© 2017. Published by The Company of Biologists Ltd.

Pib2 and EGO Complex are Both Required for Activation of TORC1 Natalia V. Varlakhanova1, Michael Mihalevic2, Kara A. Bernstein2 & Marijn G. J. Ford1 1 Department of Cell Biology and Physiology University of Pittsburgh School of Medicine 3500 Terrace Street Pittsburgh, PA 15261 2 Department of Microbiology and Molecular Genetics University of Pittsburgh School of Medicine 5117 Centre Avenue Pittsburgh, PA 15213 Correspondence: marijn@pitt.edu

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JCS Advance Online Article. Posted on 9 October 2017

Abstract

The TORC1 complex is a key regulator of cell growth and metabolism in

Saccharomyces cerevisiae. The vacuole-associated EGO Complex couples

activation of TORC1 to the availability of amino acids, specifically glutamine and

leucine. EGO Complex is also essential for reactivation of TORC1 following

rapamycin-induced growth arrest and for its distribution on the vacuolar membrane.

Pib2, a FYVE-containing PI3P-binding protein, is a newly-discovered and poorly

characterized activator of TORC1. Here, we show that Pib2 is required for

reactivation of TORC1 following rapamycin-induced growth arrest. Pib2 is required

for EGO Complex-mediated activation of TORC1 by glutamine and leucine as well

as for redistribution of Tor1 on the vacuolar membrane. Therefore, Pib2 and the

EGO Complex cooperate to activate TORC1 and connect PI3K signaling and

TORC1 activity.

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Introduction

The Target of Rapamycin Complex I (TORC1) couples multiple nutritional cues to

orchestrate an appropriate cellular growth response. Nutrients, in particular amino

acids, activate TORC1 signaling which results in a multi-pronged anabolic response,

including ribosome and protein synthesis, increase of biomass and growth. On

nutrient starvation, TORC1 is inactivated, which leads to a coordinated starvation

response, including amino acid permease synthesis and transport, amino acid

biosynthesis and induction of macroautophagy (Broach, 2012; Loewith et al., 2002;

Neufeld, 2010).

TORC1 is a multisubunit complex of ~2 mDa and consists of either Tor1 or Tor2, a

PIK-like kinase, and the accessory subunits Kog1, Lst8 and the non-essential Tco89

(Loewith et al., 2002; Wedaman et al., 2003). TORC1 appears to be constitutively

associated with the vacuolar membrane, independently of nutrient status, though

some sequestration to peri-vacuolar foci has been observed (Kira et al., 2014;

Sturgill et al., 2008). TORC1 exerts its growth effects via several downstream

signaling branches that together constitute the anabolic or catabolic response.

TORC1 stimulates protein and ribosome synthesis through several downstream

effector kinases including Sch9 and Ypk3 (Gonzalez et al., 2015; Urban et al., 2007).

Simultaneously, active TORC1 inhibits PP2A (Pph3, Pph21 and Pph22) and PP2A-

related (Ppg1 and Sit4) phosphatases, whose downstream effects include responses

to nitrogen starvation (Loewith and Hall, 2011). Furthermore, TORC1 inhibits

macroautophagy (Kamada et al., 2010). In addition to these main effector branches,

TORC1 directly interacts with an extensive array of kinases and phosphatases

(Breitkreutz et al., 2010). This includes Npr1, a kinase involved in regulating

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trafficking and localization of amino acid permeases (MacGurn et al., 2011; Merhi

and Andre, 2012; Schmidt et al., 1998) and Nnk1, implicated in nitrogen metabolism

(Breitkreutz et al., 2010).

Amino acids regulate TORC1 via several mechanisms which largely depend on the

Escape from Rapamycin-induced Growth Arrest Complex (EGO Complex) (Peli-Gulli

et al., 2015). EGO Complex consists of 2 small GTPases, Gtr1 and Gtr2, which are

recruited to the vacuolar membrane by a scaffold subcomplex (Powis et al., 2015)

consisting of Meh1 (also known as Ego1), Ego2 and Slm4 (also known as Ego3).

EGO Complex is highly conserved and the Gtrs have homologs in higher eukaryotes

known as the Rag GTPases. The Gtrs form a constitutive heterodimer whose activity

depends on their nucleotide binding status: the heterodimer is active when Gtr1 is

GTP-bound and Gtr2 is GDP-bound (Binda et al., 2009; Jeong et al., 2012;

Nakashima et al., 1999) and inactive in the opposite configuration. The nucleotide

state of the GTRs is regulated by several complexes that impinge on GTP

hydrolysis, loading or dissociation: Vam6, a component of the HOPS complex

involved in vacuolar fusion, was demonstrated to be a GEF for Gtr1 (Binda et al.,

2009); Lst4/Lst7 is a GAP for Gtr2, which results in activation of TORC1 (Peli-Gulli et

al., 2015) and the SEA complex is a GAP for Gtr1, which inactivates it (Neklesa and

Davis, 2009; Panchaud et al., 2013).

Particularly potent activators of TORC1 via EGO Complex are the amino acids

leucine and glutamine. Leucine promotes interaction between GTP-loaded Gtr1

(GTR1GTP) with Meh1 (Ego1) (Binda et al., 2009) and the leucyl tRNA synthetase

Cdc60 was shown to directly interact with Gtr1 in a leucine-dependent manner

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(Bonfils et al., 2012). Glutamine stimulates interaction of the GAP Lst4-Lst7 with

Gtr2, thereby promoting formation of Gtr2GDP, the active form that can activate

TORC1 (Peli-Gulli et al., 2015). Active Gtrs stimulate TORC1 via direct physical

interactions: Gtr1GTP interacts with Tco89 (Binda et al., 2009) and the active

heterodimer itself interacts with Kog1 (Sekiguchi et al., 2014).

In addition to GTPases, TORC1 is also regulated by signaling via the PI-3 kinase

Vps34 and its product PI3P, in both yeast and mammalian cells. In mammalian cells,

hVps34-dependent signaling is well characterized: amino acids activate hVps34,

which results in an elevation of PI3P levels (Nobukuni et al., 2005), which, in turn,

leads to activation of mTORC1 (Byfield et al., 2005; Yoon et al., 2011). Importantly,

the hVps34 pathway is also necessary for the activation of mTORC1 by the

mammalian homologs of the Gtr GTPases (the Rag GTPases). Hence, amino acids

activate mTORC1 via two necessary mutually-interdependent pathways: Rag

GTPases and hVps34. In yeast, deletion of Vps34 also results in a strong inhibition

of TORC1 (Bridges et al., 2012) but the downstream effectors of Vps34 in activation

of TORC1 are unknown. It is also currently unknown how Vps34-dependent and Gtr-

dependent activation of TORC1 are integrated.

Recent work has identified Pib2 (Phosphatidyl Inositol-3-Phosphate Binding 2) as an

additional activator of TORC1 (Kim and Cunningham, 2015; Michel et al., 2017;

Tanigawa and Maeda, 2017). Pib2 was initially identified, together with several

components of EGO Complex, as a hit in a screen for factors unable to recover from

rapamycin exposure (Dubouloz et al., 2005). Later, Pib2 was reported to be required

for TORC1 activation and lysosomal membrane permeabilization in the presence of

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ER stress (Kim and Cunningham, 2015). It has a FYVE domain, a conserved C-

terminal tail motif and a series of conserved stretches of amino acids in a region

otherwise predicted to be unstructured. The N-terminal region harbors a TORC1

inhibitory function whereas the C-terminal region is important for activation of

TORC1 (Michel et al., 2017). Pib2 interacts with vacuoles via its FYVE domain in a

PI3P-dependent manner and this depends on Vps34 (Kim and Cunningham, 2015).

Thus, we hypothesize that Pib2 integrates Vps34 signaling into Gtr-dependent

activation of TORC1.

Here, we report that Pib2 indeed genetically interacts with components of EGO

Complex and TORC1 signaling. Pib2 deletion phenocopies simultaneous loss of

Gtr1 and Gtr2 in TORC1 reactivation after rapamycin exposure, microautophagy and

Gtr-dependent relocalization of Tor1 to perivacuolar foci. Furthermore, Pib2 and the

Gtrs are reciprocally required for activation of TORC1 by glutamine and leucine. Our

data suggest that Pib2 and EGO Complex function in the same molecular pathway

that leads to activation of TORC1. Therefore, our findings provide evidence for a role

for Pib2, together with EGO Complex, in the reactivation of TORC1, thus offering

insight into how PI3P signaling might be coupled with Gtr-dependent activation of

TORC1.

