Posts Tagged ‘cell death pathway’

Cell Death Pathway Insights

Larry H. Bernstein, MD, FCAP, Curator


Phosphorylation and activation of ubiquitin-specific protease-14 by Akt regulates the 1 ubiquitin-proteasome system

Daichao Xu1,2, Bing Shan1,4, Byung-Hoon Lee3,4, Kezhou Zhu1,4, Tao Zhang1,4, Huawang Sun1, 4 Min Liu1, Linyu Shi1, Wei Liang1, et al.
eLife 2015;10.7554/eLife.10510    DOI:

In this study, we report that USP14 is an Akt substrate and that this phosphorylation activates the DUB activity of USP14 both in vitro and in cells. We also demonstrate that phosphorylation of USP14 is critical for Akt to control UPS and consequentially global protein degradation via the UPS.

Regulation of ubiquitin-proteasome system (UPS), which controls the turnover of short-lived proteins in eukaryotic cells, is critical in maintaining cellular proteostasis. Here we show that 40 USP14, a major deubiquitinating enzyme that regulates the UPS, is a substrate of Akt, a serine/threonine-specific protein kinase critical in mediating intracellular signaling transducer for growth factors. We report that Akt-mediated phosphorylation of USP14 at Ser432, which normally blocks its catalytic site in the inactive conformation, activates its deubiquitinating activity in vitro and in cells. We also demonstrate that phosphorylation of USP14 is critical for Akt to regulate proteasome activity and consequently global protein degradation. Since Akt can be activated by a wide range of growth factors and is under negative control by phosphoinosotide phosphatase PTEN, we suggest that regulation of UPS by Akt-mediated phosphorylation of USP14 may provide a common mechanism for growth factors to control global proteostasis and for promoting tumorigenesis in PTEN-negative cancer cells.

The ubiquitin-proteasome system (UPS), a major degradative mechanism in eukaryotic cells, is involved in the degradation of short-lived proteins as well as misfolded and damaged proteins 69 (Komander and Rape, 2012). The 26S proteasome specifically targets and degrades proteins conjugated to ubiquitin. Regulation of protein deubiquitination by deubiquitinating enzymes (DUBs) is recognized as an important regulatory step in the ubiquitin-proteasome system. USP14, a deubiquitinating enzyme reversibly associated with the proteasome, negatively regulates the activity of proteasomes by trimming ubiquitin chains on proteasome-bound substrates (Borodovsky et al., 2001; Koulich et al., 2008; Lee et al., 2010). Purified recombinant USP14 is largely inactive and can be highly activated when in association with proteasome (Hu 76 et al., 2005; Koulich et al., 2008; Lee et al., 2010). However, a significant fraction of USP14 is present intracellularly in a proteasome-free state (Koulich et al., 2008) and it is not clear if and how proteasome-free USP14 might serve a significant physiological function. Akt, a serine/threonine-specific protein kinase and an important intracellular signaling transducer for growth factors such as insulin, is involved in regulating cell proliferation, metabolism, transcription, migration and apoptosis (Manning and Cantley, 2007). The activity of Akt is regulated by PI(3,4,5)P3, a lipid product of the phosphoinositide 3-kinases (PI3Ks). The intracellular levels of PI(3,4,5)P3 are negatively regulated by phosphatases such as SHIP1/2 and PTEN. The latter, a phosphoinoside phosphatase, is encoded by a tumor suppressor gene that is mutated in human cancers at high frequency (Cantley and Neel, 1999). Akt has been reported to mediate the phosphorylation of many substrates that in turn regulate cell proliferation, metabolism, transcription, migration and apoptosis. However, very little is known about its role in the UPS, and furthermore no mechanistic link between Akt and UPS has been elucidated.

Two forms of USP14 have been determined crystallographically: the inactive free form and an adduct between Ub-aldehyde (Ubal) and USP14, which provides insight into the catalytically active state (Hu et al., 2005). The key difference between these two structures is in the position of the blocking loops, BL1 and BL2, which project over the catalytic cleft of USP14 and block the access of the C-terminal residues of ubiquitin in the inactive form (Figure 1A). In  Ubal-modified USP14, BL1 and BL2 are rearranged, thus exposing the cleft. In particular, Ser432, located within BL2, shifts its position over a distance of 3-5Å between the two states (Hu et al., 2005) (Figure 1B). Since Ser432 residue is located very close to a highly negatively charged patch (Figure 107 1C), we reasoned that when Ser432 residue was phosphorylated, the negatively charged phosphate group might induce a repulsive force, thereby inducing rearrangement of the BL2 loop and removing the inhibitory effect of this loop on the activity of USP14. The amino acid 110 sequences around Ser432 are highly evolutionarily conserved among USP14 orthologues 111 (Figure 1D) and Ser432 is predicted to be an Akt substrate by Scansite ( We therefore tested the possibility that USP14 might be a substrate of activated Akt. We first examined the interaction between USP14 and Akt using a co-immunoprecipitation assay. As shown in Figure1-figure supplement 1A, when USP14 and Akt were overexpressed in HEK293T cells, their interaction was readily detectable. To test whether Akt could phosphorylate USP14, we overexpressed USP14 and an activated Akt (Myr-Akt) in HEK293T cells, and performed a quantitative phosphoproteomic analysis (Figure 118 1-figure supplement 1B). We identified four phosphorylation sites on USP14 when it was 119 expressed alone: Ser143, Ser230, Thr235, and Ser432 (Figure 1-figure supplement 1C-D). Notably, the phosphorylation levels of two of the four sites, Ser143 and Ser432, were increased considerably in cells expressing activated Akt (Figure 1E).

Figure 1. Structural basis of USP14 activation by phosphorylation of Ser432. (A) Detailed view of blocking loop 2 (BL2), which occludes the active site of USP14 (PDB access code 2AYN). The BL2 loop, which contains Ser432, is shown in stick model, in the apo form. (B) Combined ribbon representation and stick model showing a comparison of the conformations of the BL2 loop containing in the apo form (blue, PDB access code 2AYN) and in the USP14-Ubal adduct (orange, PDB access code 2AYO). In this drawing, the Ser432 and Cys114 residues are 504 shown in stick model, and the bound Ubal (a ubiquitin derivative in which the C-terminal 505 carboxylate is replaced by an aldehyde) in the complex is drawn in green. (C) A surface charge potential representation (contoured at ±7 kT/eV; blue/red) of USP14 (PDB accession 2AYN) showing that the S432 residue is very close to a highly negatively charged patch mainly formed by the acidic E188, D199 and E202 residues. When S432 is phosphorylated, the negatively charged phosphate group may induce a repulsive force, thereby relieving inhibition of the catalytic activity of USP14. (D) USP14 domain organization and sequence alignment of the Akt 511 phosphorylation site within USP14 orthologues from different species. Two blocking loops (BL1 512 and BL2) covering the USP14 active site are shown. The Akt phosphorylation site in USP14 from different species as predicted by Scansite. (E) S432 is the major phosphorylation site in USP14. HEK293T cells were treated as in Figure 1-figure supplement 1B, followed by ESI-MS analysis. Spectral counts were determined by ESI-MS. (F) Akt phosphorylates USP14 in vitro. Bacterially expressed and purified USP14 was incubated with active Akt in the presence of ATP. Reaction products were resolved by SDS-PAGE, and phosphorylated species were detected by a phospho-Ser antibody.

Since Ser432 residue is located very close to a highly negatively charged patch (Figure 107 1C), we reasoned that when Ser432 residue was phosphorylated, the negatively charged phosphate group might induce a repulsive force, thereby inducing rearrangement of the BL2 loop and removing the inhibitory effect of this loop on the activity of USP14. The amino acid sequences around Ser432 are highly evolutionarily conserved among USP14 orthologues (Figure 1D) and Ser432 is predicted to be an Akt substrate by Scansite ( We therefore tested the possibility that USP14 might be a substrate of activated Akt. We first examined the interaction between USP14 and Akt using a co-immunoprecipitation assay. As shown in Figure1-figure supplement 1A, when USP14 and Akt were overexpressed in HEK293T cells, their interaction was readily detectable. To test whether Akt could phosphorylate USP14, we overexpressed USP14 and an activated Akt  (Myr-Akt) in HEK293T cells, and performed a quantitative phosphoproteomic analysis (Figure 1-figure supplement 1B). We identified four phosphorylation sites on USP14 when it was expressed alone: Ser143, Ser230, Thr235, and Ser432 (Figure 1-figure supplement 1C-D). Notably, the phosphorylation levels of two of the four sites, Ser143 and Ser432, were increased considerably in cells expressing activated Akt (Figure 1E).

To examine whether USP14 is a direct substrate for Akt, we conducted an in vitro kinase 123 assay using activated recombinant Akt and purified recombinant USP14 expressed in E. coli. We 124 found that co-incubation of USP14 and Akt led to modification of USP14 as detected by a pan phospho-Ser antibody (Figure 1F), suggesting that USP14 is a substrate for Akt.

Figure 1-figure supplement 1. Akt phosphorylates USP14. (A) Akt interacts with USP14. HEK293T cells were transfected with indicated plasmids for 24 h. The cell lysates were collected for co-immunoprecipitation and western blotting analysis. (B) Schematic representation of mass spectrometry assay to determine USP14 phosphorylation sites by Akt. (C) Four phosphorylation sites of USP14 were determined by mass spectrometry. (D) The representative MS/MS spectrum of phosphorylated tryptic peptide ‘SSSphosSGHYVSWVK’ of human USP14 protein. The peptide sequence ‘SSSphosSGHYVSWVK’ containing phosphorylated S432 was identified by shotgun analysis using mass spectrometry when USP14 was co-expressed with Myr-Akt in HEK293T cells. Fragmentation ion of the amide bond of the peptide result in formation of ‘b’ ion and ‘y’ ion series corresponding to the N-terminal and C-terminal fragments respectively. Representative ions with phosphorylation and H2O loss were manually labeled in red on the spectrum.

To determine if Ser143 and Ser432 were indeed phosphorylated by Akt, we used this pan phospho-Ser antibody as above and found phosphorylation of WT USP14, but not of S143A/S432A mutant USP14, after incubating with activated Akt in a kinase assay (Figure 2A). To differentiate the relative importance of Ser143 and Ser432 as phosphorylation sites by Akt, we overexpressed activated Akt (Myr-Akt) in HEK293T cells with WT, S143A, S432A or double S143A/S432A (AA) mutants. We found that S143A mutant showed partially reduced phosphorylation as compared to that of WT, whereas phosphorylation of the USP14 S432A mutant was significantly decreased and that of AA double mutant was completely eliminated (Figure 2B). These results suggested S432 as a major and S143 as a minor phosphorylation site of Akt.

The phosphorylation of USP14 by Akt was further confirmed using an Akt phosphorylation-consensus motif (R××S/T) antibody (Figure 2-figure supplement 1A). The reactivity of USP14 with pan phospho-Ser antibody was eliminated after incubation with lambda phosphatase (Figure 2C). Notably, the phosphorylation levels of USP14 were decreased in cells when treated with MK2206, an inhibitor of Akt (Figure 2D), or when serum deprived, a condition known to inactivate endogenous Akt (Zhang et al., 2015) (Figure 2D).

To further verify the phosphorylation of USP14 S432 by Akt, we developed a phospho-Ser432 specific antibody. Phosphorylation of S432 can be detected after incubation of WT, but not S432A mutant USP14, with recombinant activated Akt in a kinase reaction (Figure 145 2E). This was further confirmed by using phos-tag electrophoresis which can specifically retard the migration of phosphorylated protein species (Kinoshita et al., 2009) (Figure 2E). Expression of Myr-Akt also led to S342 phosphorylation of endogenous USP14 (Figure 2F). Treatment with  either MK2206 or AZD5363, two structurally unrelated Akt inhibitors, led to decrease of USP14 S432 phosphorylation levels (Figure 2-figure supplement 1B-C). Moreover, treatment with PI3K inhibitors, either Wortmannin or GDC0941, but not ERK1/2 inhibitor U0126, also significantly decreased the phosphorylation levels of USP14 S432 (Figure 2-figure supplement 152 1D-E). In addition, we tested growth factors such as IGF-1 or EGF, both of which are known to promote activation of Akt. We found that the treatment of IGF-1 or EGF resulted in phosphorylation of USP14 S432, which was blocked in cells pre-treated with MK2206 (Figure 155 2G-H). Finally, USP14 S432 is dramatically more phosphorylated in PTEN knockout mouse embryonic fibroblasts (MEFs), which carry high levels of Akt activity, than that of WT MEFs as determined by western blotting using the phospho-USP14(S432) antibody and phos-tag electrophoresis (Figure 2I), and the phosphorylation of USP14 S432 was blocked by Akt inhibitors (Figure 2-figure supplement 1F). From these results, we conclude that Ser432 of USP14 is a major phosphorylation site by Akt.

Figure 2. USP14 is phosphorylated at Ser432 by activated Akt. (A) In vitro phosphorylation 521 of USP14 at S432 by Akt. Bacterially expressed and purified wide type USP14 or AA mutant incubated with active Akt in the presence of ATP. Reaction products were resolved by SDS-PAGE, and phosphorylation was detected by the phospho-Ser antibody. (B) Akt phosphorylates USP14 at S432 in vivo. Western blot analysis of whole cell lysate and immunoprecipitates derived from HEK293T cells transfected with wild type USP14, USP14 S143A, USP14 S432A and USP14 S143A/S432A (AA) constructs using the phospho-Ser antibody. L.E., long exposure. (C) Immunoprecipitation (IP) and IB analysis of HEK293T cells transfected with HA-USP14 and Myr-Akt and preincubated with or without λ-phosphatase as indicated. (D) Inhibition of Akt decreased exogenous USP14 phosphorylation. HEK293T cells were transfected with Myc-USP14 for 20 h then treated with 1 μM MK2206 or deprived of serum for another 4 h before harvest. (E) In vitro kinase assay to detect Akt phosphorylation of USP14 by phospho-Ser432 specific antibody and phos-tag-containing gels. Bacterially expressed and purified wide type USP14 or S432A mutant was incubated with active Akt in the presence of ATP. The reaction products were resolved by SDS-PAGE, and USP14 phosphorylation was detected using an antibody that specifically recognizes Ser432 phosphorylation of USP14 or determined by differential migration on phos-tag gels. (F) In vivo detection of endogenous USP14 Ser432 phosphorylation by anti-p-Ser432 specific antibody. Western blot analysis of immunoprecipitates derived from H4 cells transfected with or without Myr-Akt plasmids using the anti-p-Ser432 specific antibody. (G, H) Phosphorylation of endogenous USP14 S432 upon 540 stimulation with IGF-1 or EGF. HEK293T cells were serum-starved and pre-treated with Akt inhibitor MK2206 (1 μM) for 30 min before stimulation with IGF-1 (100 ng/mL) for 30 min (G) or EGF (100 ng/mL) for 1 h (H). The cell lysates were immunoprecipitated with USP14 antibody and western-blotted with anti-p-S432 antibody. (I) Phosphorylation of endogenous USP14 S432 in Pten knockout cells with high activity of Akt. Lysates from MEFs with indicated genotypes 545 were immunoprecipitated with USP14 antibody and then western-blotted with p-S432 antibody. The differential migration of phospho-USP14 on phos-tag-containing gels was determined as shown in the bottom panel.

Activation of USP14 by Akt mediated phosphorylation Because bacterially expressed and purified USP14 protein exhibits very low catalytic activity (Lee et al., 2010), we tested whether Akt-mediated phosphorylation might activate the DUB activity of USP14. We compared the activity of recombinant USP14 in a Ub-AMC (ubiquitin-7-amido-4-methylcoumarin, a fluorogenic substrate) hydrolysis assay in the presence or absence of Akt. Bacterially expressed and purified USP14 (Figure 3-figure supplement 1) showed trace hydrolyzing activity towards Ub-AMC as reported (Lee et al., 2010), while USP14 incubated with Akt showed high activity (Figure 3A). To validate Akt-mediated activation of USP14 in cells, we co-expressed USP14 and Myr-Akt in HEK293T cells. USP14 immunoprecipitated from cells co-expressing activated Akt showed higher activity in Ub-AMC assay than that expressed alone (Figure 3B). On the other hand, USP14 isolated from HEK293T cells incubated with Akt inhibitor MK2206 showed reduced activity in Ub-AMC assay (Figure 3C). Moreover, USP14 isolated from HEK293T cells stimulated with IGF-1 showed higher
activity, which was suppressed when cells were pre-treated with MK2206 (Figure 3D). To determine the specific contribution of Ser432, we compared the activity of USP14 S432A mutant protein in Ub-AMC assay with that of WT in the presence of Akt, and found that the stimulating effect of Akt on the hydrolyzing activity of USP14 was largely blocked by S432A mutation (Figure 3E), but not by S143A mutation (Figure 3-figure supplement 2B).

To further characterize the effect of Ser432 phosphorylation, we expressed and purified recombinant S432E USP14 protein, which mimics the phosphorylation state of USP14, from E. coli (Figure 3-figure supplement 1) and analyzed its activity by Ub-AMC assay. Interestingly, we found that USP14 S432E mutant protein alone showed high levels of Ub-AMC hydrolyzing activity (Figure 3F). Consistent with S432 as the major phosphorylation site by Akt, double E mutant (S143E/S432E) showed almost the same levels of hydrolyzing activity as that of S432E single mutant and S143E mutation had no significant impact on the activity of USP14 (Figure 3-figure supplement 2C-D). To determine its enzyme kinetics, we incubated USP14 S432E mutant protein with increasing amounts of Ub-AMC (Figure 3-figure supplement 2E) and determined the Km value (Km = 26 μM) from the slope of a Lineweaver-Burk plot (Figure 3G).

We characterized the distributions of p-S432 USP14 and total USP14 with that of proteasome in Pten-/- MEFs using glycerol gradient centrifugation (Koulich et al., 2008). We found that majority of p-S432 USP14 was distributed in the fractions with lower molecular weight proteins and distinguishable from the fractions where larger protein complexes, such as proteasomes, were localized. On the other hand, unphosphorylated USP14 was found in the fractions where larger molecular weight complexes, such as proteasome, are known to be localized (Figure 3-figure supplement 2F). Thus, S432 phosphorylated and unphosphorylated USP14 might be distributed differently in the cells. We next determined whether phospho-mimetic mutant of USP14 could be further activated by interacting with proteasome. Interestingly, we found that the Ub-AMC hydrolytic activity of S432E mutant could be further 200 activated when incubated with proteasome in vitro (Figure 3H). Taken together, these results suggest that S432 phosphorylation and intraction with proteasome may be two different
regulatory mechanisms for USP14.

Figure 3. Phosphorylation of USP14 by Akt activates USP14 DUB activity. (A) Akt activates USP14 DUB activity in vitro. USP14 protein (1μg) was incubated with or without active Akt (1 μg) in kinase assay buffer in a total volume of 50 μL for 1 h at 30oC, then the reaction mixtures were subjected to Ub-AMC assay. RFU, relative fluorescence units. (B, C) Akt activates USP14 in cells. USP14 was immunoprecipitated from HEK293T cells co-expressed with activated Akt  (B) or treated with 10 μM MK2206 for 4h (C) and then eluted with HA-peptide following Ub-AMC hydrolysis assay. (D) Activation of USP14 by stimulating cells with IGF-1. HEK293T cells were serum-starved and pre-treated with or without Akt inhibitor MK2206 (1 μM) for 30 min before stimulation with IGF-1 (100ng/mL) for 30 min. USP14 was then immunoprecipitated and eluted with HA-peptide. The activity of USP14 was determined using Ub-AMC hydrolysis assay. (E) USP14 activation by Akt is blocked by S432A mutation. Ub-AMC hydrolysis assay of wide type USP14 or S432A mutant in the presence or absence of active Akt. (F) Ub-AMC hydrolysis assay of bacterially expressed and purified wide type USP14 or S432E mutant. (G) Lineweaver-Burk analysis of USP14 S432E, obtained by measuring the initial rates at varying Ub-AMC concentrations (see Figure 3-figure supplement 2E for reference). (H) The activity of phospho-mimetic USP14 mutant can be further stimulated by the presence of proteasome. Ub-AMC hydrolysis assay of wild type USP14 or S432E mutant in the presence or absence of Ub-VS-treated human proteasome [VS-proteasome (see Lee et al., 2010); 1 nM]. Ptsm, 26S proteasome.

Phosphorylation of USP14 promotes both K48 and K63 deubiquitination activity  To assess the impact of USP14 phosphorylation on its selectivity towards different types of 206 ubiquitin linkages, we incubated USP14 WT and S432E mutant protein with diubiquitin species of K48, K63 and linear linkages. Conversion to monomeric Ub was monitored via SDS-PAGE followed by western blotting. We observed significantly increased hydrolytic activity of S432E mutant, as compared to that of WT, towards both Lys48 and Lys63 diubiquitin, while linear diubiquitin was not readily cleaved by WT or mutant USP14 (Figure 4A-B and Figure 4-figure supplement 1A). Similarly, immunoprecipitated USP14 from cells showed significant activity toward both Lys48 and Lys63 diubiquitin, but not linear diubiquitin (Figure 4-figure supplement 1B-C). In contrast, S432A mutant immunoprecipitated from cells showed lower activity towards both Lys48 and Lys63 diubiquitin than that of WT (Figure 4C). Regulation of ubiquitin-proteasome system by Akt depends on phosphorylation of USP14. Since USP14 is a negative regulator of the UPS (Koulich et al., 2008; Lee et al., 2010; Lee et al., 2011) and we found USP14 can be phosphorylated and activated by Akt, we reasoned that 219 Akt-mediated activation of USP14 might lead to inhibition of the ubiquitin-proteasome system (UPS) and generally enhance the stability of many proteins. To this end, we generated a stable 221 cell line expressing GFP-CL1 (also known as GFPu), an engineered ubiquitin-dependent proteasome substrate widely used as a reporter for UPS activity (Bence et al., 2001; Kelly et al., 2007; Li et al., 2013; Liu et al., 2014) (Figure 5-figure supplement 1A-C). Treatment of cells with Akt inhibitors or serum deprivation or PI3K inhibitor, all of which can block Akt activity (Zhang et al., 2015), led to reduced level of GFP-CL1 as detected by both western blotting and fluorescence microscopy (Figure 5A-C and Figure 5-figure supplement 1D). Conversely, the expression of activated Akt (Myr-Akt) led to increased levels of GFP-CL1 protein. Treatment of WT H4 cells with IGF-1 or EGF also led to increased levels of GFP-CL1 protein (Figure 5D-G and Figure 5-figure supplement 1E). In contrast, in USP14 knockout H4 cells (generated using CRISPR/Cas9 technology, Figure 5-figure supplement 2A-D), the expression of Myr-Akt did not affect the levels of GFP-CL1 (Figure 5H). From these results, we conclude that Akt 232 negatively regulates the UPS in an USP14-dependent manner.

We next tested the importance of USP14 phosphorylation for Akt to regulate UPS. We found that in contrast to USP14 WT reconstituted H4 cells, USP14 AA mutant reconstituted H4 cells showed no increase in the accumulation of GFP-CL1 in response to the expression of activated Akt (Figure 5-figure supplement 2E and Figure 5I). As a control, we found that the expression of Akt had no effect on a ubiquitin-independent substrate of the proteasome, C-terminal ornithine decarboxylase-GFP (GFP-cODC) (Hoyt et al., 2005; Kelly et al., 2007; Lee et al., 2010) (Figure 5-figure supplement 2F-G), suggesting that Akt does not inhibit the UPS through a general inhibition of the proteasome itself. Taken together, these data show that 241 phosphorylation of USP14 by Akt is important for this kinase to negatively regulate the UPS in a ubiquitin-dependent manner.

Phosphorylation of USP14 regulates global protein degradation To further understand the physiological roles of Akt-mediated USP14 phosphorylation and subsequently activation, we sought to study the impact of USP14 phosphorylation on global protein degradation. Since the loss of USP14 accelerates cellular proteolysis (Koulich et al., 2008; Lee et al., 2010), we performed a quantitative proteomic analysis to determine the levels of proteins in WT H4 cells, H4 USP14-KO cells, and H4 USP14-KO cells complemented with WT USP14, S143A/S432A (AA) or S143D/S432D (DD) mutants. Using an isobaric TMT labeling approach, our mass spectrometry analysis identified 18,400 peptides with high confidence (q<0.01), corresponding to 3,648 proteins with a minimum of two peptides from each protein. 2,763 proteins, which were quantified in at least 2 replicates, were subjected to further analysis. We found the global protein patterns of H4 USP14-KO cells were similar to those of H4 USP14 KO-AA cells, but distinct from those of WT H4 cells. We identified a common set of 87 proteins that were reduced in H4 KO cells as compared to H4 WT cells or to H4 KO cells complemented with WT USP14 (KO-WT) (Figure 6, Lane1-2). The levels of these proteins were also significantly reduced in H4 KO-AA cells (Figure 6, Lane 3). Importantly, the levels of this set of 87 proteins in H4 KO-DD cells were significantly higher than that of H4 KO-AA cells (Figure 6, Lane 4).

Figure 6. Phosphorylation of USP14 regulates global protein degradation. The quantitative 605 analysis of proteome change in USP14 knockout or USP14 mutant cells were performed by 606 TMT-isobaric labeling followed by shotgun analysis. The heat map was plotted based on the set of 87 proteins that are down-regulated greater than or equal to 1.2 fold in H4 KO cells compared to H4 WT cells or to H4 KO cells complemented with WT USP14 (KO-WT). The log base 2 of average ratios was plotted as indicated.

To verify that the identified changes in protein abundance were due to proteasomal degradation, we treated H4 KO-AA cells with proteasome inhibitor MG132 and analyzed the protein level change of these 87 proteins. We found that the levels of these proteins increased significantly in MG132-treated KO-AA cells compared to that of control KO-AA cells (Figure 6, Lane 5), suggesting that these proteins were indeed subject to an increased rate of proteasome degradation with expression of non-phosphorylatable USP14. Interestingly, the top hit on this list of 87 proteins that were differentially regulated upon the loss of USP14 is mTOR, a central established regulator of cellular metabolism and tumorigenesis. We confirmed the role of USP14 on the levels of mTOR by western blotting. We found that the levels of mTOR were reduced inH4 KO and H4 KO cells complemented with USP14 AA mutant, but restored upon the expression of USP14 DD mutant (Figure 6-figure supplement 1). Taken together, our results suggest that phosphorylation of USP14 may provide a mechanism for Akt to regulate global protein degradation through the proteasome, which in turn may control key cellular pathways involved in regulating metabolism and tumorigenesis.

NF-kB-Independent Role of IKKa/IKKb in Preventing RIPK1 Kinase-Dependent Apoptotic and Necroptotic Cell Death during TNF Signaling

Yves Dondelinger, Sandrine Jouan-Lanhouet, Tatyana Divert, …, Emmanuel Dejardin, Peter Vandenabeele, Mathieu J.M. Bertrand
Molecular Cell 2015; 60, 1–14


  1.  IKKa/IKKb prevent RIPK1 kinase-dependent death independently of NF-kB activation
  2.  IKKa/IKKb directly phosphorylate RIPK1 in TNFR1 complex I
  3.  Impaired phosphorylation of RIPK1 correlates with enhanced binding to FADD/caspase-8
  4.  IKK kinase inhibition induces TNF-mediated RIPK1 kinasedependent cell death in vivo

In Brief Dondelinger et al. describe an unexpected NF-kB-independent function of the IKK complex in protecting against TNF-induced RIPK1 kinase-dependent cell death. In TNFR1 complex I, IKKa/ IKKb directly phosphorylates RIPK1, leading to a reduction in RIPK1’s ability to bind FADD/caspase-8 and to induce apoptosis.

TNF is a master pro-inflammatory cytokine. Activation of TNFR1 by TNF can result in both RIPK1-independent apoptosis and RIPK1 kinase-dependent apoptosisornecroptosis.Thesecelldeathoutcomes are regulated by two distinct checkpoints during TNFR1 signaling. TNF-mediated NF-kB-dependent induction of pro-survival or anti-apoptotic molecules is a well-known late checkpoint in the pathway, protecting cells from RIPK1-independent death. On the other hand, the molecular mechanism regulating the contribution of RIPK1 to cell death is far less understood. We demonstrate here that the IKK complex phosphorylates RIPK1 at TNFR1 complex I and protects cells from RIPK1 kinase-dependent death, independent of its function in NF-kB activation. We provide in vitro and in vivo evidence that inhibition of IKKa/IKKb or its upstream activators sensitizes cells to death by inducing RIPK1 kinase-dependent apoptosis or necroptosis. We therefore report on an unexpected, NF-kB-independent role for the IKK complex in protecting cells from RIPK1-dependent death downstream of TNFR1.

The IkB kinase (IKK) complex, composed of the regulatory subunit NEMO (also known as IKKg) and the two catalytic subunits IKKa and IKKb, plays a central role in the induction of immune and inflammatory responses as well as in promoting cell survival and tumorigenesis (Baldwin, 2012; Baud and Karin, 2009; Hayden and Ghosh, 2012; Liu et al., 2012). Its activation constitutes the ignition phase of the canonical NF-kB pathway, which
ultimately results in the translocation of NF-kB dimers to the nucleus, where they promote transcription of a myriad of genes involved in inflammation, survival, and tumorigenesis.

