TFEB degradation is regulated by an IKK/β-TrCP2 phosphorylation-ubiquitination cascade

Domain: biology
Published: 2026-04-01
Editorial outcome: major_revision

Abstract
Transcription factor EB (TFEB) is a master regulator of lysosomal biogenesis and cellular clearance pathways. TFEB activity is tightly controlled by multiple post-translational mechanisms, but the exact molecular mechanism controlling its stability has remained elusive. Here, we identify the IκB kinase (IKK) complex as a key regulator of TFEB protein stability through a phosphorylation–ubiquitination cascade. A high-content kinase inhibitor screen reveals that IKK inhibition increases TFEB protein levels, and genetic ablation of IKK components increases TFEB stability, upregulates lysosomal genes, and enhances lysosomal biogenesis and degradative capacity. Mechanistically, we show that IKK phosphorylates TFEB on a cluster of serine residues (423SPFPSLS429), generating a phosphodegron recognized by the E3 ligase β-TrCP2, which in turn targets TFEB for proteasomal degradation via ubiquitination of adjacent lysine residues (K430 and K431). Mutation of either the phosphosites or the ubiquitination sites stabilizes TFEB without impairing its ability to translocate to the nucleus, activate target gene expression, or promote tau clearance in a cell model of tauopathy. These findings establish IKK–β-TrCP2 as a core regulatory axis controlling TFEB protein turnover and levels and reveal a mechanistically distinct layer of TFEB regulation that may be leveraged to enhance lysosomal function in disease contexts.

Introduction
Transcription factor EB (TFEB) is a master regulator of the autophagy-lysosomal pathway (ALP)1. Active TFEB promotes the transcription of lysosomal and autophagy genes1-3 and participates in the induction of ALP-dependent processes such as lysosome-mediated clearance1, lysosomal proteostasis4, and lysosomal exocytosis5. TFEB also contributes to other cellular programs, including the ER stress response4, 6 and mitochondrial biogenesis7, 8. TFEB plays a pivotal role in lysosome-to-nucleus communication by integrating inputs from mechanistic target of rapamycin complex 1 (mTORC1)9-13 and other key metabolic regulators, including protein kinase B (PKB/SAKT)14, 15 and AMP-activated protein kinase (AMPK)16, all of which have specific roles at the lysosome17-19. Importantly, these and other kinases phosphorylate TFEB on distinct sets of residu es9-16, 20-23, indicating that TFEB is controlled by multiple independent regulatory layers that likely execute different branches of the cellular adaptive response or serve additional functions. Studies in cultured cells have shown that, under standard growth conditions, TFEB is predominantly cytosolic and inactive. TFEB cytosolic retention is mediated by binding of YWHA/14-3-3 proteins to TFEB phosphorylated on Ser21112, 13. Ser211 is a direct target of active mTORC112, 13, which therefore limits the ALP-enhancing functions of TFEB. Conversely, lysosomal storage, cell starvation, and other stressors promote TFEB nuclear translocation1, 2, 6, 11-13, 24. In the nucleus, TFEB binds to the “coordinated lysosomal expression and regulation” (CLEAR) motif located within the promoter regions of genes participating in lysosomal pathways and promotes their transcription1.

Active TFEB and ALP enhancement promote cellular clearance by facilitating the elimination of protein aggregates and lysosomal storage material. Building on this principle, we and others have shown that TFEB is both a component of pathogenic cascades and a candidate therapeutic target in neurodegenerative storage disorders and beyond1, 4, 5, 14, 25-30. Multiple studies have demonstrated that exogenous TFEB expression reduces aggregated tau and β-amyloid in cell lines as well as in the brains of mouse models of Alzheimer's disease26, 28, 31, underscoring the significance of TFEB as a potential target for therapeutic intervention.

While best known for their roles in Nuclear Factor kappa B (NF-κB) signaling, components of the IκB kinase (IKK) complex have more recently emerged as NF-κB-independent regulators of processes as diverse as mTORC1 signaling32-34, apoptosis35, 36, cell proliferation and differentiation37, 38, and epigenetic control of gene transcription39-41, through direct phosphorylation of key players of these processes. The IKK complex is composed of three components: the partially redundant catalytic subunits IKKα (encoded by the CHUK gene) and IKKβ (IKBKB), and a scaffold/regulatory subunit IKKγ (IKBKG or NEMO)42-45. IKK has been linked to the mTOR pathway through two converging mechanisms. IKK was previously shown to inhibit Tuberous Sclerosis Complex 1 (T SC1)32, a key negative regulator of mTORC1 signaling46. Subsequent work demonstrated that IKK subunits can associate with and phosphorylate mTOR directly, promoting mTORC1 kinase activity33, 34. Thus, active IKK functions as a positive regulator of mTORC1 via at least two independent mechanisms. IKK can be activated by various stimuli, including TNFα signaling44. For several of its targets, IKK-dependent phosphorylation generates a phosphodegron—a specific motif recognized by β-transducin repeat-containing proteins (β-TrCP), the F-box substrate adaptors of SCFβ-TrCP E3 ubiquitin ligases47-50. The mammalian members of the β-TrCP family, β-TrCP1 and β-TrCP2, share sequence similarity and biochemical properties, but may have distinct substrates47. For example, β-TrCP2 has been characterized as the dominant paralog in regulating autophagy and cell growth through starvation-induced recognition and ubiquitination of β-TrCP1 and the mTORC1 inhibitors DEPTOR and REDD151.

In this study, we show that TFEB protein levels under standard culture conditions are markedly regulated by a phosphorylation-ubiquitination cascade executed by IKK and β-TrCP2. We find that IKK phosphorylates TFEB at multiple residues, generating a phosphodegron (423SPFPSLS429) that is recognized by β-TrCP2, which in turn promotes degradative ubiquitination of TFEB at adjacent lysine residues (K430 and K431). Selective genetic disruption of this phosphorylation-ubiquitination cascade increases TFEB protein levels several fold but does not interfere with signaling pathways that regulate TFEB nuclear translocation or its capacity to enhance expression of its target genes and promote cellular clearance. This mechanism provides a molecular explanation for how TFEB is targeted for proteasomal degradation and introduces a regulatory axis that may represent a promising therapeutic target in diseases characterized by impaired cellular clearance.

Results
IKK regulates TFEB protein levels
To efficiently screen for kinases that modify TFEB levels post-translationally, we generated a TFEB-GFP fusion construct as a reporter that is independent of transcriptional regulation of gene levels and does not require incubation with antibodies. We first tested this construct by transient transfection in HeLa cells, using fluorescence microscopy to monitor both TFEB protein levels and its subcellular localization. Calculation of a TFEB nuclear translocation index (TFEB ; see Methods for details) across hundreds of cells showed that TFEB nuclear localization levels were positively associated with TFEB protein levels (Supplementary Figs.1a, b), presumably because high levels of TFEB protein saturate the systems that promote its cytosolic retention. Thus, we generated cells stably expressing TFEB-GFP (henceforth referred to as HeLa/TFEB-GFP) by selecting a non-clonal pool of cells infected with a TFEB-GFP lentivirus. Fluorescence microscopy analysis showed that TFEB was generally expressed at lower levels compared to transiently transfected cells, and that nuclear localization levels were independent of TFEB expression levels in HeLa/TFEB-GFP cells (Supplementary Figs.1c, d), indicating the suitability of these cells for screening procedures.

To identify candidate kinases that regulate TFEB stability, we carried out a human kinome screen using a library of 436 established kinase inhibitors. For each inhibitor, we applied five different combinations of incubation time and drug concentration: cells were incubated with kinase inhibitors at either 1 or 3 µM for 1 hour, or at 1, 3, or 10 µM for 6 hours. We used automated fluorescence microscopy to acquire images of hundreds to thousands of cells per condition, which we then analyzed to assess TFEB-GFP signal levels in whole cells, cytosol, and nuclei. In each plate, Torin1 (a potent catalytic inhibitor of mTORC1) was used as a positive control for its ability to alter cytosolic and nuclear TFEB levels, while DMSO was used as a vehicle control. To assess the robustness of the pipeline, we integrated the microscopy data from the five conditions to perform unsupervised hierarchical clustering of the drugs based on TFEB NTI (Supplementary Data 1). The analysis showed that the rapalogs (allosteric mTORC1 inhibitors) present in the library clustered with Torin1 (Fig. 1a, red arrow). The unsupervised hierarchical clustering procedure is effect size-independent and leverages all changes observed across the five data points to make associations amongst drugs that alter, even minimally, TFEB distribution in similar patterns. As rapalogs have been characterized as very weak modulators of TFEB nuclear translocation at best11, 12, 52, 53, yet still clustered with Torin1 (a potent activator of TFEB nuclear translocation), this result underscores the high sensitivity and robustness of the analytical pipeline.

The vast majority of the kinase inhibitors screened clustered with DMSO, indicating little to no effect on TFEB levels in the cytosol or nucleus. Interestingly, a small cluster of kinase inhibitors that increased TFEB nuclear signal (indicated by the green arrow in Fig. 1a) was distinct from the cluster of mTORC1 inhibitors, suggesting the presence of additional mechanisms contributing to changes in cytosolic and/or nuclear TFEB levels. A scatterplot of total TFEB-GFP intensity versus the product of the distances from Torin1 and vehicle in the hierarchical cluster highlighted a small group of kinases whose inhibition simultaneously increased both TFEB nuclear intensity and total TFEB intensity—IKK, Axl, JAK2, and CDK4/6 (Fig. 1b). CDK4/6 is a known kinase modifier of TFEB23, and both Axl and JAK2 are general receptor tyrosine kinases that modulate the PI3K/Akt/mTOR pathway, among others54, 55. IKK has multiple strong connections to mTORC1 signaling32-34 but has not been studied in the context of TFEB biology. Thus, we chose to focus on IKK for functional studies.

To test whether endogenous TFEB levels are regulated by IKK, we monitored TFEB protein levels in mouse embryonic fibroblast (MEF) lines in which the Chuk/IKKα, Ikbkb/IKKβ, or Nemo/IKKγ genes had been knocked out56-58. Immunoblot analyses of total cell lysates showed that deletion of any IKK subunit resulted in increased TFEB protein levels (Figs. 1c, d), confirming IKK’s control over TFEB protein levels. As a control, we monitored IκBα (inhibitor of nuclear factor kappa B), a downstream target of IKK. Knockout of either IKKβ or IKKγ increased IκBα levels (Figs. 1c, d), indicating that IKK maintains basal activity under the test conditions, even in the absence of external stimuli. RT-qPCR analysis showed that TFEB mRNA levels were not increased by deletion of either IKK subunit (Fig. 1e), indicating that the effect of IKK on TFEB must be post-translational.

