly, increased abundance of Sub2 decreases recruitment of Yra1 in yeast (30) and Xenopus oocytes (32), hence impairing mRNA export (30, 32). Thus, Sub2 ubiquitylation and proteasomal degradation are functionally important for optimal mRNA export. However, it is yet unclear whether transcriptionally engaged Sub2 is targeted for proteasomal degradation or whether it is degraded off the gene to regulate mRNA export. We find that Sub2’s abundance is increased following pharmacological and genetic inhibitions of transcription (Fig. 4 and 5). These results indicate that transcriptionally engaged Sub2 is targeted for proteasomal degradation. In agreement, Sub2’s abundance is increased at the active coding sequence in the absence of Mdm30 (30). Further, an elevated level of Sub2 was observed in the chromatin-bound fraction in the absence of Mdm30 (Fig. 6D to F, I, and J).
How does proteasomal degradation of transcriptionally engaged Sub2 promote mRNA export? As mentioned above, THO complex is cotranscriptionally recruited to the active coding sequence via carboxy-terminal domain (CTD) phosphorylation of the largest subunit of RNA polymerase II and interacts with mRNA (7–9, 71). Subsequently, the Hpr1 component of THO recruits Sub2, which interacts with Yra1 to form TREX. Yra1 is recruited by the Pcf11 component of cleavage and polyadenylation factor IA in RNA polymerase II’s CTD phosphorylation-dependent manner (37). Importantly, Sub2 competes with the Pcf11 interaction domain of Yra1 and hence transfers Yra1 to THO to form the TREX complex (37). However, Yra1 has two interaction domains (located at N and C termini) for Sub2, and each domain has the potential to interact with Sub2 independently of the other one (6). Therefore, Yra1 can potentially bind to an extra Sub2 in the TREX complex in the presence of excess Sub2. However, mRNA export receptor Mex67 recognizes Yra1 of TREX via its competitive interaction with the Sub2 interaction domain of Yra1 to promote mRNA export. Thus, binding of Sub2 to both sites of Yra1 in TREX would reduce/impair the interaction of Yra1 with Mex67 and hence mRNA export. It may be likely that extra Sub2 is ubiquitylated by Mdm30 and degraded by the 26S proteasome to promote the interaction of TREX-Yra1 with Mex67 to enhance mRNA export. Consistent with this possibility, we observed an increased level of Sub2 at the active coding sequence/chromatin and decreased mRNA export in the Δmdm30 strain (18, 30) (Fig. 6D to F, I, and J), and Sub2’s abundance is increased following pharmacological and genetic inhibitions of transcription (Fig. 4 and 5). In addition to controlling transcriptionally engaged Sub2 or Sub2 within TREX, Mdm30 might also be involved in regulation of Sub2 ubiquitylation and proteasomal degradation off the gene, as excess Sub2 might interfere with the transfer of Yra1 from Pcf11 to THO toward forming TREX complex, consistent with the fact that increased abundance of Sub2 decreases Yra1 recruitment to active gene/chromatin/RNA (18, 30, 32). Collectively, our results suggest possible mechanisms in stimulation of mRNA export via UPS regulation of Sub2, which remain to be further elucidated. Nonetheless, our results decipher new regulation of mRNA export (and hence gene expression) and set the stage for future mechanistic studies to understand how increased abundance of Sub2 impairs mRNA export.
Since Mdm30 targets transcriptionally engaged Sub2 for proteasomal degradation, it is expected to be observed at the active coding sequence or 3′ end. However, our ChIP analysis did not detect Mdm30 at the active coding sequence or 3′ end (18). This could be due to transient interaction of Mdm30 with Sub2 or weak cross-linking of Mdm30 with DNA and/or Sub2, analogous to the fact that Mediator was not found to be associated with the promoters of several Mediator-dependent highly transcribing genes, using the ChIP assay (72). Even though Mdm30 was undetectable at the active coding sequence or 3′ end in the ChIP assay, the proteasome was shown to be associated with the active coding sequence or 3′ end (73), where TREX is found to be formed (30, 33–37).