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Results

PIB2 genetically interacts with components of EGO Complex and TORC1

Recent studies have identified Pib2 as a regulator of TORC1 but the mechanism of

Pib2 action remains unclear (Michel et al., 2017; Tanigawa and Maeda, 2017). To

identify functional interaction partners of Pib2 at the genomic level, we performed a

synthetic dosage lethality (SDL) screen by overexpressing Pib2 in each member of

the non-essential yeast gene deletion collection (Giaever and Nislow, 2014). The

premise of SDL is that overexpression of a gene of interest, when combined with a

mutant of a functional interaction partner, results in a measurable fitness defect, or,

in the extreme case, lethality (Kroll et al., 1996). In contrast, overexpression of the

same gene of interest in a wild-type background may result in no observable

phenotype. SDL has been used to screen the non-essential deletion collection for

novel participants in various cellular processes (Measday et al., 2005). We used

selective ploidy ablation (SPA) to efficiently introduce the Pib2 overexpression

plasmid, or appropriate controls, into each haploid member of the non-essential gene

deletion collection (Reid et al., 2011). The result is rapid introduction of

overexpression plasmids into haploid members of the deletion collection. We

obtained several strong SDL hits (p < 0.0001, when the deletion strain

overexpressing Pib2 is compared to the same deletion strain expressing an empty

vector or overexpressing EGFP), which included meh1 ego1) and tor1 (Fig.1A

and Supplementary Table 1). Since Meh1 (Ego1) is a vacuolar membrane anchor

for both Gtr1 and Gtr2, these newly uncovered genetic interactions demonstrate that

Pib2 is functionally related to EGO Complex. An additional strong hit (p < 7.8E-7)

was par32, a component the PP2A signaling branch downstream of TORC1, as

well as the deletion of YDL172C, which overlaps with the coding sequence of PAR32

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(Fig.1A and Supplementary Table 1). We also identified a set of genes enriched in

endosomal structure and function (for example vps30, vps27, vps28) (Fig.1A

and Supplementary Table 1). These results are consistent with an enrichment of

Pib2 in PI3P-containing endosomal/vacuolar membranes (Burd and Emr, 1998; Kim

and Cunningham, 2015). We also identified several hits in genes known to be

involved in regulation of the cell cycle and amino acid biosynthesis. In this work, we

pursued further characterization of the connections between Pib2 and EGO Complex

and TORC1.

Pib2 is required for reactivation of TORC1 after treatment with rapamycin

Pib2 was initially identified as a hit in a screen for cells that were impaired in

recovery from rapamycin, together with constituents of the EGO Complex (Dubouloz

et al., 2005). EGO Complex was shown to be required for reactivation of TORC1

after inactivation by rapamycin (Binda et al., 2009) as well as for a poorly understood

subclass of autophagy known as microautophagy (Dubouloz et al., 2005). Given that

Pib2 genetically interacts with EGO Complex and Tor1, we compared the

phenotypes of cells lacking Pib2 with those lacking components of EGO Complex,

specifically the Rag family GTPases Gtr1 and Gtr2. Like cells lacking Gtr1 or Gtr2

(Fig. 1B and S1), pib2 cells do not recover from exposure to rapamycin and fail to

resume growth after rapamycin-induced growth arrest. By contrast, cells lacking

Atg7, which have a defect downstream of TORC1 (cannot undergo macroautophagy

(Xie and Klionsky, 2007)), recover from exposure to rapamycin like W303A cells

(Fig. 1B).

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To assess TORC1 activity, we monitored the phosphorylation status of a well-

characterized target, Rps6 (ribosomal protein S6). Yeast Rps6 is phosphorylated at

two serine residues at its C-terminus (S232 and S233) in a TORC1-dependent

manner (Gonzalez et al., 2015). Hence, the phospho-status of Rps6 at these sites

can be used as a faithful readout of TORC1 activity. Rapamycin treatment virtually

eliminated phosphorylation of Rps6 at these sites in both wild type and ∆pib2 cells,

as expected on TORC1 inactivation (Fig. 1C). Following recovery from rapamycin

exposure, an increase in Rps6 phosphorylation was observed in wild type cells, to

levels comparable to that seen in untreated cells. By contrast, Rps6 remained

dephosphorylated at S232 and S233 in ∆pib2 cells, even after 24 hours of recovery

(Fig. 1C, 1D, recovering to ~3.5% of the wild type untreated control, p < 0.01). This

suggests that cells lacking Pib2 fail to reactivate TORC1 during recovery.

To determine whether the growth defect of pib2 cells on recovery from rapamycin

exposure is due to a defect in Gtr activation, we introduced constitutively active

forms of both Gtr1 and Gtr2 (Gtr1 Q65L, constitutively GTP-bound and Gtr2 S23L,

constitutively GDP-bound) (Gao and Kaiser, 2006) into pib2 cells. Cells lacking

Pib2 could not be rescued by introduction of constitutively active Gtrs (Fig. 1B). As a

control, cells lacking Gtr1 or Gtr2 were fully rescued by introduction of active Gtrs

(Fig. S1). Therefore, activation of Gtrs is not the underlying cause of the defect in

pib2 cells. To eliminate the possibility that Pib2 is required for the recruitment of

Gtrs to the vacuolar membrane, or that the Gtrs are mislocalized away from the

vacuolar membrane in pib2 cells and thus cannot activate TORC1, we compared

the localization of Gtr1, Gtr2 and Meh1 (Ego1) in W303A and pib2 cells. The

cellular distribution of Gtr1, Gtr2 and Meh1 (Ego1) was unchanged in pib2

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compared to W303A cells (Fig. S2 and data not shown). Hence, Pib2 is not required

for vacuolar localization of the Gtrs.

The recovery defect in pib2 cells was TORC1-dependent as introduction of a TOR1

mutant allele (L2134M, within the kinase domain), previously shown to render Tor1

hyperactive regardless of Gtr activation (Kingsbury et al., 2014; Takahara and

Maeda, 2012), into pib2 cells resulted in recovery and growth similar to wild type

cells (Fig. 1B). Vector alone controls are provided in Fig. S3A. Since pib2 cells

could not be rescued by constitutively active Gtrs, the defect in pib2 is not due to a

defect in activation of Gtrs. This result also suggests that activated Gtrs require Pib2

for activation of TORC1.

Mutants in components of the EGO Complex display a striking vacuolar phenotype

after exposure to rapamycin: grossly enlarged vacuoles that cannot return to their

pre-exposure size after removal of rapamycin (Dubouloz et al., 2005). This was

proposed to be due to a defect in microautophagy. We next asked whether pib2

cells display a similar vacuolar morphology defect. We evaluated the size of

vacuoles in W303A, pib2 and gtr1 gtr2 cells before, during and after rapamycin

treatment. On rapamycin exposure, vacuoles of wild type cells increased in size, as

expected, as a consequence of increased macroautophagy (Chan and Marshall,

2014) (Fig. 1E). During recovery, the vacuolar size returned to pre-exposure levels

after 48 hr. By contrast, vacuoles of gtr1 gtr2 cells enlarged on rapamycin

exposure and did not recover. Vacuoles of ∆pib2 cells likewise enlarged on

rapamycin treatment but continued expanding, even during recovery from rapamycin

exposure, similar to gtr1 gtr2 cells (Fig. 1E). We quantified these observations by

calculating the ratio of the maximal vacuolar cross-sectional area to the maximal

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cellular cross-sectional area (vac:cell area), to normalize to cell size (Fig. 1F).

Untreated W303A cells had a vac:cell area ratio of 0.23 0.05, which increased to

0.47 0.11 after rapamycin treatment (p < 0.01), before recovering to 0.21 0.03

after 48 hours. gtr1 gtr2 cells had an untreated vac:cell ratio of 0.36 0.12.

Rapamycin treatment increased this to 0.54 0.07 (p < 0.01), which increased

further to 0.70 0.08 after 48 hr recovery (p < 0.01). Similarly, pib2 cells had an

untreated vac:cell ratio of 0.30 0.12 which increased to 0.52 0.06 after rapamycin

exposure (p < 0.01). As was the case for cells lacking Gtrs, this ratio increased to

0.80 0.07 after 48 hr recovery (p < 0.01). Hence, vacuolar size and cell:vac scaling

ratio does not recover after rapamycin treatment in gtr1 gtr2 or pib2 cells. These

results demonstrate that loss of Pib2 phenocopies loss of the Gtrs. Pib2 is therefore,

like components of EGO Complex, involved in vacuolar dynamics and

microautophagy.