TNF is a master pro-inflammatory cytokine, and inappropriate TNF signaling has been demonstrated to drive many inflammatory diseases. Activation of TNFR1 by TNF promotes inflammation either directly by activating the canonical NF-kB pathway or indirectly by promoting cell death, which exacerbates inflammation by releasing damage-associated molecular patterns (DAMPs) as well as by affecting the permeability of the bodily barriers to microbes (Pasparakis and Vandenabeele, 2015). In most cell types, activation of TNFR1 does not induce death but triggers canonical NF-kB-dependent transcriptional upregulation of genes encoding pro-survival and pro-inflammatory molecules. Ligation of TNF to trimeric TNFR1 induces the rapid assembly of a plasma membrane-bound signaling complex, known as complex I, that contains TRADD, RIPK1, and the E3 ubiquitin ligases TRAF2, cIAP1, cIAP2, and LUBAC (Walczak, 2011). The conjugation of ubiquitin chains to RIPK1 by cIAP1/ cIAP2 generates binding sites for TAB2/TAB3 and NEMO and allows further recruitment and activation of TAK1 and IKKa/ IKKb (Bertrand et al., 2008; Ea et al., 2006; Gerlach et al., 2011; Kanayama et al., 2004; Mahoney et al., 2008; Wu et al., 2006). TAK1 activates the IKK complex by phosphorylation, resulting in the rapid and selective IKK-mediated phosphorylation of IkBa and in its subsequent ubiquitylation-dependent proteasomal degradation. IkBa degradation then permits translocation of the NF-kB heterodimer p50/p65 to the nucleus, where it induces transcription of multiple responsive genes, including pro-survival genes such as cFLIP (Hayden and Ghosh, 2014). The anti-apoptotic potential of cFLIP resides in its ability to counteract activation of caspase-8 from a cytosolic TRADD-FADD-caspase-8 cytosolic complex, named complex IIa, which is believed to originate from complex I internalization (Irmler et al., 1997; Micheau and Tschopp, 2003; Wang et al., 2008; Wilson et al., 2009). Accordingly, TNFR1-mediated RIPK1-independent apoptosis requires inhibition of the NF-kB response (Van Antwerp et al., 1996), commonly obtained in vitro by the use of pharmacological inhibitors of transcription or translation, respectively, Actinomycin D (ActD) and cycloheximide (CHX).

The NF-kB-mediated induction of pro-survival/anti-apoptotic molecules is, however, not the only cell death checkpoint in the TNFR1 pathway (O’Donnell and Ting, 2011). Indeed, altering activation of the canonical NF-kB pathway by inhibiting components located upstream of IkBa, namely, cIAP1/cIAP2, TAK1, and NEMO, was reported to further sensitize cells to death by additionally inducing RIPK1-dependent death (Dondelinger et al., 2013; Legarda-Addison et al., 2009; O’Donnell et al., 2012). Depending on the cellular context, activated RIPK1 accelerates cell death either by promoting assembly of a RIPK1FADD-caspase-8 cytosolic apoptotic complex, referred to as complex IIb (Wilson et al., 2009), or by promoting necroptosis via activation of the RIPK3-MLKL pathway (Pasparakis and Vandenabeele, 2015). Although initiated by cIAP1/cIAP2-mediated ubiquitylation of RIPK1 in complex I, the last molecular step in the regulation of this early RIPK1 kinase-dependent cell death checkpoint is currently unknown. In this study, we demonstrate that RIPK1 is a bona fide substrate of IKKa and IKKb and that IKKa/IKKb-mediated phosphorylation of RIPK1 in complex I protects cells from RIPK1 kinase-dependent death.

NEMO Deficiency and IKKa/IKKb Double Deficiencies Induce TNFR1-Mediated RIPK1 Kinase-Dependent Apoptosis We previously reported that the ubiquitin chains conjugated to RIPK1 by cIAP1/cIAP2 do not constitute the ultimate step regulating the contribution of RIPK1 to TNF-induced cell death. Indeed, genetic or pharmacological inhibition of TAK1 also drivesRIPK1-dependentdeathwithoutaffectingRIPK1ubiquitylation in complex I (Dondelinger et al., 2013). In this study, we investigated the role of the IKK complex in the regulation of this cell death checkpoint. Indeed, the IKK complex lies between TAK1andIkBainthepathway,andalthoughexpressionofaproteasome-resistant form of IkBa (IkBaSR) induces RIPK1-independent apoptosis (Dondelinger et al., 2013), NEMO deficiency was reported to sensitize cells to TNF-induced death by additionally promoting RIPK1-dependent apoptosis (Legarda-Addison et al., 2009). In absence of cIAP1/cIAP2 or TAK1, TNF-mediated RIPK1-dependent apoptosis was shown to rely on RIPK1 kinase activity (Dondelinger et al., 2013; Wang et al., 2008). To test whether this is also true in absence of NEMO, we first stimulated NEMO-deficient mouse embryonic fibroblasts (MEFs) with TNF in the absence or presence of Nec-1, a RIPK1 kinase inhibitor. Interestingly, we found that Nec-1 greatly, but not entirely,protectedNemo/MEFsfromTNF-induced apoptosis, as monitored by cell permeability, caspase-3 activity, and caspase-3 and caspase-8 processing (Figures 1A–1D, 1K, and 1L). These results indicated that, similarly as cIAP1/cIAP2 and TAK1, NEMO also regulates both RIPK1 kinase-dependent and RIPK1-independent cell death checkpoints downstream of TNFR1. To test whether this protective function of NEMO
reflects its role as adaptor protein recruiting IKKa and IKKb to TNFR1 complex I, we next stimulated Ikka/, Ikkb/, and Ikka//Ikkb/ MEFs with TNF. Interestingly, while IKKa or IKKb single deficiency had little effect on apoptosis induction (Figures 1E–1H), their combined depletion mimicked the phenotype observed in the Nemo/ MEFs (Figures 1I–1L), suggesting redundant roles of IKKa and IKKb downstream of NEMO in preventing RIPK1-dependent apoptosis. To exclude the possibility that the phenotypes observed in the various MEF genotypes were originating from intrinsic defects due to clonal expansion, we confirmed our findings in Ripk1+/+ and Ripk1/ MEFs depleted of IKK proteins by siRNA (Figure S1). Of note, NEMO siRNA had little effect on cell death induction in these experiments, probably due to the poor efficiency in repressing NEMO.

Figure 1. NEMO Deficiency and IKKa/IKKb Double Deficiencies Induce TNFR1-Mediated RIPK1 Kinase-Dependent Apoptosis (A–L)MEFsoftheindicatedgenotypesweretreatedwith20ng/mlhTNFinthepresenceorabsenceofNec-1,andcelldeath(A,C,E,G,andI)andcaspaseactivity (B, D, F, H, and J) were measured in function of time, respectively, by SytoxGreen positivity and DEVD-AMC fluorescence. Protein levels were determined by immunoblotting in unstimulated cells (K) or 15 hr poststimulation with the indicated compounds (L). Forthe celldeath results, error bars represent theSEM ofthreeindependent experiments. Forthe caspase-3activity results, error bars represent SDof triplicates of one representative experiment. See also Figure S1.

IKKa and IKKb Mediate Their Protective Effect on RIPK1 via Their Enzymatic Activities Because IKKa and IKKb are serine/threonine kinases, we next evaluated the requirement of their enzymatic activities for their ability to repress RIPK1-dependent apoptosis. To do so, we tested the effect of five different IKK inhibitors on TNF-induced death and found that all of them led to a combination of RIPK1 kinase-dependent and RIPK1-independent death, as observed in the Ikka//Ikkb/ MEFs (Figures S2A and S2B). We further confirmed RIPK1 kinase-dependent apoptosis induction using TPCA-1 (Figures 2A–2C), as this inhibitor had no effect on TNF-induced death in Ikka//Ikkb/ MEFs (Figure S2B). TPCA-1 was used at 5 mM, a concentration reported to inhibit both IKKa and IKKb kinase activities (IC50 = 400 nM and 17.9 nM for IKKa and IKKb, respectively) (Podolin et al., 2005). We demonstrated that the apoptotic cell death was mostly depending on RIPK1 kinase activity by either co-incubating cells with Nec-1 (Figures 2A–2C) or by stimulating RIPK1 kinase-dead-expressing MEFs (Ripk1 K45A)(Figures 2D and 2E) (Berger et al., 2014). Importantly, Nec-1 had no effect in Ripk1 K45A MEFs, excluding any off-target effect (Figures S2E andS2F). Of note, similar results were obtained upon pharmacological inhibition of cIAP1/cIAP2 or TAK1 (Figures 2F, 2G, S2C, and S2D). In line with a role of IKKa and IKKb downstream of cIAP1/cIAP2, TAK1, and NEMO in the pathway, we tested the effect of TPCA-1 on TNF-induced death in ciap1/2/, Tak1/, and Nemo/ MEFs and found no additional effect (Figures 2H–2K). Together, these results indicate that the kinase activities of IKKa/IKKb regulate, downstream of cIAP1/cIAP2, TAK1, and NEMO, both RIPK1 kinasedependent and RIPK1-independent cell death checkpoints.

Figure 2. IKKa and IKKb Mediate Their Protective Effect on RIPK1 via Their Enzymatic Activities (A, B,and D–K)Ripk1+/+ or MEFsof theindicated genotypes weretreated with20ng/ml hTNF inthepresenceof theindicated compounds, and celldeath (A,D,F, H, I, J, K) and caspase-3 activity (B, E, G) were measured in function of time, respectively, by SytoxGreen positivity and DEVD-AMC fluorescence. (C) Protein levels in wild-type MEFs determined by immunoblotting 4 hr poststimulation. Forthecell death results,error bars represent the SEMof three independent experiments. Forthe caspase-3 activity results, error bars represent SDoftriplicates of one representative experiment. See also Figure S2.

IKKa/IKKb Protect Cells from RIPK1-Dependent Apoptosis Independently of NF-kB We previously demonstrated that, in absence of cIAP1/cIAP2 or TAK1, RIPK1 contribution to TNF-induced death is regulated independently of a defect in the canonical NF-kB-dependent upregulation of pro-survival genes (Dondelinger et al., 2013). Moreover, NEMO was also reported to inhibit RIPK1 activation in an NF-kB-independent manner (Legarda-Addison et al., 2009; O’Donnell et al., 2012). IKKa and IKKb are best known for their roles in NF-kB activation, but NF-kB-independent functions have also been reported (Hinz and Scheidereit, 2014). To confirm that IKKa and IKKb regulate RIPK1 activation independently of the canonical NF-kB response, we took two different approaches. In the first one, we tested the effect of inhibiting IKKa/IKKb in conditions where the NF-kB response is prevented by incubating the cells with the translational inhibitor CHX. In the second, we used p65/ MEFs, which are defective for canonical NF-kB activation (Beg et al., 1995). As previously reported (Wang et al., 2008), apoptosis induced by TNF+CHX occurred with a slow kinetic and independently of RIPK1 kinase activity (Figures 3A–3C). Remarkably, a pretreatment with TPCA-1 greatly sensitized cells to apoptosis, and this sensitization was prevented by Nec-1 (Figures 3A– 3C, S3A, and S3B). Similar results were obtained when stimu
lating NF-kB-deficient p65/ MEFs with TNF and TPCA-1 (Figures 3D–3F) or in combination with TAK1 and cIAP1/ cIAP2 inhibitors (Figures S3C and S3D). Together, these results demonstrate that RIPK1-independent and -dependent apoptotic pathways are regulated by two different cell death checkpoints downstream of TNFR1 and that IKKa/IKKb regulate both of them in NF-kB-dependent and -independent manners, respectively.

Figure 3. IKKa/IKKb Protect Cells from RIPK1-Dependent Apoptosis Independently of NF-kB (A,B,D,andE)Ripk1+/+ (AandB)orp65/(DandE)MEFswerestimulatedwith20ng/mlhTNFinthepresenceoftheindicatedcompounds,andcelldeath(Aand D) and caspase activity (B and E) were measured in function of time, respectively, by SytoxGreen positivity and DEVD-AMC fluorescence. (C and F) Ripk1+/+ (C) or p65/ (F) MEFs were stimulated for, respectively, 15 hr and 8 hr with the indicated compounds, and protein levels were determined by immunoblotting. Forthe celldeath results, error bars represent theSEM ofthreeindependent experiments. Forthe caspase-3activity results, error bars represent SDof triplicates of one representative experiment. See also Figure S3.

Defective RIPK1 Phosphorylation in Complex I Correlates with RIPK1 Kinase-Dependent Contribution to TNF-Induced Apoptosis Knowing that the IKK complex physically interacts with RIPK1 in complex I, we hypothesized that the kinase-dependent role of IKKa/IKKb in preventing RIPK1 kinase-dependent apoptosis results from its ability to phosphorylate RIPK1. To test this hypothesis, we analyzed whether RIPK1 is phosphorylated in complex I and whether its phosphorylation state is altered in conditions affecting activation of IKKa/IKKb, but not when the pathway is inhibited downstream of IKKa/IKKb. Because RIPK1 is highly ubiquitylated in complex I (which prevents the detection by immunoblot of potential mobility shifts resulting from its phosphorylation), we removed the ubiquitin chains conjugated to RIPK1 by incubating complex I, pulled-down using FLAG-TNF, with the deubiquitylase USP2 (Figure 4A). By doing so, we observed that the pool of deubiquitylated RIPK1 was running at a higher molecular weight than normal and confirmed, by l-phosphatase treatment, that this mobility shift was resulting from phosphorylation, but not auto-phosphorylation since it was not inhibited by Nec-1 or in Ripk1 K45A MEFs (Figures 4A–4C). Remarkably, and in line with the model of cIAP1/ cIAP2-mediated ubiquitylation-dependent recruitment of TAK1 and NEMO/IKKa/IKKb to complex I and with our cell death results, we found that RIPK1 phosphorylation in complex I is affected in ciap1/2/, Tak1/, Nemo/, and Ikka//Ikkb/, but not in Ikka/, Ikkb/, and p65/ MEFs or in MEFs preincubated with CHX (Figures 4D, 4E, 4G, and 4H). Importantly, IKK activity is greatly affected (as observed by IkBa phosphorylation) in all conditions in which we observed impaired RIPK1 phosphorylation, thereby further demonstrating the link between RIPK1 phosphorylation and IKK enzymatic activities (Figure 4E). Defective RIPK1 phosphorylation in complex I was also observed following pharmacological inhibition of cIAP1/ cIAP2, TAK1, or IKKa/IKKb (Figure 4F).

Figure 4. Defective RIPK1 Phosphorylation in Complex I Correlates with RIPK1 Kinase-Dependent Contribution to TNF-Induced Apoptosis (A–H) Ripk1+/+ MEFs (A, B, F, and H) or MEFs with the indicated genotype (C, D, E, and G) were stimulated for 5 min with 2 mg/ml FLAG-hTNF in the presence or absence of the indicated compounds. TNFR1 complex I was then FLAG immunoprecipitated, incubated with the deubiquitylating enzyme USP2 or lambda phosphatase (l PPase) when indicated, and RIPK1 ubiquitylation and phosphorylation finally analyzed by immunoblotting. * indicates an aspecific band. See also Figure S7

Direct Phosphorylation of RIPK1 by IKKa/IKKb Prevents RIPK1 from Integrating Complex IIb To test the direct contribution of IKKa/IKKb to RIPK1 phosphorylation, we next performed in vitro kinase assays using recombinant proteins and included Nec-1 in the reactions to prevent RIPK1 autophosphorylation. We found that both IKKa and IKKb directly phosphorylated full-length RIPK1 or a mutated version lacking the death and RHIM domain (RIPK11–479)(Figures 5A, 5B, and S4A). Of note, RIPK1 phosphorylation by IKKb induced a mobility shift of RIPK1 not detected when using IKKa (Figures 5A and 5B), suggesting some specificity in the residues phosphorylated by both kinases. In line with our cellular data, TPCA-1 repressed, although with different efficiencies, the direct phosphorylation of RIPK1 by IKKa and IKKb. In contrast, recombinant TAB1/TAK1 did notlead to detectable RIPK1 phosphorylation by autoradiography (Figure 5C).

We next tested the consequence of genetic or pharmacological inhibition of IKKa/IKKb, and of the resulting defective phosphorylation of RIPK1 in complex I, on the ability of RIPK1 to integrate the cytosolic caspase-8-activating complex IIb. We found, by performing FADD and caspase-8 immunoprecipitations, that inhibition of IKKa/IKKb enzymatic activities resulted in the binding of RIPK1 to FADD and caspase-8, a process relying on RIPK1 kinase activity (Figures 5D–5G). In contrast, CHX pre-treatment, which does not affect phosphorylation of RIPK1 in complex I (Figure 4H), led to much less recruitment of RIPK1 to FADD/caspase-8, and this recruitment was not inhibited by Nec-1 (Figures 5F, 5G, and S4B). Of note, association of TRADD with FADD/caspase-8 was not observed under these conditions. Together, these results suggest that IKKa/IKKb-mediated phosphorylation of RIPK1 either represses RIPK1 kinase activity or interferes with RIPK1’s ability to bind complex IIb components.

Figure 5. Direct Phosphorylation of RIPK1 by IKKa/IKKb Prevents RIPK1 from Integrating Complex IIb (A–C) Recombinant GST-IKKa, GST-IKKb, or GST-TAB1-TAK1 fusion protein was incubated with a recombinant truncated (GST-RIPK11–479) or full-length (GST-RIPK1FL) form of RIPK1 in a radioactive in vitro kinase assay in the presence of the indicated inhibitors. Phosphorylation was revealed by SDS-PAGE followed by autoradiography. (D–G) MEFs with the indicated genotype (D and E) or Ripk1+/+ MEFs (F and G) were pre-incubated with zVAD-fmk and with the indicated compounds for 30 min and then stimulated with 20 ng/ml hTNF. After 4 hr, complex II was isolated by FADD or caspase-8 immunoprecipitation and RIPK1 binding revealed by immunoblotting. See also Figure S4.

IKKa/IKKb Mediate In Vivo Protection to RIPK1 KinaseDependent Death To test the in vivo relevance of our in vitro findings, we evaluated the contribution of RIPK1 kinase activity to two different mouse models of TNF-induced death. In the first one, we injected Ripk1K45A/K45A and Ripk1+/+ littermates with TNF in association with D-galactosamine. In this well-known model of acute hepatitis, TNF-mediated hepatocyte apoptosis is reported to result from transcriptional inhibition (Decker and Keppler, 1974), thereby affecting the NF-kB pathway downstream of IKKa/ IKKb. In accordance with our in vitro results, we found that Ripk1K45A/K45A mice were not protected from TNF-induced lethality and apoptotic liver damage, as monitored by survival curves, blood levels of aspartate transaminase/alanine transaminase (AST/ALT), caspase-3 activation in the liver by DEVDase assays, and active caspase-3 staining (Figures 6A–6E). Blood levels of lactate dehydrogenase (LDH), a marker of necrosis, were not upregulated by TNF+GalN injection (Figure 6F).

In the next model, we injected Ripk1K45A/K45A and Ripk1+/+ littermates with a sub-lethal dose of TNF (5 mg) in presence or absence of TPCA-1 (10 mg/kg) to inhibit the canonical NF-kB pathway at the level of IKKa/IKKb. Remarkably, while TPCA-1 hadnotoxicity onitsown(FiguresS5AandS5B),itscombination with TNF resulted in the rapid death of all Ripk1+/+, but no Ripk1K45A/K45A, mice (Figure 6G). Accordingly, Ripk1K45A/K45A mice were protected from TNF-induced hypothermia and had no increase in serum levels of ASL/ALT or caspase-3 activation in the liver (Figures 6H–6L). In contrast to TNF+GalN injection, TNF+TPCA-1 led to a substantial increase of LDH levels in the serum that was also absent in Ripk1K45A/K45A mice, suggesting additional necroptosis induction (Figure 6M). Importantly, RIPK1 kinase inhibition by co-administration of Nec-1s (Degterev et al., 2013), a modified and more stable version of Nec-1, in C57BL/6J mice also significantly delayed the death and injury induced by TNF+TPCA-1 injection (Figures S5C–S5I).

Together, these in vivo results demonstrate TNF-mediated RIPK1-independent and RIPK1 kinase-dependent hepatocyte apoptosis in condition of NF-kB inhibition downstream or at the level of IKKa/IKKb, respectively.

Figure 6. IKKa/IKKb Mediate In Vivo Protection to RIPK1 Kinase-Dependent Death (A) Cumulative survival rates of littermate Ripk1+/+ and Ripk1K45A/K45A C57BL/6J females injected with GalN 15 min prior to injection with mTNF (n = 5). (B, C, and F) Blood AST (B), ALT (C), and LDH (F) levels determined 3 hr post-TNF injection (Ripk1+/+ n = 3, and Ripk1K45A/K45A n = 4). (D and E) Caspase-3 activity in liver samples (Ripk1+/+ n = 3, and Ripk1K45A/K45A n = 4) isolated 3 hr post-TNF injection and determined by Ac-DEVD-AMC fluorescence assay (D) or anti-cleaved caspase-3 staining (E). (G) Cumulative survival rates of littermate Ripk1+/+ and Ripk1K45A/K45A C57BL/6J females injected with TPCA-1 20 min prior to injection with mTNF (n = 5). (H) Body temperature as a function of time. (I, J, and M) Blood AST (I), ALT (J), and LDH (M) levels determined 3 hr post-TNF injection (n = 4). (K and L) Caspase-3 activity in liver samples (n = 4) isolated 3 hr post-TNF injection and determined by Ac-DEVD-AMC fluorescence assay (K) or anti-cleaved caspase-3 staining (L). Scale bar, 25 mm. Error bars represent the SEM of the indicated n values. See also Figure S5.

IKKa/IKKb Protect Cells from RIPK1 Kinase-Dependent Necroptosis Independently of NF-kB Our in vivo results suggested that TNF+TPCA-1 additionally induced necroptosis in the injected mice. To test the possibility that IKKa/IKKb also regulates RIPK1 kinase-dependent necroptosis independently of NF-kB, we in vitro stimulated MEFs with TNF+CHX in the presence of the pan caspase inhibitor zVAD-fmk and of TPCA-1. As shown in Figure 7A,TNF-mediated
necroptosis induced by TNF+CHX+zVAD is fully repressed by Nec-1 but still greatly enhanced by additionally inhibiting IKKa/ IKKb with TPCA-1 (Figures 7A and S6A). The mouse fibrosarcoma cell line L929sAhFAS is a prototypic model for necroptosis since these cells succumb by necroptosis upon single TNF stimulation. While inhibiting NF-kB by CHX sensitized these cells to necroptosis, the sensitization was again enhanced when IKKa/ IKKb was additionally inhibited by TPCA-1 (Figures 7B and S6B). Our results therefore demonstrate that IKKa/IKKb prevent RIPK1 kinase-dependent apoptosis and necroptosis downstream of TNFR1 independently of their known function in protecting the cells from death by mediating NF-kB-dependent upregulation of pro-survival/anti-death genes.

RIPK1 kinase-dependent necroptosis relies on the downstream activation of the RIPK3-MLKL pathway (Cho et al., 2009; He et al., 2009; Pasparakis and Vandenabeele, 2015;Sun et al., 2012; Zhang et al., 2009; Zhao et al., 2012). To further characterize the contribution of RIPK3 to the lethality resulting from the in vivo injection of TNF+TPCA-1, we challenged Ripk3+/+ and Ripk3/ littermates with this trigger. Contrary to Ripk1K45A/K45A mice, Ripk3/ mice were greatly, but not entirely, protected from death and hypothermia induced by TNF+TPCA-1 (Figures 7C and 7D). Interestingly, the protection was not originating from the liver, as RIPK3 deficiency did not prevent liver damage (Figures 7E–7H). Instead, RIPK3 deficiency prevented the increased of LDH levels in the blood, resulting from necrosis of undefined organ(s) (Figure 7I). These in vivo results therefore suggest that the lethality induced by TNF+TPCA-1 results from both RIPK1 kinase-dependent apoptosis and necroptosis.

Figure 7. IKKa/IKKb Protect Cells from RIPK1 Kinase-Dependent Necroptosis Independently of NF-kB (A and B) Ripk1+/+ MEFs (A) and L929sAhFas cells (B) were stimulated with hTNF (20 ng/ml in A and 33 pg/ml in B) in the presence of the indicated compounds, and cell death was measured as a function of time by SytoxGreen positivity. (C) Cumulative survival rates of littermate Ripk3+/+ and Ripk3/ C57BL/6J females injected with TPCA-1 20 min prior to injection with mTNF (Ripk3+/+ n = 4, and Ripk3/ n = 7). (D) Body temperature as a function of time. (E, F, and I) Blood AST (E), ALT (F), and LDH (I) levels determined 3 hr post-TNF injection (Ripk3+/+ n = 4, and Ripk3/ n = 3). (G and H) Caspase-3 activity in liver samples (Ripk3+/+ n = 4, and Ripk3/n = 3) isolated 3 hr post-TNF injection and determined by Ac-DEVD-AMC fluorescence assay (G) or anti-cleaved caspase-3 staining (H). Scale bar,25mm.Fortheinvitrocell death results,error bars represent the SEM of three independent experiments. For the in vivo results,error bars represent the SEM of the indicated n values

Sensing of TNF by TNFR1 at the cell surface can paradoxically result in the activation of signaling pathways with opposite consequences: cell survival or cell death. The fact that survival is the dominant outcome in most cell types indicates the existence of molecular mechanisms actively repressing TNFR1-mediated cell death.Two major mechanisms have been reported to control cell death downstream of TNFR1 (O’Donnell and Ting, 2011).The first identified one is well characterized and consists in a relatively slow process involving the NF-kB-dependent induction of pro-survival/anti-death molecules, such as cFLIP (Karin and Lin, 2002; Kreuz et al., 2001; Liu et al., 1996; Micheau et al., 2001; Panayotova-Dimitrova et al., 2013; Van Antwerp et al., 1996; Wang et al., 1998). The second one, which is less understood and more recently reported, is believed to take place at an earlier stage following TNFR1 activation and is shown to be independent of the NF-kB response (Dondelinger et al., 2013; Legarda-Addison et al., 2009; O’Donnell et al., 2007, 2012; Wang et al., 2008). Interestingly, while the first checkpoint regulates slow apoptosis by inhibiting activation of complex IIa (TRADD-FADD-caspase-8), the second one regulates the contribution of RIPK1 to cell death by either preventing RIPK1 from integrating the apoptotic complex IIb (RIPK1-FADD-casapase-8) or by limiting its contribution to the necrosome (RIPK1RIPK3-MLKL) (Cho et al., 2009; He et al., 2009; Sun et al., 2012; Vanlangenakker et al., 2011; Wang et al., 2008; Wilson et al., 2009; Zhang et al., 2009; Zhao et al., 2012). It has long been thought that IKKa/IKKb inhibits TNF-induced cell death through activation of the NF-kB pathway. In this study, we provide evidences that IKKa and IKKb also regulate cell death by direct phosphorylation of RIPK1 at the level of TNFR1 complex I.

TNF-induced RIPK1-dependent apoptosis was first described in conditions affecting cIAP1/cIAP2-mediated RIPK1 ubiquitylation (Bertrand et al., 2008; O’Donnell et al., 2007; Petersen et al., 2007; Wang et al., 2008), which led to the hypothesis that the ubiquitin chains on RIPK1 were directly preventing its binding to FADD, keeping RIPK1 in a survival modus. This‘‘direct’’effect, however, has later been challenged. Binding of the adaptor proteins TABs and NEMO to RIPK1 ubiquitin chains allows recruitment of TAK1 and of IKKa/IKKb to TNFR1 complex I (Ea et al., 2006; Li et al., 2006; Wu et al., 2006), and TAK1 inhibition was shown to result in TNF-mediated RIPK1-dependent apoptosis without affecting RIPK1 ubiquitylation status (Dondelinger et al., 2013). We show here that RIPK1 is phosphorylated in complex I and that affecting RIPK1 ubiquitylation by cIAP1/ cIAP2depletion directlyimpactsitsphosphorylation.Incontrast, TAK1, NEMO, or IKKa/IKKb depletion affects RIPK1 phosphorylation and induces RIPK1-dependent cell death but does not alter its ubiquitylation state in complex I. Together, these results indicate that ubiquitylation and phosphorylation of RIPK1 occur sequentially and that RIPK1 phosphorylation regulates its killing potential. Because activation of the IKK complex lies downstream of TAK1 but upstream of IkBa and p65, our results suggest a model in which IKKa/IKKb constitute the last step in the regulation of the RIPK1 cell death checkpoint (graphical abstract). Indeed, p65 deletion, expression of IkBaSR, or CHX pre-treatment induces TNF-mediated RIPK1-independent apoptosis and does not alter RIPK1 ubiquitylation or phosphorylation in complex I.

RIPK1 enzymatic activity is needed for the integration of RIPK1 to complex IIb and to the necrosome, which respectively drives apoptosis or necroptosis under TNF-stimulated conditions (Cho et al., 2009; Dondelinger et al., 2013; He et al., 2009; Wang et al., 2008). The precise role of RIPK1 kinase activity in these processes remains unclear but may involve autophosphorylation-driven conformational changes allowing increased binding of RIPK1 to the death complex components. The kinase activity of RIPK1 therefore requires active repression to avoid unnecessary cell death. A recent report suggests that RIPK1 phosphorylation on Ser89 suppresses its kinase activity (McQuade et al., 2013). It is therefore tempting to speculate that IKK-mediated phosphorylation of RIPK1 in complex I affects RIPK1 kinase activity. Alternatively, the phosphorylation of RIPK1 by IKKs may directly affect binding of RIPK1 to the death complex components or facilitate its dissociation from complex I. We performed mass spectrometry analysis to identify the residues of RIPK1 phosphorylated by IKKa and IKKb and found several sites, but not Ser89 (Figures S4C and S4D). Unfortunately, we were unable to demonstrate the direct physiological relevance of the identified phosphorylation sites due to the fact that all Ripk1/ reconstituted MEFs, even those with WT RIPK1 (irrespective of RIPK1 expression levels), started to succumb upon single TNF stimulation (data not shown), a problem previously reported (Gentleetal., 2011). The fact that the combined repression of IKKa and IKKb is needed to induce RIPK1 kinase-dependent death, and that the phosphorylation by each kinase results in different RIPK1 mobility shifts when run on gels, may indicate that phosphorylation on several residues is required to negatively regulate RIPK1.