In summary, the F-box protein, Mdm30, interacts with an mRNA splicing/export factor, Sub2, for its ubiquitylation and proteasomal degradation in a transcription-dependent manner to promote mRNA export independently of mRNA splicing and mitochondrial fusion/function. These results provide significant insights into the UPS regulation of mRNA export. Since UPS and Sub2 are conserved among eukaryotes, similar regulatory mechanisms of mRNA export are likely to exist in humans. Indeed, the human homologue of Sub2 is found to undergo ubiquitylation and proteasomal degradation (31), and increased abundance of the Sub2 homologue in Xenopus oocytes has been shown to impair mRNA export (32), similarly to the results in yeast (30). Thus, UPS regulation of Sub2 in orchestrating mRNA export is likely to be conserved from yeast to humans. Further, since altered mRNA export and associated factors are linked to a number of diseases (e.g., see references 1, 74 to 90), our results would be useful in studying mRNA export and factors in human cells and disease models for understanding pathogenesis and developing therapies in the future.
MATERIALS AND METHODS
Plasmids.The plasmids pFA6a-13Myc-KanMX6, pFA6a-3HA-His3MX6, and pFA6a-KanMX6-PGAL1-3HA (91) were used for genomic tagging of the proteins of interest by Myc and HA epitopes. The plasmid pRS404 (92) was used in the PCR-based gene disruption.
Yeast strains.Yeast (Saccharomyces cerevisiae) strains used in this study are listed in Table 1.
Yeast strains used in this studya
Growth media.The yeast strains were grown in YPD (yeast extract and peptone plus 2% dextrose) up to an optical density at 600 nm (OD600) of 1.0 at 30°C prior to formaldehyde-based in vivo cross-linking or harvesting for RNA analysis. For Mdm30 underexpression, a yeast strain expressing Mdm30 under the control of the GAL1 promoter was initially grown in YPG (yeast extract and peptone plus 2% galactose) at 30°C up to an OD600 of 0.6 and then transferred to YPD for 2 h. For cycloheximide chase assay, yeast cells were grown in YPD at 30°C up to an OD600 of 0.5 and then suspended in fresh YPD medium for 15 min prior to cycloheximide (50 μg/ml) treatment. Yeast cells were collected at different time points following cycloheximide treatment for Western blot analysis. For transcriptional inhibition by α-amanitin, yeast cells were grown in YPD at 30°C up to an OD600 of 0.4 and then treated with α-amanitin (6.25 μg/ml) for 3 h prior to cross-linking for the ChIP assay or harvesting for Western blot and RNA analyses.
ChIP assay.The ChIP assay was performed as done previously (93–98). Briefly, yeast cells were treated with 1% formaldehyde, collected, and resuspended in lysis buffer. Following sonication, cell lysate (400 μl lysate from 50 ml of yeast culture) was precleared by centrifugation, and then 100 μl lysate was used for each immunoprecipitation. Immunoprecipitated protein-DNA complexes were treated with proteinase K, the cross-links were reversed, and DNA was purified. Immunoprecipitated DNA was dissolved in 20 μl TE 8.0 (10 mM Tris HCl [pH 8.0] and 1 mM EDTA), and 1 μl of immunoprecipitated DNA was analyzed by PCR. PCR mixtures contained [α-32P]dATP (2.5 μCi for 25- μl reaction mixture), and the PCR products were detected by autoradiography after separation on a 6% polyacrylamide gel. As a control, “input” DNA was isolated from 5 μl lysate without going through the immunoprecipitation step and dissolved in 100 μl TE 8.0. To compare the PCR signal arising from the immunoprecipitated DNA with the input DNA, 1 μl of input DNA was used in the PCR analysis. The primers used for PCR analysis of ADH1 open reading frame (ORF) are 5′-CGGTAACAGAGCTGACACCAGAGA-3′ and 5′-ACGTATCTACCAACGATTTGACCC-3′. Autoradiograms were scanned and quantitated by the National Institutes of Health Image 1.62 program. Immunoprecipitated DNAs were quantitated as the ratio of immunoprecipitate to input. The average ChIP signal of the biologically independent experiments is reported with standard deviation (SD; Microsoft Excel).
