extfieldsofstudy: Chemistry Medicine

text:
Super-resolution microscopy reveals a preformed NEMO lattice structure that is collapsed in incontinentia pigmenti

The NF-κB pathway has critical roles in cancer, immunity and inflammatory responses. Understanding the mechanism(s) by which mutations in genes involved in the pathway cause disease has provided valuable insight into its regulation, yet many aspects remain unexplained. Several lines of evidence have led to the hypothesis that the regulatory/sensor protein NEMO acts as a biological binary switch. This hypothesis depends on the formation of a higher-order structure, which has yet to be identified using traditional molecular techniques. Here we use super-resolution microscopy to reveal the existence of higher-order NEMO lattice structures dependent on the presence of polyubiquitin chains before NF-κB activation. Such structures may permit proximity-based trans-autophosphorylation, leading to cooperative activation of the signalling cascade. We further show that NF-κB activation results in modification of these structures. Finally, we demonstrate that these structures are abrogated in cells derived from incontinentia pigmenti patients.

T he nuclear factor-kB (NF-kB) signal transduction pathway plays important roles in mammalian immune and inflammatory responses to injury and infection, as well as cell proliferation, differentiation and survival 1 . Indeed, several genetic diseases result from the functional alteration of key components of NF-kB signalling 2 . Among these are the various rare X-linked human diseases, including incontinentia pigmenti (IP, OMIM # 308300) and immunodeficiency with or without anhydrotic ectodermal dysplasia (ID OMIM# 300291).
On exposure to various stresses or external stimuli, cell surface receptors activate the inhibitor of kB kinase (IKK) complex, permitting nuclear entry of NF-kB and transcription of hundreds of target genes 3 . This classical view of NF-kB signalling has invoked the notion of sequential activation via posttranslational and conformational modifications to individual proteins. However, a shift in our understanding of the critical fine-tuning of this pathway is emerging from increasing evidence of 'open-ended' higher-order structures 4 . The close proximity of regulatory proteins in these structures would enable spatiotemporal control of signal amplification enhancing efficiency of subsequent catalytic reactions. Such cooperativity would be greatly facilitated if a lattice-like structure pre-existed within the cell, before signal induction 5 .
Higher-order structures have been shown for the tumour necrosis factor receptor-associated factor 6, responsible for the Toll-like receptor/interleukin (IL)-1-dependent signal transduction of NF-kB (ref. 6). By elucidating the crystal structure of tumour necrosis factor receptor-associated factor 6, Wu and colleagues revealed the presence of a scaffolding network of oligomerized proteins at the plasma membrane. The authors proposed that such higher-order oligomerization facilitates the regulation of signal transduction by enhanced proximity-based activation and autoubiquitination.
A similar model has been hypothesized for the IKK complex 7 , which represents a critical checkpoint in the canonical activation of NF-kB. A key factor in the IKK complex is the NF-kB essential modulator protein (NEMO, also known as IKKg), responsible for the regulation of the catalytic IKK subunits 8,9 . Indeed, hints of a higher-order oligomeric structure emerged in a previous study labelling NEMO following IL-1 stimulation 10 . However, confocal microscopy revealed no evidence of a preformed higher-order structure. Structurally, NEMO consists of an elongated and flexible coiled-coil protein, which can form a stable dimer in solution and putatively polyubiquitin-mediated higher-order oligomers in vitro (trimer, tetramer and multimers) and in vivo via its coiled-coil domains [11][12][13] . Furthermore, the catalytic IKKb subunits from two different species were recently crystallized 14,15 and both were shown to form a stable dimer with a trimodular organization. Strikingly, crystallization studies demonstrated that different oligomers of IKK kinases can be formed in the asymmetric unit of the crystal lattice through specific contacts between kinase domains, the predominant assembly forming octameric (tetramer of IKK dimers), hexameric (trimer of IKK dimers) and dimeric structures. The existence of these higherorder oligomers, which are largely undetectable in solution, has been proposed to provide a possible mechanism of cooperativity for IKK activation via IKKb catalysed autophosphorylation 7 . Their existence would implicate the higher-order structure directly in NEMO function. Furthermore, the loss of this higher-order structure could be a key mechanism by which NEMO signalling could be compromised.