Pib2 and Gtrs are both required for activation of TORC1 by glutamine and

leucine

Glutamine and leucine are known to be the most potent activators of TORC1 (Bonfils

et al., 2012; Peli-Gulli et al., 2015) and their activating stimuli require the EGO

Complex for relay to TORC1 (Binda et al., 2009; Kim et al., 2008; Sancak et al.,

2008). If Pib2 indeed acts within the same pathway as the Gtrs, we predict that we

would observe a defect in stimulation of TORC1 by glutamine and leucine in cells

lacking Pib2. We therefore compared TORC1 reactivation by glutamine and leucine

in cells lacking either Pib2 or both Gtr1 and Gtr2. When grown in nutrient-rich

medium, both pib2 and gtr1 gtr2 double mutant cells exhibit basal TORC1

activity, as determined by assessing the phosphorylation state of Rps6 (Figs. 2A-B

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and 3A-B, left-most column, no significant differences). Nitrogen starvation resulted

in loss of detectable TORC1 activity, as expected (p < 0.01 in all cases). Addition of

either glutamine (3 mM) or leucine (3 mM) for the indicated times (Figs. 2A and 3A)

evoked reactivation of TORC1 in wild type cells (5 min, p < 0.01; 30 min, p < 0.05)

but not in pib2 or gtr1 gtr2. Importantly, expressing activated Gtrs in pib2 cells

did not rescue the glutamine- or leucine-dependent activation of TORC1 (Figs. 2C-D

and 3C-D). Pib2 is therefore not required for activation of Gtrs. Active Gtrs cannot

overcome the requirement for Pib2 in activation of TORC1. These observations are

quantified in Figs 2B, D, F and 3B, D, F, for glutamine and leucine respectively. We

conclude therefore that Pib2 and the Gtrs are both required to relay the glutamine

and leucine signals to TORC1.

Of note, refeeding nitrogen-starved pib2 or gtr1 gtr2 cells with a mixture of all

amino acids results in a robust and full phosphorylation of Rps6 and thus activation

of TORC1 (Fig. S3D). The degree of phosphorylation of Rps6 was comparable in

each strain and directly comparable to W303A. This suggests the existence of an

additional amino acid signal that stimulates TORC1 in a Gtr1, Gtr2- and/or Pib2-

independent manner. This serves as a positive control for our readout that

demonstrates that the extent of the potential response in pib2 or gtr1 gtr2 cells is

comparable to the response in W303A cells, when the stimulus is not glutamine or

leucine. Therefore, the defect in TORC1 activation in pib2 or gtr1 gtr2 cells is

stimulus-specific and the activation of TORC1 by leucine and glutamine is dependent

on both Pib2 and Gtr1/2.

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Expression of activated Gtrs (Gtr1 Q65L and Gtr2 S23L) in gtr1 gtr2 cells resulted

in sustained activity of TORC1, even under starvation conditions, whereas TORC1

remains inhibited by nitrogen starvation in wild type cells overexpressing active Gtrs

(Figs. 2C-D, 3C-D compare W303A with overexpressed Gtrs to gtr1 gtr2 with

overexpressed Gtrs, p < 0.01). Since wild type cells still express endogenous Gtr1

and Gtr2, we conclude that inactive forms of Gtrs (i.e., Gtr1GDP and Gtr2GTP) are

therefore required for inhibition of TORC1 by nitrogen starvation, as previously

observed (Kira et al., 2014).

To further confirm the interdependence of Pib2 and Gtrs in TORC1 activation by

glutamine and leucine, we also evaluated the effect of overexpression of Pib2 in

gtr1 gtr2 cells. Overexpression of Pib2 in gtr1 gtr2 cells did not rescue TORC1

activity, whereas it rescued TORC1 activity in pib2 cells (Fig. 2E-F and 3E-F; 5

min, p < 0.05). These observations again suggest that Pib2-dependent TORC1

activation by glutamine or leucine requires Gtrs. Pib2 overexpression in gtr1 gtr2

cells repressed even TORC1 basal activity (p < 0.01 for both Pib2 overexpressed in

W303A vs. gtr1 gtr2 and Pib2 overexpressed in pib2 vs. gtr1 gtr2), confirming

the existence of a previously reported Gtr-independent inhibitory function of Pib2 on

TORC1 (Michel et al., 2017). Repression of TORC1 basal activity is only observed in

cells lacking Gtrs and not wild type cells. We further examined the effects of

overexpression of a truncated Pib2 construct lacking its N-terminal 164 amino acids

(Pib2 N-term) on TORC1 activation. The N-terminal 164 amino acids of Pib2 was

previously reported to harbor an inhibitory function on TORC1 (Michel et al., 2017).

Indeed, Pib2 N-term did not inhibit basal activity of TORC1 in gtr1 gtr2 cells,

confirming the importance of this domain for the observed inhibitory function of Pib2

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(Fig. S3C). Taken together, these data strongly suggest a novel dual mode of action

of Pib2 on TORC1 activity: in the presence of Gtrs, Pib2 is an activator of TORC1

whereas in their absence it is an inhibitor.

Pib2 regulates Tor1 localization on the vacuolar membrane

Previously, it has been reported that the nucleotide state of Gtr1 affects localization

of Tor1 at the vacuolar membrane. Gtr1GTP appears to be required for dispersion of

Tor1 throughout the vacuolar membrane: in its absence, Tor1 accumulates in

perivacuolar foci (Kira et al., 2016). Tor1 also localizes to perivacuolar foci in the

absence of both Gtrs (Fig. 4A) (Kira et al., 2016). The identity of the puncta remains

unknown – previous work has demonstrated that they do not co-localize with Snf7 or

Ape1 and are hence not endosomal or phagophore assembly site respectively (Kira

et al., 2014). As our data suggests that Pib2 is required to relay a signal from

activated Gtrs to TORC1, we sought to evaluate the role of Pib2 in localization of

Tor1. Our prediction was that Tor1 will redistribute to puncta in pib2 cells if Pib2

indeed relays signals from activated Gtrs.

In W303A cells grown in nutrient-rich media, GFP-Tor1, expressed under control of

its native promoter from a centromeric plasmid, had a diffuse vacuolar membrane

distribution with some foci associated with the vacuolar membrane (Fig. 4A), as has

been observed previously with an integrated genomic copy of GFP-Tor1 (Kira et al.,

2014). Simultaneous loss of Gtr1 and Gtr2 resulted in a marked redistribution of

GFP-Tor1 into puncta associated with the vacuole: the number of vacuoles with Tor1

puncta increased from 18.6 2.9 % in W303A cells to 65.6 5.0 in gtr1 gtr2 cells

(p < 0.01). Similarly, loss of Tco89, a component of TORC1 required for relay of the

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Gtr signal (Reinke et al., 2004), resulted in a redistribution of GFP-Tor1 into the

vacuole-associated puncta (63.3 2.9 2.4% of vacuoles were associated with

puncta, p < 0.01 compared to W303A cells). Loss of Pib2 also resulted in an

increase in vacuoles associated with GFP-Tor1 puncta (41.5 3.4 % of vacuoles

associated with puncta; p <0.01) (Figs. 4A-B). Of note, expressing constitutively

active forms of Gtr1 and Gtr2 in pib2 cells did not change the number of vacuoles

associated with Tor1 foci (Fig. S3D). This observation may be explained by two

scenarios: either Pib2 and Gtr1/Gtr2 act independently to regulate Tor1 localization,

or Pib2 acts directly downstream of the Gtrs in regulating Tor1 localization. Further

studies are required to distinguish these possibilities. Currently, the function of Tor1

foci formation is unknown. To determine if Tor1 foci formation impinges on TORC1

activity, we analyzed foci formation after nitrogen starvation, when TORC1 activity is

known to be repressed. No significant changes in foci formation were observed in

W303A, pib2 and gtr1 gtr2 cells (Fig. S4 A, B; compare Fig. 4B and Fig. S4B).

This suggests that Tor1 foci formation does not correlate with activity of TORC1.

Strikingly, exposure to rapamycin for 3 hr, which is also known to inhibit TORC1

activity, resulted in a complete loss of Tor1 foci in all strains, even gtr1 gtr2 (Fig.

S4 C, D). A mechanistic explanation of this observation awaits further

experimentation.

Pib2 has been reported to directly interact with Tor1 and Kog1 (Michel et al., 2017;

Tanigawa and Maeda, 2017). We asked, therefore, whether Pib2 changes its

localization in response to loss of Gtrs, as observed for components of TORC1.

Indeed, in W303A cells, Pib2 is associated with the vacuolar membrane with some

foci. In the absence of Gtrs (Fig. 4C) or Tco89 (data not shown), Pib2 distribution

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alters with an increased number of vacuoles containing foci (Fig. 4D; W303A – 15.8

4.7 % of vacuoles had foci compared to 47.5 7.4 % for vacuoles in gtr1 gtr2

cells, p < 0.001). These data indicate that Pib2 is likely to follow the Gtr-dependent

distribution of TORC1.