IKKa and IKKb are best known for their roles in NF-kB activation, but NF-kB-independent functions have also been reported, some of which are even implicated in cell fate decisions (Hinz and Scheidereit, 2014). Using pharmacological inhibition of IKKa/IKKb in a p65-deficient background, or together with CHX, we demonstrated an NF-kB-independent function of IKKa/IKKb in protecting cells from TNF-induced RIPK1 kinasedependent apoptosis and necroptosis. In vivo, we demonstrate that TNF induces apoptosis of hepatocytes independently of RIPK1 when the NF-kB pathway is affected downstream of IKKa/IKKb (TNF+GalN). In contrast, pharmacological inhibition of the NF-kB pathway at the level of IKKa/IKKb (TNF+TPCA-1) sensitizesmicetoTNF-inducedshock,whichisaccompaniedby RIPK1 kinase-dependent, but RIPK3-independent, apoptosis of hepatocytes and RIPK1/RIPK3-dependent cellular death, presumably necroptosis, in undefined organs. We can indeed not formally rule out the possibility that the increase in serum LDH levels originates from secondary necrosis of apoptotic cells. Importantly, genetic and chemical inhibition of RIPK1 enzymatic activity protected the mice from TNF-induced cellular damage and death. These results therefore demonstrate the in vivo roles of IKKa/IKKb in protecting cells from RIPK1 kinase-dependent death.

The LUBAC complex, which includes its component Sharpin, is recruited to complex I during TNF signaling, and the inactivating mouse Sharpin cpdm mutation was reported to cause multi-organinflammationresultingfromTNF-mediated RIPK1kinase-dependent death (Berger et al., 2014; Kumari et al., 2014; Rickard et al., 2014). In line with our results, we found that TNF-mediated RIPK1 kinase-dependent death of mouse dermal fibroblasts (MDFs) isolated from Sharpincpdm mice is associated with defective RIPK1 phosphorylation in complex I (Figures S7A and S7B), probably resulting from the altered recruitment of IKK proteins to complex I, as previously reported for other LUBAC components (Haas et al., 2009). The genetic disruption of Nemo, Ikka, Ikkb, orIkka/Ikkb in mice results in early lethality with massive cellular death in several organs, such as the liver, the skin, and, in the case of Ikka//b/ mice, the nervous system (Hu et al., 1999; Li et al., 1999, 2000; Rudolph et al., 2000; Takeda et al., 1999). So far, these phenotypes have exclusively been explained by defects in NF-kB activation, but our study indicates that RIPK1 activation probably contributes to these pathological conditions. In the same line, RIPK1 kinase-dependent apoptosis may drive the spontaneous development of hepatocellular carcinoma observed in mice ablated of Nemo in the liver parenchymal cells (Luedde et al., 2007). Testing the contribution of RIPK1 to those phenotypes is an exciting future challenge, which may open doors for the use of chemical inhibitors of RIPK1 in the treatment of human diseases associated with IKK malfunctions, such as incontinentia pigmenti (Conte et al., 2014).

Translocation of interleukin-1β into a vesicle intermediate in autophagy-mediated secretion
Min Zhang1, Sam Kenny2, Liang Ge1, Ke Xu2 and Randy Schekman1*
eLife 2015;10.7554/eLife.11205

In this study, we probed the organelle association and molecular requirements for the secretion of one such unconventional cargo protein, IL-1β. Using surrogate cell lines rather than macrophages to reconstitute autophagy-mediated secretion of IL-1β (Figure 1), we find mature IL-1β localized to the lumen of the membrane in early intermediates and mature autophagosomes (Figures 2-4, 6). This surprising location may help to explain how mature IL-1β is secreted in a soluble form to the cell surface (Figure 9C). However, localization to the lumen between the two membranes of the autophagosome would require that IL-1β is translocated from the cytoplasm across the membrane precursor of a phagophore, rather than being engulfed as the phagophore membrane matures by closure into an autophagosome.

The exact route by which the autophagosome delivers mature IL-1β to the cell surface as well as how it avoids fusion with degradative lysosome remains obscure, possibly involving interaction with the multi-vesicular body or some form of lysosome as a prelude to fusion at the cell surface (Figure 9C), and this process may require selective recruitment of membrane sorting and targeting factors such as Rabs and SNAREs.

Recent evidence suggests that autophagy facilitates the unconventional secretion of the pro-inflammatory cytokine interleukin 1β (IL-1β). Here, we reconstituted an autophagy-regulated secretion of mature IL-1β (m-IL-1β) in non-macrophage cells. We found that cytoplasmic IL-1β associates with the autophagosome and m-IL-1β enters into the lumen of a vesicle intermediate but not into the cytoplasmic interior formed by engulfment of the autophagic membrane. In advance of secretion, m-IL-1β appears to be translocated across a membrane in an event that may require m-IL-1β to be unfolded or remain conformationally flexible and is dependent on two KFERQ-like motifs essential for the association of IL-1β with HSP90. A vesicle, possibly a precursor of the phagophore, contains translocated m-IL-1β and later turns into an autophagosome in which m-IL-1β resides within the intermembrane space of the double-membrane structure. Completion of IL-1β secretion requires Golgi reassembly and stacking proteins (GRASPs) and multi-vesicular body (MVB) formation.

Most eukaryotic secretory proteins with an N-terminal signal peptide are delivered through the classical secretion pathway involving an endoplasmic reticulum (ER)-to-Golgi apparatus itinerary (Lee et al., 2004; Schatz and Dobberstein, 1996). However, a substantial number of secretory proteins lack a classical signal peptide, called leaderless cargoes, and are released by unconventional means of secretion (Nickel and Rabouille, 2009; Nickel and Seedorf, 2008). The range of unconventional secretory cargoes encompasses angiogenic growth factors, inflammatory cytokines and extracellular matrix components etc. most of which play essential roles for development, immune surveillance and tissue organization (Nickel, 2003; Rabouille et al., 2012). Unlike a unified route for classical protein secretion, leaderless cargoes undergoing unconventional secretion employ multiple means of protein delivery, the details of which are largely unknown (Ding et al., 2012; Nickel, 2010; Rabouille et al., 2012; Zhang and Schekman, 2013).

IL-1β is one of the most intensely investigated cargoes of unconventional secretion. A biologically inactive 31 kDa precursor, pro-IL-1β, is made following initiation of the NF-κB signaling cascade. Pro-IL-1β is subsequently converted into the active form, the 17 kDa mature IL-1β, by the pro-inflammatory protease caspase-1 which is activated, in response to extracellular stimuli, after its recruitment to a multi-protein complex called the inflammasome (Burns et al., 2003; Cerretti et al., 1992; Rathinam et al., 2012; Thornberry et al., 1992). Interpretation of the mechanism of unconventional secretion of IL-1β is complicated by the fact that one of the physiologic reservoirs of this cytokine, macrophages, undergoes pyroptotic death and cell lysis under conditions of inflammasome activation of caspase-1. Indeed, many reports including two recent publications make the case for cell lysis as a means of release of mature IL-1β (Liu et al., 2014; Shirasaki et al., 2014). In contrast, other reports demonstrate proper secretion of mature IL-1β without cell lysis in, for example, neutrophils, which are nonetheless dependent on the inflammasome response to activate caspase-1 and secrete mature IL-1β (Chen et al., 2014).

Macroautophagy (hereafter autophagy) is a fundamental mechanism for bulk turnover of intracellular components in response to stresses such as starvation, oxidative stress and pathogen invasion (Mizushima and Levine, 2010; Yang and Klionsky, 2010). The process is characterized by the formation of a double-membrane vesicle, called the autophagosome, through the elongation and closure of a cup-shaped membrane precursor, termed the phagophore, to engulf cytoplasmic cargoes (Hamasaki et al., 2013; Lamb et al., 2013). Completion of autophagosome formation requires a sophisticated protein-vesicle network organized by autophagic factors, such as autophagy-related (ATG) proteins, and target membranes (Feng et al., 2014; Mizushima et al., 2011). Besides the degradative function, autophagy or ATG proteins have recently been implicated in multiple secretory pathways including the delivery of leaderless cargoes undergoing unconventional secretion, such as the mammalian pro-inflammatory cytokines IL-1β and IL-18, the nuclear factor HMGB1, and the yeast acyl coenzyme A-binding protein Acb1, to the extracellular space (Bruns et al., 2011; Dupont et al., 2011; Duran et al., 2010; Manjithaya and Subramani, 2011; Pfeffer, 2010; Subramani and Malhotra, 2013). The Golgi reassembly and stacking protein(s) GRASP(s) (GRASP55 and GRASP65 in mammals, dGRASP in Drosophila, GrpA in Dictyostelium and Grh1 in yeast) are required for autophagy-regulated unconventional secretion (Giuliani et al., 2011; Kinseth et al., 2007; Levi and Glick, 2007; Manjithaya et al., 2010).

Dupont et al. (2011) documented a role for autophagy in the secretion of mature IL-1β (Dupont et al., 2011), but how a protein sequestered within an autophagosome could be exported as a soluble protein was unexplained. Here, we sought to understand how conditions of starvation-induced autophagy could localize IL-1β into an autophagosomal membrane. We reconstituted the autophagy-regulated secretion of IL-1β in cultured cell lines and detected a vesicle intermediate, possibly an autophagosome precursor, containing mature IL-1β. Three-dimensional (3D) Stochastic Optical Reconstruction Microscopy (STORM) demonstrated that, after entering into the autophagosome, IL-1β colocalizes with LC3 on the autophagosomal membrane, which, together with an antibody accessibility assay and observations from biochemical assays, implies a topological distribution in the intermembrane space of the autophagosome. This distribution of IL-1β explains the mechanism accounting for its secretion as a soluble protein through either a direct fusion of autophagosome with the plasma membrane or via the MVB pathway. Quite aside from the possible complication of cell lysis, another body of work has suggested an unconventional pathway for the proper secretion of IL-1β. Pro-IL-1β lacks a typical signal peptide and the propeptide is processed in the cytosol rather than the ER (Rubartelli et al., 1990; Singer et al., 1988). Although mature IL-1β appears to be incorporated into a vesicular transport system, secretion is not blocked by Brefeldin A, a drug that blocks the traffic of standard secretory proteins form the Golgi apparatus (Rubartelli et al., 1990). Multiple mechanisms have been implicated in the unconventional secretion of IL-1β, including autophagy, secretory lysosomes, multi-vesicular body (MVB) formation and micro-vesicle shedding (Andrei et al., 1999; Andrei et al., 2004; Brough et al., 2003; Lopez-Castejon and Brough, 2011; MacKenzie et al., 2001; Qu et al., 2007; Verhoef et al., 2003). However, a clear demonstration of the mechanism for the entry of IL-1β into a vesicular carrier, e.g. the autophagosome, is lacking.

Reconstitution of autophagy-regulated IL-1β secretion

A dual effect of autophagy has been proposed on the secretion of IL-1β in macrophages (Deretic et al., 2012; Jiang et al., 2013). On one hand, induction of autophagy directly promotes IL-1β secretion after inflammasome activation by incorporating it into the autophagosomal carrier (Dupont et al., 2011). On the other hand, autophagy indirectly dampens IL-1β secretion by degrading components of the inflammasome as well as reducing endogenous triggers for inflammasome assembly, including reactive oxygen species (ROS) and damaged components, which are required for the activation of caspase-1 and the production of active IL-1β (Harris et al., 2011; Nakahira et al., 2011; Shi et al., 2012; Zhou et al., 2011).

To focus our study specifically on the role of autophagy in IL-1β secretion, we reconstituted a stage of IL-1β secretion downstream of inflammasome activation by co-expressing pro-IL-1β (p-IL-1β) and pro-caspase-1 (p-caspase-1) in non-macrophage cells. As shown in Figure 1A, the generation and secretion (~5%) of mature IL-1β (m-IL-1β) was achieved by co-expression of p-IL-1β and p-caspase-1 in HEK293T cells. Mature IL-1β was not produced or secreted without p-caspase-1, whereas a low level of secreted p-IL-1β (~0.2%) was detected with or without the expression of p-caspase-1. Furthermore, little cell lysis occurred during the treatment we used to induce IL-1β secretion: Much less precursor than mature IL-1β and little cytoplasmic tubulin was detected released into the cell supernatant during the 2 h incubation in starvation medium (Figure 1A). Starvation, a condition that stimulates autophagy, enhanced IL-1β secretion (~3 fold) and reduced the level of IL-1β in the cell lysates (Figure 1A, B). Inhibition of autophagy by the phosphatidylinositol 3-kinase (PI3K) inhibitors 3-methyladenine (3-MA) or wortmannin (Wtm) blocked IL-1β secretion activated by starvation and caused the accumulation of mature IL-1β in the cell (Figure 1B). Likewise, in an autophagy-deficient cell line, Atg5 knockout (KO) mouse embryo fibroblasts (MEFs) (Mizushima et al., 2001), IL-1β secretion was reduced and failed to respond to starvation (Figure 1C). Moreover, IL-1β secretion was also inhibited in a dose-dependent manner in the presence of an ATG4B mutant (C74A) (Fujita et al., 2008), or after the depletion of ATG2A and B (Velikkakath et al., 2012), or FIP200 (Hara et al., 2008), which block autophagosome biogenesis at different stages (Figure 1D-F). Therefore, the reconstituted system recapitulates the autophagy-regulated secretion of IL-1β.

In macrophages, MVB formation and GRASP proteins are required for IL-1β secretion (Dupont et al., 2011; Qu et al., 2007). Inhibiting MVB formation by depletion of the ESCRT components, hepatocyte growth factor receptor substrate (Hrs) or TSG101, compromised secretion of IL-1β and CD63, an exosome marker (Figure 1-figure supplement 1A). Knockdown of the GRASP55 or GRASP65 also led to the reduction of IL-1β secretion (Figure 1- figure supplement 1B). Therefore, in addition to functions required for autophagy, the secretion of IL-1β in HEK293T cells depends on GRASP proteins and at least two proteins implicated in MVB formation, as reported previously (Dupont et al., 2011; Qu et al., 2007).

Figure 1 Reconstitution of autophagy-regulated IL-1β secretion in cultured cells (A) Reconstitution of starvation-induced IL-1β secretion in HEK293T cells. HEK293T cells were transfected with a single plasmid encoding p-IL-1β or together with the p-caspase-1 plasmid. After transfection (24 h), the cells were either treated in regular (DMEM) or starvation (EBSS) medium for 2 h. The medium and cells were collected separately and immunoblot was performed to determine the level of indicated proteins. (B) PI3K inhibitors 3-methyladenine (3-MA) or wortmannin (Wtm) inhibit IL-1β secretion. HEK293T cells transfected with p-IL-1β and p-caspase-1 plasmids were cultured in DMEM, EBSS, or EBSS containing 10 mM 3-MA or 20 nM wortmannin for 2 h. The medium and cells were collected separately and immunoblot was performed as shown in (A). (C) IL-1β secretion is blocked in Atg5 KO MEFs. Control WT or Atg5 KO MEFs were transfected with p-IL-1β and p-caspase-1 plasmids. After transfection (24 h), the cells were either cultured in DMEM or EBSS for 2 h followed by immunoblot as shown in (A). (D) IL-1β secretion is inhibited by the ATG4B mutant (C74A). HEK293T cells were transfected with plasmids encoding p-IL-1β, p-caspase-1 and different amounts of ATG4B (C74A) plasmid DNA as indicated. After transfection (24 h), cells were starved in EBSS for 2 h followed by immunoblot as shown in (A). (E) Knockdown of Atg2 reduces IL-1β secretion. HEK293T cells were transfected with control siRNA or siRNAs against Atg2A, Atg2B alone or both. After transfection (48 h), the cells were transfected with p-IL-1β and p-caspase-1 plasmids. After another 24 h, the cells were starved in EBSS for 2 h followed by immunoblot as shown in (A). (F) Knockdown of FIP200 reduces IL-1β secretion. HEK293T cells were transfected with control siRNA or FIP200 siRNA. IL-1β secretion under starvation conditions was determined as shown in (E). Quantification of IL-1β secretion was calculated as the ratio between the amount of IL-1β in the medium and the total amount (the sum of IL-1β in both medium and lysate).

Figure 1- figure supplement 1  Depletion of ESCRT or GRASPs affects IL-1β secretion HEK293T cells were transfected with indicated siRNAs (Hrs (ESCRT-0) (A), Tsg101 (ESCRT-I) (A), GRASP55 (B) or GRASP65 (B)). After transfection (48 h), the cells were transfected with p-IL-1β and p-caspase-1 plasmids. After another 24 h, the cells were starved in EBSS for 2 h followed by immunoblot as shown in Figure 1A. Quantification of IL-1β secretion was calculated as the ratio between the amount of IL-1β in the medium and the total amount (the sum of IL-1β in both medium and lysate).

IL-1β transits through an autophagosomal carrier during secretion.

To study if autophagy directly regulates IL-1β secretion, we employed a three-step membrane fractionation procedure as described previously (Figure 2A)(Ge et al., 2013). We first performed a differential centrifugation to obtain 3k, 25k and 100k membrane pellet fractions. Both IL-1β and the lipidated form of LC3 (LC3-II), a protein marker of autophagosome, were mainly enriched in the 25k membrane fraction (Figure 2B). We then separated the 25k membrane through a sucrose step gradient ultracentrifugation where both IL-1β and LC3-II co-distributed in the L fraction at the boundary between 0.25 M and 1.1 M layer of sucrose (Figure 2B). Further fractionation of the L fraction using an OptiPrep gradient showed co-fractionation of IL-1β with LC3-II (Figure 2C). To confirm the presence of IL-1β in the autophagosome, we performed immunoisolation of LC3-positive autophagosomes from the 25k fraction and found that IL-1β, especially the mature form, co-sedimented with autophagosomes (Figure 2D). Consistent with our observations, a recent study also showed a colocalization of IL-1β and LC3 in the form of puncta in macrophages (Dupont et al., 2011). These data demonstrate that at least a fraction of intracellular mature IL-1β associates with the autophagosome, possibly related to its role in IL-1β secretion.

Figure 2 IL-1β vesicles co-fractionate with LC3 vesicles (A) Membrane fractionation scheme. Briefly, HEK293T cells transfected with p-IL-1β and p-caspase-1 plasmids were starved in EBSS for 2 h, collected and homogenized. Cell lysates were subjected to differential centrifugations at 3,000×g (3k), 25,000×g (25k) and 100,000×g (100k). The level of IL-1β in each membrane fraction was determined by immunoblot. The 25k pellet, in which IL-1β was mainly enriched, was selected and a sucrose gradient ultracentrifugation was performed to separate membranes in the 25k pellet to the L (light) and P (pellet) fractions. The L fraction, which contained the majority of IL-1β, was further resolved on an OptiPrep gradient after which ten fractions from the top were collected. (B, C) Immunoblot was performed to examine the distribution of IL-1β, LC3 as well as the indicated membrane markers in the indicated membrane fractions. T, top; B, bottom (D) HEK293T cells transfected with p-IL-1β, p-caspase-1 and FLAG-tagged LC3-I plasmids were starved in EBSS for 2 h. LC3 positive membranes were immunoisolated with anti-FLAG agarose from the 25k pellet and the presence of IL-1β was determined by immunoblot analysis. FT, flowthrough.

To determine if IL-1β is localized to the phagophore in the absence of autophagosome completion, we fractionated membranes from ATG2-depleted cells, which are deficient in phagophore elongation and therefore fail to form mature autophagosomes (Velikkakath et al., 2012), and examined the distribution of LC3-II, which remains attached to immature phagophore membranes, and mature and precursor IL-1β. We performed the three-step fractionation described above. In control cells, IL-1β co-distributed with LC3-II in all three steps (Figure 3). Depletion of ATG2 did not affect the co-fractionation of IL-1β and LC3-II (Figure 3), indicating that IL-1β enters into the phagophore membrane before the completion of the autophagosome.

Figure 3 IL-1β co-distributes with LC3 in Atg2-depleted cells (A) HEK293T cells were transfected with siRNAs against Atg2A and Atg2B followed with p-IL-1β and 739 p-caspase-1 plasmids as shown in Figure 1E. The cells were starved in EBSS for 2 h. Membrane fractions (3k, 25k, 100k (×g), L and P) were separated from the post-nuclear supernatant as depicted in Figure 2B. (B) Ten membrane fractions were collected from the OptiPrep gradient ultracentrifugation as depicted in Figure 2C. Immunoblot was performed to examine the distribution of IL-1β, LC3 as well as the indicated membrane markers. T, top; B, bottom.

Autophagosome formation is not required for entry of IL-1β into vesicles

We asked how IL-1β enters into the autophagosome. One possibility is engulfment through the closure of the phagophore membrane during autophagosome maturation as in the capture of autophagic cargo. In this scenario, closure of the phagophore to complete autophagosome formation would be required to sequester IL-1β away from the cytoplasm. Alternatively, we considered the possibility that IL-1β may be translocated through a membrane into the lumen of the phagophore envelope and be sequestered from the cytoplasm even before the mature autophagosome is sealed. To test this possibility, we performed proteinase K protection experiments with the membranes from ATG2-depleted cells (Figure 4A). In control cells, p62 (an autophagic cargo) and a fraction of LC3-II (which was encapsulated after autophagosome completion), as well as mature IL-1β, were largely resistant to proteinase K digestion similar to the ER luminal protein, protein disulfide isomerase (PDI). In contrast, SEC22B, a membrane anchored SNARE protein exposed to the cytoplasm, was sensitive to proteinase K digestion (Figure 4A). Triton X-100 treatment permeabilized the membrane and rendered all proteins tested sensitive to proteinase K digestion (Figure 4A). This demonstrated that the majority of membrane localized IL-1β was sequestered within an organelle, likely the autophagosome, as demonstrated by the fractionation results of Figures 2 and 3. However, the result did not pinpoint where within the autophagosome IL-1β was housed. In ATG2-depleted cells, p62 and LC3-II remained sensitive to proteinase K digestion, consistent with the hypothesis that ATG2 is essential for maturation and closure of the autophagosome (Figure 4A). However, in the same samples the majority of IL-1β resisted degradation by proteinase K treatment (Figure 4A), except on addition of Triton X-100 to permeabilize membranes. Although the precursor form of IL-1β remained associated with isolated autophagosome and phagophore membranes (Figure 3), the protein was degraded when membranes from normal and ATG2-depleted cells were treated with protease in the presence or absence of Triton X-100 (data not shown). Thus, the mature but not the precursor IL-1β appears to be transported into the phagophore.

A most recent study showed that small, closed double-membrane structures could be observed in ATG2-depleted cells (Kishi-Itakura et al., 2014). To rule out the possibility that IL-1β was engulfed by the small closed autophagosomes, we employed Atg5 KO MEFs in which the phagophore could not be closed (Kishi-Itakura et al., 2014; Mizushima et al., 2001). Similar to what we observed in ATG2-depleted cells, IL-1β was protected from proteinase K digestion in membranes from Atg5 KO MEFs (Figure 4B). In addition, IL-1β was sequestered within vesicles in FIP200 (another early factor in phagophore development (Hara et al., 2008)) knockdown cells (Figure 4C). These data indicate that the entry of IL-1β into the vesicle carrier is not dependent on the formation of the autophagosome. These results are inconsistent with a role for engulfment of IL-1β by the maturing phagophore and suggest instead that IL-1β may be translocated across a membrane into a vesicle precursor of the phagophore, possibly at a very early stage in the development of the organelle.

Figure 4 Closure of the autophagosome is not required for the entry of IL-1β into vesicles (A) HEK293T cells were transfected with siRNAs against Atg2A and Atg2B followed by transfection with p-IL-1β and p-caspase-1 plasmids as shown in Figure 1E. The cells were starved in EBSS for 2 h and proteinase K digestion was performed with the 25k membrane fractions. (B) Atg5 WT, KO MEFs were transfected with p-IL-1β and p-caspase-1 plasmids as shown in Figure 1B. The cells were starved in EBSS for 2 h followed by proteinase K digestion as shown in (A). 752 (C) HEK293T cells were transfected with siRNA against FIP200 followed by analysis of membrane entry of 753 IL-1β as shown in (A). The level of proteinase K protection was calculated as the percentage of the total protein. Error bars represent standard deviations of at least three experiments.

Entry of IL-1β into the vesicle carrier requires protein conformational flexibility

We then sought to test if IL-1β could directly translocate across the membrane of a vesicle carrier. As protein unfolding is usually required for protein translocation, we adopted an approach used in many other circumstances wherein a targeted protein is fused to dihydrofolate reductase (DHFR), an enzyme whose three-dimensional structure is stabilized by the folate derivative aminopterin, hence providing a chemical ligand to impede the unfolding process (Backhaus et al., 2004; Eilers and Schatz, 1986; Wienhues et al., 1991). We first determined the secretion of the DHFR-fused IL-1β. As shown in Figure 5A, secretion of a mature IL-1β-DHFR fusion protein was enhanced by starvation similar to the untagged counterpart. Importantly, IL-1β-DHFR secretion was reduced in a dose-dependent manner in the presence of aminopterin (Figure 5B). Of notice, treatment of aminopterin did not completely abolish IL-1β secretion perhaps due to a cell death-induced release of IL-1β at high concentrations of aminopterin, as indicated by the release of a low level of tubulin into the medium fraction (Figure 5B). As a control, aminopterin did not reduce the secretion of untagged IL-1β, confirming its specific effect on DHFR (Figure 5- figure supplement 1). Fractionation of cells 185 incubated with aminopterin showed a reduced level of IL-1β in the membrane fraction with a corresponding 186 increase in the cytosol fraction (Figure 5C). The residual DHFR-tagged IL-1β associated with membranes from aminopterin-treated cells was sensitive to proteinase K digestion (Figure 5D), indicating that this pool of membrane-associated IL-1β did not translocate into the lumen of the vesicle. The data suggest that entry of IL-1β into a vesicle carrier involves a process of protein unfolding and translocation.

Figure 5  Protein unfolding is required for the entry of IL-1β into vesicles (A) Secretion of DHFR-tagged IL-1β. HEK293T cells were transfected with p-IL-1β-DHFR and p-caspase-1 plasmids. After transfection (24 h), the cells were treated with DMEM or EBSS for 2 h. Release of IL-1β was determined as shown in Figure 1. (B) Secretion of IL-1β-DHFR was inhibited by aminopterin. HEK293T cells were transfected with p-IL-1β-DHFR and p-caspase-1 plasmids. After transfection (24 h), the cells were treated with EBSS, or EBSS containing different concentrations of aminopterin as indicated for 15 min followed by determination of IL-1β secretion as shown in (A). Quantification of IL-1β secretion was calculated as the ratio between the amount of IL-1β in the medium and the total amount (the sum of IL-1β in both medium and lysate). (C) Less IL-1β enters into membrane in the presence of aminopterin. HEK293T cells were transfected with p-IL-1β-DHFR and p-caspase-1 plasmids. After transfection (24 h), the cells were either untreated or treated with 5 μM aminopterin in EBSS for 2 h. The membrane fraction was collected from the top fractions of a Nycodenz density gradient resolved from membranes in a 25k pellet as described in Material and Methods. The cytosolic fraction was collected as the supernatant after 100k×g centrifugation. All fractions were analyzed by immunoblotting using indicated antibodies. (D) IL-1β-DHFR is not protected from proteinase K in the presence of aminopterin. Nycodenz -floated membrane fraction collected as shown in (C) was subjected to proteinase K digestion and then analyzed by immunoblotting using indicated antibodies.

Figure 5- figure supplement 1 Secretion of IL-1β is not affected by aminopterin HEK293T cells were transfected with p-IL-1β and p-caspase-1 plasmids. After transfection (24 h), the cells were treated with EBSS, or EBSS containing different concentrations of aminopterin as indicated for 15 min followed by determination of IL-1β secretion as shown in Figure 1 (A).

IL-1β colocalizes with LC3 on the autophagosome envelope

If IL-1β is directly translocated across the membrane of a vesicle intermediate, fusion of these vesicles to form a double-membrane autophagosome would deposit IL-1β in the lumen between the two membranes of the autophagosome. To visualize the subcellular localization of IL-1β, we employed U2OS cells, which formed 194 large and distinct autophagosomes after starvation. U2OS cells co-expressing p-IL-1β and p-caspase-1 secreted IL-1β in a starvation-enhanced and PI3K-dependent manner similar to HEK293T cells (Figure 6- figure supplement 1). To prepare for the subsequent fluorescence imaging, we also employed a FLAG-tagged m-IL-1β, which allowed us to directly determine the topological localization of the m-IL-1β. Secretion of m-IL-1β-FLAG from U2OS cells was stimulated by starvation and dependent on PI3K (Figure 6- figure supplement 1).

To determine the topological distribution of IL-1β, we first performed confocal immunofluorescence labeling experiments. After starvation, cells were exposed to 40 μg/ml of digitonin to permeabilize the plasma membrane, harvested and washed with cold PBS to remove the excess cytosolic m-IL-1β-FLAG. In cells expressing either p-IL-1β and p-caspase-1, or m-IL-1β alone, LC3 and IL-1β were observed by confocal microscopy to localize together or adjacent to one another on the edge of ring-shaped autophagosomes (Figure 6- figure supplement 2). To further resolve these ring structures, we employed 3D STORM (Huang et al., 2008; Rust et al., 2006) super-resolution microscopy (Hell, 2007; Huang et al., 2010) (Figure 6 and Figure 6- figure supplements 3, 4 and Videos 1 and 2). Ring-shaped autophagosomes positive for LC3 (cyan) formed after starvation. Some IL-1β (magenta) also organized in ring-shaped structures that co-localized with LC3 (Figure 6 and Figure 6- figure supplement 3). Around 18 ring structures of IL-1β accounting for ~5% of the total IL-1β signal were observed in each cell. A 3D virtual Z-stack analysis confirmed the spatial co-distribution of LC3 and IL-1β on a ball-shaped vesicle (Video 1 and 2). The diameter of the structures double-labeled with LC3 and IL-1β are ~700 nm (larger structures up to 2 μm in diameter were also found) which is comparable to the size of the autophagosome. Occasionally, we also found IL-1β localized in the center of the ring structure, where cytoplasmic autophagic cargoes fill, surrounded by LC3 (Figure 6-figure supplement 4). This portion of IL-1β was possibly being engulfed by the autophagosome.