We and others have shown that hypomorphic mutations in NEMO that lead to impairment, but not abolition of NF-kB signalling, are associated with the less clinically severe anhydrotic ectodermal dysplasia syndrome, whereas amorphic NEMO mutations that completely abolish NF-kB activation are associated with IP in females and in utero lethality in males 16 . Most IP patients (70-80%) bear a complex genomic rearrangement, leading to the deletion of exons 4-10 (ref. 17). This recurrent NEMO rearrangement results in the synthesis of a truncated non-functional protein, which is devoid of its two carboxy-terminal ubiquitin binding domains (NOA/ubiquitinbinding domains in ABIN and NEMO (UBAN) and zinc finger (ZF)), but still able to interact with the IKKs. Other IP-associated mutations, in contrast, appear more subtle, such as the A323P missense mutation. Yet, this single base-pair change confers a severe defect to the NEMO protein, affecting both its ability to oligomerize and bind polyubiquitin 18 . An open question is whether these IP causing mutations compromise the putative higher-order structure of NEMO and thus interfere with the essential mechanism required for functional NEMO signalling.
With several lines of evidence pointing towards a functionally relevant higher-order structure of NEMO, it remains to be identified in cellulo, due in part to the limited ability of experimental techniques at our disposal.
The existence of a higher-order structure would be best established by directly visualizing its nanoarchitecture in cells. However, cryo-electron microscopy techniques involve complex sample manipulations that may result in the collapse of these fragile structures making the approach unsuitable for visualization. In light microscopes, the diffraction limit of light restricts the spatial resolution preventing the accurate observation of molecular assemblies below 200 nm. However, recent advances in super-resolution methods have allowed the classical limit to be breached and resolutions of 10-20 nm to be routinely achieved via fluorescent light microscopy 19 .
Here we use super-resolution microscopy to reveal the putative higher-order structure of NEMO in non-stimulated cells, both in live-cell and fixed conditions. We demonstrate that abrogation of non-covalent polyubiquitin chains or IKK:NEMO binding leads to the disruption of these higher-order structures. Surprisingly, the largest structures represent a very small fraction of total NEMO higher-order structures in cells, yet even a modest removal of this subset results in defective NEMO signalling. Furthermore, we provide quantitative evidence for the modification of these structures on NF-kB stimulation. Finally, we show that IP patient-derived cells lack such higher-order lattices, providing a molecular understanding of their pathology. Collectively, our data demonstrate the importance of higherorder oligomerization of NEMO in the NF-kB signalling response and emphasize how super-resolution microscopy can provide novel insights into the single-molecule mechanisms of genetic disorders.

Results
Super-resolution reveals a higher-order structure of NEMO. Biochemical and structural data reveal that NEMO forms an elongated dimer 20 , which can act as a regulatory scaffold to activate the catalytic IKK subunits in a polyubiquitin bindingdependent manner (Fig. 1a). X-ray crystallography data 14 from IKKb had further suggested that this higher-order NEMO-IKK platform would form a lattice organization in the order of 500 nm or greater, composed of B68 nm base units. We reasoned that given the resistance of the proposed higher-order structure to biochemical isolation, it would be best visualized directly using super-resolution localization microscopy (SRLM) techniques such as photoactivated localization microscopy (PALM) 21 and stochastic optical reconstruction microscopy (STORM) 22  pathogenesis 24 . Therefore, we believed this to be the best approach to investigate whether the NEMO-IKK unit is assembled into higher-order structures. For high-precision fixed cell super resolution, we opted for direct STORM (dSTORM) 25 allowing us to survey endogenous NEMO proteins minimizing possible structure anomalies when compared with overexpressed fluorophore-tagged proteins that may hinder the natural organization and interactions that form higher-order structures. For live-cell super resolution, we opted for a super-resolution optical fluctuation imaging (SOFI)-based method, enabling super resolution at low-intensity illuminations-orders of magnitude lower than comparable STORM methods-considerably reducing cell phototoxicity when compared with live dSTORM approaches, although providing a lesser resolution improvement (80-100 nm) and relying on fluorescent protein fusion constructs 26 . We used total internal reflection fluorescence (TIRF)-based illumination in tandem with SRLM to super-resolve the immunolabelled NEMO nanoarchitecture near the surface of the cell membrane. This approach allowed us to observe the presence of large lattice structures in the sub-membrane region of U2OS cells (Fig. 1b). Consistent with a model of NEMO-IKK assembly predicted from IKK X-ray crystallographic data, we observed lattice-like structures composed of round clusters of NEMO of 50-100 nm, connected to each other by elongated bridges also composed of NEMO spanning over 400 nm in maximum diameter. These structures could not be accurately resolved using conventional methods including highly sensitive TIRF approaches as well as maximum intensity projections in widefield (Fig. 1b).