Pib2 is not required for and does not regulate macroautophagy

PI3P is required for macroautophagy and removing Vps34, which is the sole PI3

kinase in yeast, results in its inhibition (Burman and Ktistakis, 2010). Since Pib2 is a

PI3P-binding protein and since its recruitment is Vps34-dependent (Kim and

Cunningham, 2015), we sought to determine whether Pib2 was an effector of PI3P in

regulating autophagy. We therefore used the well-established GFP-Atg8 processing

and flux assay in both W303A and pib2 cells expressing GFP-Atg8 from its native

promoter (Kirisako et al., 1999; Shintani and Klionsky, 2004). Basal GFP-Atg8

expression levels were directly comparable in W303A and pib2 cells. On rapamycin

exposure, similar increased expression levels of GFP-Atg8 were observed in both

W303A and pib2 cells, a consequence of enhanced microautophagic flux. Likewise,

comparable elevated amounts of free GFP, reflecting processed GFP-Atg8, were

observed in both W303A and pib2 (Fig. S5). Hence, Pib2 is not required for GFP-

Atg expression or processing and cells lacking Pib2 are not impaired in

macroautophagy.

Npr1 is constitutively active in pib2 cells

TORC1 directly interacts with, and phosphorylates, Npr1, which inhibits it

(Breitkreutz et al., 2010; MacGurn et al., 2011; Schmidt et al., 1998). Inhibition of

TORC1 activity, by rapamycin treatment or nitrogen starvation, therefore leads to

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activation of Npr1 that results in a number of downstream effects, including inhibition

of Ldb19 (Art1) (MacGurn et al., 2011), phosphorylation of Bul1 and Bul2 (Merhi and

Andre, 2012) and trafficking of the tryptophan permease Tat2 from the surface of the

cell to the vacuole for degradation (Schmidt et al., 1998). One additional target of

active Npr1 is the poorly characterized protein Par32. Active Npr1 results in

extensive phosphorylation of Par32 at multiple sites, which leads to a significant

change in migration rate (Boeckstaens et al., 2015). Therefore, migration rate of

Par32 can be used as a readout to evaluate the activity of Npr1.

par32 was a hit in our SDL screen using overexpressed Pib2 (p < 7.82E-07). We

therefore evaluated phosphorylation of Par32 as a readout of Npr1 activity in W303A

and pib2 cells. As expected, we observe that all Par32-3xHA, expressed in W303A

cells from the native PAR32 promoter, is quantitatively shifted to a slower-migrating

form on rapamycin treatment or nitrogen starvation, which would be consistent with

extensive posttranslational modification (Fig. 5A-C). Essentially all of the shift in

migration depends on the presence of Npr1. In pib2 cells, the steady-state

distribution of Par32 is shifted towards the slower migrating species than in W303A

cells, indicating that Npr1 is more active than in controls. Treatment with rapamycin

maximally shifted Par32-3xHA in pib2 cells, indicating further activation of Npr1

(Fig. 5C). All of the shifts in Par32-3xHA migration, in W303A or in pib2 cells, were

dependent on the presence of Npr1 (Fig. 5C and data not shown). In summary, at

steady state, Npr1 is partially active in pib2 cells, but not in W303A cells, and can

be further activated by additional inhibition of TORC1. The increased

phosphorylation of Par32 observed in ∆pib2 cells could not be completely reversed

by expression of the hyperactive mutant allele of Tor1 (L2134M) (Fig. 5D). Note that

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in both W303A cells and ∆pib2 cells, the extent of enhancement in TORC1 activity

on expression of Tor1 L2134M is directly comparable. Thus, Pib2 has an additional

function of repressing Npr1 activity independently of TORC1.

Deletion of Npr1 was reported to suppress the defect in exit from rapamycin-induced

growth arrest of various EGO mutants, including Meh1 (Ego1), Slm4 (Ego3) and

Gtr2 (Dubouloz et al., 2005). We therefore tested if deletion of Npr1 also suppresses

the defect in recovery from rapamycin of pib2 cells. npr1 cells displayed enhanced

growth compared to W303A cells on recovery from rapamycin (Fig. 5E). As before,

pib2 cells did not recover from treatment with rapamycin. However, simultaneous

deletion of Pib2 and Npr1 resulted in recovery from rapamycin (Fig. 5E). Hence,

activated Npr1 after rapamycin exposure contributes to the lack of growth of pib2

cells, in the same way as was previously observed in meh1 (ego1), slm4

(ego3) and gtr2 cells. Taken together these findings indicate that Pib2 has a

function in downregulation of Npr1 activity, which negatively affects recovery of

growth after rapamycin treatment (Fig. 6).

Discussion

In this work, we provide a detailed characterization of Pib2 and a comparison of its

function to that of EGO Complex. We demonstrate that Pib2, whose mechanism of

action was ill-defined, is required together with the Gtrs for activation of TORC1. We

identified strong genetic interactions by SDL between Pib2 and components of the

EGO Complex-TORC1 network. pib2 cells behaved identically to cells lacking both

Gtrs in many aspects, including recovery from exposure to rapamycin, vacuolar

dynamics, response to amino acids and distribution of GFP-Tor1 on the vacuolar

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surface. We therefore conclude that these responses of TORC1 require both Pib2

and EGO Complex (Fig. 6).

Previous reports demonstrated an ablated response to glutamine in cells lacking

Pib2 (Michel et al., 2017; Tanigawa and Maeda, 2017). Our data showed that cells

lacking Pib2 are unable to activate TORC1 in response to glutamine or leucine.

Importantly, the presence of mutants of Gtr1 and Gtr2 that are restricted to activated

states did not override the requirement for Pib2, suggesting that the role of Pib2 is

not activation of Gtrs. In mammalian cells, leucine is sensed by leucyl tRNA-

synthetase (LRS), which activates mTORC1 via two mutually necessary

mechanisms: LRS has GAP activity for RagD (a mammalian homolog of Gtr2) (Han

et al., 2012) and LRS directly interacts with and activates Vps34, thus mediating

TORC1 activation via Vps34-PLD1 branch (Yoon et al., 2016). Thus, leucine-

dependent activation of TORC1 integrates PI3P- and Rag-dependent signaling

pathways. The yeast homolog of LRS, Cdc60, has been reported to regulate the

activities of the Gtrs in response to amino acids (Bonfils et al., 2012). However, a

connection between PI3P signaling and leucine has not yet been established. We

speculate that Pib2 is an integral part of the PI3P signaling pathway that connects

leucine stimulation to TORC1 activation.

Overexpression of Pib2 in gtr1 gtr2 cells did not rescue the response to glutamine

or leucine, further highlighting the co-dependence of Pib2 and the Gtrs in activation

of TORC1. Previous models of Pib2 function suggested a Gtr-independent role in

activation of TORC1 (Kim and Cunningham, 2015; Tanigawa and Maeda, 2017),

based on the observations of knockouts of Gtr1 alone and synthetic lethality between

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PIB2 and components of EGO Complex. In these reports, residual activation of

TORC1 was detected in gtr1 cells. This residual activity was attributed to Pib2,

since it is a known activator of TORC1. Based on the fact that we do not detect

residual TORC1 activation in gtr1 gtr2 cells, it may be that the residual activation

detected in the single knockout stems from the action of the remaining component of

the Gtr dimer. Gtr1 and Gtr2, and their Rag homologs in higher eukaryotes, form

heterodimers that, when asymmetrically loaded with GTP and GDP respectively,

activate TORC1 (Hatakeyama and De Virgilio, 2016). It is possible that in gtr1 cells

the presence of Gtr2, combined with endogenous Pib2, and/or the absence of

Gtr1GDP, could have a residual activity on TORC1. It is known that Gtr1 can form

homodimers (Nakashima et al., 1999) and it would be interesting to see if the same

is true of Gtr2, especially in cells lacking Gtr1.

A prediction of independent pathways of TORC1 activation by Pib2 or Gtr1/2 would

be the existence of intermediate response levels of TORC1 to glutamine or leucine in

cells lacking either Pib2 or Gtr1/2. Under our conditions, we do not observe this and

we observe activation only in the presence of both Pib2 and Gtr1/Gtr2. One

possibility is that TORC1 is generally impaired in either pib2 or gtr1 gtr2 cells,

which may dampen an intermediate TORC1 activation response below detection

thresholds. Our data suggests otherwise for two reasons. First, basal TORC1 activity

is not impaired in either pib2 or gtr1 gtr2 cells (Figs 2, 3, 5D). Second, we show

that pib2 or gtr1 gtr2 cells can activate TORC1 to the same extent as wild type

cells (using a different, Pib2- and Gtr1/2- independent stimulus as reported in Fig.

S3B). This serves as a positive control for our readout that demonstrates that the

extent of the potential response in pib2 or gtr1 gtr2 cells is comparable to the

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response of the wild type (W303A) cells, when the stimulus is different. This argues

against a generally reduced/impaired TORC1 activity in the knockout strains, either

at the basal level or on activation by different stimuli.