Cell fractionation followed by immunoblotting showed that deletion of IKK subunits did not alter TFEB subcellular distribution (Supplementary Figs. 2a, b), a result confirmed by siRNA-mediated silencing of all IKK subunits in HeLa/TFEB-GFP cells (Supplementary Figs. 2c, d). Additional experiments clarified that IKK-16, the IKK inhibitor used in the screen, also inhibits mTORC1 in HeLa/TFEB-GFP (Supplementary Fig. 2e), which could be an off-target effect. We verified that, when mTORC1 is rendered constitutively active by expression of modified RagB and RagD constructs, IKK-16-induced nuclear localization is indeed abolished, demonstrating that this effect is mediated by mTORC1 (Supplementary Figs. 2f, h). Conversely, constitutively active mTORC1 did not antagonize the observed increases in TFEB signal intensity (Supplementary Figs. 2f, g-i), and Torin1, differently from IKK-16, was unable to increase TFEB levels (Supplementary Fig. 2j), indicating that IKK-16–induced increases in TFEB levels are not mediated by mTORC1 inhibition. Taken together, these data demonstrate that IKK deletion or inhibition increases TFEB levels through a mechanism that is uncoupled from mTORC1-mediated control of TFEB nuclear translocation. We finally tested whether the IKK-16–induced increase in TFEB expression requires IKK. To this aim, we treated WT and IKK-knockout MEF lines with DMSO or IKK-16 and evaluated TFEB levels by immunoblotting. The results showed that IKK-16 increased TFEB levels in WT cells but failed to do so in any of the IKK-knockout lines (Supplementary Figs. 2k-l), demonstrating that the observed effect of IKK-16 on TFEB protein levels is mediated by IKK.

Next, we tested whether IKK deletion caused any changes in lysosomal biogenesis and function. Staining the MEF lines with an antibody against LAMP1 (lysosomal-associated membrane protein 1), a lysosomal marker, showed that knockout of any IKK subunit resulted in a significant expansion of lysosomal compartments, indicating increased lysosomal biogenesis (Figs. 1f-h). We also carried out an analysis of lysosomal proteolytic function using DQ BSA, a self-quenched albumin probe that fluoresces upon lysosomal degradation. Cells were loaded with DQ BSA for 16 h before fixation and imaging. Torin 1 was used as a positive control of activation of lysosomal proteolytic function. Microscopy experiments showed that the knockout MEF lines exhibited enhanced DQ BSA fluorescence compared to WT MEFs (Figs. 1i-j). Together, these data show that knockout of IKK expands both the lysosomal compartment and its degradative capacity. Acute suppression of IKK function in HEK293T cells (Figs. 1k-m) and HeLa cells (Supplementary Figs. 3a-c) through siRNA-mediated silencing of any IKK subunit resulted in transcriptional upregulation of TFEB target genes, consistent with a role for IKK in suppressing TFEB levels.

IKK stimulation by TNFα or IL-1β in MEF cells resulted in a rapid decrease in TFEB protein levels, which was not observed in any of the IKK gene knockout lines (Figs. 2a-f). We also tested lipopolysaccharide (LPS), an activator of inflammatory responses previously shown to reduce TFEB levels by unknown mechanisms21, 59. The results showed that LPS also requires IKK proteins to reduce TFEB protein levels (Figs. 2g-i). To assess whether chronic LPS-mediated suppression of TFEB impacts lysosomal biogenesis, we treated WT and IKK-knockout MEF lines with LPS for 72 h prior to fixation and imaging. Confocal imaging showed that prolonged LPS exposure led to a marked reduction of lysosomal compartments in WT cells, whereas no such reduction was observed in any IKK-knockout line (Figs. 2j-k). Collectively, these results demonstrate that IKK activity controls TFEB stability and the abundance of lysosomal compartments under both basal and stimulated conditions.

IKK phosphorylates TFEB at multiple serine residues
IKKα and IKKβ are structurally similar, have partially redundant functions, and have been reported to show some degree of substate selectivity45, 60, 61. We tested whether IKKα and IKKβ phosphorylate TFEB by performing phos-tag analyses. In phos-tag gels, the rate of protein migration depends on the number of phosphorylated residues, and the presence of multiple bands indicates multiple phosphorylation sites on the same protein62. 3xFlag-tagged TFEB was transiently expressed in HeLa cells, immunoprecipitated, and dephosphorylated in vitro using 𝜆 phosphatase before being incubated with commercially available IKKα or IKKβ for phos-tag analysis. The results showed that both IKKα and IKKβ phosphorylated TFEB, with IKKβ displaying higher efficacy and generating multiple phosphorylated bands (Fig. 3a). As a control, in vitro phosphorylation of TFEB by Akt using the same procedure produced a single phosphorylated band, which was completely abolished by alanine substitution of serine 467, as expected14 (Supplementary Fig. 4a). Thus, the presence of multiple phosphorylated bands after incubation with IKKβ indicates that IKKβ mediates multiple phosphorylation events on TFEB. Of note, the reconstituted canonical IKK complex displayed higher phosphorylation efficiency than IKKβ alone (Fig. 3b). This result indicates that the IKK complex as a whole mediates TFEB regulation, consistent with the data from the knock-out lines and gene silencing experiments shown above.

In vitro peptide screens to define IKKα/β substrate preferences, along with the identification of several bona fide substrates, have shown that IKKα and IKKβ do not have strict requirements for the amino acids surrounding their target serine, making prediction of candidate substrate sites difficult61, 63, 64. To identify the TFEB residues phosphorylated by IKK, we performed experiments of mass spectrometry coupled with mutagenesis. For simplicity and owing to its ability to phosphorylate TFEB in vitro even in absence of the other IKK subunits, IKKβ was used in these experiments. Mass-spectrometry analyses of intact TFEB showed abundant phosphorylation localized to serine 427, supported by multiple fragment ions, including fragments containing intact phosphoserine (Figs. 3c-d). Specifically, we detected an intact phosphorylated peptide spanning residues 405-430, with an m/z 881.7311 (-1.99 ppm error; expected mass, 881.732851 m/z) (Supplementary Fig. 4b). MS2 analysis (where precursor peptide ions are fragmented and analyzed to determine sequence and PTM localization) produced multiple ion types supporting phosphorylation at S427. In phos-tag analyses, alanine substitution of S427 altered the pattern of phospho-TFEB bands and reduced the phosphorylation efficiency of TFEB, resulting in higher levels of unphosphorylated TFEB-S427A compared to wild-type (WT) TFEB (Fig. 3e). To narrow down the additional sites targeted by IKK, we first tested S3—the only serine present within the N-terminal domain of TFEB—and found that alanine substitution did not affect the TFEB phosphorylation pattern (Fig. 3f). Therefore, we excluded S3 as a candidate site. We then separately analyzed the phosphorylation of TFEB fragments spanning residues 53-218 and 219-476 (Supplementary Fig. 4c). The results showed that TFEB (53-218) was not phosphorylated by IKKβ, while TFEB (219-476) exhibited multiple phosphorylation bands (Fig. 3f). These findings indicate that the additional IKK target sites are also located within the 219-476 region.

To identify these sites, we generated a library of TFEB mutants containing single alanine substitutions for each of the 24 serine residues present within the 219-476 region, and evaluated the phosphorylation status of each mutant by in vitro kinase assay. The results showed that alanine substitution of S401 and S429 also altered the phosphorylation pattern of TFEB (Supplementary Fig. 4d). The S401A/S427A double mutant exhibited a significant reduction in phosphorylation efficiency compared to WT TFEB, although a small fraction of phosphorylated protein was still detected (Fig. 3g). Alanine substitution of four additional adjacent serine residues (S397, S399, S423, and S429), together with S401 and S427 (henceforth referred to as TFEB-6xA), completely abolished TFEB phosphorylation by IKKβ (Fig. 3g) or by the reconstituted IKK complex (Supplementary Fig. 4e)

In parallel, we generated non-clonal pools of HeLa cells stably expressing 3xFlag-tagged WT TFEB (henceforth referred to as HeLa/TFEB). We immunoprecipitated TFEB under basal conditions or after a 15-min stimulus with 20 ng/mL IL-1β and proceeded with mass spectrometry analysis. The results showed abundant monophosphorylation of TFEB at the SPFPSLS motif within the 405AA-431AA peptide after a 15-min stimulus with IL-1β (Figs. 3h-j). MSFragger65 localized this phosphorylation event to S423 with high confidence. However, manual inspection revealed that while S423 may be the most likely single site of localization, phosphorylation at S427 or S429 is not excluded by the same evidence, and both sites also show weaker but independent supporting evidence, consistent with the mutational analyses described above. Under basal conditions, the signal for the phosphorylated 405–431 aa peptide was below the detection limit. Together, the data from TFEB phosphorylation assays in vitro and in intact cells indicate phosphorylation events at neighboring serines clustered in two small regions. Interestingly, these six sites form two similar modules, each approximately the size of a typical kinase interaction surface: S397/399/401 (FSHSLSF) and S423/427/429 (SPFPSLS). Thus, we used TFEB-6xA, which abolishes all IKK-mediated phosphorylation of TFEB, to carry out functional analyses of this pathway.

IKK phosphorylation-resistant TFEB displays increased expression levels and stability.
To investigate the functional consequences of abolishing IKK-mediated TFEB phosphorylation on TFEB stability, we generated non-clonal pools of HeLa cells stably expressing 3xFlag-tagged TFEB-6xA (henceforth referred to as HeLa/TFEB-6xA). Immunoblot analyses showed that HeLa/TFEB-6xA displayed substantially higher TFEB levels than HeLa/TFEB (Figs. 4a, b), despite no significant differences in RNA expression levels between the two constructs (Fig. 4c). To test whether this effect was due to differences in protein stability, we used cycloheximide to block protein synthesis and measured degradation rates for each TFEB construct. The results showed that WT TFEB levels rapidly declined following inhibition of protein synthesis, whereas TFEB-6xA levels remained stable throughout the monitored time frame (Figs. 4d, e). These data indicate that loss of IKK phosphorylation sites increases TFEB stability. We next examined the stability of WT and 6xA TFEB under TNFα-induced activation of IKK. WT TFEB levels were reduced by 30 min of TNFα treatment, while TFEB-6xA levels remained unchanged (Figs. 4f, g). Together, these findings demonstrate that abolishment of IKK target sites increases TFEB protein expression levels and stability, indicating that IKK phosphorylation of TFEB promotes TFEB degradation.

β-TrCP2 mediates TFEB ubiquitination and stability downstream of IKK phosphorylation
IKK-mediated phosphorylation primes certain protein substrates for degradative ubiquitination by generating a phosphodegron recognized by the E3 ubiquitin ligases β-TrCP1/248-50. To investigate whether β-TrCPs mediate TFEB ubiquitination, we assessed TFEB ubiquitination levels upon exogenous expression, in HEK293T cells, of β-TrCP1, β-TrCP2, or the E3 ubiquitin ligases STUB1 (STIP1 homology and U‐Box containing protein 1) and DCAF7 (DDB1 and CUL4 associated factor 7), which had previously been associated with TFEB degradation66, 67. Coexpression with HA-tagged ubiquitin and TFEB-3xFlag followed by TFEB immunoprecipitation and HA immunoblotting showed that, β-TrCP2—but not β-TrCP1—promoted TFEB ubiquitination (Fig. 4h). Next, we silenced β-TrCP2 to test whether it mediates TFEB ubiquitination. Inhibition of proteasomal degradation using low concentrations of carfilzomib (CFZ) resulted in accumulation of ubiquitinated TFEB, which was completely prevented by β-TrCP2 (FBXW11) silencing (Fig. 4i and Supplementary Fig. 5a). These data show that β-TrCP2 functions as a ‘default’ TFEB ubiquitin ligase and indicate that TFEB tagging for proteasomal degradation is a continuously occurring process in unstimulated, standard cell culture conditions. Due to the unavailability of suitable antibodies against endogenous β-TrCP2, silencing efficiency was evaluated by monitoring endogenous FBXW11 mRNA levels (Supplementary Fig. 5a) and the protein levels of a HA-β-TrCP2 construct transiently transfected into HEK293T cells (Supplementary Fig. 5b).