The visual detection of IL-1β localized to ring-shaped autophagosomes is consistent with our biochemical assays that place IL-1β in the intermembrane space between the outer and inner membrane of the autophagosome. We devised a further visual test of this conclusion using selective permeabilization of cell surface and intracellular membranes with digitonin and saponin, respectively (Figure 6-figure supplement 5). We compared antibody accessibility to IL-1β and DFCP1, a marker located on the cytosolic surface of the omegasome (a harbor for the phagophore) in both WT and Atg5 KO cells. Consistent with a cytosolic surface localization, DFCP1 was readily labeled in cells treated with digitonin alone (selectively permeabilizes the plasma membrane) in both WT and Atg5 KO cells (Figure 6-figure supplement 5A-E). In contrast, IL-1β was accessible to the antibody only after treatment with digitonin and saponin (gently permeabilizes the endomembrane) (Figure 6-figure supplement 5A, B and F) in WT cells. This by itself would not distinguish localization of IL-1β to the intermembrane space vs the cytoplasmic enclosed space of a mature autophagosome. However, in Atg5 KO cells where the phagophore precursor envelope remains open and exposed to the cytosol, saponin treatment was necessary to expose IL-1β to antibody and roughly half of the labeled structures coincided with the phagophore marker DFCP1 (Figure 6-figure supplement 5C, D and F). This visual assay further confirms the intermembrane localization of IL-1β in the phagophore and 231 autophagosome.

Figure 6 Topological localization of IL-1β in the autophagosomal carrier determined by STORM U2OS cells were transfected with a plasmid containing the expression cassette of FLAG-tagged mature IL-1β (m-IL-1β-FLAG). After transfection (24 h), the cells were starved in EBSS for 1 h followed by immunofluorescence labeling with mouse monoclonal anti-LC3 and rabbit polyclonal anti-FLAG antibodies. STORM analysis imaging and data analysis were performed as described in Materials and Methods. Cyan, LC3; Magenta, IL-1β; Bars: 2 μm (original image) and 500 nm (magnified inset)

Figure 6- figure supplement 1  Secretion of IL-1β in U2OS cells 795 U2OS cells were transfected with plasmids encoding the p-IL-1β and p-caspase-1 (first 4 lanes) or m-IL-1β-FLAG (last 4 lanes). After transfection (24 h), the cells were untreated or starved in the absence or presence of indicated PI3K inhibitors (3-MA or wortmannin (Wtm)) followed by measurement of secretion as indicated in Figure 1 (A) and (B). α-m-IL-1β, IL-1b antibody; α-FLAG, FLAG antibody

Figure 6- figure supplement 2 Localization of IL-1β determined by confocal microscopy U2OS cells were transfected with plasmids encoding the p-IL-1β and p-caspase-1 (A) or m-IL-1β-FLAG (B). After transfection (24 h), the cells were starved for 1 h followed by immunofluorescence labeling and confocal 804 microscopy analysis. Bar: 10 μm

Figure 6- figure supplement 3  Extra images for Figure 6  Bars: 2 μm (original image) and 500 nm (magnified inset)

Figure 6- figure supplement 4  A minority of IL-1β engulfed by autophagosome  U2OS cells were transfected and treated followed by STORM analysis as shown in Figure 6. Arrow head points to the autophagosome with engulfed IL-1β. Bar: 2 μm

Figure 6- figure supplement 5  Determination of the topological localization of IL-1β in the autophagosome and phagophore  (A, C) Diagrams of autophagosome (A)/phagophore (B) and omegasome, antibody accessibility for each possible situation of IL-1β localization, and summaries of the antibody accessibility of m-IL-1β-FLAG (red) and EGFP-DFCP1 (green) are illustrated. (B, D) U2OS cells (B) and Atg5 KO MEFs (D) were transfected with plasmids encoding the m-IL-1β-FLAG and EGFP-DFCP1. After transfection (24 h), the cells were starved in EBSS for 1 h followed by digitonin treatment and fixation (see Materials and Methods). The cells were either labeled with anti-FLAG (to label IL-1β) and anti-EGFP (to label EGFP-DFCP1) antibodies (Digitonin) or further treated with Saponin followed by antibody labeling (Digitonin+Saponin). Images were acquired by confocal microscopy. Bar: 10 μm 825 (E) Quantification of the percentage of EGFP-DFCP1 labeled by EGFP antibody. Percentage was counted by 826 the ratio of puncta numbers of antibody labeled EGFP-DFCP1 and EGFP-DFCP1 according to the EGFP signal. Error bars are standard deviations of more than 50 cells in two independent experiments. (F) Quantification of the puncta number for m-IL-1β-FLAG puncta (red) and those colocalized with DFCP1 (yellow). Error bars are standard deviations of more than 50 cells in two independent experiments.

Video 1 832 3D section of the magnified structure in Figure 6 (upper one) 833 The virtual Z-section thickness is 150 nm, and the step size is 50 nm. Cyan, LC3; Magenta, IL-1β; Bar 500 nm 834 835 Video 2 836 3D section of the magnified structure in Figure 6 (lower one) 837 The virtual Z-section thickness is 150 nm, and the step size is 50 nm. Cyan, LC3; Magenta, IL-1β; Bar 500 nm

Two KFERQ-like motifs are required for the entry of IL-1β into the vesicle carrier

In chaperone-mediated autophagy (CMA), cargoes are recognized by a KFERQ sequence motif for transport into the lysosome (Dice et al., 1986; Kaushik and Cuervo, 2012). We analyzed the primary sequence of IL-1β and found three KFERQ-like motifs on IL-1β including 127LRDEQ131, 132QKSLV136 and 198QLESV202 (Figure 7A). We mutated the glutamine, which has been shown to be essential for the function of the motif, as well as an adjacent amino acid in each motif (E130Q131, Q132K133 and Q198L199) to alanines and examined the secretion efficiency of these mutants. The 130-131AA mutant did not affect secretion of IL-1β (Figure 7B). However, the Q132K133 and Q198L199 mutations were both defective in secretion of mature IL-1β which instead accumulated in the cytoplasmic fraction (Figure 7B). A low level of release of the pro-forms persisted as seen with WT and mutant protein (Figure 7B). The cytoplasmic mature forms of the mutant proteins were less abundant in the membrane fraction compared with the WT mature IL-1β (Figure 7C, compare the lanes without proteinase K treatment). In addition, the membrane associated mutant IL-1β remained proteinase K accessible (less than 10% of protection compared with ~45% of WT IL-1β), demonstrating that these two KFERQ-like motifs are required for the membrane translocation of IL-1β (Figure 7C). Equal amounts of WT and mutant p-IL-1β associated with the membrane but both remained largely proteinase K accessible (Figure 7C).

Figure 7 Mutation of the KFERQ-like motif affects IL-1β secretion and entry into vesicles (A) Protein sequence of IL-1β. The yellow region indicates mature IL-1β. Three KFERQ-like motifs (aa127-131, aa132-136 and aa198-202) are highlighted in red underlined bold. (B) Secretion of IL-1β mutants. HEK293T cells were transfected with p-IL-1β-DHFR and p-caspase-1 plasmids. After transfection (24 h), the cells were either treated with DMEM or EBSS for 2 h. Secretion of 845 IL-1β mutant proteins was detected by immunoblot. (C) IL-1β mutant 132-133AA or 198-199AA is accessible to proteinase K digestion. HEK293T cells were transfected with plasmids encoding p-caspase-1 and IL-1β mutant 132-133AA or 198-199AA. After transfection (24 h), the cells were treated with EBSS for 2 h. The 25k membrane fraction was collected and subjected to proteinase K digestion assay and then analyzed by immunoblot using indicated antibodies. The level of proteinase K protection was calculated as the percentage of the total protein. Error bars represent standard deviations of at least three experiments.

HSP90 is required for the entry of IL-1β into the vesicle intermediate 

The chaperone protein HSC70 and HSP90 have been reported to function in chaperone-mediated autophagy (CMA) (Kaushik and Cuervo, 2012; Majeski and Dice, 2004). HSP70 has also been implicated in autophagy and stress responses (Murphy, 2013). We performed shRNA-mediated knockdown of the three chaperone proteins to assess their potential role in the membrane translocation of IL-1β. Knockdown of Hsp90, but not of Hsp70 or Hsc70 substantially reduced IL-1β secretion (Figure 8A). As a control, knockdown of Hsc70 compromised CMA as indicated by the stabilization of a CMA cargo, GAPDH (Figure 8-figure supplement 1A). Moreover, secretion of mature IL-1β was inhibited in a dose-dependent manner by an HSP90 inhibitor geldanamycin (Figure 8B). In both experiments, mature IL-1β accumulated in the cytosol fraction at the expense of secretion. Knockdown of Hsp90 also rendered IL-1β accessible to proteinase K digestion (Figure 8C), consistent with a role for HSP90 in the translocation of IL-1β as opposed to some later secretion event. Furthermore, in a co-immunoprecipitation assay, HSP90 associated with m-IL-1β but not the translocation-deficient mutants Q132K133 and Q198L199 (Figure 8D). Although p-IL-1β also formed a complex with HSP90, the efficiency appeared lower than for m-IL-1β. These results suggest that HSP90 binds to a region of the mature IL-1β, including the essential residues Q132K133 and Q198L199, to promote the translocation event. Cleavage of p-IL-1β by caspase-1 may potentiate the recruitment of HSP90 to the mature form of IL-1β however chaperone binding is not required for this proteolytic event (Figures 8D and 7B).

In the CMA pathway, HSC70 and HSP90 play different roles. HSC70 binds to cargoes and delivers them into 266 the lysosome as well as disassembling LAMP2A oligomers, whereas HSP90 is required for the oligomerization and stability of LAMP2A (Bandyopadhyay et al., 2008; Chiang et al., 1989). Co-immunoprecipitation indicated that IL-1β associates with HSP90 but not HSC70 (Figure 8-figure supplement 1B). In addition, knockdown of Lamp2A compromised CMA but did not affect the secretion of IL-1β, and disruption of the lysosome did not result in the release of IL-1β from the membrane carrier (Figure 8-figure supplement 1C-E). These data suggest that the translocation of IL-1β into the vesicle carrier is mechanistically distinct from CMA.

Figure 8 HSP90 is involved in the entry of IL-1β into vesicles (A) Knockdown of Hsp90 inhibits IL-1β secretion. HEK293T cells were transduced with lentivirus carrying control (Ctrl) shRNA or shRNA against Hsc70, Hsp90 or Hsp70. Then the cells were transfected with p-IL-1β and p-caspase-1 plasmids. After transfection (24 h), the cells were cultured in EBSS for 2 h followed by determination of IL-1β secretion by immunoblot. (B) IL-1β secretion is reduced in the presence of HSP90 inhibitor geldanamycin. HEK293T cells were transfected with p-IL-1β and p-caspase-1 plasmids. After transfection (24 h), the cells were treated with EBSS containing different concentrations of geldanamycin as indicated. Immunoblot was performed as shown in Figure 1. Quantification of IL-1β secretion was calculated as the ratio between the amount of IL-1β in the medium and the total amount (the sum of IL-1β in both medium and lysate). (C) IL-1β remains accessible to proteinase K in Hsp90 knockdown cells. HEK293T cells were transduced with lentivirus carrying control (Ctrl) shRNA or shRNA against Hsp90. Then the cells were transfected with p-IL-1β and p-caspase-1 plasmids. After transfection (24 h), the cells were cultured in EBSS for 2 h. The 25k membrane fraction was collected and digested with proteinase K and then analyzed by immunoblotting using indicated antibodies. (D) Association of HSP90 with IL-1β WT and mutants. HEK293T cells transfected with p-caspase-1 and IL-1β mutant 132-133AA or 198-199AA were starved in EBSS for 2 h. Immunoprecipitation (IP) with anti-HSP90 antibody coupled to protein G-agarose was performed, followed by an immunoblot with anti-IL-1β and anti-HSP90 antibodies.

Figure 8- figure supplement 1 Translocation of IL-1β is mechanistically different from CMA (A) Knockdown of Hsc70 reduces CMA. HEK293T cells transduced with lentivirus carrying control (Ctrl) shRNA or shRNA against Hsc70 were incubated with regular medium (-CMA) or DMEM (+CMA) in the presence of 20 μg/ml cycloheximide for 24 h. The cells were lysed and analyzed by immunoblotting using indicated antibodies. For quantification, the ratio of GAPDH and tubulin was calculated and normalized by that in control (-CMA) treatment which was set as one. (B) Co-immunoprecipitation of HSC70 or HSP90 with IL-1β. HEK293T cells transfected with m-IL-1β-FLAG were starved in EBSS for 2 h. Immunoprecipitation (IP) with anti-HSC70 or anti-HSP90 antibody coupled to protein A/G-agarose was performed, followed by an immunoblot with indicated antibodies. (C) Knockdown of Lamp2 blocks CMA. HEK293T cells were transfected with control or LAMP2 siRNA. After transfection (48 h), the cells were trypsinized and plated. After 24 h, siRNA transfection was repeated. After another 48 h, the cells were trypsinized and plated. After 24 h, the cells were incubated with regular medium (-CMA) or DMEM (+CMA) in the presence of 20 μg/ml cycloheximide for 24 h. The cells were lysed and analyzed by immunoblotting using indicated antibodies. For quantification, the ratio of GAPDH and Tubulin was calculated and normalized by that in control (-CMA) treatment which was set as one. (D) Knockdown of LAMP2 does not affect IL-1β secretion. HEK293T cells were transfected with control or LAMP2 siRNA as show in (C). After the second siRNA transfection (24h), the cells were transfected with m-IL-1β-FLAG plasmid. After transfection (24 h), the cells were either cultured in DMEM or EBSS for 2 h followed by determination of IL-1β secretion by immunoblot as shown in Figure 1A. Quantification of IL-1β secretion was calculated as the ratio between the amount of IL-1β in the medium and the total amount (the sum of IL-1β in both medium and lysate). (E) Level of IL-1β in the membrane fraction was not affected by lysosome disruption. HEK293T cells 897 transfected with m-IL-1β were cultured in EBSS for 2 h and then treated with DMSO or 0.5 mM glycyl-L-phenylalanine-2-naphthylamide (GPN) for 10 min. The membrane fraction was collected from the top fractions of a Nycodenz density gradient resolved from membranes in a 25k pellet as described in Material and Methods. Both membrane fraction and cell lysis were analyzed by immunoblotting using indicated antibodies.

We next asked if starvation regulated the association between HSP90 and IL-1β. We performed an HSP90 co-immunoprecipitation experiment with cytosol prepared from cells grown in nutrient-rich or starvation conditions (Figure 9A). Starvation led to a ~2.5 fold increase of the association of HSP90 and IL-1β (Figure 9A). This increase was likely not due to starvation-stimulated processing of p-IL-1β because starvation had no effect on the cleavage of mutant forms of IL-1β unable to bind HSP90 (Figure 7B). Starvation led to a ~ 2 fold increase in the membrane localization and cytosolic depletion of mature IL-1β (Figure 9B). Starvation may stimulate the recruitment of a complex of m-IL-1β/HSP90 to the membrane responsible for IL-1β translocation (Figure 9B).

Figure 9 Induction of autophagy enhances the membrane incorporation of IL-1β (A) Starvation enhances the association of IL-1β with HSP90. HEK293T cells transfected with p-IL-1β and p-caspase-1 were cultured in DMEM or EBSS for 2 h. Immunoprecipitation with anti-HSP90 antibody was performed followed by an immunoblot with anti-IL-1β and anti-HSP90 antibodies. (B) Starvation promotes the entry of IL-1β into the membrane fraction. HEK293T cells transfected with p-IL-1β and p-caspase-1 were cultured in DMEM or EBSS for 2 h. The membrane fraction was collected from the top fractions of a Nycodenz density gradient resolved from membranes in a 25k pellet as described in Material and Methods. The cytosolic fraction was collected as the supernatant after 100k×g centrifugation. Immunoblot was performed to determine the levels of IL-1β in both fractions. (C) A proposed model for autophagy-mediated IL-1β secretion. Cytosolic IL-1β associates with HSP90 which facilitates the translocation of IL-1β into the lumen of a vesicle carrier which later either turns into a  phagophore and an autophagosome or fuses with them. IL-1β localizes between the outer and inner membrane after the double membrane autophagosome forms. The topological distribution ensures the secretion of IL-1β in a soluble form. The IL-1β-containing autophagosome may directly fuse with the plasma membrane or further fuse with a MVB followed by fusion with the plasma membrane.

Genetic and cell biological studies have implicated autophagy in the transport of several leaderless cargoes to the extracellular space (Bruns et al., 2011; Dupont et al., 2011; Duran et al., 2010; Manjithaya et al., 2010). Unconventional secretory cargoes, such as IL-1β and Acb1, have been shown to have overlapping requirements with formation of the autophagosome or its precursor suggesting that the autophagosome may physically convey these cargo proteins to the cell surface. A key question is if and how these cargoes engage the autophagosome and how this structure exports soluble cargo molecules. In this study, we probed the organelle association and molecular requirements for the secretion of one such unconventional cargo protein, IL-1β. Using surrogate cell lines rather than macrophages to reconstitute autophagy-mediated secretion of IL-1β (Figure 1), we find mature IL-1β localized to the lumen of the membrane in early intermediates and mature autophagosomes (Figures 2-4, 6). This surprising location may help to explain how mature IL-1β is secreted in a soluble form to the cell surface (Figure 9C). However, localization to the lumen between the two membranes of the autophagosome would require that IL-1β is translocated from the cytoplasm across the membrane precursor of a phagophore, rather than being engulfed as the phagophore membrane matures by closure into an autophagosome. Our evidence suggests that IL-1β must unfold or be held in an unfolded state to promote membrane translocation (Figure 5) and that a complex sorting signal in the mature portion of IL-1β interacts with HSP90 to deliver the chaperone and its cargo to a site on a phagophore precursor membrane where the mature species is translocated (Figures 7-9).

The unconventional secretory cargo fibroblast growth factor 2 (FGF2) has been shown to directly translocate across the plasma membrane as a folded protein without the apparent aid of chaperones (Backhaus et al., 2004; Steringer et al., 2015). Unlike FGF2, the entry of IL-1β into the autophagosomal carrier appears to be dependent on protein unfolding in a conformational state that may be fostered by the association of HSP90 with two KFERQ-like sequences within the mature portion of IL-1β (Figure 5 and 8). This translocation mechanism appears superficially similar to another delivery process termed HSC70-dependent CMA in which autophagic cargoes bearing KFERQ targeting motifs are directed into the lysosome for degradation. Indeed, using a cell-free approach to study the import of CMA cargo into isolated lysosomes, Salvador et al. (2000) reported that DHFR fused to a CMA cargo is blocked in translocation by addition of methotrexate, a drug that stabilizes DHFR to unfolding, just as we have shown that IL-1β fused to DHFR is blocked in cells treated with a cell permeable folate analog, aminopterin (Wei et al., 2013). In our fractionation study, IL-1β distributed in LC3-positive autophagosomal carriers that were separated from the lysosome marker LAMP2, the proposed receptor or channel for uptake of CMA cargo (Kaushik and Cuervo, 2012)(Figure 2B). This observation, together with the involvement of a different chaperone i.e. HSP90, suggests distinct routes for IL-1β and cargoes of the CMA pathway.

The target membrane for IL-1β translocation may be a vesicle that could fuse with or form the autophagosome. We find that mature IL-1β can be detected within protease inaccessible membranes in cells blocked early in the autophagic pathway (e.g. ATG5 null cells and cells depleted of FIP200, both of which block at a stage prior to the lipidation of LC3). The identity of the vesicle carrier is unknown and could be any one of those reported to supply membrane to the formation of the autophagosome (Ge et al., 2014a; Lamb et al., 2013). Although we have ruled out the involvement of LAMP2A IL-1β translocation, it is likely that a membrane receptor locating on the membrane of the vesicle carrier, perhaps a functional equivalent of LAMP2A, recruits the protein complex of HSP90 and IL-1β, therefore designating the correct membrane targeting of IL-1β. In addition, a protein conducting channel may be involved in the translocation of IL-1β into the membrane. It seems unlikely that a standard translocation channel, such as SEC61, is involved in this import process, but no current evidence bears on this point.

The exact route by which the autophagosome delivers mature IL-1β to the cell surface as well as how it avoids fusion with degradative lysosome remains obscure, possibly involving interaction with the multi-vesicular body or some form of lysosome as a prelude to fusion at the cell surface (Figure 9C), and this process may require selective recruitment of membrane sorting and targeting factors such as Rabs and SNAREs. Fusion of the autophagosome directly with the plasma membrane would lead to the release of soluble IL-1β available to trigger an inflammatory response in the surrounding tissue. If mature IL-1β were engulfed within the cytoplasmic interior of the autophagosome, fusion of this organelle at the cell surface might release an intact vesicle corresponding to the inner membrane-enclosed cytoplasmic compartment of the autophagosome. We found mature IL-1β secreted by macrophages or in our surrogate cell system to be completely soluble, thus inconsistent with the engulfment model (data not shown). An alternative possibility may be that the autophagosome fuses with another intracellular organelle such as the MVB or the lysosome under conditions where the inner membrane of the autophagosome is degraded. If so, mature IL-1β would be available for secretion if the combined organelle (amphisome, Figure 9C) fused with the plasma membrane. However, for this model to be viable, the mature IL-1β released on dissolution of the autophagosome inner membrane would have to withstand proteolytic attack such as may be encountered in an amphisome or lysosome. Because mature IL-1β is clearly sensitive to proteolysis (Figure 4), thus any compartment engaged in presenting autophagosomal content to the cell surface must be depleted of proteases. The nature of the organelle that delivers autophagosome content to the plasma membrane may be probed by selective ablation of different Rab proteins, e.g. Rab11, Rab27 and Rab35, which appear to be required for fusion of the MVB with the cell surface (Hsu et al., 2010; Ostrowski et al., 2010; Savina et al., 2002), or Rab27a and Rab38, implicated in the fusion of lysosomes at the cell surface (Blott and Griffiths, 2002; Hume et al., 2001; Jager et al., 2000.


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microglia and brain maintenance

Larry H. Bernstein, MD, FCAP, Curator



Mapping mosaicism: Tracing subtle mutations in our brains

Posted on January 14, 2015 by Nancy Fliesler

Posted in All PostsInformation technology

More On: brain developmentDNA sequencinggeneticsmosaicismneurosciencesomatic mutations

DNA sequences were once thought to be the same in every cell, but the story is now known to be more complicated than that. The brain is a case in point: Mutations can arise at different times in brain development and affect only a percentage of neurons, forming a mosaic pattern.

Now, thanks to new technology described last week in Neuron, these subtle “somatic” brain mutations can be mapped spatially across the brain and even have their ancestry traced.

Like my family, who lived in Eastern Europe, migrated to lower Manhattan and branched off to Boston, California and elsewhere, brain mutations can be followed from the original mutant cells as they divide and migrate to their various brain destinations, carrying their altered DNA with them.

“Some mutations may occur on one side of the brain and not the other,” says Christopher Walsh, MD, PhD, chief of Genetics and Genomics at Boston Children’s Hospital and co-senior author on the paper. “Some may be ‘clumped,’ affecting just one gyrus [fold] of the brain, disrupting just a little part of the cortex at a time.”

This tracking capability represents a significant advance for genetics research. And for neuroscientists, it provides a new way to study both the normal brain and brain disorders like epilepsy, autism and intellectual disability.

Walsh and colleagues studied normal brain tissue from a teenage boy who had passed away from other causes. Sampling in more than 30 brain locations, they used deep, highly sensitive, whole-genome sequencing of one neuron at a time—unlike usual methods, which sequence thousands or millions of cells mixed together and simply read out an average.×735.jpg

The blue and green boxes indicate different degrees of mosaicism (based on proportion of cells affected) in the left half of this teen’s normal brain. The blue shaded area indicates that retrotransposon mutation #1 (blue boxes) is limited to a focal area in the middle frontal gyrus. The empty boxes indicate areas where mutation #1 was not detected. (Courtesy Gilad Evrony, PhD, Boston Children’s Hospital)

Next, using technology developed by Alice (Eunjung) Lee in the lab of Peter Park, PhD, at Harvard Medical School’s Center for Biomedical Informatics, they zeroed in on inserted bits of DNA caused by retrotransposons, one type of mutation that can arise as the brain develops. These essentially served as markers that allowed cell lineages to be traced.

“Our findings are intriguing because they suggest that every normal brain may in fact be a mosaic patchwork of focal somatic mutations, though in normal individuals most are likely silent or harmless,” says Gilad Evrony, PhD, in the Walsh Lab.×509.jpg

This model illustrates the origins of two somatic retrotransposon mutations during prenatal development and their subsequent dissemination in the brain. Insertion #2 (in green) occurred soon after conception; #1 (in blue) happened sometime later during brain development. The ‘pie slices’ show a closeup of the layers of the cerebral cortex. Later in development, additional somatic mutations occurred inside insertions #1 and #2, creating new, smaller sublineages of cells. (Courtesy Gilad Evrony, PhD)

A parallel study from Walsh’s lab in 2014 used single-neuron sequencing to find copy number variants— a different type of mutation affecting the number of copies of chromosomes or chromosome fragments. It, too, found the mutations to be present in normal brains as well as neurologically diseased brains.

Walsh and others speculate that some somatic brain mutations might play a role in autism, epilepsy, schizophrenia and other unsolved neuropsychiatric diseases whose causes are mostly still a mystery.

“It is possible that a whole new class of brain disorders may exist that has not been previously recognized,” says Evrony. “In such disorders, a somatic mutation may subtly affect only one small part of the brain involved in a specific ability, for example language, while sparing the rest of the brain.”

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Tracking subtle brain mutations, systematicallyTool can trace and spatially map “mosaic” mutations in the brain

BOSTON, Jan. 7, 2015 /PRNewswire-USNewswire/ — DNA sequences were once thought to be identical from cell to cell, but it’s increasingly understood that mutations can arise during brain development that affect only certain groups of brain cells. A technique developed at Boston Children’s Hospital allows these subtle mutation patterns to be traced and mapped spatially for the first time. This capability is a significant advance for genetics research and provides a new way to study both the normal brain and brain disorders such as epilepsy and autism.

Described in the January 7th issue of Neuron, the technique uses “deep,” highly sensitive whole-genome sequencing of single neurons and a new technology that identifies inserted bits of DNA caused by retrotransposons, one of several kinds of so-called somatic mutations that can arise as the brain develops.

The technique picks up somatic mutations that affect just a fraction of the brain’s cells, in a “mosaic” pattern. It also allows “lineage tracing,” showing when during brain development the mutations arise and how they spread through brain tissue as the mutated cells grow, replicate and migrate, carrying the mutation with them.

“There is a lot of genetic diversity from one neuron to the other, and this work gets at how somatic mutations are distributed in the brain,” says Christopher Walsh, MD, PhD, chief of Genetics and Genomics at Boston Children’s and co-senior author on the paper. “Some mutations may occur on one side of the brain and not the other. Some may be ‘clumped,’ affecting just one gyrus [fold] of the brain, disrupting just a little part of the cortex at a time.”

The study examined brain tissue from a deceased 17-year-old who had been neurologically normal, sampling in more than 30 brain locations. It builds on work published by the Walsh lab in 2012, which developed methods to sequence the genomes of single neurons, and represents the first time single neurons have been sequenced in their entirety. The single-cell technique is better at detecting subtle mosaicism than usual DNA sequencing methods, which sequence many thousands or millions of cells mixed together and read out an average for the sample.

Somatic brain mutations, affecting just pockets of cells, can be harmful, and have been suggested as a possible cause of neurodevelopmental disorders such as autism, epilepsy or intellectual disability (see this review article for further background). But they also can be completely benign or have just a subtle effect.

“Our findings are intriguing because they suggest that every normal brain may in fact be a mosaic patchwork of focal somatic mutations, though in normal individuals most are likely silent or harmless,” says Gilad Evrony, PhD, in the Walsh Lab, co-first author on the Neuron paper. “These same technologies can now be used to study the brains of people who died from unexplained neuropsychiatric diseases to determine whether somatic mutations may be the cause.”

Finally, says Evrony, the findings provide a proof-of-principle for a systematic way of studying how brain cells disperse and migrate during development, “something that has not been possible to do before in humans,” he says.

Co-first author Alice Eunjung Lee, PhD, from the lab of Peter Park, PhD, at the Center for Biomedical Informatics at Harvard Medical School, developed the study’s retrotransposon analysis tool, which detects somatic retrotransposon mutations in single-cell sequencing data.

Mirroring these findings, study published by Walsh’s lab in 2014 used single-neuron sequencing to detect copy number variants—another type of mutation affecting the number of copies of chromosomes or chromosome fragments. The study found that these mutations can occur in both normal and neurologically diseased brains.

Evrony and Lee are first authors on the Neuron paper; Walsh and Park are senior authors. The research was supported by the National Institutes of Health (MSTP grant T32GM007753), the National Institute of Neurological Disorders and Stroke (R01 NS079277 and R01 NS032457), the Louis Lange III Scholarship in Translational Research, the Eleanor and Miles Shore Fellowship, the Research Connection and the Manton Center for Orphan Disease Research at Boston Children’s Hospital, the Paul G. Allen Family Foundation and the Howard Hughes Medical Institute.