The mechanism of TORC1 activation by Pib2 and the Gtr1/2 may be explained by

two overarching models: dependent and independent (Fig. S6). The prediction for

the independent model (model C in Fig. S6) is that intermediate levels of TORC1

activation would be detected. Model A (dependent hierarchical) postulates that

activation of TORC1 requires both Pib2 and the Gtrs that act in some hierarchical

manner (upstream/downstream of each other). A prediction of this model is that no

intermediate activation of TORC1 will be detected when Pib2 or the Gtrs are missing.

A small modification of model A is model B (dependent threshold). In this case,

activation of TORC1 depends, again, on both Pib2 and the Gtrs. However, the extent

of activation by either Pib2 alone or the Gtrs alone, either does not exist, or is so low

that it cannot be detected by multiple assays. However, a potentiation occurs

between Pib2 and the Gtrs which would result in a full response. Potentiation implies

dependence. Based on our results of activation of TORC1 by glutamine or leucine

after nitrogen starvation, we suggest that models A or B are most plausible. Model C

might be supported by the observed synthetic lethality between PIB2 and

components of EGO Complex. However, synthetic lethality is not necessarily

inconsistent with a dependent mechanism of action of Pib2 and Gtr1/2 on TORC1

activation (models A and B). Here, we report that Pib2 has an additional TORC1-

independent inhibitory function on Npr1 (Fig. 5). This could provide an alternative

explanation for the observed synthetic lethality between PIB2 and components of

EGO Complex. Cells lacking both Pib2 and components of EGO Complex will have

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constitutive Npr1 activity that is toxic (Schmidt et al., 1998). Taken together, the lack

of an intermediate response to glutamine and leucine in pib2 or gtr1 gtr2 cells,

as well as our detection of an additional function of Pib2 in regulating Npr1, which is

an alternative explanation for synthetic lethality, favors a dependent model of Pib2

action on TORC1 (models A or B).

Overexpression of Pib2 in gtr1 gtr2 cells not only failed to rescue TORC1 activity

in response to amino acids but also significantly dampened the basal response of

TORC1, suggesting Pib2 has an additional inhibitory function on TORC1, that is

unmasked in the absence of the Gtrs. This inhibitory function is Gtr-independent.

This supports previous work that identified an inhibitory region at the N-terminus of

Pib2 (Michel et al., 2017). Taken together, our observations suggest that Pib2 has

two antagonistic functions: activation of TORC1 that is co-dependent on the Gtrs and

Gtr-independent inhibition of TORC1. Intriguingly, mammalian cells have two PI3P-

binding homologs of Pib2 that are yet to be implicated in the regulation of mTORC1:

Phafin-1 and Phafin-2. These both lack the N-terminal supposedly inhibitory regions

as compared to Pib2, and have, instead a PH domain. Currently a link between the

Phafins and mTORC1 has not yet been established and thus it is of immediate

interest to determine if indeed Phafins play a role in mTORC1 signaling and, if so,

how their mechanism of action differs from Pib2.

Although the vacuolar localization of Tor1 in yeast is independent of the nutritional

status of the cell, TORC1 complex distribution is dynamically regulated by the

nucleotide bound state of Gtr1 and Gtr2 (Kira et al., 2016). Absence of the active

form of Gtr1 or of the Gtrs altogether lead to the accumulation of Tor1 in perivacuolar

foci. In pib2 cells, we observe a similar accumulation of Tor1 in foci suggesting that

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Pib2 also plays a role in Tor1 localization on the vacuolar membrane. In gtr1 gtr2

cells, Pib2 similarly accumulates in perivacuolar foci. Pib2 physically interacts with

Tor1 and Kog1 (Michel et al., 2017; Tanigawa and Maeda, 2017). Pib2 likely

therefore associates with TORC1 and follows its distribution in response to signaling

via Gtrs.

We observed that cells lacking Pib2 have partially activated Npr1, as assessed by

monitoring the phosphorylation status of the direct Npr1 effector Par32. We also

observed that Pib2 inactivates Npr1 in parallel to TORC1. Furthermore, loss of Npr1

resulted in growth resumption in cells lacking Pib2 after rapamycin exposure, as has

been previously observed for cells lacking components of EGO Complex (Dubouloz

et al., 2005). Thus, loss of Npr1 overrides the requirement for Pib2 or EGO Complex

in reactivation of TORC1 after rapamycin exposure. This suggests that sustained

Npr1 activity during recovery from rapamycin makes reactivation of TORC1

completely dependent on EGO Complex and Pib2. One potential mechanism for this

is that Npr1 directly phosphorylates TORC1 components or regulators, preventing

activation by all other activators except for activated Gtrs and Pib2. In this context, it

is of interest that Npr1 interacts with multiple components of TORC1 (Breitkreutz et

al., 2010). Alternatively, the mechanism for Npr1 suppression of the phenotypes of

loss of Pib2 and EGO Components could be more complex and indirect. Npr1 is a

known regulator of the stability, localization and transport of several permeases,

including the tryptophan permease Tat2 (Schmidt et al., 1998), the arginine and

uracil transporters Can1 and Fur4 (MacGurn et al., 2011) and the general amino acid

permease Gap1 (Merhi and Andre, 2012; O'Donnell et al., 2010; Shimobayashi et

al., 2013). Hence Npr1 may regulate the stability or activity of a permease that

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supplies an amino acid or other nutrient which is capable of activating TORC1 in an

EGO Complex and Pib2 independent manner after rapamycin treatment.

In summary, we establish a function for Pib2, a FYVE domain-containing PI3P

binding protein, in Gtr-dependent activation of TORC1, identifying a molecular bridge

between PI3P signaling and the EGO Complex. Future work will focus on the

conservation of function of the Pib2 homologs in mammalian cells.

Materials and methods

Yeast genetic manipulation and molecular biology

Strains used in this work are listed in Supplementary Table 2. Gene deletions were

generated in W303A/ diploids by homologous recombination and complete

replacement of the target open reading frame using cassettes amplified from pFA6a-

kanMX6, pFA6a-His3MX6 (Longtine et al., 1998) or pFA6-natMX4 (Goldstein and

McCusker, 1999) flanked with sequence (30 nt) proximal to the coding sequence of

the target gene. Diploids were subsequently sporulated by starvation in SPO

medium. Following manual tetrad dissection, knockout haploids were validated by

colony PCR, microscopy and, in some cases, sequencing. Strains harboring more

than 1 genomic modification were generated by mating and sporulation of

appropriate parental strains, followed by extensive revalidation. The standard PEG

3,350/lithium acetate/single-stranded carrier DNA protocol was used for yeast

transformation (Gietz and Schiestl, 2007).

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Media

YPD (2 % yeast extract, 1 % peptone, 2 % glucose, supplemented with L-tryptophan

and adenine) was used for routine growth. Synthetic Complete (SC; yeast nitrogen

base, ammonium sulfate, 2 % glucose, amino acids) or synthetic defined (SD; yeast

nitrogen base, ammonium sulfate, 2 % glucose, appropriate amino acid dropout)

media were used prior to microscopy or to maintain plasmid selection as indicated.

For sporulation, cells were successively cultured in YPA (2 % potassium acetate, 2%

peptone, 1 % yeast extract) and SPO (1 % potassium acetate, 0.1 % yeast extract,

0.05 % glucose). For starvation, cells were grown in SD –N (0.17 % yeast nitrogen

base without amino acids and ammonium sulfate, 2 % glucose). For stimulation,

cells were treated with SD –N supplemented with glutamine (Glu, 3 mM), leucine

(Leu, 3 mM) or supplemented with a complete dropout mix and were incubated for

the indicated times prior to lysis and processing.

Cloning and Plasmids

Plasmids used in this work are listed in Supplementary Table 3. GFP-S cer. PIB2

was generated by amplifying the PIB2 promoter and a fragment containing the PIB2

coding sequence and terminator from genomic DNA, prepared from W303a/

diploids using the Yeast DNA Extraction kit (Thermo Fisher Scientific, Pittsburgh),

using appropriate primers. The fragments were assembled with an additional

fragment encoding EGFP by overlap extension PCR. The resulting construct was

introduced into pRS316, previously linearized with SacI and ClaI, by Gibson

Assembly. S cer. PAR32-3xHA and GFP-TOR1 were amplified from genomic DNA

and were cloned using a similar approach.

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GTR1 Q65L, GTR2 S23L, and TOR1 L2134M, with their respective promoters and

terminators, were cloned by overlap extension and Gibson Assembly after

amplification from W303a/ genomic DNA. All point mutants described in this work

were constructed by overlap extension PCR at the site of the mutation using

appropriate primers followed by Gibson Assembly into the linearized target vector.

All primer sequences used in this work are available on request.

Dosage Lethality Screening

Selective ploidy ablation was used to introduce a control or Pib2 overexpression

plasmid into each strain in the non-essential haploid deletion collection (Thermo

Fisher Scientific) (Reid et al., 2011). In brief, the plasmid of interest (PGAL1-S cer.