Silencing of β-TrCP2 in unstimulated HEK293T cells resulted in a significant increase in endogenous TFEB levels (Figs. 5a-c), similar to the silencing of IKK subunits (Supplementary Figs. 6a-c), indicating that IKK and β-TrCP2 participate in TFEB turnover in multiple cell types. In contrast to TFEB protein levels, TFEB mRNA levels remained unchanged upon β-TrCP2 or IKK subunit silencing (Fig. 5d and Supplementary Fig. 6d), confirming that the IKK–β-TrCP2 axis affects TFEB post-transcriptionally. In contrast, DCAF7 silencing did not affect endogenous TFEB levels under the same conditions (Supplementary Figs. 6e-g). Similarly, knockdown of β-TrCP2 in WT MEFs significantly increased endogenous TFEB levels, an effect that did not occur in any of the IKK-knockout lines (Figs. 5e-g). These results demonstrate that β-TrCP2–mediated regulation of TFEB occurs downstream of IKK and that the IKK–β-TrCP2 axis regulates TFEB turnover in multiple cell types.

Next, we investigated mechanistically whether β-TrCP2-mediated modulation of TFEB levels requires the IKK-targeted phosphodegron by testing the effects of β-TrCP2 silencing on the IKK-resistant TFEB-6xA construct. To enable comparison with WT TFEB, we silenced β-TrCP2 in both HeLa/TFEB and HeLa/TFEB-6xA. The results showed that β-TrCP2 silencing nearly doubled WT TFEB levels in standard cell culture conditions, while having only a minimal effect on TFEB-6xA levels (Figs. 5h-j). We next tested whether IKK-mediated phosphorylation of TFEB is necessary for its ubiquitination by β-TrCP2. To this aim, we performed sequential transfections by first introducing either WT TFEB or TFEB-6xA constructs along with HA-tagged ubiquitin, and subsequently transfecting the cells with siRNAs targeting FBXW11 or scrambled controls. Immunoprecipitation of TFEB followed by HA immunoblotting showed that IKK-resistant TFEB-6xA exhibited markedly lower levels of ubiquitination compared to WT TFEB (Fig. 5k). As expected, β-TrCP2 silencing abated the ubiquitination of WT TFEB but had no effect on TFEB-6xA (Fig. 5k and Supplementary Fig. 5c). Remarkably, TFEB-6xA treated with scramble siRNAs displayed ubiquitination levels similar to those of WT TFEB upon β-TrCP2 silencing, indicating that removal of IKK-targeted phosphorylation sites from TFEB mimics a lack of engagement of β-TrCP2. Furthermore, β-TrCP2 silencing abrogated TNFα-induced degradation of TFEB (Figs. 5l-m) as well as IKK-16-induced increase in TFEB levels (Figs. 5n-o), further demonstrating that IKK regulates TFEB degradation in a β-TrCP2–dependent manner.

Collectively, these findings demonstrate that β-TrCP2 ubiquitinates TFEB downstream of IKK phosphorylation in both unstimulated and stimulated cells, presumably to maintain tight control of TFEB levels.

TFEB ubiquitination and degradation is mediated by phosphodegron-adjacent lysines.
β-TrCP proteins interact with their substrates via a phosphodegron that contains two phosphorylated serine residues separated by three or four amino acids [pSxxx(x)pS]68. The canonical phosphodegron targeted by β-TrCP has a DpSGxx(x)pS consensus sequence68, but β-TrCP also interacts with non-canonical phosphodegrons featuring different amino acids around the two phosphoserines69-71. Lysine residues that undergo ubiquitination are typically located in proximity to their cognate phosphodegrons72-74. Although TFEB does not contain the canonical β-TrCP phosphodegron, serines 423 and 427 are separated by three amino acids (423SPFPS427) and lie adjacent to two consecutive lysines, K430 and K431 (Fig. 6a). Mass-spec analysis confirmed ubiquitination at K430 and K431 in cells expressing TFEB-3×Flag and ubiquitin (Supplementary Figs. 7a-e). To test whether these lysines are the sites of β-TrCP-mediated ubiquitination, we generated HeLa cells stably expressing a TFEB mutant in which both lysines were replaced by arginine residues (HeLa/TFEB-KKRR). Notably, compared with HeLa/TFEB, HeLa/TFEB-KKRR showed markedly elevated TFEB expression (Supplementary Figs. 8a-b), despite no differences in mRNA levels between the two transgenes (Supplementary Fig. 8c). Thus, similar to the IKK-resistant TFEB-6xA construct, the increase in TFEB-KKRR protein levels must result from a post-translational mechanism.

A cycloheximide chase assay demonstrated that mutation of the phosphodegron-adjacent lysines increased TFEB protein stability (Supplementary Fig. 8d), again mimicking the effect of mutating the serines within TFEB’s phosphodegron (see Fig. 4d). A comparison of protein levels from WT TFEB, TFEB-6xA, TFEB-KKRR, and a phospho-mimetic TFEB-6xE mutant showed that 6xA and KKRR protein levels were similar to each other, whereas the levels of the phospho-mimetic mutant were comparable to those of WT TFEB—albeit slightly increased, presumably because the glutamic acid residues do not fully mimic phosphorylation (Supplementary Fig. 8e). IKK inhibition increased the levels of WT TFEB but had no effect on any of the mutants, consistent with the conclusion that IKK acts through the identified phosphorylation and ubiquitination sites (Supplementary Figs. 8e, f).

Notably, TFEB-KKRR was insensitive to β-TrCP2 silencing (Figs. 6b-d), mimicking the behavior of TFEB-6xA. Overexpression of β-TrCP2 decreased TFEB protein levels, and this effect was abolished by arginine substitution of the two lysines (Figs. 6e-f). In contrast, overexpression of DCAF7 had no effect on the protein levels of either construct (Supplementary Figure 9a-b). A ubiquitination assay showed that arginine substitution also reduced TFEB ubiquitination to levels comparable to those observed for WT TFEB upon β-TrCP2 silencing (Figs. 6g-h). Furthermore, neither silencing nor overexpression of β-TrCP2 altered the ubiquitination levels of TFEB-KKRR (Figs. 6g-i), confirming that β-TrCP2-mediated ubiquitination of TFEB requires these phosphodegron-adjacent lysines. In comparison, arginine substitution of the two lysines did not affect DCAF7-induced changes in TFEB ubiquitination levels (Supplementary Fig. 9c). Of note, both TFEB-6xA and TFEB-KKRR exhibited minimal ubiquitination upon proteasome blockade (Supplementary Fig. 10), indicating that the IKK–β-TrCP2-mediated phosphorylation-ubiquitination cascade is the primary pathway regulating TFEB proteasomal degradation.

To directly link IKK-mediated phosphorylation of TFEB to regulation of its stability via the phosphodegron-adjacent lysines, we monitored the protein levels of WT TFEB and TFEB-KKRR following siRNA-mediated silencing of IKK. Immunoblotting showed WT TFEB levels increased upon silencing of IKKγ and IKKβ in unstimulated cells, whereas TFEB-KKRR levels remained unchanged (Figs. 7a-b; Supplementary Figs. 8g-h). IKK activation by IL-1β induced a rapid decrease in WT TFEB protein levels, an effect that was not observed for either TFEB-6xA or TFEB-KKRR (Figs. 7c-d). Furthermore, the IL-1β–induced acute decrease in TFEB levels was abolished by proteasomal inhibition, indicating that IL-1β promotes TFEB degradation through the proteasome (Figs. 7e-f). Similarly, LPS treatment reduced WT TFEB levels but had no effect on TFEB-6xA or TFEB-KKRR (Figs. 7g-h). It is worth noting that LPS treatment also decreased IκBα levels (Fig. 7i), confirming IKK activation. These results indicate that IL-1β and LPS-induced degradation of TFEB require the IKK-targeted phosphodegron.

Together, these results show that TFEB levels are regulated by a phosphorylation-ubiquitination cascade mediated by IKK and β-TrCP2 under both basal and IKK-activating conditions.

Inactivation of TFEB degron does not interfere with TFEB nuclear translocation and activation of clearance pathways
Next, we investigated whether degron-dependent modifications of TFEB affect its nuclear translocation or its ability to modulate clearance pathways. To this aim, we first tested whether TFEB-6xA and TFEB-KKRR remain responsive to starvation and mTORC1 inhibition, both of which are known to promote TFEB nuclear translocation. Confocal microscopy of HeLa/TFEB, HeLa/TFEB-6xA, and HeLa/TFEB-KKRR cells showed that inactivation of either component of the TFEB degron did not alter its responsiveness to starvation or mTORC1 inhibition (Figs. 8a, b)—a result confirmed by immunoblotting following subcellular fractionation (Fig. 8c).

Next, we tested the ability of the three constructs to promote transcription of TFEB target genes. To allow a fair comparison, we did not use the stable cell lines, as they express TFEB proteins at markedly different levels despite similar mRNA levels, as shown above. We noticed that transiently expressed WT TFEB, TFEB-6xA, and TFEB-KKRR exhibited similar protein levels when standard amounts of plasmids were transfected (Fig. 8d), whereas the protein levels of the IKK- and β-TrCP2-insensitive TFEB mutants were far greater than those of WT TFEB when lower amounts of plasmid were used (Supplementary Figs. 11a-b). This difference likely reflects saturation of IKK /β-TrCP2 or their associated machineries under conditions of high protein expression, as standard transient transfection results in higher TFEB levels than those achieved in stable clones (see Supplementary Fig. 1). Thus, to compare the transcriptional responses of the three TFEB constructs, we transfected standard amounts of plasmids. RT-qPCR analysis of RNA from transiently transfected cells showed that all three constructs similarly enhanced the transcription of TFEB target genes, with TFEB-KKRR exhibiting a slightly stronger effect (Figs. 8d, e).

We have previously shown that exogenous TFEB expression enhances tau fibril uptake and reduces tau spreading in cell lines and mouse models of Alzheimer’s disease28, 31. To assess the potential of phosphorylation/ubiquitination-resistant TFEB constructs to attenuate tauopathy, we cotransfected HEK293T cells with a V5-tagged tau-P301L construct along with WT TFEB or either TFEB mutant construct. Quantification of total tau and phospho-tau (PHF1) levels under comparable expression levels of the three TFEB constructs showed similar overall efficiencies in promoting the degradation of both total tau and phospho-tau, with the two TFEB mutants being slightly more effective at reducing total tau levels (Figs. 8f, g). Taken together, these results indicate that the TFEB modifications involved in the phosphorylation-ubiquitination cascade are functionally distinct from those that control TFEB nuclear translocation, transcriptional activity, and clearance function.

Methods
Eukaryotic cell lines. HeLa cells (female; human cervical carcinoma; ATCC, CCL-2) and HEK293T cells (female; human embryonic kidney; ATCC, CRL-3216) were purchased from ATCC. Wildtype, IKKα−/−, IKKβ−/− and IKKγ–/–MEFs were kindly provided by Dr. Michael Karin’s laboratory (University of California at San Diego), the sex of these MEF lines was not determined. HeLa/TFEB-GFP, HeLa/TFEB, HeLa/TFEB-6xA and HeLa/TFEB-KKRR stable cell lines were generated from parental HeLa cells using the corresponding expression constructs following standard stable cell line generation protocols. MEF/TFEB, MEF/TFEB-6xA and MEF/TFEB-KKRR stable cell lines were generated analogously from parental WT MEFs.