SOURCE Boston Children’s Hospital


Beth Stevens: A transformative thinker in neuroscience

Posted on September 29, 2015 by Nancy Fliesler

Posted in All PostsDrug discoveryProfiles


More On: Alzheimer’s diseaseautismFM Kirby Neurobiology Centerglial cellsneurosciencesynapse development

When 2015 MacArthur “genius” grant winner Beth Stevens, PhD, began studying the role of glia in the brain in the 1990s, these cells—“glue” from the Greek—weren’t given much thought. Traditionally, glia were thought to merely protect and support neurons, the brain’s real players.

But Stevens, from the Department of Neurology and the F.M. Kirby Neurobiology Center at Boston Children’s Hospital, has made the case that glia are key actors in the brain, not just caretakers. Her work—at the interface between the nervous and immune systems—is helping transform how neurologic disorders like autism, amyotrophic lateral sclerosis (ALS), Alzheimer’s disease and schizophrenia are viewed.

Soon after college graduation in 1993, without prior experience in neuroscience, she helped discoveran interplay between neurons and glial cells known as Schwann cells that controlled production of the nerve insulation known as myelin It was one of the early pieces of evidence that glia and neurons talk to each other.

In 2007, while still a postdoctoral fellow, Stevens showed how star-shaped glial cells called astrocytes influence the development of synapses, or brain connections. Studying neurons, her lab showed that a gene called C1q was markedly more active when astrocytes were present. C1q is an immune gene, one nobody had expected to see in a normal brain. In the context of disease, it initiates the complement cascade, an immunologic pathway for tagging unwanted cells and debris for clearance by other immune cells.

But in healthy developing brains, Stevens showed, C1q was concentrated at developing synapses, or brain connections, apparently marking certain synapses for pruning.

Then in 2012, the Stevens lab showed that microglia—another type of glia usually thought of as immune cells themselves—actively sculpt the brain’s wiring. They literally trim away unwanted, inappropriate synapses by eating them—in the same way they’d engulf and destroy invading bacteria.

That paper was cited by the journal Neuron as the year’s most influential paper.

The same year, she received a Presidential Early Career Award for Scientists and Engineers, honoring her innovative research and scientific leadership.

Stevens’s current investigations are looking at synapse loss—a hallmark of neurodegenerative conditions such as Alzheimer’s—and trying to understand why it occurs. Her lab’s recent work suggests that normal pruning mechanisms that are active during early brain development get re-activated later in life. Intervening with this activation could lead to a new treatment approach, she believes.

Stevens isn’t the only brain researcher at Boston Children’s to become a MacArthur fellow. Neurosurgeon Benjamin Warf, MD, received the honor in 2012.

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Immune cells “sculpt” brain circuits — by eating excess connections

Posted on June 5, 2012 by Nancy Fliesler

Posted in All PostsDrug discoveryPediatrics

More On: ALSAlzheimer’s diseaseautismbrain developmentepilepsyglaucomaHuntington’s diseaseLou Gehrig’s disease,Parkinson’s diseasesynapse development

The above movie shows an immune cell caught in the act of tending the brain—it’s just eaten away unnecessary connections, or synapses, between neurons.

That’s not something these cells, known as microglia, were previously thought to do. As immune cells, it was thought that their job was to rid the body of unwanted pathogens and debris, by engulfing and digesting them.

The involvement of microglia in the brain’s development has started to be recognized only recently. The latest research finds that microglia tune into the brain’s cues, akin to the way they survey their environment for invading microbes, and get rid of excess synapses the same way they’d dispatch these invaders—by eating them.

It’s a whole other way of understanding how the healthy brain develops—at the hands of cells that were once thought to be merely nerve “glue” (the literal meaning of “glia” from the Greek), playing a protective role to neurons, say investigators Beth Stevens, PhD, and Dori Schafer, PhD, of the F.M. Kirby Neurobiology Center at Boston Children’s Hospital.

“In the field of neuroscience, glia have often been ignored,” says Stevens. “But glia aren’t the nerve glue, they’re actively communicating with neurons. People have gotten a new respect for glia and are hungry to know more about them.”

Such knowledge could eventually shed light on brain disorders ranging from autism to Alzheimer’s.

The “eat me” sign

We’re all born with more brain connections than we need. As we begin to encounter our world, they’re trimmed back to fine-tune our circuitry. It’s a bit of an oversimplification, but Stevens and Schafer demonstrated last week in the journal Neuron that when two neurons start talking to each other less – because their connection is no longer important to our lives– the microglia notice that and prune the synapse away.

To study microglia’s pruning activity, Stevens and Schafer used a time-honored model: the visual system. When you cover one eye soon after birth, you force the brain to rewire: Brain connections with the covered eye weaken and those synapses eventually get eliminated.

Using this model, Stevens and Schafer showed that microglia take their cues from a set of signals also used by the immune system, known as the complement cascade. Specifically, microglia carry receptors that recognize the complement protein C3—the same protein found on synapses that are destined for elimination.

“We think that weaker synapses are being tagged with C3, and that microglia are eliminating them just as macrophages would eliminate bacteria,” says Schafer.  “C3 is like an ‘eat me’ signal.”

As a postdoctoral fellow in 2007, Stevens showed that neurons are loaded with complement proteins soon after birth, just when pruning is at its peak. In the new study, she and Schafer deliberately disrupted complement signaling in mice—stripping the microglia of C3 receptors, or blocking those receptors with a drug. When they did so, pruning of irrelevant synapses didn’t occur.

Stevens thinks their findings might have relevance for brain disorders. Developmental brain disorders such as autism, epilepsy or schizophrenia are increasingly seen as disorders of synapse development, and some data suggest that microglia and/or the complement cascade are involved.

At the other end of the spectrum, scientists have noted that microglia—normally in a resting state in adults—are activated in neurodegenerative diseases like glaucoma, Alzheimer’s disease, Lou Gehrig’s disease, Huntington’s disease and Parkinson’s disease. Subtle changes have been found in synapses that might cause them to be targeted for elimination.

So could targeting microglia or the complement cascade prevent synapse loss or alter pruning in these diseases?  “All this is still very speculative,” Stevens cautions. “We first need to understand normal brain development.”


Beth Stevens


Assistant Professor of Neurology, F. M. Kirby Neurobiology Center, Boston Children’s Hospital

Department of Neurology, Harvard Medical School

Boston, Massachusetts

Age: 45

Published September 28, 2015

Beth Stevens is a neuroscientist whose research on microglial cells is prompting a significant shift in thinking about neuron communication in the healthy brain and the origins of adult neurological diseases. Until recently, it was believed that the primary function of microglia was immunological; they protected the brain by reducing inflammation and removing foreign bodies.

Stevens identified an additional, yet critical, role: the microglia are responsible for the “pruning” or removal of synaptic cells during brain development. Synapses form the connections, or means of communication, between nerve cells, and these pathways are the basis for all functions or jobs the brain performs. Using a novel model system that allows direct visualization of synapse pruning at various stages of brain development, Stevens demonstrated that the microglia’s pruning depends on the level of activity of neural pathways. She identified immune proteins called complement that “tag” (or bind) excess synapses with an “eat me” signal in the healthy developing brain. Through a process of phagocytosis, the microglia engulf or “eat” the synapses identified for elimination. This pruning optimizes the brain’s synaptic arrangements, ensuring that it has the most efficient “wiring.”

Stevens’s discoveries indicate that our adult neural circuitry is determined not only by the nerve cells but also by the brain’s immune cells. Her work suggests that adult diseases caused by deficient neural architecture (such as autism and schizophrenia) or states of neurodegeneration (such as Alzheimer’s or Huntington’s disease) may be the result of impaired microglial function and abnormal activation of this pruning pathway. Stevens is redefining our understanding of how the wiring in the brain occurs and changes in early life and shedding new light on how the nervous and immune systems interact in the brain, both in health and disease.

Beth Stevens received B.S. (1993) from Northeastern University and a Ph.D. (2003) from the University of Maryland. She was a postdoctoral fellow (2005–2008) at Stanford University and is currently an assistant professor in the Department of Neurology at Harvard Medical School and the F. M. Kirby Neurobiology Center at Boston Children’s Hospital. She is also an Institute Member of the Broad Institute of MIT and Harvard. Her scientific papers have appeared in such journals as NeuronScienceProceedings of the National Academy of Sciences, and Nature Neuroscience, among others.

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Portraits of scientists who are making a mark on autism research.

Beth Stevens: Casting immune cells as brain sculptors


Shortly after Beth Stevens launched her lab at Boston Children’s Hospital in 2008, she invited students from the Newton Montessori School, in a nearby suburb, to come for a visit. The children peered at mouse and rat brains bobbing in fluid-filled jars. They also learned how to position delicate slices of brain tissue on glass slides and inspect them with a microscope.

This visit sparked a running relationship with the school, with a steady stream of students visiting the growing lab each year. Soon it became too complicated to bring so many children to the lab, so Stevens decided to take her neuroscience lessons on the road, visiting a number of local elementary schools each year. Last year, she dropped in on the classrooms of her 5- and 8-year-old daughters, Zoe and Riley.

“The kids got really excited,” Stevens says. “It’s become such a thing that the principal wants me to come back for the whole school.”

Stevens’ enthusiasm for science has left a lasting impression on researchers, too. Her pioneering work points to a surprise role in brain development formicroglia, a type of cell once considered to simply be the brain’s immune defense system, cleaning up cellular debris, damaged tissue and pathogens. But thanks to Stevens, researchers now appreciate that these non-neuronal cells also play a critical role in shaping brain circuits.

In a 2012 discovery that created a buzz among autism researchers, Stevens and her colleagues discovered that microglia prune neuronal connections, calledsynapses, in the developing mouse brain. The trimming of synapses is thought to go awry in autism. And indeed, emerging work from Stevens’ lab hints at a role for microglia in the disorder.

Stevens has already earned praise and several prizes for her work. In 2012, shereceived the Presidential Early Career Award for Scientists and Engineers, the most prestigious award that the U.S. government bestows on young scientists. And in October, she’ll deliver one of four presidential lectures at the world’s largest gathering of neuroscientists — the annual meeting of the Society for Neuroscience — an honor she shares with three neuroscience heavyweights, including two Nobel laureates.

“The field is probably expecting a lot from Beth,” says Jonathan Kipnis, professor of neuroscience at the University of Virginia. Stevens has put microglia at the forefront, Kipnis says. “What used to be a stepchild of neuroscience research is now getting a lot of attention, and I think in part it’s due to her research.”

Curious mind:

Stevens was born in 1970 in Brockton, Massachusetts, where her mother taught elementary school and her father was the school’s principal. As a child, she was deeply inquisitive, eager to understand how things work. She enjoyed collecting bugs and worms, and would analyze these precious specimens in makeshift labs in her backyard.

But a career in science wasn’t on her radar until high school, when she took a biology class with an inspiring teacher named Anthony Cabral. “He totally made me realize that this could be a career, that I could be a scientist,” Stevens says. “It was that one class that changed it, and I’m like, ‘Okay, I’m going to do this.’”

In 1988, she began studying biology at Northeastern University in Boston, which offered an unusual opportunity. It had a unique cooperative education program that allowed Stevens to spend several semesters working full time in medical labs after finishing her coursework.

After that experience, Stevens knew she wanted to find a job in a research lab. After graduating in 1993, she joined her then-boyfriend Rob Graham, now her husband, in Washington, D.C., where he had landed a job in the U.S. Senate. Stevens headed to the National Institutes of Health (NIH) in Rockville, Maryland, to apply for a job as a research assistant.

At around the same time, neuroscientist R. Douglas Fields was launching his lab at the NIH. He studied how neural impulses influence glia — a class of non-neuronal cells that includes microglia — and shape the structure of the developing brain. Fields readily hired Stevens despite her lack of expertise in neuroscience. “I was impressed with her work ethic, energy and drive,” he says.

Stimulating research:

In Fields’ lab, Stevens used a multi-compartment cell culture system to investigate whether stimulating neurons influences the activity of Schwann cells, glial cells that produce a fatty substance called myelin, which insulates nerves1. She discovered that patterns of neural impulses similar to those that occur during early development influence the maturation of Schwann cells and the production of myelin.

The findings added to mounting evidence that glia and neurons communicate with each other, a newly emerging concept at the time.

“What I loved about the glia research was that there were so few neuroscientists studying it; it was such a mysterious part of neuroscience,” Stevens says. “Those years in Doug’s lab were really exciting because it was a new field.”

Stevens spent five years in Fields’ lab. “She was doing extraordinary work,” Fields says. “She had the potential and the interest to do neuroscience research, and I recommended that she should consider going to graduate school.”

But Stevens didn’t want to give up her position in the lab, and at that time, the NIH did not allow its researchers to have graduate students. So she and Fields convinced the University of Maryland, College Park, just 10 miles away, to allow her to take graduate courses in neuroscience while completing the necessary research for her Ph.D. in Fields’ lab.

In 2000, less than two years after starting graduate school, Stevens published a paper in Science showing that nerves in the peripheral nervous system (located outside the brain and spinal cord) use chemical signals to communicate with Schwann cells2. Two years later, she reported in Neuron that a similar form of communication occurs in the brain, between neurons and oligodendrocytes, the myelin-producing cells in the brain3.

As she was closing in on her Ph.D., Stevens sought career advice from Story Landis, then-director of the National Institute of Neurological Disorders and Stroke. Landis turned Stevens on to the possibility of starting her own lab one day. “I convinced her that she really had the abilities and energy and intelligence to run an independent research program,” Landis says.

In 2004, Stevens sought a postdoctoral fellowship with neurobiologist Ben Barres at Stanford University. “She was already seen as a leading researcher in the glial field,” recalls Barres, who promptly hired her. “She had done all sorts of beautiful work on glia.”

In Barres’ lab, Stevens continued to explore the dialogue between neurons and glia, turning her attention to star-shaped glia called astrocytes. Barres and his team had discovered that astrocytes help neurons form synapses4. To get a better handle on this process, Stevens examined how astrocytes influence gene expression in neurons in the developing mouse brain.

To her surprise, she found that astrocytes trigger neurons to produce a ‘complement’ protein that is best known for its role in the immune system. There, the protein serves as an ‘eat me’ signal, flagging pathogens and debris for removal. She found that neurons deposit this tag around immature synapses, but not mature ones, in mouse brain tissue, and mice that lack this protein have too many immature synapses. The findings suggested that astrocytes might help eliminate synapses by triggering the complement cascade5.

Young recruit: Beth Stevens’ daughter Riley inspects brain tissue during a visit to her mother’s lab. | Courtesy of Beth Stevens

But it was still unclear exactly how the tagged synapses are cleared. The prime suspects were microglia, the only cells in the brain known to have the receptor for the ‘eat me’ signal.

Stevens set out to test this hypothesis in her own lab: After four years as a postdoc, she had decided to branch out on her own. In 2008, neuroscientist Michael Greenberg — chair of the neurobiology department at Harvard — recruited her to the Harvard-affiliated Boston Children’s Hospital. Even when her lab was in its infancy, she had little trouble convincing new staff to join her.

“A lot of people might be a little hesitant to join a new lab,” says Dorothy Schafer, a former postdoctoral fellow in Stevens’ lab who is now assistant professor of neurobiology at the University of Massachusetts-Worcester. “But I was so excited by the research, and she was so energetic and extremely positive, and just seemed like a very nice person.”

One decision Stevens made early on was to continue to studying microglia in mice rather than experiment with new model systems. “You’ll never see her working on songbirds, because she has this aversion to birds,” Schafer says. “I think they think her curly blond hair is a nest or something, and she’s had really bad experiences with many types of birds dive-bombing her head.”

Just four years into her foray as an independent researcher, Stevens found the proof she had been looking for. In 2012, her team published evidence that microglia eat synapses, especially those that are weak and unused6.

The findings pinned down a new role for microglia in wiring the brain. They also helped to explain how the brain, which starts out with a surplus of neurons, trims some of the excess away. Neuron named the paper its most influential publication of 2012.

Stevens continues to study the function of microglia in the healthy brain, most recently uncovering preliminary evidence that a certain protein serves as a ‘don’t eat me’ tag that protects synapses from being engulfed by microglia. She is also exploring the role of microglia in disorders such as autism.

Several studies suggest that microglia are more active and more numerous in the brains of people with autism than in controls. Stevens and her team are looking at whether the activity of microglia is altered during brain development in mouse models of autism.


Immunodulatory Thalidomides in ~ conjugants unleash proteasome degradation on ~ oncoproteins with distinct mechanisms- BRD4,MYC & PIM1 & little collteral damage to 7429 other proteins!

Imagine being able to specifically target a cancer protein for immediate destruction, slipping Robert Louis Stevenson’s notorious black spot into a crevice in the secondary structure and spelling imminent death. Well, this is what Winter et al. (2015) describe in a recent drug discovery report for Science.1 Using phthalimide conjugation, the researchers not only specifically marked BRD4, a transcriptional coactivator important in MYC oncogene upregulation, for proteasomal degradation, but also achieved reduced tumor burdens in vivo.

The research team combined two drugs, thalidomide and JQ1, exploiting the properties of each to create a bifunctional compound, dBET1, that drives the proteasomal degradation of BRD4. JQ1, which in itself is anti-oncogenic, selectively binds BET bromodomains on the transcription factor, thus competitively inhibiting BRD4 activity on chromatin. Thalidomide, a phthalimide-based drug with immunomodulatory properties, binds cereblon (CRBN) in the cullin-RING ubiquitin ligase (CRL) complex, which is important in proteasomal protein degradation.

After confirming that the new phthalimide conjugate, dBET1, retained affinity for BRD4 and that this binding was specific, the team used a human acute myelocytic leukemia (AML) cell line, MV4;11, to show that treatment with the conjugate over 18 hours reduced BRD4 abundance. The researchers also found this with dBET1 treatment of other human cancer cell lines (SUM159, MOLM13). Following this, Winter et al. investigated the mechanisms by which dBET1 inhibits BRD4. By focusing primarily on proteasome function, the researchers determined that the reduction in BRD4 abundance in MV4;11 cells is proteasomal and dependent on CRBN binding activity.

Having established targeted proteasomal degradation using the dBET1 conjugate, Winter et al. then investigated the proteomic consequences of treatment in MV4;11 cells. Scientists at the Thermo Fisher Scientific Center for Multiplexed Proteomics (Harvard Medical School) used quantitative proteomics analysis with an isobaric tagging approach to compare the immediate effects of dBET1 treatment following two hours of incubation with the responses to JQ1 and vehicle control. Spectral data analysis identified 7,429 proteins with few differences in response to either treatment. JQ1 treatment reduced levels of MYC and oncoprotein PIM1 similarly to the response following dBET1 incubation. However, treatment with the latter also reduced BRD2, BRD3 and BRD4 abundance, findings that the research team confirmed with specific immunoblotting. Measuring expression of mRNA showed that both treatments reduced levels of MYC and PIM1 abundance. However, Winter et al. found no difference in BRD3 and BRD4, suggesting that dBET1 reduces the protein levels by post-transcriptional regulation.

Investigating the antiproliferative potential of the phthalimide conjugate, dBET1, Winter and coauthors examined apoptotic response in both MV4;11 and DHL4 lymphoma cells, and in primary human AML blast cultures. Compared to JQ1 treatment, dBET1 stimulated a profound and prolonged apoptotic response in both cell lines, suggesting that targeted degradation could be a more effective treatment than target inhibition.

shapes of proteins as they shift from one stable shape to a different, folded one Protein-structural-changes

shapes of proteins as they shift from one stable shape to a different, folded one Protein-structural-changes

Orchestrating the unfolded protein response in health and disease

Randal J. Kaufman Department of Biological Chemistry,
Howard Hughes Medical Institute, University of Michigan Medical Center, Ann Arbor, Michigan, USA J. Clin. Invest. 110:1389–1398 (2002).

The endoplasmic reticulum (ER), the entrance site for proteins destined to reside in the secretory pathway or the extracellular environment, is also the site of biosynthesis for steroids and for cholesterol and many lipids. Given the considerable number of resident structural proteins and biosynthetic enzymes and the high expression of many secreted proteins, the total concentration of proteins in the this organelle can reach 100 mg/ml. The ER relies on an efficient system of protein chaperones that prevent the accumulation of unfolded or aggregated proteins and correct misfolded proteins that are caught in low-energy kinetic traps (see Horwich, this Perspective series, ref. 1).

These chaperone-mediated processes expend metabolic energy to ensure high-fidelity protein folding in the lumen of the ER. For example, the most abundant ER chaperone, BiP/GRP78, uses the energy from ATP hydrolysis to promote folding and prevent aggregation of proteins within the ER. In addition, the oxidizing environment of the ER creates a constant demand for cellular protein disulfide isomerases to catalyze and monitor disulfide bond formation in a regulated and ordered manner. Operating in parallel with chaperone dependent protein folding are several “quality control” mechanisms, which ensure that, of all proteins translocated into the ER lumen, only those that are properly folded transit to the Golgi compartment. Proteins that are misfolded in the ER are retained until they reach their native conformation or are retrotranslocated back into the cytosol for degradation by the 26S proteasome. The ER has evolved highly specific signaling pathways to ensure that its protein-folding capacity is not overwhelmed. These pathways, collectively termed the unfolded protein response (UPR), are required if the cell is to survive the ER stress (see Ron, this Perspective series, ref. 2) that can result from perturbation in calcium homeostasis or redox status, elevated secretory protein synthesis, expression of misfolded proteins, sugar/glucose deprivation, or altered glycosylation. Upon accumulation of unfolded proteins in the ER lumen, the UPR is activated, reducing the amount of new protein translocated into the ER lumen, increasing retrotranslocation and degradation of ER-localized proteins, and bolstering the protein-folding capacity of the ER. The UPR is orchestrated by the coordinate transcriptional activation of multiple genes, a general decrease in translation initiation, and a concomitant shift in the mRNAs that are translated.

The recent discovery of the mechanisms of ER stress signaling, coupled with the ability to genetically engineer model organisms, has led to major new insights into the diverse cellular and physiological processes that are regulated by the UPR. Here, I summarize current discoveries that have offered insights into the complex regulation of the UPR and its relevance to human physiology and disease.

Glucose and protein folding Early studies demonstrated that both viral transformation and glucose depletion induce transcription of a set of related genes that were termed glucose-regulated proteins (GRPs) (3). Since viral transformation increases both the cellular metabolic rate and ATP utilization, it became evident that, in both cases, this signal emanates from the ER as a consequence of energy deprivation. Because proteins have different ATP requirements for protein folding prior to export, it has been proposed that the threshold for UPR activation might differ among various cell types, depending on their energy stores and the amount and nature of the secretory proteins they produce (4). Glucose not only provides the metabolic energy needed by cells but also participates directly in glycoprotein folding as a component of oligosaccharide structures.

The recognition and modification of oligosaccharide structures in the lumen of the ER is intimately coupled to polypeptide folding (5). As the growing nascent chain is translocated into the lumen of the ER, a 14-oligosaccharide core (GlcNAc2Man9Glc3) is added to consensus asparagine residues. Immediately after the addition of this core, the three terminal glucose residues are cleaved by the sequential action of glucosidases I and II to yield a GlcNAc2Man9 structure. If the polypeptide is not folded properly, a UDP-glucose:glycoprotein glucosyltransferase (UGGT) recognizes the unfolded nature of the glycoprotein and reglucosylates the core structure to re-establish the glucose-α(1, 3)–mannose glycosidic linkage. Monoglucosylated oligosaccharides containing this bond bind to the ER-resident protein chaperones calnexin and calreticulin.

This quality control process ensures that unfolded glycoproteins do not exit the ER. Treatment of cells with castanospermine, a transition-state analogue inhibitor of glucosidases I and II, inhibits this monoglucosylation cycle, prevents interaction of unfolded glycoproteins with calnexin and calreticulin, and activates the UPR. Genetic alterations that reduce the nucleotide sugar precursor pool or glycosyltransferase reactions likewise activate the UPR (6). Therefore, the recognition of altered carbohydrate structures is in some manner linked to UPR activation.

The UPR in yeast and higher eukaryotes On a cellular level, the accumulation of unfolded proteins in the ER lumen induces the transcription of a large set of genes whose products increase the ER’s volume or its capacity for protein folding or promote the degradation of misfolded proteins through the process of ER-associated protein degradation (ERAD) (7). For example, transcription of the ER protein chaperone BiP is a classical marker for UPR activation in yeast and mammalian cells (8). BiP binds hydrophobic exposed patches on the surfaces of unfolded proteins and interactive sites on unassembled protein subunits, and it releases its polypeptide substrates upon ATP binding.

In parallel, as Ron (this Perspective series, ref. 2) details in his accompanying article, translation is attenuated to decrease the protein-folding load. The complex network of physiological responses to ER stress is regulated by only a few ER transmembrane proteins: IRE1, PERK, and ATF6 (9). IRE1, PERK, and ATF6 are proximal sensors that regulate the production and/or quality of basic leucine zipper–containing (bZIP-containing) transcription factors that may form homo- and heterodimers. Combinatorial interactions of these factors generate diversity in responses for different subsets of UPRresponsive genes. In multicellular organisms, if these adaptive responses are not sufficient to relieve ER stress, the cell dies through apoptosis or necrosis.

IRE1-dependent splicing The UPR-signaling pathway was first described less than ten years ago in the budding yeast Saccharomyces cerevisiae. Elegant studies identified IRE1 as the sensor of unfolded proteins in the ER lumen. IRE1 is a type 1 transmembrane Ser/Thr protein kinase that also has a site-specific endoribonuclease (RNase) activity. The presence of unfolded proteins in the ER lumen promotes dimerization and trans-autophosphorylation, rendering IRE1 active as an RNase, and allowing it to cleave a 252-base intron from the mRNA encoding the transcription factor HAC1 (10). The 5′ and 3′ ends of HAC1 mRNA are spliced together by tRNA ligase in a process that is independent of the spliceosome and the usual intranuclear machinery for mRNA splicing. Splicing of HAC1 mRNA increases its translational efficiency and alters sequence of the encoded HAC1 protein, yielding a potent transcriptional activator (11) that can bind and activate the UPR elements (UPREs) upstream of many UPR-inducible genes. In S. cerevisiae, the UPR activates transcription of approximately 381 genes (7).

All eukaryotic cells appear to have maintained the essential and unique properties of the UPR present in S. cerevisiae, but higher eukaryotes possess additional sensors that generate diverse, coordinately regulated responses that promote stress adaptation or cell death. The mammalian genome contains two homologues of yeast IRE1 — IRE1α and IRE1β. Whereas IRE1α is expressed in most cells and tissues, with high-level expression in the pancreas and placenta (12), IRE1β expression is prominent only in intestinal epithelial cells (13). Both IRE1 molecules respond to the accumulation of unfolded proteins in the ER, which activate their kinase and, thereby, their RNase activities. The cleavage specificities of IRE1α and IRE1β are similar, if not identical, suggesting that they do not recognize different sets of substrates but rather generate temporally specific and tissue-specific expression (14, 15).

Searching for transcription factors that mediate the UPR, Yoshida et al. defined a mammalian ER stress response element [ERSEI; CCAAT(N9)CCACG] that is necessary and sufficient for UPR gene activation. Using a yeast one-hybrid screen, these authors isolated XBP1, a bZIP transcription factor X-box DNA binding protein (16). Subsequently, several groups demonstrated that XBP1 mRNA is a substrate for mammalian IRE1, much as the HAC1 mRNA in S. cerevisiae is processed by the yeast IRE1; this pathway is also conserved in Caenorhabditis elegans (17–20). On activation of the UPR, XBP1 mRNA is cleaved by IRE1 to remove a 26-nucleotide intron and generate a translational frameshift. As expected given the precedent of HAC1 regulation in yeast, the resulting processed mRNA encodes a protein with a novel carboxy-terminus that acts as a potent transcriptional activator.

Overexpression of either IRE1α or IRE1β is sufficient to activate transcription from a BiP promoter reporter construct (15). Analysis of a minimal UPRE motif (TGACGTGC/A) (21) uncovered a transcriptional defect in IRE1α-null mouse embryo fibroblasts that could be complemented by expression of spliced XBP1 mRNA (20), and Yoshida et al. (unpublished data) recently identified a UPR-inducible gene that uniquely requires IRE1α-mediated splicing of XBP1 mRNA. However, neither IRE1α nor IRE1β is necessary for transcriptional activation of the BiP gene, as judged by the phenotype of IRE1α/β–deleted murine cells (20, 22, 23). These results indicate that a subset of UPR targets require IRE1 but that at least one IRE1-independent pathway exists for UPR-mediated transcriptional induction. Deletion of IRE1α causes embryonic lethality at embryonic day 10.5 (E10.5) (20, 22, 23). Therefore, although IRE1α is not required for the UPR, it is clearly required for mammalian embryogenesis. XBP1 deletion also causes embryonic lethality, but the mutant embryos can survive up to day E14.5, consistent with the notion that XBP1 acts downstream of IRE1α. XBP1 deletion causes cardiomyopathy and liver hypoplasia (24, 25). In contrast, IRE1β-null mice develop normally but exhibit increased susceptibility to experimentally induced colitis, a phenotype that is consistent with the specific expression of this kinase in the intestinal epithelium (26).

Activation of ATF6 and PERK by ER stress The activating transcription factor ATF6 (16) has been identified as another regulatory protein that, like XBP1, can bind ERSEI elements in the promoters of UPRresponsive genes. There are two forms of ATF6, both synthesized as ER transmembrane proteins. ATF6α (90 kDa) and ATF6β (110 kDa, also known as CREB-RP) both require the presence of the transcription factor CBF (also called NF-Y) to bind ERSEI (27–30).