PIB2 or the control PGAL1) is introduced into a Universal Donor Strain (UDS), where

all chromosomes are conditionally unstable, by standard transformation. Each

chromosome in the UDS has both a galactose-inducible promoter and a URA3

counter-selectable marker adjacent to its centromere. The UDS containing the

plasmid of interest is mated to each member of the non-essential deletion collection.

UDS chromosomes are subsequently eliminated from the diploids by centromere

destabilization followed by counter-selection (Reid et al., 2011). Destabilization and

Pib2 overexpression are simultaneously induced by switching to galactose as a

carbon source. After induction of Pib2 overexpression, colony sizes are measured,

compared to the same strain containing either of two control plasmids (PGAL1,

containing only the galatose promoter, or PGAL1-EGFP) and subjected to further

analysis.

Yeast colony manipulations were performed using a BM3 Colony Processing Robot

(S&P Robotics Inc., Toronto). The non-essential haploid deletion collection was

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reformatted into a density of 4x 384 colonies per plate, as 32 x 48 grids, such that

each member of the deletion array was present as a tetrad of 4 colonies. MAT

UDS, containing the plasmid of interest, was pinned into grids of 32 x 48 colonies per

plate on SC –LEU, followed by overnight growth at 30 ºC. UDS colonies were pinned

onto deletion array colonies, followed by 24 hr incubation at 30 ºC, to allow thorough

mating.

The diploids were repinned onto SC –LEU + galactose to induce overexpression of

Pib2, or the EGFP control, and simultaneous destabilization of the UDS

chromosomes. After ~48 hrs, the colonies were repinned onto SC –LEU + galactose

+ 5-fluoro-orotic acid (5-FOA, Toronto Research Chemicals Inc., Toronto) and

incubated at 30 ºC. After 72 hr, the colonies were repinned onto SC –LEU +

galactose + 5-FOA (Toronto Research Chemicals Inc., Toronto), followed by an

additional ~72 hr incubation at 30 ºC, prior to colony size measurement.

SDL Data Analysis

Colony sizes from high-resolution photographs of plates were measured using

SGAtools (Wagih et al., 2013). Colony size data were then visualized using the web

interface of the Data Review Engine in ScreenMill (Dittmar et al., 2010), to enable

manual checking of colonies flagged for attention due to potential pinning errors or

those colonies within individual 2x2 arrays that may be suspect. Data Review Engine

was also used to normalize colony sizes to the plate median for every plate

analyzed, to allow direct comparison of colony sizes between control and

experimental plates. Subsequently, the normalized growth values were used to

calculate Z-scores and p-values for each member of the deletion collection

overexpressing either Pib2 or containing the control plasmid. The results were then

analyzed using the Statistics Visualization Engine of ScreenMill. All experimental

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strains with a growth difference compared to the control strains with an implied p-

value of < 0.0001 were examined further.

Analysis of growth by serial dilution

Following overnight growth in YPD, target cells were diluted and regrown to mid-

logarithmic phase in YPD at 30 °C (OD600 0.6-0.8). Cells were then diluted to 0.5

OD600/ml and 5-fold serial dilutions were made in water. 2 l of each dilution was

spotted onto YPD or YPD + 2.5 ng/ml rapamycin plates. Where relevant, cells were

incubated for the indicated times with YPD supplemented with 200 ng/ml rapamycin

at 30 °C. After extensive washing, cells were resuspended in fresh YPD and

recovered at 30 °C for the indicated time prior to plating on YPD. Plates were then

incubated at 30 °C for 3 days prior to imaging.

Preparation of yeast for microscopy

Cells were grown overnight in YPD or synthetic defined medium appropriately

supplemented to maintain plasmid selection. Cells were then diluted in YPD and

grown to mid-logarithmic phase. Vacuolar membranes were stained with 10 M FM

4-64 (Thermo Fisher Scientific) for 45 min, followed by washing and incubation in

YPD medium without dye for 1 hr. For rapamycin treatment, cells in YPD were

treated for the indicated time with a final concentration of 200 ng/ml rapamycin

(Thermo Fisher Scientific). For recovery from rapamycin exposure, cells were

extensively washed and resuspended in fresh YPD and incubated as indicated. Cells

were plated onto No. 1.5 glass-bottomed coverdishes (MatTek Corporation, Ashland)

previously treated with 15 l 2 mg/ml concanavalin-A (Sigma-Aldrich, St. Louis).

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Western Blotting

Protein extracts for western blotting were obtained as described (Millen et al., 2009).

Briefly, cells were lysed on ice by resuspension in 1 ml ice-cold H2O supplemented

with 150 l 1.85 M NaOH and 7.5% (v/v) β-mercaptoethanol. Protein was

precipitated by addition of 150 l 50% (w/v) trichloracetic acid. Pellets were washed

twice with acetone, resuspended in 150 l 1× SDS-PAGE buffer and incubated for

30 min at 30 °C followed by 2 min at 95 °C. Antibodies used were as follows: anti-

Rps6 (ab40820, Abcam, Cambridge), anti-PGK1 (ab113687, Abcam), anti-EGFP

(ab290, Abcam), Phospho-Rps6 (4858, Cell Signaling Technology, Danvers) and

anti-HA (ab9110, Abcam). Labeled secondary antibodies were IRDye 680RD goat

anti-Rabbit antibody (926-68171, Li-Cor, Lincoln) and IRDye 680RD Goat anti-

mouse (926-68070, Li-Cor). These were detected using the Odyssey system (Li-

Cor). Bands were integrated and quantified using the Fiji distribution of ImageJ

(Schindelin et al., 2012).

Confocal microscopy and image analysis

Confocal images were acquired on a Nikon (Melville, NY) A1 confocal, with a 100x

Plan Apo 100x oil objective. NIS Elements Imaging software was used to control

acquisition. Images were further processed using Fiji or NIS Elements.

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Acknowledgements

The authors would like to thank Suzanne Hoppins and Jeff Brodsky for extensive

discussion and John Dittmar for assistance with ScreenMill.

Competing Interests

The authors declare no competing interests.

Funding

This work was supported by the National Institutes of Health grants 1R01GM120102-

01 (M.G.J.F) and ES024872 (K.A.B).

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Fig. 1. Pib2 is required for exit from rapamycin-induced growth arrest. (A)

Representative quartets from matched control and Pib2 overexpressing strains in the

SDL screen. Overexpression of Pib2 results in synthetic lethality with meh1

(ego1), tor1, par32, ydl172c and vps30 but not with avo2, which is shown

here as a non-interacting control. (B) Growth of W303A, atg7, pib2 and gtr1

expressing the indicated constructs on YPD during recovery from exposure to

rapamycin. Exponentially-growing cells (OD600 0.6-0.8) were treated with 200 ng/ml

rapamycin in YPD at 30 C for 5 hr. After washing, cells were plated on YPD and

were incubated for 3 days at 30 C. The left-most spot in each case corresponds to 2

l of a culture with OD600 0.5. Spots to the right of this correspond to 2 l of

sequential 5-fold dilutions. (C) Evaluation of the phosphorylation levels of S232 and

S233 of Rps6 in W303A and pib2. Cells as indicated were treated with rapamycin

as in (B). Total Rps6 and Pgk1 levels were used as loading controls. (D)

Quantification of the data presented in (C). Ratios of phosphorylated Rps6 to Pgk1

for each measurement (n = 3 in each case) were normalized to the mean ratio of

phosphorylated Rps6 to Pgk1 for untreated W303A cells. A two-way ANOVA was

conducted to determine the effects of genetic background (W303A and pib2) and

treatment (untreated, rapamycin treated and recovery) on Rps6 phosphorylation

levels. There was a significant interaction effect of background and treatment on

Rps6 phosphorylation levels (F2,12 = 9.46 hence p = 0.0034). Selected pairs of

values significant by the post-hoc Tukey HSD test (**, p < 0.01) are shown. (E)

W303A or the indicated knockout strains were stained with FM 4-64 for 45 min,

washed and chased in YPD for 1 hr prior to visualization. Where indicated, cells

were treated with rapamycin (200 ng/ml) for 3 hr. For recovery, cells were thoroughly

washed and were incubated for 48 hr in YPD. Scale – 5 m. (F) Quantification of the

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increase in vacuolar scaling for the cells shown in (E). The maximal vacuolar cross-

sectional area was divided by the maximal cellular cross-sectional area. For cells

where more than 1 vacuolar lobe existed (usually only W303A untreated or at 48 hrs

recovery), the maximal cross sectional area of each lobe was determined. 10-14

vacuoles and cells were measured for untreated and rapamycin-treated cells and 5-

10 for cells after recovery. For W303A and the knockout strains, the means of the

untreated, treated and recovery measurements were determined to be significantly

heterogenous (one-way ANOVA: W303A F2,31 = 45.25 hence p < 6.39E-10; gtr1

gtr2 F2,34 36.62 hence p < 7.26E-9; pib2 F2,26 = 55.40 hence p < 1.01 E-9).