Cell Culture. HeLa and HEK were cultured in DMEM (Cat# 10013CV, Corning™), supplemented with 10% inactivated FBS (Cat# 16140071, Gibco), 2 mM L-glutamine (Cat# 25030081, Life Technologies), and Penicillin/streptomycin (100 U/mL) (Cat# 15140122, Life Technologies). MEFs were cultured in DMEM supplemented with 15% inactivated FBS, 2 mM L-glutamine and Penicillin/streptomycin (100 U/mL). All cells were maintained at 37°C and 5% CO2.

Lentivirus infection and stable cell lines. HEK293T Cells were plated at a density of 3.5 × 106 per 60-mm diameter dish 18 h prior to transfection. HEK293T packaging cells were cotransfected with pLenti expression construct, packaging vector psPAX2 (Addgene#12260) and envelope vector pMD2.G (Addgene #12259) using Lipofectamine™ 3000 Transfection Reagent (Cat# L3000001, Thermo Fisher Scientific). The lentiviruses were harvested 52hr after transfection. Virus particles were purified by centrifugation at 2,000 × g for 10 minutes, followed by filtration through a 0.22 μm pore size syringe filter. For viral transduction, HeLa cells and MEFs with C57BL/6 background (Dr. Mark Sands’ laboratory, Washington University in St. Louis) were seeded at 50–60% confluence and treated overnight with the medium containing freshly harvested lentivirus in the presence of 8 μg/mL polybrene (Cat# TR-1003-G, MilliporeSigma). Selection of stable clones was carried out using puromycin (Cat# A1113802, Thermo Fisher Scientific).

Antibodies. Primary antibodies: Antibodies to TFEB (E5P9M) (Cat# 83010S) (1:1000), Flag (D6W5B) (Cat# 14793) (1:1000), β-actin (Cat# 3700S) (1:20,000), HA (C29F4) (Cat# 3724S) (1:1000), α-tubulin (Cat# 2144S) (1:2000), Histone H3 (Cat #9715S) (1:30,000), IKKγ Kinase (DA10-12) (Cat# 2695T) (1:1000), Phospho-S6 Ribosomal Protein (Ser240/244) (Cat# 2215S) (1:10,000), S6 Ribosomal Protein (5G10) (Cat# 2217S) (1:2000), 4E-BP1 (53H11) Rabbit Monoclonal Antibody (Cat #9644) (1:1000) and phospho-4E-BP1 (Ser65) Antibody (Cat #9451S) (1:1000) were from Cell Signaling Technology; antibodies to IKKβ Kinase (Cat# ab124957) (1:5000), IKKα Kinase (Cat# ab32041) (1:5000), IKKγ Kinase [EPR16629] (Cat# ab178872) (1:5000) and WD R68 (Cat# ab138490) (1:5000) were from Abcam; antibodies to GAPDH (Cat# 60004-1-Ig) (1:150,000) and IκBα (Cat# 10268-1-AP) (1:2000) were from Proteintech; antibodies to Flag (produced in mouse) (Cat# F1804) (1:5000), Flag (produced in Rabbit) (Cat# F7425) (1:2000) and c-Myc (Cat# C3956) (1:1000) were from Sigma-Aldrich; antibodies to TrCP1 (C-6) (Cat# sc-390629) (1:1000), STUB1 (Cat# sc-133066) (1:1000) and Lamp1 (Cat# sc-19992) (1:1000) were from Santa Cruz; antibodies to V5 (Cat# MCA1360GA) (1:1000) was from Bio-Rad; antibodies to PHF1 antibody (1:1000) was a gift from Dr. Peter Davies (Feinstein Institute for Medical Research).

Secondary antibodies: Goat anti-mouse HRP-conjugated antibody (Cat# 1705047) (1:2000) and Goat anti-rabbit HRP-conjugated antibody (Cat# 1705046) (1:2000) were from Biorad, IRDye® 680RD Goat anti-Mouse IgG Secondary Antibody (Cat #926-68070) (1:10000) and IRDye® 800CW Goat anti-Rabbit IgG Secondary Antibody (Cat #926-32211) (1:10000) were from LI-COR Biosciences, Goat anti-mouse antibody, Alexa Fluor™ 555 (Cat# A-21422) (1:400) and Goat anti-mouse antibody Alexa Fluor™ 488 (Cat# A-11001) (1:400) were from Invitrogen.

Reagents. Torin 1 (Cat# 424710) was from Tocris; DMSO (Cat# D8418), Tumor Necrosis Factor-α (TNFα) (Cat# H8916), cycloheximide (Cat# 01810-1G) and lipopolysaccharide (LPS) from Escherichia coli O111:B4 (Cat# L4391-1MG) were from Sigma-Aldrich; carfilzomib (Cat# 17554) was from Cayman Chemical; Protease Inhibitor Cocktail (Cat# P3100-010) and Phosphotase Inhibitor Cocktail (Cat# P3200-010) were from Gendepot. Opti-MEM (Cat# 31985070) and HBSS (Cat# 14025092) were from Gibco. Recombinant mouse IL-1β/IL-1F2 protein (Cat# 401-ML-005/CF), recombinant mouse TNFα protein (Cat# 410-MT-010), recombinant human TNFα protein (Cat# 210-TA-005) and recombinant human IL-1β/IL-1F2 protein (Cat# 201-LB-005/CF) were from R&D systems. DQ Green BSA (Cat #D12050) was from Life Technologies.

Primers. RT–qPCR primers were obtained from published studies or designed using the PrimerQuest™ Tool (Integrated DNA Technologies, IDT) based on reference gene sequences. Primer specificity was verified using NCBI Primer-BLAST. Primers for In-Fusion cloning were designed using the In-Fusion Cloning Primer Design Tool (Takara Bio), and mutagenesis primers were designed using NEBaseChanger (New England Biolabs). All oligonucleotides were synthesized by MilliporeSigma. Sequences of all primers are provided in Supplementary Data 2.

Plasmids. CMV-TFEB-3×Flag: Human TFEB was expressed from a pcDNA3.1(+)-based mammalian expression vector containing a C-terminal 3×Flag tag under control of the CMV promoter. The vector confers ampicillin resistance for bacterial selection and neomycin resistance for mammalian selection. pCDH-EF1-TFEB-GFP: A third-generation self-inactivating lentiviral transfer vector containing the EF-1α promoter (with intron A) was used to express human TFEB as a C-terminal Myc–EGFP fusion. CMV-TFEB (aa53–218)-3×Flag, CMV-TFEB (aa219–476)-3×Flag, Lenti-TFEB-3×Flag, β-TrCP2, DCAF7, and STUB1 were synthesized by Twist Bioscience. TFEB_6xA-3×Flag and TFEB_6xE-3×Flag were synthesized by Twist Bioscience and cloned into a pcDNA3.1-based expression vector using In-Fusion® Snap Assembly Master Mix (Cat# 638948, Takara Bio). Myc-β-TrCP2, Lenti-TFEB-6xA-3×Flag, and Lenti-TFEB-KKRR-3×Flag were generated using In-Fusion® Snap Assembly Master Mix (Cat# 638948, Takara Bio). Other TFEB mutants were generated from the CMV-WT-TFEB-3×Flag construct using the Q5® Site-Directed Mutagenesis Kit (Cat# E0554, New England Biolabs) according to the manufacturer’s instructions. All plasmids were verified by Sanger sequencing prior to use. Detailed information on all plasmids is provided in Supplementary Data 2.

Human kinome screen. All drug treatment, washing, fixing and Hoechst staining steps were performed using the SAMI-integrated automated screening system (Beckman Coulter). HeLa/TFEB-GFP were seeded at a density of 7,000 cells per well in 96-well plates using a Multidrop and incubated at 37°C in a humidified 5% CO incubator for 12 hours prior to treatment. On each plate, four wells were seeded with parental HeLa cells for imaging background controls, eight wells of cells were treated with 0.1% DMSO as drug vehicle controls, and four wells of cells were treated with 250 nM Torin1 as nuclear translocation positive controls. For the remaining wells, each well was treated with one of the 436 drugs within the L1200 library from SelleckChem. Each drug was applied under five different conditions: 10 μM for 6 hours, 3 μM for 6 hours, 1 μM for 6 hours, 10 μM for 1 hour, and 3 μM for 1 hour. After treatment, cells were fixed with 4% paraformaldehyde at room temperature for 20 minutes and incubated with 2 μM Hoechst 33342 overnight at 4°C. The cells were washed with PBS, and the plates were sealed. Imaging was performed using the IN Cell 2000 Analyzer (GE Healthcare Life Sciences) with FITC (for TFEB-GFP) and DAPI (for nuclei) channels at 20X magnification. Sixteen fields were captured for each well. The acquired images were analyzed with the IN Cell 1000 Workstation analyzer software. DAPI channel images were used to segment the nuclei and identify individual cells for TFEB-GFP intensity measurements. Intensity analysis was conducted by measuring the average intensity of the whole cell. Drugs that yielded intensities greater than the average intensity for all 436 drugs plus four times the standard deviation were considered as outliers and were excluded from further analysis. The Multi Target Analysis Module was used to measure the average nuclear intensity (N) and the average intensity within a 2 µm collar (C) surrounding the nucleus, as the cytoplasmic sampling region.

Kinome Data analysis. TFEB nuclear translocation index (NT index) was calculated as N/(N+C), which ranges from 0 to 1, and was divided into 100 equal intervals. The frequencies of cells falling into each interval were calculated, resulting in 100 data points for each drug, which constitute the distribution curve for each drug and are referred to as the frequency data points. To assess the similarity of the 436 distribution curves to the control distribution curves of DMSO and Torin1, a heatmap was generated using python. The similarity was quantified using the formula: D * D , where D represents the DMSO Torin1 DMSO similarity to the DMSO control curves and DTorin1 represents the similarity to the Torin1 control curves. In addition, intensity analysis was conducted by measuring the average intensity of the whole cell. To cluster components based on the frequency data points, several steps were taken to ensure proper normalization and clustering. Initially, min-max normalization was applied to each drug's frequency data points. This step scaled the values to the range of 0 to 1. Subsequently, we applied quantile normalization to the control components' data (DMSO and Torin1), which rescaled the frequency data points to a unified scale among all controls and mitigated variations in measurement scales. Following this, frequency data points of all other drugs were adjusted. This adjustment was performed based on the average impact of the quantile normalization on the control data for the associated plates. We then combined the data from different conditions into a single data table for ease of analysis.

To identify the relationships between drugs, we computed cosine similarities (distances). These values were utilized in hierarchical clustering, where we implemented single-linkage clustering. In this methodology, the distance between clusters is defined as the shortest distance between any two points in the clusters. In the resulting matrix, each entry signifies the correlation value between the adjusted frequency data points of the two corresponding drugs. Moreover, we calculated the distances between each drug and the mean frequency data points of corresponding Torin1 and DMSO values. These distances were computed using the correlation values that were previously calculated for each drug in relation to all other drugs.

Transient transfection. Cells were seeded in six-well plates at 70% confluence 12 h prior to transfection. Transfection was performed using Jetprime Transfection Reagent (Cat# 101000027, Polyplus) according to the manufacturer’s protocols. Cells were harvested after 24-48 hours.