To ascertain whether these structures visible in fixed cells represented bona fide structures in live cells, we performed SOFIbased live super resolution using NEMO-defective cells 17 transiently transfected with a GFP-NEMO fusion construct. Video acquisition shows that cells positive for green fluorescent protein (GFP) formed similar lattice structures (Supplementary Movie 1), to those observed with endogenous NEMO labelling in wild-type (WT) cells. In addition, SOFI high-speed acquisition indicated that these structures are highly dynamic.
We sought to confirm the presence of the lattice structure in primary cells. SRLM analysis revealed similar extensive lattice structures in human dermal fibroblasts (HDFs) approaching lengths of 1-3 mm (Fig. 1c). As with the U2OS cells, no distinguishable structures were observed in corresponding non-super-resolved TIRF or wide-field maximum intensity projection images (Fig. 1c).
To eliminate the possibility of a technical artefact, several control experiments were performed. An alternative antibody targeting NEMO revealed lattice structures of highly similar dimensions ( Supplementary Fig. 1a,b). Potential artefacts due to nonspecific staining were excluded by performing SRLM on cells transfected with a small interfering RNA (siRNA) targeting NEMO and on cells immunolabelled with the secondary antibody alone. No structures of significant size were observed ( Supplementary Fig. 1c,d). Further modifications to fixation and permeabilization showed no impact on the detected structures ( Supplementary Fig. 1e,f). Finally, SRLM on a control protein, Calnexin, revealed no lattice-like structure ( Supplementary  Fig. 1g). These data, together with similar observations in live super-resolution microscopy convincingly indicated that the identified lattice structures are not a technical artefact.
Inhibition of polyubiquitin chain binding abrogates lattices. In addition to the structural evidence of oligomerization of the IKK complex, further connections of oligomers may be facilitated by non-covalent binding via polyubiquitin chains. Several studies have revealed that the binding of K63 and Met1 (linear) polyubiquitin chains to two ubiquitin-binding domains, the NOA (also called UBAN) and ZF domains, located at the C-terminal extremity, are essential for NF-kB activation [27][28][29][30][31][32][33][34] (see Fig. 1a). Although NEMO oligomerization and its binding properties to ubiquitin oligomers have been shown to be essential in canonical NF-kB signalling 11,12,35,36 , no report has yet demonstrated the extent to which these oligomers may be linked to each other via polyubiquitin chains to form a preformed higher-order structure (see Fig. 1a).
As polyubiquitin chains are constitutively present in the cell (reviewed in ref. 37) and NEMO contains ubiquitin-binding domains, we theorized that NEMO would be able to co-opt these polyubiquitin chains to facilitate the assembly of the lattice structure before cytokine induction. To test this hypothesis we performed SRLM on U2OS cells transfected with plasmids expressing the de-ubiquitinases (DUBs) CYLD, (cylandromatosis DUB) or OTULIN (ovarian tumour DUB with linear linkage specificity; also known as FAM105B). Although CYLD has been shown to hydrolyse both K63 and linear polyubiquitin chains 38,39 , the specificity of OTULIN is solely targeted towards linear polyubiquitin chains 40 .
SRLM imaging of the CYLD-transfected cells revealed the removal of previously identified large lattice structures (defined as those over 400 nm in length, unless specified otherwise), although small clusters of NEMO were still apparent (Fig. 2a). As expected, we observed no structural alterations using conventional immunofluorescence at the same magnification ( Fig. 2a, lower micrographs, insets) compared with untransfected cells. SRLM revealed a similar pattern of smaller structures in U2OS cells expressing OTULIN (Fig. 2b). These data strongly suggest that preformed higher-order NEMO structures are dependent both on linear and K63 polyubiquitin chains. This result was further validated by expressing CYLD in HDFs, which also revealed a similar reduction of large lattice structures coupled with an inability to translocate p65 to the nucleus following IL-1 stimulation ( Supplementary Fig. 2).