On activation of the UPR, both forms of ATF6 are processed to generate 50- to 60-kDa cytosolic, bZIP containing transcription factors that migrate to the nucleus (27). Processing of ATF6 by site-1 protease (S1P) and site-2 protease (S2P) occurs within the transmembrane segment and at an adjacent site exposed to the ER lumen. S1P and S2P are the processing enzymes that cleave the ER-associated transmembrane sterolresponse element–binding protein (SREBP) upon cholesterol deprivation (31). The cytosolic fragment of cleaved SREBP migrates to the nucleus to activate transcription of genes required for sterol biosynthesis. Interestingly, although the mechanism regulating ATF6 processing is similar to that regulating SREBP processing (32), the UPR only elicits ATF6 processing, whereas sterol deprivation alone induces SREBP processing. The SREBP cleavage–activating protein (SCAP) confers specificity for SREBP transport to the Golgi compartment, and consequently cleavage in response to sterol deprivation (33). It is unknown whether another cleavage-activating protein, analogous to SCAP but active only following induction of the UPR, promotes the specific cleavage and activation of ATF6 by S1P and S2P.

Transcription of UPR-responsive genes is induced when the cleaved form of ATF6 activates the XBP1 promoter. Therefore, signaling through ATF6 and IRE1 merges to induce XBP1 transcription and mRNA splicing, respectively (Figure 1, a and b). ATF6 increases XBP1 transcription to produce more substrate for IRE1- mediated splicing that generates more active XBP1, providing a positive feedback for UPR activation. However, cells that lack either IRE1α or ATF6 cleavage can induce XBP1 mRNA (20). These two pathways may thus provide parallel signaling pathways for XBP1 transcriptional induction. Alternatively, another pathway — possibly mediated by the ER-localized protein kinase PERK (see Ron, this Perspective series, ref. 2) — may also contribute to induction of XBP1 mRNA. The binding specificities of XBP1 and ATF6 are similar, although ATF6 binding requires CBF binding to an adjacent site, whereas XBP1 binds independently (17, 20, 21, 34). These binding specificities provide another avenue for complementary interaction between the IRE1-XBP1 and ATF6 pathways at the level of transcriptional activation. In addition, these transcription factors might regulate transcription from a second ERSE (ERSEII), which also contains a CCACG motif (35).

In parallel with the activation of ATF6 processing and the consequent changes in gene transcription, the accumulation of unfolded proteins in the ER also alters cellular patterns of translation. The protein kinase PERK has been implicated in this aspect of the ER stress response (see Ron, this Perspective series, ref. 2). Activated PERK phosphorylates the α subunit of eukaryotic translation initiation factor 2 (eIF2α) and attenuates general protein synthesis. Inactivation of the PERK-eIF2α phosphorylation pathway decreases cells’ ability to survive ER stress (36, 37). The PERK pathway promotes cell survival not only by limiting the protein-folding load on the ER, but also by inducing transcription of UPR- activated genes, one-third of which require phosphorylation of eIF2α for their induction (36). Preferential translation of the transcription factor ATF4 allows for continued activation of these genes under conditions of stress, when general protein synthesis is inhibited (36, 37).

A coordinated mechanism for activation One puzzling question about the UPR is how three independent sensors are activated by a common stimulus, the accumulation of unfolded proteins in the ER lumen. BiP, which negatively regulates the UPR, interacts with all three sensors, IRE1, PERK, and ATF6, under nonstressed conditions and may indeed be the master regulator of UPR activation.

Upon accumulation of unfolded proteins in the ER, BiP is released from IRE1, PERK, and ATF6. It is believed that the unfolded proteins bind BiP and sequester it from interacting with IRE1, PERK, and ATF6 to elicit their activation. In this manner, BiP senses both the level of unfolded proteins and the energy (ATP) level in the cell in regulating the UPR. Following release from BiP, IRE1 and PERK are each free to undergo spontaneous homodimerization mediated by their lumenal domains and to become phosphorylated by their endogenous kinase activities (38, 39). BiP interaction with ATF6 prevents trafficking of ATF6 to the Golgi compartment. For this reason, BiP release permits ATF6 transport to the Golgi compartment, where it gains access to S1P and S2P proteases (32). The regulation of signaling through the free level of BiP is an attractive hypothesis providing a direct mechanism by which all three ER stress sensors could be activated by the same stimulus. In addition, the increase in BiP during the UPR would provide a negative feedback to turn off UPR signaling. However, in certain cells, different stress conditions can selectively activate only one or two of the ER stress sensors. For example, in pancreatic β cells, glucose limitation appears to activate PERK prior to activation of IRE1 (D. Scheuner and R.J. Kaufman, unpublished results). It will be important to elucidate how general BiP repression permits the selective activation of individual components of the UPR that mediate various downstream effects.

Signaling the UPR in eukaryotes

Signaling the UPR in eukaryotes

Figure 1 Signaling the UPR in eukaryotes.

Three proximal sensors, IRE1, PERK, and ATF6, coordinately regulate the UPR through their various signaling pathways. Whereas IRE1 and PERK are dispensable for many aspects of the response, ATF6 cleavage is required for UPR transcriptional induction and appears to be the most significant of these effectors in mammalian cells. BiP negatively regulates these pathways. BiP interacts with ATF6 to prevent its transport to the Golgi compartment (a). BiP binds to the lumenal domains of IRE1 (b) and PERK (c) to prevent their dimerization. As unfolded proteins accumulate, they bind BiP and reduce the amount of BiP available to bind and inhibit activation of IRE1, PERK, and ATF6. (a) BiP release from ATF6 permits transport to the Golgi compartment. In the Golgi, ATF6 is cleaved by S1P and S2P proteases to yield a cytosolic fragment that migrates to the nucleus to activate transcription of responsive genes, including XBP1. (b) BiP release from IRE1 permits dimerization to activate its kinase and RNase activities to initiate XBP1 mRNA splicing. XBP1 splicing removes a 26-base intron, creating a translational frameshift to yield a more potent transcriptional activator. (c) BiP release permits PERK dimerization and activation to phosphorylate Ser51 on eIF2α to reduce the frequency of AUG initiation codon recognition. As eIF2α phosphorylation reduces the functional level of eIF2, the general rate of translation initiation is reduced. However, selective mRNAs, such as ATF4 mRNA, are preferentially translated under these conditions, possibly by the presence of open reading frames within the 5′ untranslated region of the mRNA. Upon recovery from the UPR, GADD34 targets PP1 to dephosphorylate eIF2α and increase protein translation.

The UPR as a mediator of programmed cell death In contrast to UPR-signaling adaptation in response to ER stress, prolonged UPR activation leads to apoptotic cell death (Figure 2). The roles of several death-promoting signaling pathways have been shown by analysis of specific gene-deleted cells. Activated IRE1 recruits c-Jun-N-terminal inhibitory kinase (JIK) and the cytosolic adaptor TRAF2 to the ER membrane (22, 40). TRAF2 activates the apoptosis-signaling kinase 1 (ASK1), a mitogen-activated protein kinase kinase kinase (MAPKKK) (41). Activated ASK1 leads to activation of the JNK protein kinase and mitochondriadependent caspase activation (40–42).

ER insults lead to caspase activation by mitochondria/APAF-1–dependent and –independent pathways. ER stress promotes cytochrome c release from mitochondria, possibly by c-ABL kinase (43) or calcium (44). However, APAF1–/– cells are susceptible to ER stress–induced apoptosis, indicating that the mitochondrial pathway is not essential (45). Caspase-12 is an ER-associated proximal effector in the caspase activation cascade, and cells lacking this enzyme are partially resistant to inducers of ER stress (46). ER stress induces TRAF2 release from procaspase 12, allowing it to bind activated IRE1. As shown in Figure 2, release of TRAF2 permits clustering of procaspase-12 at the ER membrane, leading to its activation (40). Caspase-12 can activate caspase-9, which in turn activates caspase- 3 (47). Procaspase-12 can also be activated by m-calpain in response to calcium release from the ER, although the physiological significance of this pathway is not known (48). In addition, upon ER stress, procaspase-7 is activated and recruited to the ER membrane (49). These findings support the notion that ER stress leads to several redundant pathways for caspase activation.

A second death-signaling pathway activated by ER stress is mediated by transcriptional activation of genes encoding proapoptotic functions. Activation of UPR sensor IRE1, PERK, or ATF6 leads to transcriptional activation of CHOP/GADD153, a bZIP transcription factor that potentiates apoptosis (see Ron, this Perspective series, ref. 2).

The UPR in health and disease Primary amino acid sequence contains all the information for a protein to attain its final folded conformation. However, many folding intermediates exist along the folding pathway (see Horwich, this Perspective series, ref. 1), and some of these intermediates can become irreversibly trapped in low-energy states and activate the UPR. Clearance of such misfolded species requires a functional ER-associated degradation (ERAD) pathway, which is regulated by the UPR. Proteasomal degradation of ER-associated misfolded proteins is required to protect from UPR activation. Proteasomal inhibition is sufficient to activate the UPR, and, in turn, genes encoding several components of ERAD are transcriptionally induced by the UPR (7). Therefore, it is to be expected that UPR activation and impaired ERAD function might contribute to a variety of diseases and that polymorphisms affecting the UPR and ERAD responses could modify disease progression. The following examples provide the best available evidence linking the UPR pathway to the natural history of human diseases and animal models of these diseases.

The UPR and ERAD in genetic disease Many recessive inherited genetic diseases are due to loss  of-function mutations that disturb productive folding and that produce proteins that are either not secreted or not functional. In other cases, protein-folding mutations can interfere with cellular processes, resulting in a gain of function and a dominant pattern of inheritance. In several instances, UPR activation by the accumulation of unfolded proteins in the ER is known to contribute to disease progression. The distinction between these two classes of genetic disease is important, because gain-of-function protein-misfolding mutations will be less amenable to treatment by gene therapy to deliver a wild-type copy of the mutant gene.

One well-characterized protein-folding defect results from a mutation that leads to type 1 diabetes. The Akita mouse has a gain-of-function Cys96Tyr mutation in the proinsulin 2 (Ins2) gene; this mutation disrupts proinsulin folding. The mutant protein is retained in the ER of the pancreatic β cell and activates the UPR. Crucially, the progressive development of diabetes in this model is not solely due to the lack of insulin but is rather a consequence of the misfolded protein accumulation, UPR activation, and β cell death. When bred into a Chop–/–background, the Akita mutation causes a lesser degree of β cell death and delayed onset of diabetes (50), indicating that the loss of at least one downstream signaling component of the UPR can ameliorate pathogenesis in this setting.

Signaling UPR-mediated cell death

Signaling UPR-mediated cell death

Figure 2 Signaling UPR-mediated cell death.

The activation of procaspase-12 is likely the major pathway that induces apoptosis in response to ER stress. Upon activation of the UPR, c-Jun-N-terminal inhibitory kinase (JIK) release from procaspase-12 permits clustering and activation of procaspase-12. Caspase-12 activates procaspase-9 to activate procaspase-3, the executioner of cell death. In addition, activated IRE1 binds JIK and recruits TRAF2, which signals through apoptosis-signaling kinase 1 (ASK1) and JNK to promote mitochondria-dependent apoptosis. In addition, in vitro studies suggest that localized calcium release from the ER activates m-calpain to cleave and activate procaspase-12. Upon UPR activation, procaspase-7 is activated and recruited to the ER membrane. Finally, IRE1, PERK, and ATF6 induce transcription of several genes encoding apoptotic functions, including CHOP/GADD153. CSP, caspase; pCSP, procaspase.

Deficiency in α1-proteinase inhibitor (α1-PI, also known as α1-antitrypsin) results in emphysema and destructive lung disease in one out of 1,800 births. However, a subgroup of affected individuals develop chronic liver disease and hepatocellular carcinoma as a consequence of a secretion defect in the misfolded protein at the site of synthesis, the hepatocyte. This is the most common genetic cause of liver disease in children. The Z allele of the α1 gene PI (Glu342Lys mutation) produces a protein that polymerizes and is retained in the ER for degradation by the proteasome (see Lomas and Mahadeva, this Perspective series, ref. 51; and Perlmutter, this series, ref. 52). While α1-PI Z neither binds BiP nor activates the UPR, analysis of fibroblasts obtained from these patients demonstrates that individuals susceptible to liver disease have inherited a second trait that slows degradation of the misfolded protein in the ER (53), consistent with the idea that polymorphisms that reduce ERAD function can exacerbate pathogenesis of certain diseases.

There are numerous additional genetic misfolding diseases that are also likely influenced by UPR signaling. Because BiP release from IRE1, PERK, or ATF6 can activate the UPR, the expression of any wild-type or mutant protein that binds BiP can have a similar effect. In contrast, misfolded proteins that do not bind BiP are unlikely to activate the UPR. For example, cystic fibrosis is due to mutations in the cystic fibrosis transmembrane conductance regulator (CFTR) protein. Approximately 70% of patients with this disease carry a common mutation, deletion of Phe508, that results in a molecule that is retained in the ER and eventually degraded by the proteosome (see Gelman and Kopito, this Perspective series, ref. 54). Although expression of ∆508 CFTR does not activate the UPR in cultured cells, the protein does interact with calnexin, as well as HSP70, and requires ERAD function for cell survival.

Osteogenesis imperfecta (OI) results from misfolding mutations in procollagen that produce molecules that bind BiP and activate the UPR (55). Interestingly, Wolcott-Rallison syndrome is due to inactivating mutations in the PERK gene. Affected individuals, as well as mice with deletions in Perk, display osteoporosis and deficient mineralization throughout the skeletal system (56, 57), the same defects that are observed in OI. Procollagen type I accumulates to high levels and mature collagen is not detected in bone and osteoblasts from PERKnull mice. Osteoblasts from PERK-null humans and mice display fragmented and distended ER that is filled with electron-dense material (56, 57). These observations suggest that procollagen type 1 uniquely requires PERK function to maintain its transport out of the ER, processing, and secretion In this case, PERK may be required to limit procollagen synthesis so that it does not saturate the ER protein-folding capacity.

The UPR and ERAD in conformational diseases Diseases caused by expansion of polyglutamine repeats and neurodegenerative diseases, such as Alzheimer disease and Parkinson disease, represent a large class of conformational diseases associated with accumulation of abnormal protein aggregates in and around affected neurons. Recent evidence indicates that the pathogenesis of these diseases is due to a defect in proteasomal function that results in UPR activation, leading to cell death. The protein aggregates in these diseases are localized to the nucleus or the cytoplasm and would not be predicted to disturb ER function directly. Nevertheless, they have been found in some cases to activate the UPR and to promote cell death. Analysis of the polyglutamine repeat associated with the spinocerebrocellular atrophy protein (SCA3) in Machado-Joseph disease suggests that cytoplasmic accumulation of the SCA3 aggregate can inhibit proteasome function, thereby interfering with ERAD to induce the UPR and elicit caspase-12 activation (41, 58). These findings support the idea that the UPR can signal the accumulation of unfolded proteins in the cytosol via proteasomal inhibition and disruption of ERAD function.

Parkinson disease is the most common movement disorder, affecting about 1% of individuals 65 years of age or older. Autosomal recessive juvenile parkinsonism (AR-JP) results from defects in the Parkin gene (59), which encodes a ubiquitin protein ligase (E3) that functions with ubiquitin-conjugating enzyme UbcH7 or UbcH8 to tag proteins for degradation. Overexpression of Parkin suppresses cell death associated with ER stress (60). Inherited Parkinson disease is associated with the accumulation in the ER of dopaminergic neurons of PAEL-R, a putative transmembrane receptor protein that is detected in an insoluble form in the brains of AR-JP patients (61). The accumulation of PAEL-R results from defective Parkin that does not maintain the proteasome-degrading activity necessary to maintain ER function (62). Other, still-unidentified substrates of the Parkin E3 ligase may also be relevant to the pathogenesis of AR-JP.

The UPR in diabetes The metabolism of glucose is tightly controlled at the levels of synthesis and utilization through hormonal regulation. The most dramatic phenotype in Wolcott-Rallison syndrome is pancreatic β cell death with infancy onset diabetes (56). A similar defect is observed in PERK-null mice; this defect also correlated with increased apoptosis of β cells (57, 63). In addition, mice with a homozygous Ser51Ala mutation at the PERK phosphorylation site in eIF2α display an even greater β cell loss that appears in utero (36). Therefore, translational control through PERK-mediated phosphorylation of eIF2α is required to maintain β cell survival (see Ron, this Perspective series, ref. 2). The more severe β cell loss in mice harboring the Ser51Ala eIF2α mutation suggests that additional eIF2α kinases partially complement the requirement for PERK in β cell function (36)

Glucose not only promotes the secretion of insulin but also stimulates insulin transcription and translation (64–66). Our group has proposed that glucose stimulated proinsulin mRNA translation is regulated by PERK-mediated phosphorylation of eIF2α in response to UPR activation 36). As blood glucose declines, energy may become limiting for protein folding in the ER and therefore activate the UPR to promote PERK-mediated phosphorylation of eIF2α. Conversely, a rise in blood glucose would turn off the UPR so that translation would accelerate, allowing entry of new preproinsulin into the ER. In this manner, PERK mediated phosphorylation of eIF2α provides a brake on protein synthesis, including proinsulin translation. Continual elevation of blood glucose may also prolong elevated proinsulin translation, eventually activating the UPR as the secretion capacity of the ER is overwhelmed. Therefore, a delicate balance between glucose levels and eIF2α phosphorylation needs to be maintained: Disturbances in either direction may lead to excessive UPR activation, with eventual β cell death.

The insulin resistance and hyperglycemia associated with type 2 diabetes is accommodated by an increase in proinsulin translation. Under these conditions the UPR is activated to compensate for the increased protein-folding requirement in the ER. Prolonged activation of the UPR could contribute to the β cell death associated with insulin resistance. Thus, the signaling mechanisms that β cells use for sensing glucose levels, triggering insulin secretion, and rapidly controlling insulin biosynthesis may have coevolved with ER signaling pathways to support these specialized functions. Pancreatic β cells are exquisitely sensitive to physiological fluctuations in blood glucose, because, in contrast to other cell types, they lack hexokinase, an enzyme with a low affinity but a high capacity for binding glucose. Therefore, in β cells, the production of glucose 6-phosphate and the production of ATP through glycolysis are controlled by glucokinase (67), and the ratio of ATP to ADP correlates directly with the blood glucose level. Periodic decreases in blood glucose level (as occurs between meals) would decrease the ATP/ADP ratio and compromise protein folding in the ER so that the UPR may be frequently activated in these cells. Hence, when glucose levels vary within the normal physiological range, the ER compartment of the β cell may be exposed to greater energy fluctuations than is the ER of other cell types, making the β cell uniquely dependent on the UPR for survival during intermittent decreases in blood glucose levels, as happens between meals. Additionally, the high-level expression of PERK and IRE1α in the pancreas may predispose these kinases to dimerization and activation in response to intermittent stress.

The UPR in organelle expansion The UPR is required for ER expansion that occurs upon differentiation of highly specialized secretory cells, but ER membrane expansion can also proceed independently of UPR activation. Overexpression of membrane proteins, such as HMG CoA reductase or the peroxisomal protein Pex15, promotes the expansion of smooth membranes without UPR activation (68, 69), as does overexpression of the p180 ribosome acceptor in the rough ER membrane (70). Conversely, protein overexpression, even under circumstances in which secretory capacity is unchanged (as occurs following the induction of high levels of cytochrome p450), can activate the UPR to induce ER chaperone levels to match the expanded membrane area (71, 72).

During the terminal differentiation of certain secretory cells, such as those in the pancreas or liver, membrane expansion is accompanied by a dramatic increase in protein secretion. Likewise, upon B cell maturation into high-level antibody-secreting plasma cells, the ER compartment expands approximately fivefold to accommodate the large increase in Ig synthesis. The requirement for the UPR in this latter process has been demonstrated in XBP1–/– cells. Since deletion of XBP1 produces an embryonic-lethal phenotype at day E14.5, the role of XBP1 in B and T cell development had to be studied in immunoincompetent RAG1–/– mice reconstituted with XBP1–/– embryonic stem cells (73). Work in these chimeric mice demonstrated that XBP1 is required for high-level Ig production. Interestingly, the induction of Ig heavy-chain and light-chain gene rearrangement and the assembly and transport of Igµ to the surface of the B cells occurred normally. However, plasma cells were not detected, suggesting a role for XBP1 in plasma cell differentiation or survival.

These findings support the hypothesis that induction of Ig synthesis activates the UPR to induce ER expansion to accommodate the high-level antibody expression. Alternatively, activation of the UPR may be part of the differentiation program that occurs prior to induction of high-level antibody synthesis. Plasma cell differentiation is stimulated in vivo by treatment with LPS or by ligation of CD40 receptors, treatments that activate the innate immune response and have been shown to induce XBP1 mRNA splicing (19). Thus, the UPR may contribute to a programmed response to signals that increase a cell’s protein-secretory demand.

The UPR in hyperhomocysteinemia. The association between high levels of serum homocysteine and the development of ischemic heart disease and stroke is supported by substantial epidemiological data. Unfortunately, it is not known whether homocysteine is the underlying cause of atherosclerosis and thrombosis. Severe hyperhomocysteinemia is caused by mutation in the cystathionine β-synthase (CBS) gene, whose product is a vitamin B6–dependent enzyme required for the conversion of homocysteine to cysteine. Elevated homocysteine is also associated with vitamin B deficiency. In cultured vascular endothelial cells, homocysteine induces protein misfolding in the ER by interfering with disulfide bond formation, and it activates the UPR to induce expression of several ER stress response proteins, such as BiP, GRP94, CHOP, and HERP (74–76). Homocysteine also activates apoptosis in a manner that requires an intact IRE1-signaling pathway (76).

These findings suggest that homocysteine acts intracellularly to disrupt ER homoeostasis. Indeed, recent studies confirm that induction of hyperhomocysteinemia elicits UPR activation in the livers of normal or Cbs+/– mice (77). In addition, hyperhomocysteinemia activates SREBP cleavage, leading to intracellular accumulation of cholesterol (77). Increased cholesterol biosynthesis may explain the hepatic steatosis and possibly the atherosclerotic lesions associated with hyperhomocysteinemia. Finally, hyperhomocysteinemia accelerates atherosclerosis in ApoE–/– mice (78, 79), although the molecular mechanisms remain to be elucidated.

Hyperhomocysteinemia is also associated with increased amyloid production and increased amyloid-mediated neuronal death in animal models of Alzheimer disease (80). These observations suggest that the UPR may link the disease etiologies of hyperhomocysteinemia and Alzheimer disease. HERP, a homocysteine-induced ER stress–responsive gene, appears to be involved in amyloid β-protein (Aβ) accumulation, including the formation of senile plaques and vascular Aβ deposits (81), and that it interacts with both presenilin-1 (PS1) and presenilin-2 (PS2), thus regulating presenilin-mediated Aβ generation. Immunohistochemical analysis of brains from patients with Alzheimer disease reveals intense HERP staining in activated microglia in senile plaques.

The UPR in cancer Hypoxia is a common feature of solid tumors that display increased malignancy, resistance to therapy, and poor prognosis. Hypoxia in the tumor results from increased demand due to dysregulated cell growth and from vascular abnormalities associated with cancerous tissue. The importance of hypoxia has been seen in the clinic, since it predicts for poor outcome of treatments, independent of treatment modality. Hypoxia activates the UPR, whose downstream signaling events can undermine the efficacy of treatment. Tumor cells need to adapt to the increasingly hypoxic environment that surrounds them as they grow, and the induction of the UPR is key to this response. Induction of the ER stress response genes, for example BiP and GRP94, in cancerous tissue correlates with malignancy, consistent with their antiapoptotic function (82). In addition, the UPR confers resistance to topoisomerase inhibitors, such as etoposide, and some UPR-induced genes directly mediate drug resistance via the multi-drug-resistance gene MDR. Therefore, approaches to prevent UPR activation in cancerous cells may significantly improve treatment outcome.

The proteasome inhibitor PS-341 is now in earlyphase clinical evaluation for the treatment of multiple myeloma, a clonal B cell tumor of differentiated plasma cells (83). The mechanism of PS-341 function is thought to be inhibition of IκB degradation, which prevents activation of the antiapoptotic transcription factor NF-κB. However, proteasomal inhibition would also prevent ERAD. As high-level heavy- or light-chain Ig production is likely associated with a certain degree of protein misfolding, it is possible that inhibition of ERAD function may be selectively toxic to B cell myelomas through activation of the UPR and apoptosis.

The UPR and viral pathogenesis The two major mediators of the IFN-induced arm of the innate immune response are evolutionarily related to IRE1 and PERK. The kinase/endoribonuclease domain of IRE1 is homologous to RNaseL, and the protein kinase domain of PERK is related to the double-stranded RNA–activated (dsRNA-activated) eIF2α protein kinase PKR. RNase L and PKR mediate the IFN induced antiviral response of the host, which is required to limit viral protein synthesis and pathogenesis. As part of the innate immune response to viral infection, RNase L and PKR are activated by dsRNAs produced as intermediates in viral replication. In contrast to activation by dsRNA, IRE1 and PERK are activated by ER stress, which can be induced by high-level viral glycoprotein expression. All enveloped viruses produce excess glycoproteins that could elicit PERK and IRE1 activation to meet the need for increased folding and secretory capacity. More studies will be required to elucidate the role of the UPR in various viral diseases.

Hepatitis C virus (HCV) is a positive-stranded RNA virus encoding a single polyprotein. Polyprotein cleavage generates at least ten polypeptides, including two glycoproteins, E1 and E2. A large amount of E1 forms disulfide–cross-linked aggregates with E2 in the ER (84). Since the accumulation of misfolded α1-PI elicits UPR activation, with subsequent hepatocyte death and hepatocellular carcinoma, it is possible that the aggregated E1/E2 complexes in the HCV-infected hepatocyte also contribute to hepatitis and hepatocellular carcinoma. Future studies should identify whether these glycoprotein aggregates activate the UPR to mediate the hepatocyte cell death and transformation associated with the pathogenesis of HCV infection.

The UPR in tissue ischemia Finally, neuronal death due to reperfusion after ischemic injury is associated with activation of the UPR (85, 86). Immediately after reperfusion, protein synthesis is inhibited, due at least in part to phosphorylation of eIF2α; this inhibition may represent a protective mechanism to prevent further neuron damage. Recent studies support the idea that eIF2α phosphorylation in response to reperfusion injury is mediated by PERK and hence that it depends on the UPR (87). If so, UPR activation prior to ischemic injury might protect the brain and other tissues from cell death during periods of reperfusion.

Summary A variety of approaches have been employed to identify the UPR signaling components, their function, and their physiological role. Yeast genetics allowed the definition of the basic ER stress–signaling pathway. The identification of homologous and parallel signaling pathways in higher eukaryotes has produced a mechanistic framework the cell uses to sense and compensate for ER over-load and stress. The high-level tissue-specific expression patterns of several ER stress–signaling molecules indicated the pancreas and intestine as organs that require UPR for physiological function. Analysis of UPR-induced gene expression established that protein degradation is required to reduce the stress of unfolded protein accumulation in the ER. Major advances in identifying UPR function and rele vance to disease were derived from mutation of UPR signaling components in model organisms and the identification of mutations in humans.

Despite tremendous progress, our knowledge of the UPR pathway remains incomplete. Further studies promise to expand our understanding of how ER stress impacts the other cellular signaling pathways. It will be very exciting and informative to understand how the UPR varies when critical components are genetically manipulated by deletion or other types of mutations. In addition, although the accumulation of unfolded protein in the ER is now known to contribute to pathogenesis in a variety of diseases, there are still few therapeutic approaches that target these events. With a greater understanding of protein-folding processes, pharmacological intervention with chemical chaperones to promote proper folding becomes feasible, as observed with sodium phenylbutyrate for ∆508 CFTR (see Gelman and Kopito, this Perspective series, ref. 53). Future intervention should consider activation of different subpathways of the UPR or overexpression of appropriate protein chaperones, as in the case of overexpression of the J domain of cytosolic HSP70, which suppresses polyglutamine toxicity in flies (88). Treatments that activate the ERAD response may also ameliorate pathogenesis in a number of the conformational diseases.

Over the past ten years, tremendous progress has been made in understanding the mechanisms and physiological significance of the UPR. The processes of protein folding and secretion, transcriptional and translational activation, and protein degradation are intimately interconnected to maintain homeostasis in the ER. A variety of environmental insults, genetic disease, and underlying genetic modifiers of UPR function contribute to the pathogenesis of different disease states. As we gain a greater understanding of the mechanisms that control UPR activation, it should be possible to discover methods to activate or inhibit the UPR as desired for therapeutic benefit.

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Citation: Cell Death and Disease (2014) 5, e1578; doi:10.1038/cddis.2014.539
Published online 18 December 2014

An activated unfolded protein response promotes retinal degeneration and triggers an inflammatory response in the mouse retina

T Rana1, V M Shinde1, C R Starr1, A A Kruglov1, E R Boitet1, P Kotla1, S Zolotukhin2, A K Gross1 and M S Gorbatyuk1

  1. 1Department of Vision Sciences, University of Alabama at Birmingham, AL, USA
  2. 2Department of Pediatrics, University of Florida, FL, USA

Correspondence: M Gorbatyuk, Department of Vision Sciences, University of Alabama at Birmingham, 1670 University Boulevard, Birmingham, 35233 AL, USA. Tel: +1 205 934 6762; Fax: +1 205 934 3425;

Received 20 July 2014; Revised 23 October 2014; Accepted 27 November 2014

Edited by P Ekert

Recent studies on the endoplasmic reticulum stress have shown that the unfolded protein response (UPR) is involved in the pathogenesis of inherited retinal degeneration caused by mutant rhodopsin. However, the main question of whether UPR activation actually triggers retinal degeneration remains to be addressed. Thus, in this study, we created a mouse model for retinal degeneration caused by a persistently activated UPR to assess the physiological and morphological parameters associated with this disease state and to highlight a potential mechanism by which the UPR can promote retinal degeneration. We performed an intraocular injection in C57BL6 mice with a known unfolded protein response (UPR) inducer, tunicamycin (Tn) and examined animals by electroretinography (ERG), spectral domain optical coherence tomography (SD-OCT) and histological analyses. We detected a significant loss of photoreceptor function (over 60%) and retinal structure (35%) 30 days post treatment. Analysis of retinal protein extracts demonstrated a significant upregulation of inflammatory markers including interleukin-1β (IL-1β), IL-6, tumor necrosis factor-α (TNF), monocyte chemoattractant protein-1 (MCP-1) and IBA1. Similarly, we detected a strong inflammatory response in mice expressing either Ter349Glu or T17M rhodopsin (RHO). These mutant rhodopsin species induce severe retinal degeneration and T17M rhodopsin elicits UPR activation when expressed in mice. RNA and protein analysis revealed a significant upregulation of pro- and anti-inflammatory markers such as IL-1β, IL-6, p65 nuclear factor kappa B (NF-kB) and MCP-1, as well as activation of F4/80 and IBA1 microglial markers in both the retinas expressing mutant rhodopsins. We then assessed if the Tn-induced inflammatory marker IL-1β was capable of inducing retinal degeneration by injecting C57BL6 mice with a recombinant IL-1β. We observed ~19%reduction in ERG a-wave amplitudes and a 29% loss of photoreceptor cells compared with control retinas, suggesting a potential link between pro-inflammatory cytokines and retinal pathophysiological effects. Our work demonstrates that in the context of an established animal model for ocular disease, the persistent activation of the UPR could be responsible for promoting retinal degeneration via the UPR-induced pro-inflammatory cytokine IL-1β.