RNA interference and quantitative PCR. HEK293T cells were seeded in six-well plates at 30% confluence 12 h prior to transfection. Cells were transfected with a total of 20 nM siRNAs using the Lipofectamine RNAiMAX transfection reagent (Cat# 13778075, Invitrogen) according to the manufacturer’s protocols. To enhance gene silencing efficiency, two individual siRNAs were used for one gene target. Cells were harvested 48 h or 72 h after transfection. Silencer® Select Negative Control #1 siRNA (Cat# 4390843) and gene specific siRNAs for FBXW11 (β-TrCP2) (Cat# 4392420, assay ID: s23487 and s23485), IKBKB (IKKβ) (Cat# 4392420, assay ID: s223926 and s7263), CHUK (IKKα) (Cat# 4390824, assay ID: s3077 and s3078), IKBKGI (IKKγ) (Cat# 4390824, assay ID: s16186 and Cat # 4392420, assay ID: s533388), WDR68 (DCAF7) (Cat# 4427037, assay ID: s19985 and s19986) and fbxw11 (β-Trcp2) (Cat# 4390771, assay ID: s98104) were obtained from Thermo Fisher Scientific. For expression studies, RNA was extracted from cells using the NucleoSpin® RNA kit (Cat# 740955.50, Takara) according to the manufacturer's instructions. The concentration and quality of the extracted RNA were assessed using a Nanodrop ONEc (Thermo Scienfic) spectrophotometer. For cDNA synthesis, 1 µg of RNA was used per sample using the QuantiTect Reverse Transcription Kit (Cat# 205311, Qiagen). The quantitative RT-qPCR reactions were carried out using the PowerUp™ SYBR™ Green Master Mix (Cat# A25742, Applied Biosystems) and the QuantStudio-3 real-time PCR systems with the following conditions: 95ºC, 2min; (95ºC, 15 s; 55ºC, 15 s; 72ºC, 60 s) x 40. Primer sequences are listed in Supplementary Data 2. Relative RNA levels were normalized against an internal control (GAPDH or HPRT1) and were calculated as 2−ΔΔCT.

Cell lysis and immunoblotting. Following completion of the treatment, cells were washed once with cold PBS and lysed using either NP-40 lysis buffer (150 mM NaCl, 1% NP-40, 50 mM Tris-Cl pH 8.0) or RIPA buffer (150 mM NaCl, 1% Triton-X100, 50 mM Tris-Cl pH 8.0, 0.5% sodium deoxycholate and 0.1% SDS) supplemented with 1X protease and phosphatase inhibitors. The total lysates were kept at 4°C with agitation for 1 hour, followed by centrifugation at 16,000 × g for 15 minutes at 4°C to remove debris. The protein concentration was measured using the Pierce™ BCA Protein Assay Kit (Cat# 23225, Thermo Scientific), with Pierce™ Bovine Serum Albumin Standard, 2 mg/mL (Cat# 23210, Thermo Scientific) as the standard. Protein samples were prepared by adding Laemmli sample buffer (Cat# 1610737EDU, biorad) and incubating at 95°C for 5 minutes. The samples were then stored at −80°C until use. Denatured samples were separated by SDS-PAGE and subsequently transferred onto PVDF membranes (Cat# 1620177, Biorad). The membranes were incubated in blocking buffer (5%, w/v, dried skimmed milk in Tris-buffered saline, pH 7.4, and 0.1% Tween 20, TBST), followed by overnight incubation with the indicated primary antibodies, appropriately diluted in the blocking buffer. Subsequently, the membranes were washed three times with TBST and incubated with the appropriate secondary antibodies, either Goat anti-mouse HRP-conjugated antibody (Cat# 1705047, Biorad) or Goat anti-rabbit HRP-conjugated antibody (Cat# 1705046, Biorad) or IRDye® 680RD Goat anti-Mouse IgG Secondary Antibody (926-68070, 1:10000, LI-COR Biosciences) or IRDye® 800CW Goat anti- Rabbit IgG Secondary Antibody (926-32211, 1:10000, LI-COR Biosciences), diluted in blocking buffer, for 1 h at room temperature. After washing, the membranes were detected using SuperSignal™ West Dura Extended Duration Substrate reagent (Cat# A38554, ThermoFisher Scientific). Images were taken with ImageQuant LAS 4000 (GE Healthcare) and quantified by Fiji analysis software. Uncropped scans of all blots are provided in Source data file.

Immunoprecipitation. Protein extraction from cultured HEK293T and HeLa cells was performed with either RIPA buffer (50 mM Tris-HCl pH 7.4, 150 mM NaCl, 2 mM EDTA, 1% NP-40, 0.1% SDS) or NP-40 lysis buffer. Centrifugation was performed at 16,000 × g for 15 minutes at 4°C to remove cell debris. For lysates in RIPA buffer, the volume of the lysate was roughly measured, and powdered SDS was added in proportion to the volume to achieve a final concentration of 1% SDS. The lysate was then boiled at 95°C for 5 minutes to dissociate any non-covalently bound proteins. Subsequently, RIPA buffer without SDS was added to reduce the SDS concentration back to 0.1%. The lysates were then pre-cleared using dynabeads (Cat# 10003D, Invitrogen) conjugated with normal mouse IgG (Cat# 12-371, Sigma Aldrich). The pre-cleared lysates were then incubated with dynabeads conjugated to the indicated primary antibodies, rotating at 4°C for 2 hours. Subsequently, the beads were washed four times with 500 μL of RIPA lysis buffer and eluted in 2x Laemmli buffer at 95°C for 5 minutes.

Immunofluorescence. HeLa cells were seeded at 50–60% confluence on 24-well glass bottom plates (Cat# P24-1.5H-N, Cellvis) and subjected to either transfection or treatment. Cells were fixed with 4% paraformaldehyde at room temperature for 20 minutes, followed by permeabilization in 0.1% Triton X-100 in PBS for 10 min at room temperature with agitation. Next, cell samples were incubated in a blocking buffer consisting of 5% (w/v) Bovine Serum Albumin and 0.1% saposin. Overnight incubation with Flag antibodies (diluted 1:500 in the blocking buffer) was carried out. Subsequently, the cell samples were washed three times with PBS and incubated with the appropriate secondary antibodies: Goat anti-mouse antibody, Alexa Fluor™ 555 (Cat# A-21422, Invitrogen) or Goat anti-mouse antibody, Alexa Fluor™ 488 (Cat# A-11001, Invitrogen), both diluted 1:400 in blocking buffer, for 1 hour at room temperature. Following this, the cell samples were incubated with 300 nM DAPI (Cat# 28718-90-3, Roche) in PBS for 15 min and washed three times with PBS. A Zeiss cell discoverer 7 microscope and accompanying Zen software were used to acquire confocal images.

In vitro kinase assay and phos-tag analysis. 3xFlag-tagged WT and mutant TFEB were transiently transfected and immunoprecipitated from HeLa cell lysates using Flag antibody-conjugated beads. The immunoprecipitated TFEB proteins were first dephosphorylated by λ phosphatase (Cat# P9614, Sigma-Aldrich). The dephosphorylation master mix is composed of 1x λ Protein Phosphatase Buffer (Cat# L9288), 1x MnCl2 (Cat# M0564), 1x protease inhibitor (Cat# P3100-010) and about 16 units per μl λ phosphatase. Resuspend 25 μl of beads conjugated with immunoprecipitated TFEB in 50 μl of dephosphorylation master mix and incubate the reaction in a 30°C water bath for 40 minutes, pipetting the beads up and down every 10 minutes to resuspend them. Remove the master mix and wash the beads with lysis buffer. Elute the dephosphorylated TFEB using 200 mM glycine-HCl, pH 2.6. Immediately neutralize the elute with 1ul 1 M Tris, pH 10.6. Subsequently, the dephosphorylated TFEB molecules were subjected to a kinase assay using 300 nM IKKα (Cat# ab102103, Abcam) or IKKβ (Cat# 81066, Active Motif) or Akt (Cat# 1775-KS-010, R&D Systems) in the presence of 100 μM ATP, 2 mM MgCl2, 1 mM DTT, 100 μM Na3VO4, and 500 μM EDTA. The reaction mixtures were incubated for either 60 or 120 minutes at 25°C and the reaction was stopped by adding 2X Laemmli sample buffer, followed by boiling for 5 minutes.

Phos-tag analysis was performed according to the manufacturer’s instructions (Cat# F4002, APExBIO). Briefly, the resolving gel solution is composed of 10% (w/v) acrylamide, 30 μM Phosbind Acrylamide (PB-A), 0.1 mM MnCl2, 0.375 M Tris/HCl solution (pH 8.8), 0.1% (w/v) SDS solution, 0.1% TEMED (tetramethylethylenediamine), and 0.05% (w/v) ammonium persulfate solution. The stacking gel follows the standard composition of regular SDS-PAGE gels. Subsequently, the samples were separated by Phos-tag SDS-PAGE and transferred onto PVDF membranes for immunoblotting analysis.

Sample preparation and LC-MS analysis for phosphorylation detection.
For phosphorylated TFEB from the in vitro IKK kinase phosphorylation: Phosphorylated TFEB protein was prepared as described in the “in vitro kinase assay” section. Three aliquots of in vitro phosphorylated TFEB was used as n=3 replicates for this analysis. To remove MS-incompatible reagents, TFEB was isolated by acetone precipitation. Dry TFEB was reconstituted in 50 mM ammonium bicarbonate buffer, pH 7, reduced using dithiothreitol (Cat#20291, Pierce), alkylated using iodoacetamide (Cat#A3221, Sigma-Alrich), and digested using sequencing grade trypsin (Cat#V5113, Promega). TFEB tryptic peptides were dried and reconstituted in LC-MS mobile phase A (2% acetonitrile (Cat# 047138.M6, Thermo Scientific), 98% MilliQ purified water, 0.1% formic acid (Cat# 85178, Pierce)) for LC-MS analysis. We used a Thermo Fisher Ultimate RSLC3000 nanoLC system equipped with a micro silica column packed with 20 cm of ReproSil-Pur C18 resin (120Å, C18-AQ,1.9-μmresin; Cat# r119.aq, Dr. Maisch) and an Orbitrap Fusion Lumos Tribrid Mass Spectrometer. Our acquisition method includes an untargeted acquisition in parallel with a targeted acquisition for the tryptic phosphopeptides 406AA-430AA and 406AA-431AA. This approach allows a general analysis of TFEB tryptic peptides and targeted analysis of select candidate phosphopeptides for further fragmentation that ultimately enables the rigorous localization of the phosphorylation sites. The untargeted acquisition settings include Orbitrap as the detector, top 15 data dependent MS2, charge state selection 2-6, 60K and 30K was set as the Orbitrap resolution for MS1 and MS2 respectively, 250-2000 m/z scan range, 30% RF lens, 0.8 m/z isolation window, stepped HCD at 28%, 30%, 34% collision energy for fragmentation, and profile as acquired data type. The targeted acquisition involves MS2 acquisition, and the settings include a targeted precursor list of m/z 881.73 and m/z 924.43, Orbitrap as the detector, 30K Orbitrap resolution, scan range is 110-2000 m/z, 30% RF lens, stepped HCD fragmentation with 28%, 30%, and 32% collision energy. These fragmentation parameters are optimized to minimize the loss of phosphate for the targeted phosphopeptides. We used our in-house software (TDMS 7.04) to process the LC-MS data using UniProt database P19484 for TFEB122, 123. The data shows a clear presence of the intact phosphorylated peptide at 406AA-430AA with m/z 881.7311 (-1.99 ppm error, expected mass of 881.732851 m/z). The subsequent MS2 data exhibits multiple ion types supporting phosphorylation localization at S427. We also manually validated the MS2 fragment ions independently using Qualbrowser (Thermo Scientific Excalibur 4.3) to ensure confident phosphorylation site localization. No statistical tests were applied to the mass spectrometry data, as phosphorylation site localization was determined based on qualitative MS2 fragment ion evidence.