Although every effort was made to remove bias in the observation of higher-order lattice structures between conditions, we sought to make our observations as quantitative as possible. We therefore developed an algorithm to perform unbiased analysis of the size of NEMO structures revealed by SRLM data (Methods; Fig. 2c). We compared the number of larger lattice structures (defined as those over 400 nm in length) per mm 2 of cell surface area in untreated cells and those transiently transfected with DUBs. Quantitative analysis revealed the number of large lattice structures is reduced fourfold in U2OS cells transfected with DUBs, validating our observations.
The global catalytic removal of polyubiquitin chains by DUBs may have additional effects on cellular machinery and therefore contribute to an indirect loss of lattice structure. We therefore sought to specifically obstruct the binding of polyubiquitin chains to NEMO alone. The peptide A-UBI specifically blocks the binding of polyubiquitin chains to the NOA domain of NEMO 41 . Treatment of HDFs with A-UBI showed a loss of lattice structures via super-resolution imaging (Fig. 2d). Quantitative analysis revealed a threefold reduction in the prevalence of larger lattice structures. This was consistent with our hypothesis that polyubiquitin chain binding to the NOA domain was essential for the assembly of higher-order NEMO structures.
Lattices provide a mechanism for proximity enhanced activation. It is well established that IL-1 induction of the NF-kB pathway requires the formation of a large signalosome at the receptor 42 . Indeed, we have recently demonstrated that these activating centres show recruitment of NEMO in the form of large aggregates 10 , which can be visualized in live cells using diffraction-limited spinning disk fluorescence microscopy ( Fig. 3a). Therefore, to further investigate the functional significance of the preformed NEMO lattice, we stimulated U2OS cells with IL-1 and performed SRLM. On image reconstruction, we were able to distinguish between preformed lattice structures of NEMO and the more condensed IL-1-induced NEMO aggregates (Fig. 3b), the latter being absent in untreated cells (Fig. 1b). IL-1-induced aggregates were observed by TIRF and shown to be in the range of 400 nm in diameter after SRLM reconstruction. Magnified images shown in Box I and Box II show examples of both an IL-1 aggregate (a) and a preformed lattice (b) within the sub-membrane region. The IL-1 aggregates appear more dense, thus explaining their detection in conventional fluorescence imaging, whereas the preformed NEMO lattice is detectable only by SRLM. This may suggest a mechanism by which a preformed lattice compacts on itself to form a dense IL-1-induced NEMO structure, a consequence of cooperative activation of the high number of enzymatic complexes present in close proximity. Indeed, we did observe a slight reduction in the number of large lattice structures per mm 2 following IL-1 induction (Fig. 3c).
Catalytic IKK subunits are essential to lattice structure. Our previous work has shown that IL-1-induced NEMO aggregates visible using conventional microscopy colocalize with activated IKK 10 . Here we sought to investigate the degree to which the catalytic IKK subunits contribute to the formation of the preformed NEMO lattice. The Nemo-binding domain (NBD) peptide is a well-established inhibitor of the NF-kB pathway, which acts by preventing the interaction between NEMO and catalytic IKK subunits 43 . SRLM of HDFs treated with NBD peptide showed a dramatic reduction in the number of large NEMO lattices (Fig. 4a). Similarly, we saw a threefold reduction in preformed NEMO lattices in MEFs lacking IKKa and b-subunits (Fig. 4b). Interestingly, both inactive and active forms of the catalytic IKK subunits formed lattices resembling that of the NEMO structure identified using SRLM (Supplementary Fig. 3  observe whether the IKK subunits were co-localized. Unsurprisingly, preformed NEMO lattices appeared to colocalize with IKKb but not the endoplasmic reticulum (ER)associated protein, Calnexin (Fig. 4c). Together, these data strongly support an essential role for the IKKb subunits in maintaining preformed NEMO lattices.