Isolation of total and cytoplasmic RNAs.Total RNA was prepared from yeast cell culture as done previously (18, 30, 99). Briefly, 10 ml yeast culture was harvested and then suspended in 100 μl RNA preparation buffer (500 mM NaCl, 200 mM Tris-HCl, 100 mM Na2EDTA, and 1% SDS) along with 100 μl phenol-chloroform-isoamyl alcohol and a 100-μl volume equivalent of glass beads (acid washed; Sigma). Subsequently, the yeast cell suspension was vortexed with maximum speed (10 in a VWR minivortexer; catalog no. 58816-121) five times (30 s each). The cell suspension was placed in ice for 30 s between pulses. After vortexing, 150 μl RNA preparation buffer and 150 μl phenol-chloroform-isoamyl alcohol were added to the yeast cell suspension followed by vortexing for 30 s with maximum speed on a VWR minivortexer. The aqueous phase was collected following 5 min of centrifugation at maximum speed in a microcentrifuge machine. The total RNA was isolated from the aqueous phase by precipitation with ethanol.
Cytoplasmic RNA was prepared from yeast cells as described previously (18, 30, 99). Briefly, harvested yeast cells from a 10-ml culture were suspended in 400 μl sorbitol solution (0.9 M sorbitol, 0.1 M EDTA, and 14 mM β-mercaptoethanol) and then were incubated with 20 μl Zymolyase (10 mg/ml) for 25 min at 37°C followed by centrifugation for 5 to 10 s. The supernatant was carefully removed, and the spheroplast was gently suspended in 100 μl RNA preparation buffer along with a 20-μl volume equivalent of glass beads for immediate cytoplasmic fractionation with mild vortexing (around 15 s on the VWR minivortexer with a low speed of 5). Following centrifugation, the supernatant was carefully collected and was used for cytoplasmic RNA preparation following phenol-chloroform-isoamyl alcohol extraction and precipitation with ethanol. The quality of cytoplasmic fractionation was verified by the absence of nuclear DNA in the cytoplasmic fraction (Fig. 3E) (30).
RT-PCR analysis.RT-PCR analysis was performed as done previously (30). Briefly, total RNA was prepared from a 10-ml yeast culture. Ten micrograms of total RNA was used in the reverse transcription assay. RNA was treated with RNase-free DNase (M610A; Promega) and then reverse transcribed into cDNA using oligo(dT) as described in the protocol supplied by Promega (A3800; Promega). PCR was performed using synthesized first strand or cDNA as the template and the primer pairs targeted to the ORF regions of the 18S ribosomal DNA (rDNA), ADH1, and other genes. RT-PCR products were separated by 2.2% agarose gel electrophoresis and visualized by ethidium bromide staining. For splicing analysis, radioactive PCR was also performed. The primer pairs used in the PCR analysis were as follows: ADH1, 5′-CGGTAACAGAGCTGACACCAGAGA-3′ and 5′-ACGTATCTACCAACGATTTGACCC-3′; 18S rDNA, 5′-GAGTCCTTGTGGCTCTTGGC-3′ and 5′-AATACTGATGCCCCCGACC-3′; YRA1 (exon), 5′-TCCAATCGATGGAGGCAGATCAAGA-3′ and 5′-CTATTTGGACCCCTCTTTACTGGTC-3′; YRA1 (intron), 5′-TCGCCTCGACCGTGATAGTTA-3′ and 5′-GAAAGACATGTTTCCCATAGC-3′; SUS1 (exon-1/exon-3), 5′-GGATACTGCGCAATTAAAGAGTCA-3′ and 5′-CTACAATCTCTTCAAGAAATTCCC-3′; SUS1 (intron-2/exon-3), 5′-GAGAACGTTACTAACAGTGGAATT-3′ and 5′-CTACAATCTCTTCAAGAAATTCCC-3′.
WCE preparation and Western blot analysis.To analyze protein levels, yeast strains were grown as described above, harvested, lysed, and sonicated to prepare whole-cell extract (WCE) with solubilized chromatin following the protocol as described previously for the ChIP assay (93–98). The WCE was run on an SDS-polyacrylamide gel and then analyzed by Western blotting. Anti-HA (F-7; Santa Cruz Biotechnology, Inc.), anti-Myc (9E10; Santa Cruz Biotechnology, Inc.), and antiactin (A2066; Sigma) antibodies to HA-tagged and Myc-tagged proteins and actin, respectively, were used in the Western blot analysis.