For phosphorated TFEB from the IL-1β treatment: Three cell plates were used in each IL-1β treatment timepoint as n=3 replicates for this analysis. After IL-1β treatment, cultured HeLa/TFEB cells were harvested, washed in PBS, resuspended in modified RIPA buffer, incubated on Thermomixer for 15 minutes at 37 ºC at 600 rpm, sonicated using BioRuptor Pico on high power for 7 cycles (30s on, 30s off), and centrifuged at 10,000 × g for 10 minutes at 4 ºC to pellet cell membrane debris. The supernatant cell lysate was then transferred to a low bind tube for Flag-tag immunoprecipitation. For a 15-cm plate of IL-1β-treated cells, 50 µL of Anti-DYKDDDDK Mag Agarose bead (Cat# A36798, Pierce) was used. The beads were washed using 500 µl modified RIPA 3 times, and the washed buffer was discarded each time with the aid of DynaMag rack. The beads were incubated in cell lysate on rotator for 20 minutes at room temperature. Conjugated beads were collected, and the supernatant was discarded. Beads were washed with 500 µl PBS twice and with 500 µl milliQ water once. Flag-tagged TFEB was eluted from the beads using 100 µl Glycine-HCl, pH 3.0 on the thermomixer at 1400 rpm, 25 ºC for 5 minutes. DynaMag rack was used to retain the beads, while the elute was collected to a low bind tube. This was repeated once more, and all elutes were combined. The final elution was neutralized with 30 µl 1 M Tris buffer, pH 8.0. This eluate was reduced using dithiothreitol (Cat# 20291, Pierce), alkylated using iodoacetic acid (Cat#43964, Alfa Aesar), and digested using sequencing grade trypsin (Cat# V5113, Promega). TFEB tryptic peptides were dried and reconstituted in LC-MS mobile phase A (2% acetonitrile (Cat# 047138.M6, Thermo Scientific), 98% MilliQ purified water, 0.1% formic acid (Cat# 85178, Pierce)) for LC-MS analysis. The LC-MS method used here was the same to the method described above. Briefly, we used a Thermo Fisher Ultimate RSLC3000 nanoLC system equipped with a micro silica column packed with 20 cm of ReproSil-Pur C18 resin (120Å, C18-AQ,1.9-μmresin; Cat# r119.aq, Dr. Maisch) and an Orbitrap Fusion Lumos Tribrid Mass Spectrometer. Our acquisition method includes an untargeted acquisition in parallel with a targeted acquisition for the tryptic phosphopeptides 406AA-430AA and 406AA-431AA. The untargeted acquisition settings include Orbitrap as the detector, top 15 data dependent MS2, charge state selection 2-6, 60K and 30K was set as the Orbitrap resolution for MS1 and MS2 respectively, 250-2000 m/z scan range, 30% RF lens, 0.8 m/z isolation window, stepped HCD at 28%, 30%, 34% collision energy for fragmentation, and profile as acquired data type. The targeted acquisition involves MS2 acquisition, and the settings include a targeted precursor list of m/z 881.73 and m/z 924.43, Orbitrap as the detector, 30K Orbitrap resolution, scan range is 110-2000 m/z, 30% RF lens, stepped HCD fragmentation with 28%, 30%, and 32% collision energy. MS Fragger v.21.1 was used for data search against the reviewed Uniprot human proteome database (UP000005640) with preset LFQ-phospho workflow, strict-trypsin as protease at K, R residue, 3 allowed miscleavage, mass shifts for: variable oxidation modification at M residue 15.9949 Da, variable acylation modification at N terminal 42.0106 Da, variable phosphorylation modification at S , T, Y residue 79.96633 Da, fix modification at C (cysteine) from iodoacetic acid 58.005 Da. Other parameters that are not mentioned here used default values set by the workflow. MS1 quantification data was analyze and plotted using GraphPad Prism v.9.5.1 and the statistical analysis used one-tail Welch’s t-test to obtain p-value of 0.0008. Phosphorylation site localization was further evaluated by manual inspection of MS2 fragment ions.

Sample preparation and LC-MS analysis for ubiquitination detection. The samples were digested using a modified filter-aided sample preparation method124. The beads were washed four times with 1 ml of 50 mM cold ammonium bicarbonate buffer, pH 8.0 and samples were eluted and reduced with SDS buffer (4% (wt/vol) SDS, 100 mM Tris-HCl pH 8.0) containing 100 mM DTT and then the samples were boiled at 95 ºC for 10 minutes. The reduced samples were mixed with 200 µl of 100 mM Tris-HCL buffer, pH 8.5 containing 8 M urea (UA buffer) and transferred onto the top chamber of a 30,000 MWCO filtration unit (Cat# MRCF0R030, Millipore) and spun in a microcentrifuge at 14,000 x g for 10 minutes. An additional 200 µl of UA buffer was added to the top chamber of the filter unit and the filter was spun at 14,000 x g for 15 to 20 minutes in a microcentrifuge (Eppendorf 5424). The flow through was discarded and the proteins were alkylated by adding 100 µl of UA buffer containing 50 mM iodoacetamide (Cat# A39271, Pierce) to the top chamber of the filtration unit and gyrating at 550 rpm in dark at room temperature for 30 minutes using a thermomixer (Thermomixer R, Eppendorf). The filter was spun at 14,000 x g for 15 minutes and the flow through was discarded. Unreacted iodoacetamide was washed away from the filter with two sequential additions of 200 µl UA buffer and centrifugation at 14,000 x g for 15 to 20 minutes after each addition. The urea buffer was exchanged into 50 mM ammonium bicarbonate buffer, pH 8 (digestion buffer), two sequential additions of 200 µl digestion buffer with centrifugation after each addition to the top chamber was performed. The top filter units were transferred to a new collection tube and 0.2 µg of Asp-N (Cat# 9053, Pierce) was added and samples were digested at 37 °C overnight. The next day, 0.1 µg of sequencing-grade trypsin (Cat# V5113, Promega) was added and samples were digested for two hours at 37 °C. The filters were spun at 14,000 x g for 15 minutes to collect the peptides in the flow through. The filter was washed with 50 µl of 100 mM ammonium bicarbonate buffer and the wash was collected with the peptides. In preparation for desalting, peptides were acidified to 1% (vol/vol) TFA final concentration. The peptides were desalted using two micro-tips sequentially. (C4, BIOMEKNT3C04, and C18, BIOMEKNT3C18) (Glygen) on a Beckman robot (Biomek NX), as previously described125. The peptides were eluted with 60% (vol/vol) acetonitrile in 0.1% TFA (vol/vol) and dried in a Speed-Vac (Thermo Scientific, Model No. Savant DNA 120 concentrator). The peptides were dissolved in 20 µl of 1% (vol/vol) acetonitrile in water. An aliquot (10%) was removed for quantification using the Pierce Quantitative Fluorometric Peptide Assay kit (Cat# 23290, Thermo Scientific). The remaining peptides were transferred to autosampler vials (Cat# 200046, Sun-Sri), dried and stored at −80ºC for LC-MS analysis.

The peptides were analyzed using a nano-Elute chromatograph (Bruker Daltonics) coupled online to a hybrid trapped ion mobility-quadrupole time of flight mass spectrometer (timsTOF Pro2, Bruker Daltonics with a modified nano-electrospray source (CaptiveSpray, Bruker Daltonics). The mass spectrometer was operated in dda-PASEF mode126. The samples in 0.1% (vol/vol) aqueous formic acid (FA) were loaded (2 µl) onto a 75 µm i.d. × 25 cm Aurora Series column with CSI emitter (Ionopticks) on a Bruker nano-ELUTE (Bruker Daltonics). The column temperature was set to 50 °C. The column was equilibrated using constant pressure (800 bar) with 8 column volumes of solvent A (0.1% (vol/vol) aqueous FA). Sample loading was performed at constant pressure (800bar) at a volume of 1 x sample pick-up volume plus 2 µl. The peptides were eluted using the one column separation mode with a flow rate of 300 nL/min and using solvents A and B (0.1% (vol/vol) FA/MeCN): solvent A containing 2%B increased to 17% B over 60 min, to 25% B over 30 min, to 37% B over 10 min, to 80% B over 10 min and constant 80% B for 10 min.

The MS1 and MS2 spectra were recorded from m/z 100 to 1700. Suitable precursor ions for PASEF-MS/MS were selected in real time from TIMS-MS survey scans by a PASEF scheduling algorithm126. A polygon filter was applied to the m/z and ion mobility plane to select features most likely representing peptide precursors rather than singly charged background ions. The quadrupole isolation width was set to 2 Th for m/z < 700 and 3 Th for m/z > 700, and the collision energy was ramped stepwise as a function of increasing ion mobility: 52 eV for 0–19% of the ramp time; 47 eV from 19–38%; 42 eV from 38–57%; 37 eV from 57–76%; and 32 eV for the remainder. The TIMS elution voltage was calibrated linearly using the Agilent ESI-L Tuning Mix (m/z 622, 922, 1222).

Data from the mass spectrometer were converted to peak lists using DataAnalysis (version 6.1, Bruker Daltonics). The MS2 spectra with charges +2, +3 and +4 were analyzed using Mascot software 127 (Matrix Science, London, UK; version 2.8.2). Mascot was set up to search against a UniProt (ver July 2019) database of human proteins (20,667 entries), assuming the digestion enzyme was AspN with a maximum of 4 missed cleavages allowed. The searches were performed with a fragment ion mass tolerance of 50 ppm and a parent ion tolerance of 25 ppm. Carbamidomethylation of cysteine was specified in Mascot as a fixed modification. Deamidation of asparagine, deamidation of glutamine, formation of pyro-glutamic acid from N-terminal glutamine, acetylation of protein N-terminus, oxidation of methionine, and diglycine modification of lysine were specified as variable modifications. Peptides and proteins were filtered at 1% false-discovery rate (FDR) by searching against a reversed protein sequence database. Scaffold (version Scaffold_5.3.0, Proteome Software Inc., Portland, OR) was used to validate MS/MS based peptide and protein identifications. Peptide identifications were accepted if they could be established at greater than 54.0% probability to achieve an FDR less than 1.0% by the Scaffold Local FDR algorithm. Protein identifications were accepted if they could be established at greater than 99.0% probability and contained at least 2 identified peptides. Protein probabilities were assigned by the Protein Prophet algorithm (Nesvizhskii, Al et al Anal. Chem 2003;75(17):4646-58).

Subcellular fractionation. Cells were cultured to 70% confluence in a 6-well plate. Subcellular fractionation was performed using Subcellular Protein Fractionation Kit for Cultured Cells (Cat# 78840, Thermo Scientific) following the manufacturer’s protocols. Each fraction was subsequently subjected to immunoblotting analyses.