Lattice structures are absent in IP-derived patient cells. To investigate the clinical relevance of the NEMO structures, we performed super-resolution analysis on fibroblasts with NEMO mutations acquired from samples with clinically defined IP. Fibroblasts from IP GR contain a genomic rearrangement of exons 4-10 in NEMO, causative of the majority of IP cases 17 , resulting in a truncated protein 44 . Because of the concern of reduced binding sites available to the NEMO antibody in this mutant protein, we acquired fibroblasts from a second IP patient with a different disease-causing mutation in the NEMO gene. IP SS cells contain a novel splice site mutation in NEMO resulting in the deletion of exons 4-6, and a truncated protein ( Supplementary  Fig. 4a and ref. 45). Significantly, the truncated version of NEMO in IP SS cells retains the domains necessary to bind polyubiquitin chains, yet NEMO signalling is deficient in these patients. On the basis of structural modelling of the IP SS NEMO dimer, we hypothesized that this mutation confers a different orientation of the dimer of NEMO UBD domains relative to the IKKb amino-terminal binding site. This change in the spatial arrangement of UBDs is due to the modification of the left-handed superhelix of the coiled coil ( Supplementary  Fig. 4b). Importantly, the level of NEMO protein in cells from the IP SS line is as high or higher than in WT control lines as measured by western blotting (Supplementary Fig. 4c).
SRLM analysis on both these cell lines revealed the presence of small clusters of NEMO with no significant higher-order lattice structure observed (Fig. 5a,b) when compared with that of WT HDFs. Indeed, we observed an almost complete absence of preformed lattice structures over 1 mm in length (uniquely observed in HDFs compared with other cell types). Quantitative analysis confirmed our observation showing a substantial decrease in the frequency of large NEMO lattices in IP derived cells compared with their WT counterparts (Fig. 5c).
Apart from the reduced immunolabelling in IP GR -derived cells (expected from truncated proteins and thus reduced antibody binding), neither conventional immunofluorescence nor TIRF microscopy revealed any differences in NEMO staining (Fig. 5a,b). Indeed, the only structural difference observed between WT HDFs and IP-derived fibroblasts identifiable using conventional fluorescent microscopy was that of an inability of the IP-derived cells to form IL-1-induced NEMO aggregates ( Supplementary Fig. 5).
Quantitative analysis of NEMO lattice structures. Although we had provided some quantification of our SRLM data, we sought to further interrogate our algorithm data. We therefore assessed the prevalence of the larger NEMO lattices relative to the total number of structures in each cell type or condition, as well as the density of each structure in IL-1-stimulated cells.
Analysis of the maximum diameter of NEMO structures in U2OS cells across multiple conditions revealed smaller structures of 50-300 nm to be the predominant species ( Supplementary  Fig. 6a). Large lattice structures represented only 3.3% of the total number of NEMO structures in untreated U2OS cells (Fig. 6a). In contrast, cells treated with CYLD and OTULIN were revealed to have only 1.1 and 1.5% of NEMO lattices over 400 nm. This represents a three-and twofold decrease, respectively. Interestingly, we observed that the fold decrease in structures was more pronounced when comparing the largest higher-order order structures over 550 nm between conditions. In restricting our analysis to this size bin, we observed a ten-and fourfold decrease of CYLD and OTULIN-treated cells respectively compared with controls ( Supplementary Fig. 6a). This reduction was enhanced in CYLD-transfected cells compared with those transfected with OTULIN. Indeed, overexpression of CYLD led to a significant increase in the percentage of smaller structures of 20-50 nm, possibly representing NEMO reduced to its basic dimer unit. The more severe effect by CYLD is likely to be due to the combined elimination of K63 and linear chains, whereby K63 linkage represents the predominant species. Abrogation of polyubiquitin chains should reduce the lattice to its multimeric IKK/NEMO complex size of 128 nm. Although we did not observe a significant increase in lattices in the range of 100-300 nm in DUBtreated cells, this could well be explained by the transient nature of the experiment, whereby only the largest structures have been affected after 48 h. Importantly, the transient expression of either CYLD or OTULIN lead to the functional disruption of the NF-kB pathway. Thus, it appears that although large preformed higherorder lattice structures represent only a small proportion of total structures in the cell, they provide sufficient and essential higherorder platforms for a functional signal transduction response.
Analysis of NEMO structures in IL-1-stimulated cells compared with untreated cells revealed a small but general reduction in larger lattice structures, although this only achieved significance for a single bin size (500-550 nm; Supplementary  Fig. 6a and Fig. 6a). Given IL-1-induced NEMO aggregates can be observed with conventional microscopy (Fig. 3), we asked whether they were composed of a higher density of NEMO particles. Quantitative analysis revealed that there was no significant difference in the predominant species of low-density structures (o2 particle detections per nm 2 ; Supplementary  Fig. 6b). In contrast, high-intensity structures increased from 2 to 6% in IL-1-treated cells (Fig. 6b). This threefold increase was more pronounced in comparisons of higher particle density ( Supplementary Fig. 6b). Taken together, the modest reduction in large structures concurrent with the appearance of IL-1-induced NEMO aggregates may suggest a structural rearrangement of preformed lattices into IL-1 activation centres. Furthermore, the increased density of NEMO structures in IL-1-stimulated cells may lend further support to a model of proximity enhanced activation and cooperativity.