Formaldehyde-based in vivo cross-linking and co-IP assay.The co-IP assay was performed as described previously (47, 48). Briefly, a yeast strain carrying Myc-tagged Mdm30 and HA-tagged Sub2 was grown in YPD up to an OD600 of 1.0 and then cross-linked by formaldehyde. WCE was prepared by lysing and sonicating the cross-linked yeast cells. Immunoprecipitation was performed using an anti-Myc antibody and protein A/G plus agarose beads. After immunoprecipitation, the agarose beads were washed as in the ChIP assay. The washed A/G plus agarose beads were boiled in the SDS-PAGE loading buffer, and supernatant was analyzed by SDS-PAGE and Western blotting. An anti-HA-peroxidase antibody (H6533-1VL; Sigma) was used in the Western blot analysis. Following a similar experimental strategy, interaction of Myc-tagged Mdm30 with HA-tagged Mex67 was analyzed.
Immunopurification of Sub2 and Mex67.The yeast strain expressing HA-tagged Sub2 or Mex67 was grown in 100 ml YPD up to an OD600 of 1.0 and then was harvested. Subsequently, 800 μl WCE was prepared from the culture of each strain. Immunoprecipitation was performed using an anti-HA antibody and protein A/G plus agarose beads for 4 h at 4°C. Four hundred microliters WCE, 100 μl protein A/G plus agarose beads (25% slurry), and 10 μl anti-HA antibody (2 μg) were used for each immunoprecipitation. The agarose beads following immunoprecipitation were washed under high-stringency washing conditions as in the ChIP assay (93–98), but 0.5 M NaCl, instead of 1 M NaCl, was used in the second and third washes of the beads. Then, the beads were equilibrated by buffer E (50 mM Tris-HCl, 250 mM NaCl, 1% NP-40, 1 mM EDTA, pH 7.5) before elution of HA-tagged protein by HA peptide. The immobilized protein (HA-tagged Sub2 or Mex67) on A/G plus agarose beads was eluted by incubating the beads in 2 bed volumes (100 μl) of buffer E containing HA peptide with a final concentration of 1 mg/ml and aprotinin (10 μg/ml). The beads were incubated for 30 min at 25°C on a rotator. Elution was performed three times. Buffer E (100 μl) containing HA peptide and aprotinin was used for each elution.
Protein interaction assay in vitro.To analyze the interaction of Mdm30 with Sub2, the yeast strain expressing Myc-tagged Mdm30 was grown in 100 ml YPD up to an OD600 of 1.0, and subsequently 800 μl WCE was prepared. Four hundred microliters WCE was used for immunoprecipitation as in the ChIP assay using 10 μl anti-Myc antibody (2 μg) and 100 μl protein A/G plus agarose beads. Immobilized Mdm30 on beads was thoroughly washed under high-stringency washing conditions as in the ChIP assay, and then the washed beads with immobilized Mdm30 were incubated with immunopurified HA-tagged Sub2 in buffer E (50 mM Tris-HCl, 250 mM NaCl, 1% NP-40, 1 mM EDTA, pH 7.5) containing HA peptide and aprotinin for 15 min at 25°C. Subsequently, the beads were washed by buffer W (50 mM Tris-HCl, 2 mg/ml bovine serum albumin [BSA], 250 mM NaCl, 1% NP-40, 1 mM EDTA, pH 8.5) containing aprotinin four times (1 ml each time). The last wash was performed using buffer E containing aprotinin. The washed beads were then boiled in SDS-PAGE loading buffer, and supernatant was subsequently analyzed by SDS-PAGE and Western blotting to determine the interaction between Mdm30 and Sub2. An anti-HA-peroxidase antibody was used in the Western blot analysis to detect HA-tagged Sub2. Likewise, the interaction of Mex67 with Mdm30 was analyzed using Myc-tagged Mdm30 and immunopurified HA-tagged Mex67.
Growth analysis.The growth of different yeast strains was analyzed on solid growth medium containing YP (yeast extract and peptone) plus 2% dextrose, glycerol, or acetate as a sole carbon source. Wild-type, Δfzo1, and Δmdm30 strains were inoculated in liquid YPD medium and grown up to an OD600 of 0.2 at 30°C. Subsequently, yeast cells were suspended in fresh liquid YPD medium and grown up to an OD600 of 0.4 prior to spotting (3 μl) on solid growth medium. Yeast cells were spotted with serial dilutions (10-fold), grown at 30°C, and photographed after 2 or 3 days.