Significantly different pairs of means, as assessed by the post-hoc Tukey HSD test,

are indicated (**, p < 0.01). Non-significantly different means are indicated below the

W303A chart (p = 0.90).

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Fig. 2. Pib2 is required for stimulation of TORC1 activity by glutamine.

Phosphorylation levels of Rps6 were evaluated under the indicated conditions.

Untreated cells were grown in Synthetic Complete medium. Cells were nitrogen-

starved by incubating in SD –N medium for 3 hr. For stimulation, cells were treated

with SD –N supplemented with glutamine (Glu, 3 mM) and were incubated for the

indicated times prior to lysis and processing. Both total Rps6 and Pgk1 were used as

loading controls. (A) W303A, pib2, gtr1 gtr2. (B) Quantification of the data shown

in (A). Grey lines: selected statistically significant differences between means of

phospho-Rps6 (Tukey HSD; *, p < 0.05; **, p < 0.01). For each cell type, differences

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in means of phospho-Rps6 were evaluated by one-way ANOVA for each of the

treatment conditions. Black lines: selected statistically significant differences

between means of phospho-Rps6 (Tukey HSD; *, p < 0.05; **, p < 0.01). For each

treatment shown, the means of phospho-Rps6 were compared for W303A, pib2

and gtr1 gtr2 by one-way ANOVA. For quantification, the phospho-Rps6 signal

was normalized to the corresponding Pgk1 loading control. (C) Strains as in (A) but

expressing Gtr1 Q65L and Gtr2 S23L from their native promoters on centromeric

plasmids. (D) Quantification of the data shown in (C). (E) Strains as in (A) but

overexpressing Pib2 from an episomal Tet-Off plasmid. Cells were grown in

appropriate medium containing 5 g/ml doxycycline. Cells were diluted and

inoculated into doxycycline-free medium for 12 hr to allow overexpression of Pib2.

The nitrogen starvation and amino acid stimulation were then performed as in (A).

(F) Quantification of the data shown in (E).

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Fig. 3. Pib2 is required for stimulation of TORC1 activity by leucine. This work

was performed as in Fig. 2, but with leucine (leu) stimulation (3 mM) instead of

glutamine. (A) W303A, pib2, gtr1 gtr2. (B) Quantification of the data shown in

(A). (C) Strains as in (A) but expressing Gtr1 Q65L and Gtr2 S23L from their native

promoters on centromeric plasmids. (D) Quantification of the data shown in (C). (E)

Strains as in (A) but overexpressing Tet-Off PIB2 from an episomal Tet-Off plasmid.

(F) Quantification of the data shown in (E).

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Fig. 4. Pib2 regulates localization of Tor1 on vacuoles. (A) GFP-Tor1 localization

in W303A, gtr1 gtr2, tco89 and pib2 cells as indicated. The indicated strains

expressed GFP-Tor1 from its native promoter on a centromeric plasmid. Cells were

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grown in Synthetic Complete medium until they reached OD600 0.6-0.8. W303A or

the indicated knockout strains were stained with FM 4-64 for 45 min, washed and

chased in YPD for 1 hr prior to visualization. Scale – 5 m. (B) Quantification of the

numbers of vacuoles displaying GFP-Tor1 foci in each of the indicated strains. Foci

were counted on z-stacks collected for each of the strains (from 250 to 400 vacuoles

were assessed for each strain). Means of numbers of vacuoles displaying foci were

significantly heterogeneous (one-way ANOVA, F4,15 = 150.45; p < 8.77E-10). A post-

hoc Tukey HSD test for significance was performed between each of the means.

Selected significant differences between means (**, p < 0.01) are indicated on the

plot and the means showing a non-significant difference (p = 0.80) are indicated

below the plot. (C) As in (A) but with strains as indicated expressing GFP-Pib2. (D)

Quantification of the data shown in (C). Foci were counted on z-stacks collected for

each of the strains (~250 vacuoles were assessed in each strain). The means of

vacuoles displaying foci were significantly different for the two strains (two-tail t-test

with 6 degrees of freedom; t = 7.23 hence p = 0.0003).

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Fig. 5. Npr1 is active and is the underlying cause of the defect in recovery from

rapamycin exposure of pib2 cells. (A) W303A and npr1 cells expressing Par32-

3xHA were treated with rapamycin (200 ng/ml) for 3 hr as indicated. Par32-3xHA

was visualized using an anti-HA monoclonal antibody. (B) W303A and pib2 cells

expressing Par32-3xHA were nitrogen-starved for 3 hr. Par32-3xHA was then

visualized as in (A). (C) The strains as indicated were treated with rapamycin as in

(A). (D) W303A or pib2 cells expressing Par32-3xHA and Tor1 L2134M, as

indicated, were grown in Synthetic Complete medium. Par32-3xHA was then

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visualized as in (A). Relative TORC1 activity was calculated based on the

phosphorylation levels of Rps6, normalized to a Pgk1 loading control. W303A = 100

%. (E) Growth of W303A and isogenic strains containing the indicated knockout on

YPD during recovery from exposure to rapamycin. Exponentially-growing cells

(OD600 0.6-0.8) were treated with 200 ng/ml rapamycin in YPD at 30 C for 5 hr. After

washing, cells were plated on YPD and were incubated for 3 days at 30 C. The left-

most spot in each case corresponds to 2 l of a culture with OD600 0.5. Spots to the

right of this correspond to 2 l of sequential 5-fold dilutions.

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Fig. 6. Proposed model for control of TORC1 signaling by Pib2 and Gtr1/2.

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J. Cell Sci. 130: doi:10.1242/jcs.207910: Supplementary information

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Supplementary Table 2: Yeast strains used in this work

Strain Genotype Reference W303A MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;;

trp1-­1;; ura3-­1;; can1-­100 PY_126 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;;

trp1-­1;; ura3-­1;; can1-­100;; Dpib2::KAN This work

PY_96 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dgtr1::HIS3

This work

PY_100 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dgtr2::KAN

This work

PY_104 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dgtr1::HIS3;; Dgtr2::KAN

This work

PY_150 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dnpr1::NAT

This work

PY_162 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dpib2::KAN;; Dnpr1::NAT

This work

PY_160 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dgtr1::HIS3;; Dgtr2::KAN;; Dnpr1::NAT

This work

PY_108 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Datg7::HIS3

This work

PY_110 MATa;; ade2-­1;; leu2-­3,112;; his3-­11,15;; trp1-­1;; ura3-­1;; can1-­100;; Dtco89::HIS3

This work

UDS (W8164-­2B) MATa; CEN1GCS;; CEN2GCS;; CEN3GCS;; CEN4GCS;; CEN5GCS;; CEN6GCS;; CEN7GCS;; CEN8GCS;; CEN9GCS;; CEN10GCS;; CEN11GCS;; CEN12GCS;; CEN13GCS;; CEN14GCS;; CEN15GCS;; CEN16GCS;; ADE2;; can1-­100;; his3-­11,15;; leu2-­3,112;; LYS2;; met17;; trp1-­1;; ura3-­1;; RAD5

(Reid et al., 2011)

Yeast Mata Knockout Collection in BY4741

MATa;; his3D1;; leu2D0;; met15D0;; ura3D0;; DgeneX::KAN

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Supplementary Table 3: Plasmids used in this work Plasmid Reference PGAL1-­S cer. PIB2 pRS315 PGAL1-­S cer. PIB2 This work PGAL1 pRS315 PGAL1 This work PGAL1-­EGFP pRS315 PGAL1-­EGFP This work GFP-­PIB2 pRS316 GFP-­S cer. PIB2 This work PIB2 DN-­term pCM190 S cer. PIB2 D(1-­164) This work GTR1-­GFP pRS316 S cer. GTR1-­GFP This work GTR2-­GFP pRS316 S cer. GTR2-­GFP This work GTR1 Q65L pRS316 S cer. GTR1 Q65L This work GTR2 S23L pRS315 S cer. GTR2 S23L This work GFP-­TOR1 pRS316 GFP-­S cer. TOR1 This work TOR1 L2134M pRS426 S cer. TOR1 L2134M This work PAR32-­3xHA pRS316 S cer. PAR32-­3xHA This work GTR1-­GFP pRS316 S cer. GTR1-­GFP This work GTR2-­GFP pRS316 S cer. GTR2-­GFP This work Tet-­Off-­Pib2 pCM190 S cer. PIB2 This work GFP-­ATG8 GFP-­ATG8(416)/GFP-­AUT7(416) Addgene #49425

(Guan et al., 2001)

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Δgtr1

Recovery fromrapamycin

+ GTR1 Q65L

+ GTR2 S23L

+ GTR1 Q65L, GTR2 S23L

Untreated

Δgtr2+ GTR1 Q65L

+ GTR2 S23L

+ GTR1 Q65L, GTR2 S23L

Fig. S1. Rescue of growth on YPD following rapamycin exposure in Dgtr1 and Dgtr2

strains by expressing active forms of the Gtrs. Growth of Dgtr1 and Dgtr2 cells

expressing the indicated constructs on YPD during recovery from exposure to rapamycin.