ERG, electroretinography; SD-OCT, spectral domain optical coherence tomography; UPR, unfolded protein response; IL-1β, Interleukin-1β; TNF-α, tumor necrosis factor-α; MCP-1, monocyte chemoattractant protein-1; NF-kB, ; nuclear factor kappa B, ; ER, endoplasmic reticulum; ADRP, autosomal dominant retinitis pigmentosa; RHO, rhodopsin; ERAI, ER stress activated indicator; Tn, tunicamycin; ONL, outer nuclear layer; H&E, hematoxylin and eosin; ONH, optic nerve head


ER stress and neuroinflammation: connecting the unfolded protein response to JAK/STAT signaling (P5196)

Gordon Meares,1 and Etty Benveniste1

1Department of Cell, Developmental and Integrative Biology, University of Alabama at Birmingham, Birmingham, AL

J Immunol May 2013 190 (Meeting Abstract Supplement) 198.5

Neuroinflammation and endoplasmic reticulum (ER) stress are associated with many neurological diseases. ER stress is brought on by misfolded proteins. In turn, cells respond with activation of the unfolded protein response (UPR). The UPR is a highly conserved pathway that transmits both adaptive and apoptotic signals to restore homeostasis or eliminate the irreparably damaged cell. Recent evidence indicates that ER stress and inflammation are linked. In this study, we have examined the interaction between ER stress and JAK/STAT-dependent inflammation in astrocytes. The JAK/STAT pathway mediates the biological actions of many cytokines and growth factors. We have found that ER stress leads to the activation of STAT3 in a JAK1-dependent fashion. ER stress-induced activation of the JAK1/STAT3 axis leads to expression of IL-6 and several chemokines. The activation of STAT3 signaling is dependent on the protein kinase PERK, a central component of the UPR. Knockdown of PERK abrogates ER stress-induced activation of STAT3 and overexpression of PERK is sufficient to activate STAT3. Additionally, ER stressed astrocytes, via paracrine signaling, can stimulate activation of microglia leading to production of oncostatin M (OSM). OSM can then synergize with ER stress in astrocytes to drive inflammation. Together, this work describes a new PERK-JAK1-STAT3 signaling pathway that may elicit a feed-forward inflammatory loop involving astrocytes and microglia to drive neuroinflammation.


Neural Plasticity
Volume 2014 (2014), Article ID 610343, 15 pages

Review Article

Surveillance, Phagocytosis, and Inflammation: How Never-Resting Microglia Influence Adult Hippocampal Neurogenesis

Amanda Sierra,1,2,3 Sol Beccari,2,3 Irune Diaz-Aparicio,2,3 Juan M. Encinas,1,2,3 Samuel Comeau,4,5 and Marie-Ève Tremblay4,5

1Ikerbasque Foundation, 48011 Bilbao, Spain
2Achucarro Basque Center for Neuroscience, Bizkaia Science and Technology Park, 48170 Zamudio, Spain
3Department of Neurosciences, University of the Basque Country, 48940 Leioa, Spain
4Centre de Recherche du CHU de Québec, Axe Neurosciences, Canada G1P 4C7
5Département de Médecine Moléculaire, Université Laval, Canada G1V 4G2

Received 10 December 2013; Accepted 11 February 2014; Published 19 March 2014

Academic Editor: Carlos Fitzsimons

Microglia cells are the major orchestrator of the brain inflammatory response. As such, they are traditionally studied in various contexts of trauma, injury, and disease, where they are well-known for regulating a wide range of physiological processes by their release of proinflammatory cytokines, reactive oxygen species, and trophic factors, among other crucial mediators. In the last few years, however, this classical view of microglia was challenged by a series of discoveries showing their active and positive contribution to normal brain functions. In light of these discoveries, surveillant microglia are now emerging as an important effector of cellular plasticity in the healthy brain, alongside astrocytes and other types of inflammatory cells. Here, we will review the roles of microglia in adult hippocampal neurogenesis and their regulation by inflammation during chronic stress, aging, and neurodegenerative diseases, with a particular emphasis on their underlying molecular mechanisms and their functional consequences for learning and memory.

  1. Microglia: The Resident Immune Cells of the Brain

Microglia were first described in 1919 by the Spanish neuroanatomist Pío del Río Hortega, a disciple of the renowned Santiago Ramón y Cajal, almost half a century later than neurons and astrocytes and just before oligodendrocytes [1]. This delayed appearance into the neuroscience arena is still apparent today, as microglia remain one of the least understood cell types of the brain. Traditionally, microglia were simply considered as “brain macrophages” controlling the inflammatory response during acute insults and neurodegenerative conditions, and only recently was their unique origin revealed. Indeed, microglia were shown to derive from primitive myeloid progenitors of the yolk sac that invade the central nervous system (CNS) during early embryonic development (reviewed in [2]). In contrast, circulating monocytes and lymphocytes, as well as most tissue macrophages, derive from hematopoietic stem cells located initially in the foetal liver and later in the bone marrow [3]. In the adult brain, the microglial population is maintained exclusively by self-renewal during normal physiological conditions [2]. As a consequence, microglia are the only immune cells which permanently reside in the CNS parenchyma, alongside neural tube-derived neurons, astrocytes, and oligodendrocytes.

These past few years, unprecedented insights were also provided into their extreme dynamism and functional behaviour, in health as much as in disease. Indeed, microglia were revealed to be exceptional sensors of their environment, responding on a time scale of minutes to even subtle variations of their milieu, by undergoing concerted changes in morphology and gene expression [45]. During pathological insults, “activated” microglia were particularly shown to thicken and retract their processes, extend filopodia, proliferate and migrate, release factors and compounds influencing neuronal survival (such as proinflammatory cytokines, trophic factors, reactive oxygen species (ROS), etc.), and phagocytose pathogens, degenerating cells and debris, thus providing better understanding of their roles in orchestrating the inflammatory response [6]. These abilities as immune cells are also recruited during normal physiological conditions, where “surveillant” microglia further participate in the remodeling of neuronal circuits by their phagocytic elimination of synapses and their regulation of glutamatergic receptors maturation and synaptic transmission, among other previously unexpected roles [79], in addition to their crucial involvement in the phagocytic elimination of newborn cells in the context of adult neurogenesis [10].

Our review will discuss the emerging roles of microglia in adult hippocampal neurogenesis and their regulation by inflammation during chronic stress, aging, and neurodegenerative diseases, with a particular emphasis on their underlying molecular mechanisms and their functional consequences for learning and memory (Figure 1).

Figure 1: The effects of surveillant and inflammatory microglia on the adult hippocampal neurogenic cascade. During physiological conditions, surveillant microglia effectively phagocytose the excess of apoptotic newborn cells and may release antineurogenic factors such as TGF. This anti-inflammatory state is maintained by neuronal (tethered or released) fractalkine. Enriched environment drives microglia towards a phenotype supportive of neurogenesis, via the production of IGF-1. In contrast, inflammatory challenge triggered by LPS, irradiation, aging, or AD induces the production of proinflammatory cytokines such as IL-1, TNF, and IL-6 by microglia as well as resident astrocytes and infiltrating monocytes, neutrophils, and lymphocytes. These cytokines have profound detrimental effects on adult neurogenesis by reducing the proliferation, survival, integration, and differentiation of the newborn neurons and decreasing their recall during learning and memory paradigms.

  1. A Brief Overview of Adult Hippocampal Neurogenesis

Adult hippocampal neurogenesis is continuously maintained by the proliferation of neural stem cells located in the subgranular zone (SGZ) [1113]. These neuroprogenitors have been named “radial glia-like cells” (rNSCs), or type 1 cells, since they morphologically and functionally resemble the embryonic radial glia. They have also been defined as “quiescent neuroprogenitors” because only a small percentage of the population is actively dividing during normal physiological conditions. The lineage of these cells is frequently traced by using analogs of the nucleotide thymidine, such as bromodeoxyuridine (BrdU) which gets incorporated into the DNA of dividing cells during the S phase and can be detected by immunofluorescence. Alternatively, their lineage can be traced by labeling with fluorescent reporters which are delivered to dividing cells by retroviral vectors or expressed by specific cell type promoters via inducible transgenic mice (for a review of the methods commonly used to study adult neurogenesis, see [14]). The daughter cells of rNSCs, also called type 2 cells or amplifying neuroprogenitors (ANPs), rapidly expand their pool by proliferating before becoming postmitotic neuroblasts. Within a month, these neuroblasts differentiate and integrate as mature neurons into the hippocampal circuitry [15]. They however display unique electrophysiological characteristics during several months, being more excitable than mature neurons [16], and constitute a special cell population that is particularly inclined to undergo synaptic remodeling and activity-dependent plasticity [17].

These unique properties of the newborn neurons and the neurogenic cascade in general suggested that adult hippocampal neurogenesis could play an important role in hippocampal-dependent functions that require extensive neuroplasticity such as learning and memory. Indeed, activity-dependent plasticity and learning are long known for modulating adult neurogenesis in a complex, yet specific manner, with adult hippocampal neurogenesis being influenced by learning tasks which depend on the hippocampus [4445]. For instance, hippocampal-dependent learning paradigms were found to regulate the survival of newborn neurons, in a positive manner that depends on the timing between their birth and the phases of learning [4647]. Young (1.5–2 months old) newborn neurons were also shown to be preferentially activated during memory recall in a water maze task, compared to mature neurons, as determined by colabeling of BrdU with immediate early genes such as c-Fos and Arc, in which expression correlates with neuronal firing [48]. Nonetheless, it has only been in the last few years that loss-of-function and gain-of-function approaches with inducible transgenic mice were able to confirm that adult hippocampal neurogenesis is necessary for synaptic transmission and plasticity, including the induction of long-term potentiation (LTP) and long-term depression [49], as well as trace learning in conditioned protocols [50], memory retention in spatial learning tasks [5152], and encoding of overlapping input patterns, that is, pattern separation [53].

Adult hippocampal neurogenesis and its functional implications for learning and memory are however influenced negatively by a variety of conditions that are commonly associated with microglial activation and inflammation in the brain, such as chronic stress, aging, and neurodegenerative diseases, as we will review herein. Indeed, inflammation caused by irradiation produces a sustained inhibition of neurogenesis, notably by decreasing the proliferation and neuronal differentiation of the progenitors, and therefore, exposure to therapeutic doses of cranial irradiation has been widely used for modulating neurogenesis experimentally before the development of more specific approaches [54].

  1. Regulation of Adult Hippocampal Neurogenesis by Inflammation

Inflammation is a natural bodily response to damage or infection that is generally mediated by proinflammatory cytokines such as interleukin 1 beta (IL-1), interleukin 6 (IL-6), and tumour necrosis factor alpha (TNF), in addition to lipidic mediators such as prostaglandins and leukotrienes. Oftentimes, it is associated with an increased production of ROS, as well as nitric oxide (NO). Together, these proinflammatory mediators lead to an increase in local blood flow, adhesion, and extravasation of circulating monocytes, neutrophils, and lymphocytes [55]. In the brain, microglia are the main orchestrator of the neuroinflammatory response, but other resident cell types, including astrocytes, endothelial cells, mast cells, perivascular and meningeal macrophages, and even neurons, can produce proinflammatory mediators, though perhaps not to the same extent as microglia [56]. In addition, peripheral immune cells invading the CNS during inflammation can further produce proinflammatory mediators, but the respective contribution of microglia versus other cell types in the inflammatory response of the brain is poorly understood.

The harmful effects of inflammation are also widely determined by the actual levels of proinflammatory mediators released, rather than the occurrence or absence of an inflammatory response in itself. For instance, TNF regulates synaptic plasticity by potentiating the cell surface expression of AMPA glutamatergic receptors, thus resulting in a homeostatic scaling following prolonged blockage of neuronal activity during visual system development [57]. However, TNF also produces differential effects at higher concentrations,ranging from an inhibition of long-term potentiation to an enhancement of glutamate-mediated excitotoxicityin vitro [58]. Inflammation induced by chronic ventricular infusion of bacterial lipopolysaccharides (LPS; a main component of the outer membrane of Gram-negative bacteria), that is, the most widely used method for inducing an inflammatory challenge, also increases ex vivo the hippocampal levels of TNF and IL-1, thereby impairing novel place recognition, spatial learning, and memory formation, but all these cognitive deficits can be restored by pharmacological treatment with a TNF protein synthesis inhibitor, a novel analog of thalidomide, 3,6′-dithiothalidomide [59].

The impact of inflammation on adult hippocampal neurogenesis was originally discovered by Olle Lindvall and Theo Palmer’s groups in 2003, showing that systemic or intrahippocampal administration of LPS reduces the formation of newborn neurons in the adult hippocampus, an effect that is prevented by indomethacin, a nonsteroidal anti-inflammatory drug (NSAID) which inhibits the synthesis of proinflammatory prostaglandins [6061]. Similarly, inflammation can determine the increase in neurogenesis that is driven by seizures, a context in which neurogenesis can be prevented by LPS and increased by the anti-inflammatory antibiotic minocycline [60]. In these studies, hippocampal proliferation remained unaffected by LPS or minocycline and thus it is likely that inflammation targeted the survival of newborn cells [6061], as LPS is known to increase SGZ apoptosis [62]. Inflammation also has further downstream effects on the neurogenic cascade. For instance, LPS increases the number of thin dendritic spines and the expression of the excitatory synapses marker “postsynaptic density protein of 95kDa” (PSD95) in newborn neurons. LPS in addition increases the expression of GABAA receptors at early stages of synapse formation, leading to suggesting a possible imbalance of excitatory and inhibitory neurotransmission in these young neurons [63]. Finally, LPS also prevents the integration of newborn neurons into behaviourally relevant networks, including most notably their activation during spatial exploration, as determined by the percentage of BrdU cells colabeled with the immediate early gene Arc [64].

Importantly, none of these manipulations is specific to microglia and may directly or indirectly affect other brain cells involved in the inflammatory response of the brain. For instance, both LPS and minocycline affect astrocytic function in vitro and in vivo [6569]. Furthermore, LPS is known to drive infiltration of monocytes and neutrophils into the brain parenchyma [70]. Monocytes and neutrophils produce major proinflammatory mediators and could therefore act on the neurogenic cascade as well. The implication of microglia in LPS-induced decrease in neurogenesis is nonetheless supported in vivo by the negative correlation between the number of newborn neurons (BrdU+, NeuN+ cells) and the number of “activated” microglia (i.e., expressing ED1) [60]. ED1, also called CD68 or macrosialin, is a lysosomal protein which is overexpressed during inflammatory challenge. While the location of ED1 previously suggested its involvement in phagocytosis, its loss of function did not result in phagocytosis deficits and thus, its function still remains unknown (reviewed in [10]). The number of ED1-positive microglia also negatively correlates with neurogenesis during inflammation provoked by cranial irradiation [61]. While correlation does not involve causation, nor can pinpoint to the underlying mechanism, these experiments were the first to reveal a potential role for “activated” microglia in the regulation of adult hippocampal neurogenesis. More direct evidence of microglial mediation in LPS deleterious effects was obtained from in vitro experiments, as it was shown that conditioned media from LPS-challenged microglia contained IL-6, which in turn caused apoptosis of neuroblasts [61]. Nonetheless, astrocytes can also release IL-6 when stimulated with TNF or IL-1 [71] and chronic astrocytic release of IL-6 in transgenic mice reduced proliferation, survival, and differentiation of newborn cells, thus resulting in a net decrease in neurogenesis [72]. In summary, while the detrimental impact of inflammation on neurogenesis is well established, more work is needed to define the specific roles played by the various inflammatory cells populating the brain.

  1. Inflammation Associated with Chronic Stress

Across health and disease, the most prevalent condition that is associated with neuroinflammation is “chronic stress,” which commonly refers to the repeated or sustained inability to cope with stressful environmental, social, and psychological constraints. Chronic stress is characterized by an imbalanced secretion of glucocorticoids by the hypothalamic-pituitary-adrenal (HPA) axis (most notably cortisol in humans and corticosterone in rodents), which leads to an altered brain remodeling, massive loss of synapses, and compromised cognitive function [73]. In particular, an impairment of spatial learning, working memory, novelty seeking, and decision making has been associated with chronic stress [74]. Glucocorticoids are well known for their anti-inflammatory properties, as they interfere with NF-B-mediated cytokine transcription, ultimately delaying wound healing [75]. They are also potent anti-inflammatory mediators in vivo [76] and in purified microglia cultures [77]. Recently, repeated administration of high doses of glucocorticoids by intraperitoneal injection, to mimic their release by chronic stress, was also shown to induce a loss of dendritic spines in the motor cortex, while impairing learning of a motor task. A transcription-dependent pathway acting downstream of the glucocorticoid receptor GR was proposed [7879] but the particular cell types involved were not identified.

Microglia are considered to be a direct target of the glucocorticoids, as they were shown to express GR during normal physiological conditions in vivo [77]. In fact, transgenic mice lacking GR in microglia and macrophages show an increased production of proinflammatory mediators (including TNF and IL-1) and greater neuronal damage in response to an intraparenchymal injection of LPS, compared to wild-type mice [80]. In contrast, glucocorticoids are considered to be proinflammatory in the chronically stressed brain [81], where among other changes they can promote inflammation, oxidative stress, neurodegeneration, and microglial activation [82]. For example, repeated restraint stress induces microglial proliferation and morphological changes, including a hyperramification of their processes in the adult hippocampus following restraint stress [83], but a nearly complete loss of processes in the context of social defeat [84]. Prenatal restraint stress also causes an increase in the basal levels of TNF and IL-1, while increasing the proportion of microglia showing a reactive morphology in the adult hippocampus [85]. Similarly, social defeat leads to an enhanced response to the inflammatory challenge induced by intraperitoneal injection of LPS, including an increased production of TNF and IL-1, and expression of inducible NO synthase (iNOS) by microglia, accompanied by an increased infiltration of circulating monocytes [8486]. Therefore, microglia are a strong candidate for mediating some of the effects of stress on adult neurogenesis, as will be discussed below, in synergy with other types of inflammatory cells.

Chronic stress is well known for its negative effects on hippocampal neurogenesis (reviewed in [8788]), although not all stress paradigms are equally effective [89]. Several stress paradigms can decrease neuroprogenitors proliferation in the tree shrew [90] and in mice [9192], although this effect seems to be compensated by an increased survival of newborn neurons [92] and whether stress results in a net increase or decrease in neurogenesis remains controversial (reviewed in [8788]). The effects of stress on adult neurogenesis seem to be mediated at least partially by glucocorticoids, because mice lacking a single copy of the GR gene show behavioural symptoms of depression including learned helplessness, neuroendocrine alterations of the HPA axis, and impaired neurogenesis [93]. In parallel, chronic stress is associated with an increased inflammatory response, which may inhibit neurogenesis as well. For instance, serum levels of IL-1and IL-6 are significantly increased in depressed patients [94]. In mice, restraint stress leads to a widespread activation of NF-B in the hippocampus, including at the level of neuroprogenitors [95] and increased protein levels of IL-1 [96]. In addition to the direct role of glucocorticoids, IL-1 also seems to mediate some of the effects of mild chronic stress, because in vivo manipulations that block IL-1 (either pharmacologically or in null transgenic mice) prevent the anhedonic stress response and the antineurogenic effect of stress [9196]. Moreover, the corticoids and IL-1 pathways may regulate each other in a bidirectional manner because the administration of a GR antagonist can blunt the LPS-induced production of hippocampal IL-1 in stressed mice [97], whereas mice knockout for the IL-1 receptor (IL-1R1) fail to display the characteristic elevation of corticosterone induced by mild chronic stress [96]. Another stress-related cytokine, IL-6, induces depressive phenotypes and prevents the antidepressant actions of fluoxetine when administered to mice in vivo [98]. So far the effects of stress on neurogenesis via corticosteroids and inflammation have been assumed to be cell autonomous, as neuroprogenitors express both GR [99] and IL-1R1 [95]. The potential participation of microglia is yet to be determined, but there are some reports of a direct effect of stress on microglial activation. For instance, microglia acutely isolated from mice subjected to acute stress (by inescapable tail shock) showed a primed response to LPS challenge by producing higher levels of IL-1 mRNA ex vivo [100], and the specific loss of expression of GR in microglia leads to a blunted inflammatory response in vitro and to a decreased neuronal damage in vivo in response to LPS [80]. In stress paradigms, these enhanced responses of microglia to inflammatory challenges are similar to their age-related “priming” which has been associated with and is possibly due to an increased basal production of proinflammatory mediators. However, whether microglia express increased levels of IL-1 and other proinflammatory cytokines in response to stressful events is presently unclear [101]. It is thus possible that some of the antineurogenic effects of stress are exerted by means of microglial-dependent inflammation, but this hypothesis remains to be experimentally tested.

  1. Inflammation Associated with Aging and Neurodegenerative Diseases

Inflammation is also commonly associated with normal aging and neurodegenerative diseases and, therefore, could represent a putative underlying mechanism that explains their decrease in hippocampal neurogenesis. Nonetheless, inflammation is also associated with neurological diseases, such as epilepsy or stroke, where neurogenesis is thought to be increased, although the data from rodents and humans is somewhat conflictive [102]. Neurogenesis is well known to decline throughout adulthood and normal aging in rodents and humans [103104], but the decay is more pronounced and occurs later in life in mice than in humans [105]. The aging-associated decrease in neurogenesis has been shown to occur mainly as a consequence of exhaustion of the rNSC population which, after being recruited and activated, undergo three rounds of mitosis in average and then terminally differentiate into astrocytes [12106]. In addition, a reduced mitotic capacity of the neuroprogenitors could further contribute to decreasing neurogenesis [106], and moreover, an age-related increase in the levels of proinflammatory cytokines could also hinder neurogenesis in the aging brain. Serum levels of IL-1, IL-6, and TNF are elevated in elderly patients [107108]. Aged microglia express higher levels of these proinflammatory cytokines and show a greater response to LPS inflammatory challenge, that is, a “primed” response, than their younger counterparts [109]. The origin of this low-grade age-related inflammation (“inflamm-aging” [110]) remains unknown and may be related to both aging and damage to the surrounding neurons, as well as aging of the immune system per se.

At the cellular level, stress to the endoplasmic reticulum (ER) caused by various perturbations, such as nutrient depletion, disturbances in calcium or redox status, or increased levels of misfolded proteins, can induce a cell-autonomous inflammatory response to neurons. Stress to the ER, a multifunctional organelle which is involved in protein folding, lipid biosynthesis, and calcium storage triggers a homeostatic response mechanism named the unfolding protein response (UPR), aiming to clear the unfolded proteins in order to restore normal ER homeostasis [111]. However, if the ER stress cannot be resolved, the UPR also initiates inflammatory and apoptotic pathways via activation of the transcription factor NF-B which controls the expression of most proinflammatory cytokines [112]. In the brain, ER stress is often initiated by the formation of abnormal protein aggregates in several neurodegenerative diseases such as Alzheimer’s disease (AD), Parkinson’s disease (PD), amyotrophic lateral sclerosis (ALS), Huntington’s disease (HD), and prion-related disorders [113]. This neurodegeneration-associated ER stress is assumed to occur mostly in neurons, but there are some examples of microglial protein misfolding as well. For instance, both microglia and neurons overexpress CHOP (C/EBP homologous protein), a transcription factor which is activated during ER stress in human patients and mouse models of ALS [114]. Inflammation has been speculated to be a main negative contributor to the pathology of ALS [115], but a direct microglial involvement in mediating the inflammatory response to abnormal protein aggregation in ALS and other neurodegenerative conditions remains to be tested. Finally, ER stress has been linked to a variety of inflammatory conditions [116117], including chronic stress, diet-induced obesity, and drug abuse, as well as atherosclerosis and arthritis [118120]. During normal aging, a progressive decline in expression and activity of key ER molecular chaperones and folding enzymes could also compromise the adaptive response of the UPR, thereby contributing to the age-associated decline in cellular functions [118]. Therefore, aging is strongly associated with a chronic ER stress which leads to increased activation of NF-B [112]; however, the contribution of the different brain cell types to “inflamm-aging” is still poorly understood. The detrimental effects on neurogenesis of increased proinflammatory cytokines in the aging brain are not necessarily related to microglia, but also to stressed neurons. Furthermore, ER stress may also cause a cell-autonomous response in neural stem cells [121], although its impact on neurogenesis remains to be experimentally determined.

In addition, aging is accompanied by an increased level of mitochondrial oxidative stress, which in turn activates the “Inflammasome” [122], a group of multimeric proteins comprising the interleukin 1 converting enzyme (ICE, caspase 1) which serves to release the active form of the cytokine [123]. IL-1 may act directly on rNSCs (visualised by labeling with the Sox2 marker), as they express IL-1R1 in the adult hippocampus [91]. Treatment with IL-1 decreases hippocampal proliferation in young mice [91] and pharmacological inhibition of ICE partially restores the number of newborn neurons in aged mice without significantly affecting their differentiation rate [124]. Transgenic IL-1 overexpression results in chronic inflammation and depletion of doublecortin-labeled neuroblasts, thus mimicking the aging-associated depletion of neurogenesis [125]. The actual mechanism of action of IL-1 on neurogenesis in aged mice, including decreased proliferation of rNSCs/ANPs and survival of newborn neurons, remains undetermined. Microglia are a main source of IL-1in the aging brain, but the hypothesis that microglia-derived IL-1 is responsible for depleting neurogenesis in the aging brain remains to be directly tested.

The regulation of neurogenesis by IL-1 in the aging brain has been further linked to the activity of another cytokine, the chemokine fractalkine, or CX3CL1. Fractalkine has soluble and membrane-tethered forms and is exclusively expressed by neurons, while the fractalkine receptor (CX3CR1) is expressed in the brain by microglia alone [126]. This module forms a unique neuron-microglia signalling unit that controls the extent of microglial inflammation in several neurodegenerative conditions including PD, ALS [127], or AD [128]. In fact, CX3CR1 blocking antibodies increase the production of hippocampal IL-1 when administered to young adult rats [129]. Importantly, chronic treatment with fractalkine increases hippocampal proliferation and the number of neuroblasts in aged (22 months old) but not young (3 months old) or middle-aged rats (12 months old), whereas an antagonists of CX3CR1 has the opposite effects in young, but not in middle-aged nor old rats [129]. Since fractalkine expression is decreased during aging [129], a reduced neuron-microglia signalling might be releasing the brake on microglial contribution to inflammatory responses, although increased levels of fractalkine were instead reported in aged rat hippocampus by other studies [68]. Additional insights into the role of fractalkine signalling come from knock-in mice in which the endogenous CX3CR1 locus is replaced by the fluorescent reporter GFP [126]. The initial studies suggested that  (i.e., ) mice have no significant differences in brain development and functions [130], but more systematic investigations recently revealed a long list of hippocampal-dependent changes in young (3 months old)  and  mice compared to wild-type mice. These changes notably included decreased neuroprogenitors proliferation and neuroblasts number, impaired LTP, performance in contextual fear conditioning and water maze spatial learning and memory, and, importantly, increased IL-1 protein levels [131]. The signalling pathway of fractalkine-IL-1 is functionally relevant, because IL-1R1 antagonists rescued LTP and cognitive function in  mice [131]. In sum, even though neuronal fractalkine seems to be sufficient for restraining the inflammatory activity of microglia in young rats, its downregulation during aging could activate the microglial inflammatory response and thereby subsequently reduce the proliferation of remaining neuroprogenitors.

In AD, inflammatory cytokines such as IL-1 are overexpressed in the microglia associated with the amyloid beta (A) plaques of postmortem samples [132] and in transgenic mice modeling the disease [133]. The loss of synapses (from hippocampus to frontal cortex) is one of the main pathological substrates in this disease, but adult neurogenesis is also severely reduced in most mouse models of AD, possibly due to a decreased proliferation of neuroprogenitors and a decreased survival of newborn cells, even though the putative changes in the neurogenic cascade in postmortem samples remain controversial (reviewed in [102]). This lack of agreement is possibly explained by the fact that the vast majority of AD cases have a late onset over 65 years of age, when little neurogenesis remains. In contrast, in most transgenic AD mouse models, the Aaccumulation, cognitive deficits, and changes in neurogenesis are already detectable in young animals (2-3 months old). The study of AD is further hindered by the difficulty in comparing the time course and pathology across different mouse models. For instance, early treatment with minocycline can improve cognition and reduce A burden in mice expressing the human amyloid precursor protein (APP) [134]. In contrast, in mice expressing APP and a mutated form of presenilin 1 (PS1), which is part of the  secretase pathway that cleaves A, inflammation is reduced without any detectable changes in A plaques deposition [135]. Concomitantly with a decrease in tissue inflammatory cytokines and number of microglial cells, minocycline restores neurogenesis and hippocampus-dependent memory deficits in these APP/PS1 mice [135], indirectly suggesting that cognitive decay in AD may be at least in part related to a detrimental effect of inflammation on hippocampal neurogenesis. Direct evidence that neurogenesis is associated with the cognitive performance in AD is still lacking. Further research is also necessary to determine the neurogenic targets of AD-related inflammation. One central open question for future therapies aiming at increasing neurogenesis and cognition in AD is whether neuroprogenitors are spared or whether their age-induced loss becomes accelerated. Rather than increasing the proliferation and neurogenic output of the few rNSCs remaining in an old AD brain, it may be more relevant to develop strategies that prevent the age-related loss of neuroprogenitors in presymptomatic patients.

