We show that mutants lacking either the phosphatase activator Rrd1 or the phosphatase Pph3 are resistant to rapamycin and that double mutants exhibit a synergistic response. This phenotype could be related to an inability of the mutants to degrade RNA polymerase II, leading to transcription of critical genes that sustain growth.
THE yPtpa1/Rrd1 protein shares 35% identity with the human phosphotyrosyl phosphatase activator, hPTPA, which has been proposed to function as a phosphatase activator (Cayla et al. 1994; Janssens et al. 1998; Ramotar et al. 1998). Rrd1 interacts with Sit4, a catalytic subunit belonging to the PP2A Ser/Thr phosphatase family (Douville et al. 2004; Zheng and Jiang 2005), and both proteins participate in the same genetic pathway to mediate resistance to 4-nitroquinoline-1-oxide and ultraviolet A (320–400 nm) light (Douville et al. 2004). The Rrd1–Sit4 complex may function to activate gene expression, as Sit4 is required for expression of several genes, e.g., SWI4 (Fernandez-Sarabia et al. 1992).
rrd1Δ mutants are resistant to the immunosuppressant rapamycin, suggesting that Rrd1 plays a role in the cellular response to this drug (Rempola et al. 2000). In yeast, rapamycin binds to the peptidyl-prolyl isomerase Fpr1 and this complex inactivates the Tor1, -2 kinases (Heitman et al. 1991), leading to the activation of Sit4 via dissociation from an inhibitor protein Tap42 (Di Como and Arndt 1996; Jiang and Broach 1999). The activated Sit4 dephosphorylates several targets, including the nutrient-responsive transcriptional activator Gln3 (which translocates to the nucleus to activate GLN1 and MEP2 expression) and the Ser/Thr kinase Npr1, which regulates the amino acid permeases (Schmidt et al. 1998). In general, rapamycin generates a profound modification in the transcriptional profile of yeast. While some genes, e.g., the diauxic shift genes CPA2 and PYC1, are upregulated by rapamycin exposure, others, such as the ribosomal protein genes including RPS26A, RPL30, and RPL9, are downregulated (Hardwick et al. 1999; Powers and Walter 1999).
Here, we investigated the cellular response mechanism of rrd1Δ mutants to rapamycin. We first assessed whether rrd1Δ mutants respond to the initial challenge of rapamycin by examining Npr1 kinase phosphorylation status and MEP2 expression (Schmelzle et al. 2004). As shown in Figure 1, rapamycin quickly triggered Npr1 dephosphorylation, as well as MEP2 expression, in both the parent and the mutant, eliminating a role for Rrd1 in these early events. We next determined the effect of rapamycin on the growth rate of the rrd1Δ mutant. As shown in Figure 2A, the parent and rrd1Δ mutant grew at nearly the same rate in the absence of rapamycin, but growth was reduced when the strains were challenged with rapamycin. Interestingly, growth of the parent ceased by 9 hr, while the mutant continued to grow (Figure 2A) and divide (data not shown) in the presence of rapamycin. Thus, the mechanism by which rapamycin induces growth arrest appears to depend on Rrd1.
A genomewide screen revealed that the Pph3 catalytic subunit of the PP2A family is involved in rapamycin resistance (Chan et al. 2000). We therefore tested whether Rrd1 might function via Pph3. As expected, growth of the parent ceased within 9 hr, while the rrd1Δ and pph3Δ single mutants continued growing in the presence of rapamycin (Figure 2B). Importantly, the rrd1Δ pph3Δ double mutant showed a synergistic response (Figure 2B), suggesting that Rrd1 and Pph3 act separately to mediate rapamycin-induced growth arrest. Whether Rrd1 acts via another member of the PP2A family, such as Pph21 and Pph22, is not known (Stark 1996).
We next tested whether Rrd1 could influence expression of genes known to be regulated in response to rapamycin (Hardwick et al. 1999). Expression of the CPA2 and PYC1 genes was induced in the parent strain, but not in the rrd1Δ mutant (Figure 3A), suggesting that Rrd1 is required to mediate rapamycin-induced gene expression. Since rapamycin is also known to trigger the downregulation of some genes, e.g., RPS26A and RPL9A, we tested whether Rrd1 is involved in this process. The RPS26A gene was nearly completely downregulated in the parent strain, but remained partially expressed in the rrd1Δ mutant upon rapamycin exposure (Figure 3B). Since RPL9A is downregulated in both strains, it appears that a subset of rapamycin-responsive genes is under the control of Rrd1. In fact, Rrd1 is required to support activator-dependent in vitro transcription of a well-characterized reporter E4 (Wu et al. 1996; data not shown), consistent with Rrd1 involvement in gene expression.