Exponentially-­growing cells (OD600 0.6-­0.8) were treated with 200 ng/ml rapamycin in

YPD at 30 °C for 5 hr. After washing, cells were plated on YPD and were incubated for 3

days at 30 °C. The left-­most spot in each case corresponds to 2 µl of a culture with OD600

0.5. Spots to the right of this correspond to 2 µl of sequential 5-­fold dilutions.

J. Cell Sci. 130: doi:10.1242/jcs.207910: Supplementary information

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Gtr1-GFP

Gtr2-GFP

FM 4-64 Merge

FM 4-64 Merge

W303A

Δpib2

W303A

Δpib2

Fig. S2. Localization of Gtr1-­GFP and Gtr2-­GFP is unaltered in Dpib2 cells when

compared to W303A cells. The indicated strains expressed Gtr1-­GFP or Gtr2-­GFP from

their respective native promoters on centromeric plasmids. Cells were grown in Synthetic

Complete medium until they reached OD600 0.6-­0.8. Cells were then stained with FM 4-­

64 for 45 min, washed and chased in YPD for 1 hr prior to visualization. Scale – 5 µm.

J. Cell Sci. 130: doi:10.1242/jcs.207910: Supplementary information

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Recovery from rapamycinUntreated

W303A

Δpib2

Δgtr1 Δgtr2

+ pRS315

+ pRS316+ pRS315,+ pRS316

+ pRS315

+ pRS316+ pRS315,+ pRS316

+ pRS315

+ pRS316+ pRS315,+ pRS316

Pgk1

Phospho-Rps6

W303A Δpib2 Δgtr1 Δgtr2

Nitrogen starvation+ r

e-fee

ding,

30 m

in

+ re-f

eedin

g, 30

min

+ re-f

eedin

g, 30

min

Pgk1

Phospho-Rps6

Nitrogenstarvation, 3 hr

+ Gln, 3 mM

0 5 30

- + + +

Time, min

W303A+ PIB2 ΔN-term

Pgk1

Phospho-Rps6

Pgk1

Phospho-Rps6

Δpib2+ PIB2 ΔN-term

Δgtr1 Δgtr2+ PIB2 ΔN-term

Vacu

oles

with

Tor1

pun

cta,

%

0

20

40

60

80 ***

n.s.n.s.

+- +-+- + active GtrsW303A Δpib2 Δgtr1

Δgtr2

A

B

C D

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Fig. S3. (A) Vector-­only controls for recovery after exposure to rapamycin exhibit

no non-­specific effects. Growth of W303A, Dpib2 and Dgtr1 Dgtr2 cells expressing the

indicated vectors on YPD during recovery from exposure to rapamycin. Exponentially-­

growing cells (OD600 0.6-­0.8) were treated with 200 ng/ml rapamycin in YPD at 30 °C for

5 hr. After washing, cells were plated on YPD and were incubated for 2 days at 30 °C.

The left-­most spot in each case corresponds to 2 µl of a culture with OD600 0.5. Spots to

the right of this correspond to 2 µl of sequential 5-­fold d ilutions. ( B) Rps6

phosphorylation is stimulated by refeeding with complete dropout mix in W303A,

Dpib2 and Dgtr1 Dgtr2 cells. Phosphorylation levels of Rps6 were evaluated under the

indicated conditions. Untreated cells were grown in Synthetic Complete medium. Cells

were nitrogen-­starved by incubating in SD –N medium for 3 hr. For amino acid stimulation,

cells were treated with SD –N supplemented with 1x amino acid dropout mix (including

all amino acids, para amino-­benzoic acid, inositol and adenine) and were incubated for

30 min prior to lysis and processing. Pgk1 was used as a loading control. (C) Pib2 DN-­

term does not rescue TORC1 activation in Dgtr1 Dgtr2 cells. Phosphorylation levels

of Rps6 were evaluated under the indicated conditions. Cells expressing Pib2 DN-­term

were grown in Synthetic Complete medium prior to nitrogen-­starvation in SD –N medium

for 3 hr. For amino acid stimulation, cells were treated with SD –N supplemented with 3

mM glutamine for the indicated times prior to lysis and processing. Pgk1 was used as a

loading control. (D) Localization of Tor1 in Dpib2 is unchanged by the expression of

Gtr1 Q65L and Gtr2 S23L. Tor1 foci were quantified in W303A, Dpib2 and Dgtr1 Dgtr2

cells expressing Gtr1 Q65L and Gtr2 S23L (active Gtrs) as indicated. 206t 392 total

vacuoles were quantified in each case over 4 separate experiments.

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The significance of differences in numbers of vacuoles displaying foci for each strain

in the absence or presence of active Gtrs was determined using a twot tail to test with

6 degrees of freedom. Only the vacuoles in D gtr1 D gtr2 cells in the absence or

presence of active Gtrs displayed significantly different number of foci (t = 7.74 hence p =

0.0002).

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GFP-Tor1

W303A

FM 4-64 Merge

Δgtr1 Δgtr2

Δpib2

GFP-Tor1 FM 4-64 Merge

W303A

Δgtr1 Δgtr2

Δpib2

Vacu

oles

with

Tor1

pun

cta,

%

0

20

40

60

80 W303AΔgtr1 Δgtr2Δpib2

** ****

Vacu

oles

with

Tor1

pun

cta,

%

0

20

40

60

80 W303AΔgtr1 Δgtr2Δpib2

Nitrogen starvation, 3 hr

Rapamycin 200 ng/ml, 3 hr

A B

C D

Lorem ipsum

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Fig. S4. Localization of Tor1 is independent of TORC1 activity. (A) GFP-­Tor1

localization in W303A, Dgtr1 Dgtr2 and Dpib2 cells as indicated. The indicated strains

expressed GFP-­Tor1 from its native promoter on a centromeric plasmid. Cells were grown

in Synthetic Complete medium until they reached OD600 0.6-­0.8. W303A or the indicated

knockout strains were then stained with FM 4-­64 for 45 min, washed and chased in YPD

for 1 hr prior to starvation in SD –N medium for 3 hr. Scale – 5 µm. (B) Quantification of

the numbers of vacuoles displaying GFP-­Tor1 foci in each of the indicated strains. Foci

were counted on z-­stacks collected for each of the strains (~140-­190 vacuoles were

assessed for each strain). Means of numbers of vacuoles displaying foci were significantly

heterogeneous (one-­way ANOVA, F2,11 = 105.67;; p < 5.62E-­07). A post-­hoc Tukey HSD

test for significance was performed between each of the means. All significant differences

between means (**, p < 0.01) are indicated on the plot. (C) As in (A) but instead of nitrogen

starvation, cells were treated with rapamycin (200 ng/ml) for 3 hr prior to visualization. (D)

Quantification of the data shown in (C). Foci were counted on z-­stacks collected for each

of the strains (~150-­250 vacuoles were assessed in each strain). Means of numbers of

vacuoles displaying foci were not heterogeneous (one-­way ANOVA, F2,11 = 0.01;; p =

0.99).

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Untreated + rapamycin

W30

3A +

GFP-Atg8

W30

3A +

GFP-Atg8

Δpib2

+ GFP-A

tg8

Δpib2

+ GFP-A

tg8

GFP-Atg8

GFP

Pgk1

Fig. S5. Induction and flux of GFP-­Atg8 is unchanged in Dpib2 cells compared to

W303A. W303A or Dpib2 cells expressing GFP-­Atg8 were treated with rapamycin (200

ng/ml) for 3 hr before processing. GFP-­Atg8 and free GFP were detected using an anti-­

GFP polyclonal antibody.

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Mechanism of TORC1 Activation

A B C

TORC1

Gtr1/2Pib2

TORC1

Partial/intermediateactivation response

TORC1

Pib2 Gtr1/2

TORC1

Pib2 Gtr1/2

potentiation

Full activation response

TORC1

Gtr1/2Pib2

TORC1or

NO / LOW (below detection)activation response

Gtr1/2

Pib2

TORC1

NO activation responseif either Pib2 or Gtrs are absent

If both Pib2 and Gtrs are present,FULL activation response Full activation response

Hierarchical(upstream / downstream)

Parallel(threshold)

Independent

Partial/intermediateactivation response

NO / LOW (below detection)activation response

IndependentDependent integral

Fig. S6. Schematic illustrating possible models for the reactivation of TORC1 by Pib2 and Gtr1/2.

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