Chromatin fractionation.Chromatin fractionation was performed as described previously (100, 101). Briefly, 25 ml yeast cell culture was harvested at an OD600 of 1.0. Cells were washed in ice-cold STOP buffer (150 mM NaCl, 50 mM NaF, 10 mM EDTA, 1 mM NaN3, pH 8.0) twice at 400 × g for 5 min at 4°C. Cell pellet was then resuspended in PEMS buffer {100 mM PIPES [piperazine-N,N′-bis(2-ethanesulfonic acid)], 50 mM EDTA, 10 mM MgSO4, 1.2 M sorbitol, pH 6.9} with 200 μg lyticase and incubated at 37°C in a rotator for 20 min to generate spheroplasts. A small volume of this suspension was collected to be used as WCE. The remaining spheroplast suspension was pelleted down at 400 × g for 5 min at 4°C and washed twice in 1.2 M sorbitol. The spheroplasts were then resuspended in lysis buffer (HBS buffer [25 mM MOPS {morpholinepropanesulfonic acid}, 60 mM beta-glycerophosphate, 15 mM MgCl2, 15 mM EGTA, 15 mM p-nitrophenylphosphate, 0.2 mM Na3VO4, pH 7.2], 0.4 M sorbitol, 1 mM dithiothreitol [DTT], 1 cOmplete minitablet of protease inhibitors, and phenylmethylsulfonyl fluoride [PMSF]) and incubated in ice for 5 min. Triton X-100 was added to a final concentration of 1%, and the sample was put in a rotator in a cold room for 15 min to facilitate membrane solubilization. The spheroplast suspension was then mildly vortexed (∼15 s on the VWR minivortexer with a low speed of 5) and spun down at 22,000 × g for 15 min at 4°C to separate the chromatin-unbound fraction as a supernatant. This was followed by washing the pellet (chromatin-bound fraction) twice in lysis buffer. The chromatin-bound fraction was then suspended in the lysis buffer. Formaldehyde-mediated cross-linking-based chromatin fractionation was performed under the same conditions, but prior to harvesting, cells were treated with 1% formaldehyde for 15 min and subsequently quenched with 125 mM glycine for 5 min.
ACKNOWLEDGMENTS
We thank Judy Davie for providing yeast strains.
The work in the Bhaumik laboratory was supported by grants from the National Institutes of Health (1R15GM088798-01, 2R15GM088798-02, and 2R15GM088798-03), the American Heart Association (15GRNT25700298), and Southern Illinois University School of Medicine.
FOOTNOTES
- Received 11 November 2019.
- Returned for modification 9 December 2019.
- Accepted 3 January 2020.
- Accepted manuscript posted online 13 January 2020.
- Copyright © 2020 American Society for Microbiology.
REFERENCES
- 1.↵
- 2.↵
- 3.↵
- 4.↵
- 5.↵
- 6.↵
- 7.↵
- 8.↵
- 9.↵
- 10.↵
- 11.↵
- 12.↵
- 13.↵
- 14.↵
- 15.↵
- 16.↵
- 17.↵
- 18.↵
- 19.↵
- 20.↵
- 21.↵
- 22.↵
- 23.↵
- 24.↵
- 25.↵
- 26.↵
- 27.↵
- 28.↵
- 29.↵
- 30.↵
- 31.↵
- 32.↵
- 33.↵
- 34.↵
- 35.↵
- 36.↵
- 37.↵
- 38.↵
- 39.↵
- 40.↵
- 41.↵
- 42.↵
- 43.↵
- 44.↵
- 45.↵
- 46.↵
- 47.↵
- 48.↵
- 49.↵
- 50.↵
- 51.↵
- 52.↵
- 53.↵
- 54.↵
- 55.↵
- 56.↵
- 57.↵
- 58.↵
- 59.↵
- 60.↵
- 61.↵
- 62.↵
- 63.↵
- 64.↵
- 65.↵
- 66.↵
- 67.↵
- 68.↵
- 69.↵
- 70.↵
- 71.↵
- 72.↵
- 73.↵
- 74.↵
- 75.↵
- 76.↵
- 77.↵
- 78.↵
- 79.↵
- 80.↵
- 81.↵
- 82.↵
- 83.↵
- 84.↵
- 85.↵
- 86.↵
- 87.↵
- 88.↵
- 89.↵
- 90.↵
- 91.↵
- 92.↵
- 93.↵
- 94.↵
- 95.↵
- 96.↵
- 97.↵
- 98.↵
- 99.↵
- 100.↵
- 101.↵
- 102.↵
- 103.↵