Since Rrd1 interacts with Sit4 and this complex has been shown to dephosphorylate the target proteins Sap185 and Gln3 (Crespo et al. 2002; Fellner et al. 2003), it is reasonable to assume that Rrd1 could affect gene expression by modulating the phosphorylation status of the C-terminal domain (CTD) of the Rpb1 subunit of RNA polymerase II upon rapamycin exposure. As such, the parent strain, the isogenic single mutants rrd1Δ and pph3Δ, and the double mutant rrd1Δ pph3Δ were assessed for the phosphorylation status of Rpb1. As shown in Figure 4A, the high-molecular-weight bands (∼150–200 kDa), corresponding to a heterogeneous population of Rpb1 polypeptides, disappeared when the parent strain was treated with rapamycin. Interestingly, a polypeptide band of ∼70 kDa was intensified following the drug treatment (Figure 4A), suggesting that Rpb1 is not dephosphorylated, but proteolytically processed. It is noteworthy that rapamycin treatment caused degradation of both the phosphorylated and the nonphosphorylated form of Rpb1 (data not shown). When a similar experiment was conducted with the rrd1Δ mutant, the phenomenon was significantly reduced, and more dramatically in the rrd1Δ pph3Δ double mutant (Figure 4A), but not in the gln3Δ mutant, which also displays rapamycin resistance (data not shown) (Cardenas et al. 1999; Powers and Walter 1999). These data are consistent with a model whereby both Rrd1 and Pph3 contribute separately to the molecular process that acts independently of the Gln3 signaling pathway to trigger degradation of Rpb1 upon rapamycin exposure. We note that the timing of Rpb1 degradation coincided with the reported kinetics of downregulation of the ribosomal protein genes that occurred within 60 min following rapamycin treatment (Cardenas et al. 1999; Powers and Walter 1999). At least, >50% of Rpb1 was converted to the 70-kDa form after 75 min of rapamycin treatment (Figure 4B). Thus, the downregulation of the ribosomal protein genes triggered by rapamycin could be a consequence of increased degradation of Rpb1 in the parent strain.
The rationale underlying the potential degradation of Rpb1 during rapamycin treatment may be the necessity of the cell to drastically reduce its metabolism. Since rapamycin mimics starvation conditions, drug-treated cells will adapt to a specific energy-saving response involving the downregulation of several physiological processes, including transcription. In this context, destruction of an important fraction of Rpb1 would be an efficient mechanism to prevent unnecessary initiation of gene transcription. Although the half-life of Rpb1 (<75 min) correlates with the timing of the downregulation of the ribosomal protein genes triggered by rapamycin, the cell growth ceased only at a much later time, presumably due to the additional time required to dilute and turn over all the Rpb1 and various other molecules (Cardenas et al. 1999). Thus, preventing the destruction of Rpb1, as in the case of rrd1Δ mutants and the even more pronounced case of rrd1Δ pph3Δ double mutants, is expected to permit continuous expression of critical genes required to promote growth in the presence of rapamycin.
The only other protein documented to be degraded in response to rapamycin is the translation initiation factor eIF-4G (Berset et al. 1998). eIF-4G is degraded within 2–3 hr in response to rapamycin treatment, while other initiation factors, such as eIF-4E associated with eIF-4G, remain unaffected (Berset et al. 1998). The degradation of eIF-4G requires a functional Tor1–kinase signaling pathway (Berset et al. 1998). Since the time frame during which Rpb1 is degraded coincides with that of eIF-4G, it is likely that Rpb1 degradation is also via the Tor1 pathway. If so, Rrd1 might execute a function downstream of the Tor1 pathway that culminates in the activated expression of a protease (e.g., those induced by rapamycin such as Lap3, Lap 4, Pep4, and Pbr1) that could degrade key proteins involved in translation and transcription (Mewes et al. 1999). Clearly, additional experiments are needed to unravel the exact molecular function by which Rrd1 controls rapamycin resistance.
This study was supported by a grant to D.R. from the Canadian Institutes of Health Research. During the tenure of this work D.R. was supported by a career scientist award from the National Cancer Institute of Canada, and currently by a senior fellowship from the Fonds de la Recherche en Sante du Quebec.
Communicating editor: B. J. Andrews
- Received May 25, 2005.
- Accepted November 12, 2005.
- Copyright © 2006 by the Genetics Society of America