In summary, inflammation associated with a wide variety of experimental models of disease produces strong detrimental effects on hippocampal neurogenesis. These effects on human neurogenesis are however not so well described and, in vitro, IL-1 increases the proliferation of hippocampal embryonic neuroprogenitors but decreases their differentiation into neurons [136]. Novel methods to assess hippocampal neurogenesis in the living human brain, from metabolomics of neuroprogenitors to hippocampal blood brain volume (reviewed in [102]), will help to determine the contribution of inflammation to adult neurogenesis in the healthy and diseased human brain during aging.

  1. Normal Physiological Conditions

In the healthy mature brain, microglia are an essential component of the neurogenic SGZ niche, where they physically intermingle with neuroprogenitors, neuroblasts, and newborn neurons [62]. Here, surveillant microglia effectively and rapidly phagocytose the excess of newborn cells undergoing apoptosis [62]. Importantly, microglial phagocytosis in the adult SGZ is not disturbed by inflammation associated with aging or by LPS challenge, as the phagocytic index (i.e., the proportion of apoptotic cells completely engulfed by microglia) is maintained over 90% in these conditions [62]. Nonetheless, the consequences of microglial phagocytosis on adult hippocampal neurogenesis remain elusive. Treatment of mice with annexin V, which binds to the phosphatidylserine (PS) receptor and prevents the recognition of PS on the surface of apoptotic cells, presumably blocking phagocytosis, increases the number of apoptotic cells in the SGZ [40]. Concomitantly, annexin V reduces neurogenesis by decreasing the survival of neuroblasts without affecting neuroprogenitors proliferation [40]. Similar results were obtained in transgenic mice knock-out for ELMO1, a cytoplasm protein which promotes the internalization of apoptotic cells, although the effects on neurogenesis were ascribed to a decreased phagocytic activity of neuroblasts [40]. The actual phagocytic target of the neuroblasts remains undetermined, but the newborn apoptotic cells in the adult SGZ are exclusively phagocytosed by microglia, at least in physiological conditions [62]. Nevertheless, none of the above manipulations has specifically tested the role of microglial phagocytosis in hippocampal-dependent learning and memory and thus, the functional impact of microglial phagocytosis in adult neurogenic niches during normal physiological conditions remains to be elucidated.

Microglial phagocytosis of apoptotic cells is actively anti-inflammatory, at least in vitro, and thus it has been hypothesized that anti-inflammatory cytokines produced by phagocytic microglia may further regulate neurogenesis [10]. For instance, transforming growth factor beta (TGF), which is produced by phagocytic microglia in vitro [137], inhibits the proliferation of SGZ neuroprogenitors [138]. Microglia are further able to produce proneurogenic factors in vitro [139]. When primed with cytokines associated with T helper cells such as interleukin 4 (IL-4) or low doses of interferon gamma (IFN), cultured microglia support neurogenesis and oligodendrogenesis through decreased production of TNF and increased production of insulin-like growth factor 1 (IGF-1) [139], an inducer of neuroprogenitor proliferation [26]. A list of potential factors produced by microglia and known to act on neuroprogenitor proliferation can be found in Table 1. In addition, recent observations suggest that neuroprogenitor cells may not only regulate their own environment, but also influence microglial functions. For instance, vascular endothelial growth factor (VEGF) produced by cultured neuroprecursor cells directly affects microglial proliferation, migration, and phagocytosis [20]. More potential factors produced by neuroprogenitors shown to be influencing microglial activity and function can be found in Table 2. However, it has to be taken into account that most of these observations were obtained in culture and that further research is needed in order to elucidate whether those factors are also secreted and have the same regulatory responses in vivo.

Table 1: Summary of factors secreted by microglia and the potential effect they have on neuroprogenitors in vitro.
Microglia secreted
Reference Modulation of neural progenitor cells Reference
BDNF [18] Differentiation [19]
EGF [20] Survival, expansion, proliferation, differentiation [21]
FGF [22] Survival and expansion [23]
GDNF [24] Survival, migration, and differentiation [25]
IGF-1 [21] Proliferation [26]
IL-1 [27] Reduction in migration [27]
IL-6 [28] Inhibition of neurogenesis [29]
IL-7 [20] Differentiation [30]
IL-11 [20] Differentiation [30]
NT-4 [24] Differentiation [31]
PDGF [32] Expansion and differentiation [33]
TGF [34] Inhibition of proliferation [19]


Table 1: Summary of factors secreted by microglia and the potential effect they have on neuroprogenitors in vitro.



Table 2: Summary of factors secreted by neuroprogenitors and the potential effect they have on microglia in vitro.

NPC secreted factors Reference Modulation of microglia Reference
BDNF [18] Proliferation and induction of phagocytic activity [35]
Haptoglobin [24] Neuroprotection [36]
IL-1 [37] Intracellular Ca+2 elevation and proliferation [22]
IL-6 [37] Increase in proliferation [38]
M-CSF [20] Mitogen [39]
NGF [40] Decrease in LPS-induced NO [41]
TGF [37] Inhibition of TNF secretion [42]
TNF [37] Upregulation of IL-10 secretion [43]
VEGF [20] Induction of chemotaxis and proliferation [20]

Table 2: Summary of factors secreted by neuroprogenitors and the potential effect they have on microglia in vitro.

In addition, microglial capacity to remodel and eliminate synaptic structures during normal physiological conditions has suggested that microglia could also control the synaptic integration of the newborn neurons generated during adult hippocampal neurogenesis [140]. Three main mechanisms were proposed: (1) the phagocytic elimination of nonapoptotic axon terminals and dendritic spines, (2) the proteolytic remodeling of the perisynaptic environment, and (3) the concomitant structural remodeling of dendritic spines [7140]. Indeed, microglial contacts with synaptic elements are frequently observed in the cortex during normal physiological conditions, sometimes accompanied by their engulfment and phagocytic elimination [141143], as in the developing retinogeniculate system [144]. Microglial cells are distinctively surrounded by pockets of extracellular space, contrarily to all the other cellular elements [142], suggesting that microglia could remodel the volume and geometry of the extracellular space, and thus the concentration of various ions, neurotransmitters, and signalling molecules in the synaptic environment. Whether microglia create the pockets of extracellular space themselves or not remains unknown, but these pockets could result from microglial release of extracellular proteases such as metalloproteinases and cathepsins [145], which are well known for influencing the formation, structural remodeling, and elimination of dendritic spines in situ and also experience-dependent plasticity in vivo [7146]. More recently, microglial phagocytosis of synaptic components was also observed in the developing hippocampus, in the unique time window of synaptogenesis, a process which is notably regulated by fractalkine-CX3CR1 signalling [147]. Therefore, the attractive hypothesis that microglial sculpts the circuitry of newborn cells in the adult hippocampus deserves further attention.

Lastly, microglia were also involved in increasing adult hippocampal neurogenesis in the enriched environment (EE) experimental paradigm. EE is a paradigm mimicking some features of the normal living circumstances of wild animals, as it gives them access to social interactions, toys, running wheels, and edible treats. EE has long been known to enhance neurogenesis by acting on newborn cells survival, resulting ultimately in an enlargement of the dentate gyrus [148]. Functionally, these changes are accompanied by enhanced spatial learning and memory formation with the water maze paradigm [149]. Similar increases in neurogenesis are obtained by subjecting mice to voluntary running paradigms, although in this case the effect is mediated by increased neuroprogenitor proliferation [150]. During inflammatory conditions, EE is antiapoptotic and neuroprotective [151] and it limits the hippocampal response to LPS challenge by decreasing the expression of several cytokines and chemokines, including IL1- and TNF [152]. In fact, EE is believed to counteract the inflammatory environment and rescue the decreased number of neuroblasts in mice compared to wild-type mice [153]. The effects of EE are independent of the IL-1 signalling pathway, as it increases neurogenesis in mice that are null for IL-1R1 [154]. EE also induces microglial proliferation and expression of the proneurogenic IGF-1 [155], but the full phenotype of microglia in EE compared to standard housing and its impact on the neurogenic cascade remains to be determined.

The mechanisms behind the anti-inflammatory actions of EE are unknown, but they were suggested to involve microglial interactions with T lymphocytes through an increased expression of the major histocompatibility complex of class II (MHC-II) during EE [155]. MHC-II is responsible for presenting the phagocytosed and degraded antigens to the antibodies expressed on the surface of a subtype of T lymphocytes (T helper or CD4+ cells), thus initiating their activation and production of antigen-specific antibodies. Severe combined immunodeficient (SCID) mice lacking either T and B lymphocytes or nude mice lacking only T cells have impaired proliferation and neurogenesis in normal and EE housing compared to wild-type mice [155], as well as impaired performance in the water maze [156]. Similarly, antibody-based depletion of T helper lymphocytes impairs basal and exercise-induced proliferation and neurogenesis [157]. Furthermore, a genetic study in heterogeneous stock mice, which descend from eight inbred progenitor strains, has found a significant positive correlation between genetic loci associated to hippocampal proliferation and to the proportion of CD4+ cells among blood CD3+ lymphocytes [158]. Additional experiments are needed to fully determine the possible interactions between microglia and T cells in neurogenesis, because, at least in normal physiological conditions, (1) T cell surveillance of the brain parenchyma is minimal, (2) microglia are poor antigen presenting cells, and (3) antigen presentation by means of MHC-II family of molecules is thought to occur outside the brain, that is, in the meninges and choroid plexus [159]. In fact, during voluntary exercise, there are no significant changes in T cell surveillance of the hippocampus, nor a direct interaction between T cells and microglia, nor any changes in the gene expression profile of microglia, including that of IGF-1, IL-1, and TNF [160]. The number of microglia is also inversely correlated with the number of hippocampal proliferating cells, rNSCs, and neuroblasts in aged (8 months) mice subjected to voluntary running, as well asin vitro cocultures of microglia and neuroprogenitors, which has been interpreted as resulting from an overall inhibitory effect of microglia on adult neurogenesis [161]. Even though EE is clearly a more complex environmental factor than voluntary running, further research is necessary to disregard nonspecific or indirect effects of genetic or antibody-based T cells depletion on microglia and other brain cell populations, including rNSCs. For instance, adoptive transfer of T helper cells treated with glatiramer acetate, a synthetic analog of myelin basic protein (MBP) approved for the treatment of multiple sclerosis, produces a bystander effect on resident astrocytes and microglia by increasing their expression of anti-inflammatory cytokines such as TGF[162]. Alternatively, it has been suggested that T cells may mediate an indirect effect on adult hippocampal neurogenesis by increasing the production of brain-derived neurotrophic factor (BDNF) [157], which is involved in the proneurogenic actions of EE [163]. Whether BDNF can counteract the detrimental effects of T cell depletion on neurogenesis remains unknown. Overall, the roles of microglia in EE and running-induced neurogenesis are unclear and have to be addressed with more precise experimental designs. In summary, surveillant microglia are part of the physical niche surrounding the neural stem cells and newborn neurons of the mature hippocampus, where they continuously phagocytose the excess of newborn cells. Microglia were also linked to the proneurogenic and anti-inflammatory effects of voluntary running and EE, but direct evidence is missing. The overall contribution of microglia to neurogenesis and learning and memory in normal physiological conditions remains largely unexplored at this early stage in the field.

  1. Conclusion

In light of these observations, microglia are now emerging as important effector cells during normal brain development and functions, including adult hippocampal neurogenesis. Microglia can exert a positive or negative influence on the proliferation, survival, or differentiation of newborn cells, depending on the inflammatory context. For instance, microglia can compromise the neurogenic cascade during chronic stress, aging, and neurodegenerative diseases, by their release of proinflammatory cytokines such as IL-1, IL-6, and TNF. A reduced fractalkine signalling between neurons and microglia could also be involved during normal aging. However, microglia are not necessarily the only cell type implicated because astrocytes, endothelial cells, mast cells, perivascular and meningeal macrophages, and to a lesser extent neurons and invading peripheral immune cells could further contribute by releasing proinflammatory mediators.

Additionally, microglia were shown to phagocytose the excess of newborn neurons undergoing apoptosis in the hippocampal neurogenic niche during normal physiological conditions, while a similar role in the synaptic integration of newborn cells was also proposed in light of their capacity to phagocytose synaptic elements. Lastly, microglial interactions with T cells, leading to the release of anti-inflammatory cytokines, neurotrophic factors, and other proneurogenic mediators (notably during EE and voluntary running), could counteract the detrimental effects of inflammation on adult hippocampal neurogenesis and their functional implications for learning and memory.

However, further research is necessary to assess the relative contribution of microglia versus other types of resident and infiltrating inflammatory cells and to determine the nature of the effector cytokines and other inflammatory mediators involved, as well as their cellular and molecular targets in the neurogenic cascade. Such research will undoubtedly help to develop novel strategies aiming at protecting the neurogenic potential and ultimately its essential contribution to learning and memory.


AD: Alzheimer’s disease
ANPs: Amplifying neuroprogenitors
APP: Amyloid precursor protein
A: Amyloid beta
BDNF: Brain-derived neurotrophic factor
BrdU: 5-Bromo-2′-Deoxyuridine
CX3CL1: Fractalkine
CX3CR1: Fractalkine receptor
EAE: Experimental acute encephalomyelitis
EE: Enriched environment
EGF: Epidermal growth factor
FGFb: Basic fibroblast growth factor
GDNF: Glial cell line-derived neurotrophic factor
GFAP: Glial fibrillary acidic protein
GR: Glucocorticoid receptor
HPA: Hypothalamic-pituitary-adrenal axis
ICE: Interleukin 1 converting enzyme
IL-1: Interleukin 1 beta
IL-1R1: Interleukin 1 beta receptor
IL-4: Interleukin 4
IL-6: Interleukin 6
IL-7: Interleukin 7
IL-11: Interleukin 11
IFN: Interferon gamma
IGF-1: Insulin-like growth factor 1
iNOS: Inducible nitric oxide synthase
LPS: Bacterial lipopolysaccharides
LTP: Long term potentiation
M-CSF: Macrophage colony-stimulating factor
MBP: Myelin basic protein
MHC-II: Major histocompatibility complex class II
MOG: Myelin oligodendrocyte glycoprotein
NF-B: Nuclear factor kappa-light-chain-enhancer of activated B cells
NGF: Nerve growth factor
NO: Nitric oxide
NSAID: Nonsteroidal anti-inflammatory drug
NT-4: Neurotrophin-4
PDGF: Platelet-derived growth factor
PS: Phosphatidylserine
PS1: Presenilin 1
ROS: Radical oxygen species
SCID: Severe combined immunodeficiency
SGZ: Subgranular zone
TGF: Transforming growth factor beta
TNF: Tumor necrosis factor alpha
VEGF: Vascular endothelial growth factor.

Conflict of Interests

The authors declare that there is no conflict of interests regarding the publication of this paper.


This work was supported by grants from the Spanish Ministry of Economy and Competitiveness to Amanda Sierra (BFU2012-32089) and Juan M. Encinas (SAF2012-40085), from Basque Government (Saiotek S-PC 12UN014) and Ikerbasque start-up funds to Juan M. Encinas and Amanda Sierra, and from The Banting Research Foundation, the Scottish Rite Charitable Foundation of Canada, and start-up funds from Université Laval and Centre de recherche du CHU de Québec to Marie-Ève Tremblay.


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Nature Reviews Molecular Cell Biology 8, 519-529 (July 2007) | doi:10.1038/nrm2199

Signal integration in the endoplasmic reticulum unfolded protein response

David Ron & Peter Walter

The endoplasmic reticulum (ER) responds to the accumulation of unfolded proteins in its lumen (ER stress) by activating intracellular signal transduction pathways — cumulatively called the unfolded protein response (UPR). Together, at least three mechanistically distinct arms of the UPR regulate the expression of numerous genes that function within the secretory pathway but also affect broad aspects of cell fate and the metabolism of proteins, amino acids and lipids. The arms of the UPR are integrated to provide a response that remodels the secretory apparatus and aligns cellular physiology to the demands imposed by ER stress.


Figure 1: The unfolded protein response (UPR) signalling pathways.

FromThe impact of the endoplasmic reticulum protein-folding environment on cancer development

Nature Reviews Cancer 14, 581–597 (2014)

(UPR) signalling pathways

(UPR) signalling pathways

Upon endoplasmic reticulum (ER) stress, unfolded and misfolded proteins bind and sequester immunoglobulin heavy-chain binding protein (BIP), thereby activating the UPR. The UPR comprises three parallel signalling branches: PRKR-like ER kinase (PERK)–eukaryotic translation initiation factor 2α (eIF2α), inositol-requiring protein 1α (IRE1α)–X-box binding protein 1 (XBP1) and activating transcription factor 6α (ATF6α). The outcome of UPR activation increases protein folding, transport and ER-associated protein degradation (ERAD), while attenuating protein synthesis. If protein misfolding is not resolved, cells enter apoptosis. CHOP, C/EBP homologous protein; GADD34, growth arrest and DNA damage-inducible protein 34; JNK, JUN N-terminal kinase; P, phosphorylation; RIDD, regulated IRE1-dependent decay; ROS, reactive oxygen species; XBP1s, transcriptionally active XBP1; XBP1u, unspliced XBP1.

Figure 3: The unfolded protein response (UPR) and inflammation.

(UPR) and inflammation

(UPR) and inflammation

The three UPR pathways augment the production of reactive oxygen species (ROS) and activate nuclear factor-κB (NF-κB) and activator protein 1 (AP1) pathways, thereby leading to inflammation. NF-κB, which is a master transcriptional regulator of pro-inflammatory pathways, can be activated through binding to the inositol-requiring protein 1α (IRE1α)–TNF receptor-associated factor 2 (TRAF2) complex in response to endoplasmic reticulum (ER) stress, leading to recruitment of the IκB kinase (IKK), IκB phosphorylation (P) and degradation, and nuclear translocation of NF-κB196. Moreover, the IRE1α–TRAF2 complex can recruit apoptosis signal-regulating kinase 1 (ASK1) and activate JUN N-terminal kinase (JNK), increasing the expression of pro-inflammatory genes through enhanced AP1 activity197. The PRKR-like ER kinase (PERK)–eukaryotic translation initiation factor 2α (eIF2α) and activating transcription factor 6α (ATF6α) branches of the UPR activate NF-κB through different mechanisms. Engaging PERK–eIF2α signalling halts overall protein synthesis and increases the ratio of NF-κB to IκB, owing to the short half-life of IκB, thereby freeing NF-κB for nuclear translocation198199. ATF6α activation following exposure to the bacterial subtilase cytotoxin that cleaves immunoglobulin heavy-chain binding protein (BIP) leads to AKT phosphorylation and consequent NF-κB activation109200.



Figure 4

The cancer-supporting role of the unfolded protein response (UPR).

cancer-supporting role of the unfolded protein response

cancer-supporting role of the unfolded protein response

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Eppendorf Award for Young European Investigators

Curator: Larry H. Bernstein, MD, FCAP

Series E. 2; 8.11

The independent Eppendorf Award Jury chaired by Prof. Reinhard Jahn selected Dr. Thomas Wollert (Research Group Leader at the Max Planck Institute of Biochemistry in Martinsried, Germany) as the 2015 winner of the Eppendorf Award for Young European Investigators.

Thomas receives the €20,000 prize for his groundbreaking work in reconstituting complex intracellular membrane events in the test tube using artificial membranes and purified components. His experiments have paved the way for understanding key steps in autophagy, a fundamental process required for the clearance of damaged cell parts in all eukaryotic cells.

Listen to a podcast with Thomas Wollert and learn more about his work, and read excerpts from the interview in a Q&A feature article.

Presented in partnership with Nature The Eppendorf Award for Young European Investigators was established in 1995 to recognize outstanding work in biomedical science. It also provides the opportunity for European researchers to showcase their work and communicate their research to a scientific audience. Nature is pleased to partner with Eppendorf to promote the award and celebrate the winner’s work in print and online. Nature’s Julie Gould talks to the 2015 winner Thomas Wollert (Max Planck Institute of Biochemistry, Germany) about his work — which looks at the complex molecular process that cells use to remove their waste — and how it felt to win the award.
To listen to the full interview, visit:

About the Award Thomas Wollert is the twentieth recipient of the Eppendorf Award for Young European Investigators, which recognizes talented young individuals working in the field of biomedical research in Europe. The Eppendorf Award is presented in partnership with Nature. The winner is selected by an independent jury of scientists under the chairmanship of Reinhard Jahn, Director at the Max Planck Institute for Biophysical Chemistry in Göttingen, Germany. Nature and Eppendorf do not influence the selection. For more information see:

Julie Gould: Congratulations on being awarded this year’s prize. How did it feel when you found out that you had won?

Thomas Wollert: That came as a big surprise to me. It’s a great honor and it’s of course a major recognition of our work; not only my work, but also the work that my laboratory has done over the past five years. So this is very important to me.

JG: Tell us a little bit about the research you are working on.

TW: The cells in our bodies recycle almost everything — they do not waste much. The question in the past has been: how is this achieved? The process needs to be highly regulated. You don’t want to degrade something that you still need, but you do want to get rid of dangerous material that accumulates in the cell. We became interested in one pathway that is involved in transporting this sort of trash, or unwanted material, to recycling stations in the cell. We are particularly interested in how the molecular mechanism is driven.

JG: What sort of molecular trash are we talking about?

TW: Everything that needs to be degraded in a cell has to end up at a recycling station, one of which is called the lysosome. What ends up there is chemically degraded, and the building blocks are reused by the cell to build material. Proteins that become aggregated, big material or composite structures, and everything else in the cell cytoplasm (such as mitochondria) need to be transported to the lysosome. There is a specialized pathway to do that — this has been called autophagy for self-digestion. During autophagy, crescent-shaped membranes are formed, which expand and capture cytoplasmic components. These structures become autophagosomes, which are like entire organelles and are the containers that transport the trash to the lysosomes for degradation.

JG: How do these autophagosomes form in the cell?

TW: In yeast the system is fairly well understood. Small membrane vesicles are recruited and fuse to form the crescents haped autophagic precursor membrane. This membrane then surrounds and captures material, and, after sealing, the full autophagosome is formed and finally fuses with the lysosome. There are 40 different proteins in yeast that have been identified as those that have an essential function in autophagy — they are specific to the autophagy pathway. The question was, what are they doing with the membrane and what is their molecular function? And that was the major interest of my lab.

JG: What did you discover? TW:

We analysed two important steps in autophagy. The first is initiation and the second is expansion.

An autophagosome is built from small vesicles, which come together and fuse. This process is driven by one big complex called the Atg1-kinase complex. This complex is known to be involved in recruiting the donor vesicles that create the autophagosome. We recently published work on the expansion step. This is an interesting step that involves a small ubiquitin-like molecule, Atg8. The unique feature of this particular molecule is that it becomes covalently attached to autophagic precursor membranes. Many Atg8 molecules get conjugated to these membranes, so the question has been: why is there so much Atg8 on the membrane and what is its job there? To answer this, we analyzed the proteins independently of the complex cellular environment. We produced recombinant molecular machines that drive the formation of autophagosomes and analyzed their function in the test tube. The test-tube components include the protein subunits of these molecular machines and model membranes that serve as the platform for proteins to assemble into large complexes. What we realized — and what came as a surprise to us — was that the molecular machine that drives conjugation of Atg8 stays with Atg8 at the membrane, rather than leaving after conjugation. We predicted that something needs to happen, some bigger structure needs to form on the membrane to keep the conjugation machine there. Using high-resolution approaches, we observed that Atg8 forms together with its conjugation machine, a protein shell on membranes. It’s like a meshwork that sits on top of the membrane and stabilizes the forming autophagosome. Presumably.

JG: Why presumably?

TW: Because the details of how this expansion is driven by the scaffold is something that we are investigating.

JG: Will you be following this up over the next few years?

TW: Yes. This is an interesting question, but not an easy one to answer. We need to understand the direct relationship of how this really works in vivo.

JG: How does the autophagosome capture material from cells?

TW: The selection of cargo comes in two flavours. Under normal conditions, when the cell is happy, it only wants to degrade unwanted material or something damaged. It chooses these materials quite selectively. For example, it might only want to degrade dysfunctional mitochondria, the cell’s power plants. The membrane then wraps tightly around these structures. However, if a cell becomes stressed or starved, it can use autophagy to degrade anything that’s around. That means bulk cytoplasm without any selectivity. Imagine a big happy cell that is starved and goes on a low-value nutritional diet. The cell will shrink, but it survives. If nutritional conditions improve, it can grow again.

JG: What big impacts will this research have?

TW: The research focus at the moment is neurodegenerative disease and cancer. In certain neurodegenerative diseases, some proteins can accumulate in cells. There are a couple of diseases, such as Huntington’s disease, in which particular genetic modifications lead to alterations in proteins, which then tend to aggregate. In other diseases, such as Alzheimer’s disease, proteins also accumulate, and those protein oligomers, or aggregates, are toxic to the cell. In some neurodegenerative diseases, it has been observed that increasing autophagy is beneficial for cells, and thus patients, because increasing autophagy increases the removal of the toxic material. Neurodegenerative disease is usually not observed until the later stages, when this material has already accumulated. If you could remove this harmful material from cells, you could maybe rescue some neurons from dying. This is one application where you would really want to increase autophagy. In cancer, it has already been shown that combining chemotherapy with an inhibitor of autophagy is beneficial because autophagy just counteracts chemotherapy.

JG: What is it about this field that you find so interesting?

TW: What excites me the most is that you can use a minimal system, combining a few components and then trying to get them to work in a test tube. Our major goal, and our holy grail in this research, is to have the full autophagy pathway in a test tube, combining the autophagy components, step by step, to produce an autophagosome from small membranes, and to have some material wrapped in the autophagosome.

Award Winners

2015 Winner

In 2015 Eppendorf AG is presenting the Eppendorf Award for Young European Investigators for the 20th time. The independent Eppendorf Award Jury chaired by Prof. Reinhard Jahn selected Dr. Thomas Wollert (Research Group Leader Molecular Membrane and Organelle Biology at the Max Planck Institute of Biochemistry in Martinsried, Germany) as the 2015 winner of the Eppendorf Award for Young European Investigators. Thomas Wollert, born 1979, receives the €20,000 prize for his groundbreaking work in reconstituting complex intracellular membrane events in the test tube using artificial membranes and purified components. Thomas talks about his work in this Award Feature

The official prize ceremony took place at the EMBL Advanced Training Centre in Heidelberg, Germany, on June 25, 2015.

To hear an interview with prize winner Thomas, listen here.

2014 Winner

The independent Eppendorf Award Jury chaired by Prof. Reinhard Jahn selected Madeline Lancaster, Ph.D., of the Institute of Molecular Biotechnology of the Austrian Academy of Sciences, Vienna, Austria, as the 2014 winner of the Eppendorf Award for Young European Investigators. Madeline Lancaster, born 1982, receives the € 15,000 research prize for her work showing that complex neuronal tissues resembling early states of fetal human brain can be created in vitro from pluripotent stem cells. Madeline talks about her work in this Award Feature

To hear an interview with prize winner Madeline, listen here or watch the video from the award ceremony.

2013 Winner

The independent Eppendorf Award Jury chaired by Prof. Reinhard Jahn selected Ben Lehner, Ph.D., of the Centre de Regulació Genòmica, Barcelona, Spain, as the 2013 winner of the Eppendorf Award for Young European Investigators. Ben, born 1978, receives the € 15,000 research prize for his discoveries concerning the fundamental question why mutations in the genome result in variable phenotypes. Ben talks about his work in this Award Feature.

To hear an interview with prize winner Ben, listen here or watch the video from the award ceremony.

2012 Winner

The 2012 prize was awarded to Elizabeth Murchison, Ph.D. (Wellcome Trust Sanger Institute, Cambridge, United Kingdom) for her discoveries concerning a deadly cancer that is spreading among the endemic population of Tasmanian devils in Tasmania and threatening the survival of the species. Elizabeth talks about her work in this Award Feature.

To hear an interview with prize winner Elizabeth, listen here or watch the video from the award ceremony in Heidelberg.

2011 Winner

The 2011 Eppendorf Young European Investigator Award goes to Suzan Rooijakkers for her contribution to discovering how Staphylococcus aureus evades immune attack. Suzan talks about her work on this Award Feature.

To hear an interview with prize winner Suzan, listen here.

Listen here to the podcast from the award ceremony in Heidelberg.

2009 Winner

In 2009 the prize was awarded to Óscar Fernández-Capetillo, head of the Genomic Instability Group at the Spanish National Cancer Center. Read the highlights of his interview with Nature in this Award Feature.

Listen here to learn about the impact the Award had on his career.

2008 Winner

The 2008 prize was awarded to Dr. Simon Boulton of the London Research Institute. Read the highlights of his interview with Nature in this Award Feature.

Listen here to learn about the impact the Award had on his career.

2007 Winner

Dr Mónica Bettencourt-Dias is the 2007 winner of the Eppendorf Young European Investigator Award. Monica gives a personal account of her research and the Eppendorf Award in an Award Feature forNature.

Listen here to learn more about the impact the award had on her career.

2006 Winner

Dr Luca Scorrano won the award in 2006. Read more about his research on the Eppendorf Young Investigator website.

Listen here to learn more about Dr Scorrano’s work and the impact the award has had on his career.   2015   2014     2013    2012

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