Having identified that transient expression of DUBs could significantly decrease the proportion of preformed NEMO lattices, we asked whether cells lacking catalytic IKK subunits showed a similar reduction. We found a threefold decrease in lattice prevalence in MEF IKK À / À cells (Fig. 6c). Interestingly, this corresponded with a robust and significant increase in base unit NEMO dimers (o100 nm) in IKK À / À cells ( Supplementary  Fig. 6c). Alhough we did not observe a similar increase in the prevalence of smaller structures post DUB treatment, this is probably explained by the prolonged absence of IKK subunits in these stable knockout cell lines rather than a transient removal of polyubiquitin chains.
Lastly, we compared the maximum diameter of structures from WT HDFs and IP-derived cells (Fig. 6d, expanded data in Supplementary Fig. 6d). Our analysis validated our observations of a significant reduction in preformed lattice structures in IP-derived cells, with a two-to threefold decrease in the prevalence of large structures compared with control cells. This effect was exacerbated in the more severe IP GR patient cells. As fibroblasts exhibit larger structures than U2OS cells, we further compared even larger higher-order lattices over 800 nm in length ( Supplementary Fig. 6d). These structures represented 0.6% of all structures in the cell, but only 0.05% in IP SS cells. We observed no structures of this size in IP GR cells. Although these numbers represent a small proportion of the total structures, those over 400 nm in size occupy over 50% of total NEMO structure area in WT HDFs, but only 20 and 13% in IP SS and IP GR cells, respectively ( Supplementary Fig. 6e). We further observed a significant increase in the prevalence of smaller structures of 200-300 nm in IP-derived cells, reinforcing the notion that these cells are unable to form NEMO lattices of significant size, but yield an increased proportion of NEMO conglomerates of a non-functional nature. The shift in the distribution of structure size was more severe in IP GR cells, although this is likely to be due to the more severe nature of the mutation. Although the increased proportion of small structures in IP patient cells within the range of 200-300 nm does not directly equate to the NEMO dimer 'base unit', these structures may represent non-functional conglomerates of 'sticky' NEMO protein.
Although they are still able to bind IKK subunits, this alone cannot facilitate significant lattice formation without polyubiquitin chains.
The combination of these data are summarized in the schematic representation in Fig. 6e. Under normal conditions, unstimulated cells contain preformed lattices extending from 400 nm to over 1 mm in length held together by delicate bridges o100 nm in width. Importantly, cells stimulated with IL-1 largely maintain these lattices but reveal highly dense smaller NEMO aggregates 200-400 nm in size, which can be visualized by conventional microscopy. In contrast, cells in which the interactions of NEMO with polyubiquitin chains or catalytic IKK subunits are perturbed form small non-functional NEMO aggregates. Importantly, the presence or absence of preformed lattices is only distinguishable by SRLM.

Discussion
In this report we use SRLM to demonstrate the existence and, indeed, extent of a preformed higher-order lattice structure of NEMO, as well as its functional significance in the context of a genetic disease (Fig. 6). Although such a higher-order structure has been hypothesized and inferred from biochemical data, to the best of our knowledge this is the first time it has been visualized in both live-cell and fixed-cell conditions, and therefore the strongest evidence that exist to date. Despite the size of the NEMO higher-order lattice surpassing the diffraction limit of light, the delicate NEMO bridges along which it is connected appear to be within the range of 20-100 nm (cell type dependent), thus obscuring their observation using diffraction-limited microscopy. Alternative techniques have been used to reveal similar nanoscale structures, including electron microscopy and PALM. However, to preserve the structural integrity of the delicate lattice within the context of endogenous NEMO expression levels, we believed a dSTORM strategy was best suited to this application, although a SOFI-based method was also used to demonstrate that the lattices could still be observed in live-cell conditions.