Data analysis. Data are presented as mean ± SEM. unless otherwise stated. Statistical comparisons between two groups were performed using two-tailed unpaired Student’s t-tests. Comparisons among multiple groups were performed using one-way analysis of variance (ANOVA) followed by Tukey’s multiple-comparisons test, one-way ANOVA followed by Dunnett’s multiple-comparisons test, or Welch’s one-way ANOVA followed by Dunnett’s T3 multiple-comparisons test, as indicated. In analyses involving multiple pairwise comparisons, multiple two-sided t-tests with Benjamini–Hochberg false discovery rate (FDR) correction were applied. Where specified, one-tailed Welch’s t-tests were used. A p value < 0.05 was considered statistically significant.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Xiong et al. investigated the mechanisms regulating TFEB protein turnover. By performing a human kinome screening, the authors found that IKK inhibition strongly stabilized TFEB and promoted its nuclear translocation. They also found that silencing of IKK subunits resulted in increased TFEB protein levels. IKK was able to phosphorylate TFEB in vitro at multiple serine residues. The authors propose that multisite TFEB phosphorylation by IKK primes TFEB for  -TrCP2-mediated ubiquitination and subsequent degradation. Although results of this work may be interesting in principle, there are several issues that require further clarification. First, the physiological relevance of IKK-mediated degradation of TFEB is unclear. The authors propose that IKK regulates basal TFEB protein turnover. However, the IKK complex is mainly inactive under basal conditions and its activity is well-known to be induced by a plethora of stimuli, including inflammatory inputs (e.g. cytokines, PAMPs, etc). It is therefore quite puzzling that TFEB stability is controlled by an inactive kinase. In addition, how the IKK complex is involved in the degradation of TFEB upon IKK-activating stimuli has not been properly addressed. Furthermore, the authors show that IKK inhibition results not only in increased stability but also in TFEB nuclear translocation. However, the relationship between these two processes has not been investigated in detail. Finally, there are major issues with the interpretation of some of the data, as listed below. Specific points: 1. As stated above, the physiological relevance for the role of IKK in TFEB degradation is unclear. The authors propose that the IKK/ -TrCP2 axis regulates basal TFEB protein turnover. For instance, on page 12 the authors state that “ -TrCP2 functions as a default TFEB ubiquitin ligase”. However, the IKK complex is mainly inactive under basal conditions and its activity is well-known to be induced by several stimuli, including inflammatory inputs (e.g. cytokines, PAMPs, etc). How can basal TFEB stability be controlled by an inactive kinase? For instance, IKKbeta- and IKKgamma-KO cells are characterized by impaired degradation of its main substrate IkBalpha in response to IKK-activating stimuli, such as TNFalpha, whereas basal IkBalpha levels are unaffected in IKK-KO cells. The authors should check basal IkBalpha levels in their IKK-silenced cells. Also, the use of IKKbeta- and IKKgamma-KO cells is needed in order to check basal TFEB (and IkBalpha) levels in a more relevant cellular system. 2. Related to the previous point, the role of IKK-mediated TFEB degradation has not been properly addressed in physiologically relevant conditions. For instance, despite being a well-known IKK-activating stimulus, TNFalpha does not seem to induce a significant TFEB degradation (Figure 3e), as opposite to LPS stimulation which promotes a strong reduction in TFEB levels (Figure 3f). This is in contrast with a role of IKK as a key mediator of TFEB degradation. Do other well-known IKK-activating stimuli, such as IL-1, promote TFEB degradation? Furthermore, to define whether IKK is indeed a physiological mediator of TFEB phosphorylation/degradation, the authors should assess whether the levels of TFEB are affected in IKKbeta- and IKKgamma-KO cells in response to LPS, TFNalpha and IL-1 stimulation, as compared to WT-stimulated cells. 3. What is the relationship between IKK-mediated TFEB phosphorylation and nuclear translocation? The authors show that the IKK-16 inhibitor stabilized TFEB levels but also strongly promoted its nuclear translocation (Figure 1c). Does IKK silencing also promote TFEB nuclear translocation? Also, what is the subcellular localization of TFEB in IKK-KO cells? The authors showed that the use of IKK-16 also results in mTORC1 inhibition (Supplementary Figure 2a, b), but stated that “the two processes are uncoupled”. In order to determine whether the effect of IKK-16 on TFEB stabilization is uncoupled from mTORC1 inhibition, the authors should use this inhibitor in cells with constitutive mTORC1 activity (i.e. cells expressing constitutively active Rag GTPases) and assess TFEB levels and subcellular localization. 4. I have major issues with the interpretation of some results. The authors state that, opposite to WT TFEB, the levels of TFEB-6xA remained stable upon cycloheximide or LPS treatment (Figures 3d and 3f). However, the starting levels of TFEB-6xA are much higher compared to WT-TFEB, which makes very difficult any comparison between the two constructs. Despite this, I do see a clear decrease in TFEB-6xA levels upon these treatments (Figures 3d and 3f), thus arguing against the conclusions made by the authors. Similar issues apply to Figure 5e, when comparing the levels of WT-TFEB to TFEB-6xA upon  -TrCP2 silencing, and to Figure 6i, when using the TFEB-KKRR mutant. The authors should use similar starting protein levels and a similar exposure to make any comparisons between the two plasmids. 5. It is quite puzzling that, when performing transient transfections, no differences are observed in the levels of wt, 6xA-, and KKRR-TFEB (Supplementary Figure 7a, b). The authors state that this puzzling difference, compared to the stable cell lines, occurs “presumably because the very high protein levels associated with transient transfection by themselves overwhelm the processing abilities of IKK/b-TrCP2…”. It is surprising that the authors did not test this hypothesis, as it could be easily tested by transfecting lower amounts of wt and mutant TFEB. When doing so, is the stability of transiently transfected TFEB mutants affected? Do they show a different degradation pattern in response to LPS or TNFalpha stimulation? 6. The authors should generate and use a 6xD/6xE TFEB phospho-mimetic mutant. Does IKK-16 or IKK depletion stabilize this construct? 7. The rationale for focusing on  -TrCP2 as a putative TFEB ubiquitin ligase is unclear. In Figure 4a, DCAF7 appears to have a strong effect on TFEB ubiquitination. Yet, the authors only focused on  -TrCP2. What is the effect of DCAF7 depletion on TFEB stability? What is the effect of either DCAF7 or  -TrCP2 depletion on LPS-mediated TFEB degradation? 8. I found surprising that the authors only assessed the levels of the TFEB-KKRR mutant in response to  -TrCP2 silencing. It is indeed crucial to test the effect of  -TrCP2 or DCAF7 overexpression on TFEB-KKRR protein levels and ubiquitination. Furthermore, the effect of both IKK-16 and IKKbeta and IKKgamma depletion on this mutant should also be assessed. 9. The authors state that Torin1 did not influence TFEB stability. However, previous evidence showed the opposite (PMID: 27268034). In addition, the authors found that rapalogs induce TFEB nuclear translocation. However, TFEB is a well-known rapamycin-insensitive mTORC1 substrate (PMID: 22692423, 22343943, 21617040). How do the authors explain the discrepancy of their findings with previous work? These data raise serious concerns on the cellular systems and/or methods that the authors used to assess TFEB localization and stability. 10. TFEB ubiquitination experiments need to be performed at higher standards. To avoid the presence of confounding contaminants and properly assess TFEB ubiquitination, extracts need to be boiled in 1% SDS prior immunoprecipitation, which ensures the dissociation of any non-covalently bound proteins. Please see PMID: 17991829 (or many other references on the topic) for a more detailed protocol. 11. Detailed information on the protocol used for the kinase assays should be provided in the methods section.", "reviewer_2": "To authors The authors report a potential regulation of the lysosomal activity and autophagy-controlling transcription factor TFEB by the IκB kinase (IKK)-proteasome pathway, which results in subsequent phosphorylation, βTrCP2-mediated ubiquitination and proteolytic degradation of TFEB following cellular stimulation. Serine cluster and potential ubiquitination sites were identified using tagged transfected components. These degron residues can be removed without affecting the transcriptional activity or nuclear translocation of TFEB. The analyses were almost exclusively performed in in vitro systems and/or by using transfected molecules. Technical or conceptional flaws are seen in several experiments. The potential biological significance remains unclear: Why should all physiological NF-κB-activating stimuli, which result in IKK activation, cause a degradation of TFEB? No physiological evidence is provided to support a hypothesis for IKK to TFEB signaling. Only HeLa and 293T cells have been used. The authors should have included non-cancerous/transformed cell types and analysis of the endogenous proteins and multiple IKK stimuli and provide genetic evidence (e.g. mouse models) for their hypothesis. The analysis of the degron and ubiquitination does not provide significant advance over the analysis of other already known IKK substrates. The paper does not provide sufficient advance to be considered for Nature Communications. Some specific points: Fig. 1b. This is a debatable strategy to use kinase inhibitors because perhaps none of these inhibitors is selective for the presumed target kinase and IKK requires cellular stimulation with specific agents to be activated. In non-stimulated cells IKK has very low basal activity. Why then should an IKK inhibitor be effective on TFEB-GFP in this experiment? As said, the inhibitor IKK-16 is not selective and also inhibits LRRK2. How can indirect effects of the drugs on cell viability, affecting apparent TFEB levels, and on cell cycle be controlled? Fig. 2c. The effect of IKK-16 on TFEB nuclear translocation should be confirmed by using a additional, different IKK inhibitor and/or siRNA-mediated silencing of IKKβ or γ. Fig. 1e. Why are there two main bands for each, IKKα and IKKβ and also for IKKγ? Both bands for α and β (and γ) are affected by the respective siRNA, but the migration of at least one them does not fit to the known molecular weight. It is completely unexpected, and contradicts comprehensive literature knowledge, that siIKBKG strongly affects IKKα protein level. Why should this be? The effect of siRNAs on endogenous TFEB is not convincing. Fig. 2. Recombinant TFEB-3xFlag was expressed in HeLa cells, isolated, dephosphorylated by λ-phosphatase, phosphorylated in vitro by purified IKKβ (or IKKα). This approach likely results in artificial substrate phosphorylation events, which are meaningless in intact cells. Furthermore, substrate recognition in intact cells may involve association of IKKβ with NEMO/IKKγ. The problem of their approach is consistent when comparing the sites with intact cell-derived data in PhosphoSitePlus: Ser401 and 427 are not indicated as phosphorylated human TFEB. Human S427 is not even conserved in mouse or rat. These results are thus questionable. The proposed site SPFPSLS does not correspond to well characterized IKK phosphorylation sites. A determination of phosphorylation sites by mass spectrometry on endogenous TFEB after stimulation with IKK-activating agents is required. Fig. 3. The rapid and almost complete degradation of transfected tagged TFEB by LPS and much weaker or even marginally so by TNFα (panels e and f) is surprising since IKK is activated more strongly by TNFα, as many studies have shown. The IκB control is missing in panel f. The tag may interfere with degradation. It is important that degradation of endogenous TFEB is shown. Fig. 3d. That TFEB-6xA is stabilized in CHX treated cells rather suggest that kinases other than IKK phosphorylate these residues in the “WT” version, since IKK is not active in non-stimulated cells. Fig. 4 and S6. In some cases, proteasomal inhibition is needed to show elevated HA-ubiquitinated species, in others not. A mass-spec-based determination of TFEB ubiquitination is needed, since the HA-Ub approach may result in physiologically non-relevant results. For other substrates, βTrCP1 and βTrCP2 are interchangeable.", "reviewer_3": "This manuscript describes how TFEB degradation is regulated by both phosphorylation and ubiquitylation. I have been asked to comment specifically on the phosphoproteomics experiments that were done. To this end, my review is very short. I have examined all the spectra provided and I conclude that they are accurately interpreted and that the localization of the phosphorylation event has strong support in the MS/MS"}, "author_response": "We sincerely thank the Reviewers for the time and effort dedicated to reviewing this manuscript. We are pleased to share that they suggestions have led to substantial refinements in the presentation of the data – most notably, direct evidence of TFEB’s serine 423 as a target of IKK phosphorylation and of TFEB ubiquitination at lysines 430 and 431, mechanistic evidence of the uncoupling between IKK-mediated control over TFEB levels and mTORC1-mediated control over TFEB nuclear translocation, and the use of cells lines with individual knock-out of the IKK components which substantiate the model. Point-by-point answers to the Reviewers’ queries and suggestions are reported below. Because of overlaps between some of the Reviewers’ comments, our answers are redundant in certain points. We have preferred to report all added experiments in each Reviewer’s point-by-point reply rather than referring the Reviewer to questions from other Reviewers, which could have been confusing. [Detailed point-by-point responses follow...]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "The authors failed to address most of my concerns. In general, the data provided and the effects shown on TFEB degradation by the IKK complex remain very weak to support the authors’ conclusions. As an example, although the authors claim that IKKbeta is a major modulator of TFEB stability, silencing of IKKbeta shows no effects on TFEB levels in Figure 3e. This raises significant concerns about the validity of the conclusions of the manuscript. Furthermore, I have serious concerns regarding the authors’ data, experimental setup, and interpretation of results related to this manuscript or to previously published work: first, the IKK inhibitor IKK-16, which was used in the initial screening as the starting point of the manuscript and was shown to strongly increase TFEB levels and nuclear translocation, turned out to also inhibit mTORC1 activity. Indeed, in response to point 3, the authors state that “mTORC1 inhibition may be an off-target effect of IKK-16”. It is concerning that the significantly increased levels and nuclear localization of TFEB observed in IKK-16-treated cells, which are attributed to an off-target effect, are still presented in Figure 1C-D and used as a rationale to study the IKK complex. Second, in response to point 4 of the rebuttal, where I requested the use of similar starting protein levels when comparing the stability of different TFEB mutants, the authors did not repeat the experiments using cell lines expressing lower amounts of mutant plasmids to ensure comparable starting levels of wild-type and mutant proteins. Instead, they loaded different amounts of protein extracts, which however displayed very high differences in the expression of wild-type versus mutant protein. This approach is flawed, as differing starting levels will naturally lead to different degradation rates. Accordingly, in response to point 5, the authors themselves acknowledge this issue by transfecting increasing amounts of TFEB and concluding that “this difference presumably is a consequence of an overwhelmed ability of IKK/β-TrCP2 or their associated machineries to process the very high protein levels associated with standard transient transfection”. It is perplexing that the authors applied two different arguments and approaches to address essentially the same issue. Finally, I am deeply concerned with the authors’ response to point 9, in which they attempt to justify their puzzling observation that rapamycin promotes TFEB nuclear translocation. First, they claim that publications PMID 22692423 (Fig. 3A) and PMID 21617040 (Fig. 2E), which I cited as evidence showing no TFEB nuclear translocation in response to rapamycin, “have no quantitative assessment of TFEB nuclear translocation”. However, while direct quantification is not provided, the first study explicitly states that the IF images are representative of three independent experiments, with at least 30 cells imaged per experiment. Similarly, the second study includes nucleo/cytosolic fractionation and western blotting, which are inherently quantitative. Even more puzzlingly, the authors refer to the article PMID 22343943 - which I cited as further evidence of TFEB insensitivity to rapamycin – stating: “Fig. 2D shows, with a dose-response analysis, that TFEB is responsive to rapamycin”. However, they appear to have misinterpreted the data. In Fig. 2D of the cited manuscript, the EC50 of rapamycin is reported as 104322 nM, whereas the standard working concentration of rapamycin is 25 nM - a 4000-fold difference. Accordingly, PMID 22343943 states: “our data indicate that mTOR is a potent inhibitor of TFEB nuclear translocation and that TFEB is a rapamycin-resistant substrate of mTORC1”. To further support their argument, the authors also cite a study by Haoxing Xu and colleagues (PMID 31112550), asserting that they “used microscopy-based quantitative assessment to find that rapamycin promotes TFEB nuclear translocation”. However, I believe they have also misinterpreted this study. In Fig. 4A, Xu and colleagues show that TFEB nuclear translocation occurs only in cells overexpressing TRPML1 treated with very high concentrations of rapamycin (5000 nM). Even at these concentrations, and consistent with previous observations, no nuclear translocation of TFEB was observed upon rapamycin treatment in WT cells. This is quantified in Fig. 4B. Altogether, these issues raise serious concerns about the experimental setup, robustness of data, and the validity of the model proposed by the authors.", "reviewer_2": "The authors have considerably improved their work by inclusion of new experiments, which have addressed most of my previous concerns. Particularly important is the use of fibroblasts in addition to transformed cell lines and analysis of the endogenous system, including new determination of phosphorylation and ubiquitination sites. While mainly of the new data are convincing, several technical problems again emerge and need to be addressed either by experimental and/or suitable text modifications: New figures 1e and f: The claimed increase of TFEB upon IKKβ silencing is not convincing. Only silencing of subunits α or γ causes a clear increase. In contrast, IκBα levels are affected, as expected, upon silencing of IKKβ or IKKγ, but not IKKα. New figures 1g-j: Which of the double bands (criticized before) in the IKK subunit blots (also seen for IKKβ, but not IKKα, in MEFs, Suppl Fig. 2c) was used for quantification? For subunit γ most likely both, as they are closely spaced. For IKKα, both bands react equivalently under the different conditions, but for IKKβ, in lanes 2 and 4, this is not the case. In figure 1h, IKKβ silencing results in an apparent 2-fold increase in IKKγ. But this is not visible in panel 1e. Rather, IKKγ silencing results in increased levels of IKKβ (upper band). This differs from the results in KO cells (Suppl. Figure 2c). New Suppl. Fig 4a, while convincingly showing that silencing of any IKK subunit leads to abolished TFEB decline upon TNF stimulation, while the effect on IκBα is limited to loss of IKKβ and γ, again some technical problems emerge: Why do IKKα (and to some extent IKKβ) intensities increase upon TNF stimulation? The effect on IKKα is also evident after IL1 stimulation (Suppl. Fig 4d). Is this a gel/blotting effect, seen in multiple experiments?"}, "author_response": "We sincerely thank the Reviewers for the time and effort dedicated to reviewing this manuscript. We are pleased to share that they suggestions have led to substantial refinements in the presentation of the data – most notably, direct evidence of TFEB’s serine 423 as a target of IKK phosphorylation and of TFEB ubiquitination at lysines 430 and 431, mechanistic evidence of the uncoupling between IKK-mediated control over TFEB levels and mTORC1-mediated control over TFEB nuclear translocation, and the use of cells lines with individual knock-out of the IKK components which substantiate the model. Point-by-point answers to the Reviewers’ queries and suggestions are reported below. [Detailed point-by-point responses follow...]"}, {"round": 3, "reviewer_comments": {"reviewer_1": "The revision has not adequately addressed my main concerns. No new or conclusive experimental evidence has been provided to address my preivous points and to support the authors’ conclusions. Thus, several inconsistencies in the data remain unresolved. In light of the persistent issues, I believe the revised manuscript does not meet the standard required for publication.", "reviewer_2": "The authors have satisfactorily addressed my remaining concerns. I now suggest acceptance of the manuscript for publication.", "reviewer_4": "The paper by Xiong et al. argues that the stability of TFEB is controlled by constitutive and stimulated activation of IKK complexes that phosphorylate a novel degron in TFEB, then recognized by the β-TrCP2 ubiquitin ligase. Through revision, the authors attribute several internal inconsistencies to redundancy among IKK subunits or to cell-type variation, however, the more immediate issue is that key experiments are not rigorously controlled, which impairs coherent integration of the results. Consequently, the evidence presented does not yet substantiate the proposed model across perturbations and contexts. A focused set of targeted experiments, outlined below, should directly address specificity and reconcile divergent phenotypes. As is, I am not convinced of the final model presented. Major points • The human kinome screen is presented in the responses as merely an entry point to implicate IKK in TFEB regulation, yet it is in fact a major point of the paper. The pharmacologic–genetic link needs to be tightened: IKK-16 should be tested in IKKβ/γ-KO cells to assess effects on TFEB expression, establishing specificity. Likewise, because β-TrCP2 is identified (by silencing) as the TFEB ubiquitin ligase, those β-TrCP2–silenced cells should be treated with IKK-16, and β-TrCP2 should be silenced in IKK-deleted backgrounds (not in TFEB-6xA or TFEB-KKRR overexpression contexts, which are removed from the initial findings). • The manuscript argues partial redundancy among IKK subunits, yet deletion of each subunit yields distinct, at times opposing phenotypes. This is evident from the outset (IκBα baselines in the chosen control readout). IKKβ deletion shows minor effects on lysosome expansion and LAMP1 levels but similar increases in DQ-BSA dequenching as deletion of IKKa and IKKg. The DQ-BSA data are not convincing since after 16 h loading, the signal appears far lower than expected relative to lysosomal staining. Additionally, Torin-1 for only 1 h as a “biogenesis” positive control is questionable as one would expect lysosomal biogenesis to require longer treatment. • IKKα deletion has little effect on TFEB at rest (Fig. 2a, 2d) but protects TFEB from TNF/LPS-induced degradation. This raises basic questions: what activates IKK at baseline in the other lines but not in MEFs? More fundamentally, inflammatory cues are often associated with increased lysosomal biogenesis. Do the authors actually observe decreases in lysosome abundance or LAMP1 in MEFs upon LPS? The Discussion focuses on autophagy under inflammation, but more simply, throughout the manuscript lysosomal numbers are largely not determined. • Transcriptional analyses of lysosomal genes are shown only for IKKγ and in a single cell type. The pS423-TFEB signal that appears with IL-1β but not basally also needs clarification; if TFEB is being degraded, this should be tested with cycloheximide (CHX) to establish turnover. Moreover, in experiments with transient transfection (e.g., Fig. 4 in HEK cells), pathway activation is likely (e.g., endotoxin or transfection-related), so I would hesitate to describe those as “normal growth” conditions. Other points • Fig. 4d: The TFEB-6xA band after 4 h CHX shows either substantial degradation or a consistent downward shift in apparent molecular weight; this needs to be explained. • The in vitro TFEB phosphorylation assays use IKKα/IKKβ, yet the strongest cellular effects appear with IKKγ. This mismatch should be addressed potentially by testing IKKγ dependence in cellular assays and/or using reconstituted canonical complexes in vitro."}, "author_response": "We thank the reviewer for their careful evaluation and for the constructive suggestions, which prompted a series of targeted experiments that substantially strengthened the mechanistic framework of the manuscript. Below, we address each point and detail the new data added in response. [Detailed point-by-point responses follow...]"}, {"round": 4, "reviewer_comments": {"reviewer_4": "The authors have sufficiently addressed my comments, which has ultimately provided a more in depth set of clear controls that support their model. The article is ready for publication in this reviewer's opinion."}, "author_response": ""}]

Decision letter