Notably, our results contribute significantly to the mechanistic understanding of how a regulatory protein can so efficiently and exquisitely react to external stimuli. We determined that a small number of preformed open-ended NEMO lattice structures linked by catalytic IKK subunits and polyubiquitin chain binding are sufficient to rapidly activate the NF-kB cascade. Although it has been shown that polyubiquitin chains bind NEMO on IL-1 stimulation 46 , our data strongly suggest both K63 and linear chains are necessary and exist within a preformed NEMO structure. An explanation for this inconsistency between our and previous experimental data may lie in the fact that this delicate non-covalent association of reticulated NEMO may be destroyed In untreated or healthy cell types, only SRLM reveals extensive NEMO lattice structures shown with an accompanying cross-sectional plot profiles and associated particle density as measured by metric analysis. In stimulated cells, conventional microscopy reveals punctate NEMO aggregates, which can be distinguished from preformed lattice structures in SRLM. Cells unable to bind catalytic IKK subunits and polyubiquitin chains cannot form extensive lattice structures, nor high-density structures. Each representative image is associated with its plot profile, indicating it's maximum length on the x axis as shown by a dashed red or blue line across the structure in question. Scale bar, 500 nm. n is the number of whole cells analysed per condition. Error bars are s.e.m. *P-value of o0.05 after a two-tailed t-test.
by insensitive biochemical techniques. Interestingly, the proportion of preformed structures varied across different cell types, although this may be attributed to the well-documented variation in the concentration of polyubiquitin chains between cell types 47 . We observed that the appearance of a few highly dense NEMO aggregates in IL-1-stimulated cells correlated with a very small reduction in open-ended NEMO lattices. Although we cannot exclude additional protein factors, the data presented here supports a mechanism by which a small number of lattices are primed to swiftly respond to external stimuli and 'fold-in' on each other to create compact NEMO-IKK signalosomes leading to a proximity-enhanced response. Such a mechanism would require a highly dynamic structure. Both our live-cell data and a previous report 20 provide strong experimental evidence to show the highly dynamic nature of NEMO oligomers. Indeed, this may further explain the modest reduction (Fig. 6a) in open-ended lattices in IL-1-treated cells, in which we would expect to observe some latency in the reformation of the lattice. Instead, a rapid and highly dynamic feedback mechanism is implicated in the reformation of the NEMO lattice structure on IL-1 induction. It would therefore be highly valuable to further investigate the nature of this dynamic interaction between inactive and activated cellular states with the advancement of such technologies in a broader study. In particular, NEMO ubiquitination-which corresponds to a covalent attachment of non-degradative polyubiquitin chains (linear and K63) to NEMO-could considerably enhance the conversion of preformed open-ended NEMO lattice structures into activating centres, where specific ubiquitin ligases are presumably present. Importantly, the small proportion of higher-order NEMO lattices and subsequent IL-1-induced signalosomes further provides a mechanism by which the signalling response could 'ignore' biological noise. This property, in conjunction with an ability to mediate proximity-enhanced activation, would effectively facilitate a binary biological switch.
The importance of such a higher-order lattice of NEMO is apparent from our data showing the near elimination of these structures in IP-derived cells. At first glance, the NEMO mutation in IP SS cells confers no apparent detriment to the function of the protein, apart from a reduction in length. Importantly, the domains necessary for IKKb and polyubiquitin chain binding are still present in these cells. Thus, NEMO should be able to facilitate activation of the signalling cascade. Yet, our analysis of structural data ( Supplementary Fig. 4) indicates that the IP SS mutation results in the ubiquitin-binding domains being misconfigured, leading to incorrectly positioned polyubiquitin chains, obstructing their ability to form a higherorder lattice structure. Therefore, in demonstrating that cells with this mutation lack higher-order structures before stimulation, we show the extent to which such accurate structural oligomerization and cooperativity is required for NF-kB signalling. Indeed, our data provide evidence that despite the broad spectrum of IP causing mutations, the common molecular cause for disease may be an inability to form higher-order lattices of significant size and number. These structures permit the signal transduction cascade through cooperativity, to achieve rapid Hill function responses once the appropriate threshold is achieved.
Higher-order structures constitute a very small proportion of total structure number, yet are absent in IP patient-derived cells, which lack a functional NF-kB response. This suggests that only a small number of large higher-order structures are necessary to efficiently regulate reliable signal transmission and interpretation by balancing signal intensity, while acting as a noise filtering mechanism. This concept bolsters previous observations where it has been shown that NF-kB signalling is activated in a digital manner, rapidly reaching a stimulus saturation point, exemplified in rapid Hill function responses 48 .
Although there exist simple methods for diagnosis of IP, the demonstration here that IP patients harbouring different mutations have a common molecular defect in the higher-order structure of NEMO protein may bypass the mutational screening and characterization required for identifying NEMO mutations. It is possible to speculate that this method may be useful for prenatal diagnosis in families in which IP is suspected but not fully confirmed in males but also, as evident by our study, in females in which skewed X-inactivation has not occurred. In our analysis we were able to show a statistical difference with only six cells from different clinical samples, suggesting the isolation and limited expansion of amniocytes may be useful in augmenting such a diagnosis. The expansion of such super-resolution techniques may therefore provide not only invaluable mechanistic insight into disease but also facilitate the genetic diagnosis of a clinically severe disorder. and 4BWN, residues 257-344) and ZF (PDB ID: 2JVX, residues 392-419). All regular heptad phases (abcdefg) n within the predicted fragment of NEMO (residues 49-364), as well as coiled-coil discontinuities such as stutters (deletion of three residues) and stammers (deletion of four residues), were correctly predicted using PAIRCOIL2 when compared with the heptad phases identified from available crystal structures by the coiled-coil analysis programme SOCKET 51 . This includes, for instance, one stutter and one stammer at positions 103 and 295, respectively. Two other highly predictable discontinuities at position 203 (stutter) and 256 (deletion of two residues) were taken into account to accommodate the heptad phases at the KD/HLX2 and HLX2/CC2-LZ junctions. A first model was built using O 52 by assembly of crystal and NMR NEMO fragments with the following modelled regions: 110-192 (CC1, 82 aa), 250-256 (7 aa), 345-364 (20 aa) and 365-391 (27 aa). The long WT CC1 (110-192) and the short mutant CC1 (110-133) fragments were generated using CCBuilder 53 with a radius of 5.07 Å, a superhelical pitch of 225.8 Å and an interface angle of 26.42°. The fragment building corresponding to the disordered region between CC2-LZ and ZF domains (residues 365-391) was based on the circular dichroism (CD) spectra analyses of the 248-344, 248-419 and 392-419 constructs of human NEMO (CD spectra available upon request). This disordered fragment was manually generated and refined from initial atomic coordinates derived from a I-TASSER-generated model based on threading algorithm 53 . The NEMO:IKKb:K63-di-Ub complexes (WT and Mu) were built by homology modelling using as structural templates the crystal structure of the mouse CC2-LZ in complex with a K63 di-Ub chain (PDB ID: 3JSV 54 ) or the modelled NEMO ZF in complex with a K63 di-Ub chain 55 . This structural modelling of NEMO is consistent with a recent in vitro EPR study showing that NEMO forms a parallel coiled-coil dimer in solution 20 . The incomplete crystal structure of the human IKKb (residues 16-665, PDB ID:4E3C) and its short C-terminal fragment bound to the NEMO N terminus (NDB, residues S705-Q743, PDB ID:3BRV) were previously described 15,56 . PyMOL (version 1.7, Schrödinger) was used for molecular display and structural figures.
Cell culture and transfection conditions. All patient material was gathered, after receiving informed consent from participants, under protocols approved by the Declaration of Helsinki, which was approved by the ethics committee at Hôpital Necker. U2OS cells and WT fibroblasts (WT HDFs) were obtained from Lonza (USA). WT MEFS were obtained from ATCC (USA). MEF IKK À / À cells were a kind gift of I.M. Verma (The Salk Institute Institute for Biological studies, La Jolla, CA). All cells were grown in complete RPMI medium (Gibco) containing 10% fetal bovine serum (PAA) and were screened for mycoplasma before experimental use. Cells were authenticated by western blotting where appropriate. Plasmids and siRNA were transfected into U2OS cells using Fugene HD (Roche) and into fibroblasts using the Invitrogen Neon Transfection system. Live-cell experiments were performed as previously described 10 using IP GR NEMO defective cells. Transiently transfected cells were imaged 48 h after transfection. U2OS cells and fibroblasts were stimulated with 10 ng ml À 1 IL-1b (H2691, Sigma) for 5 and 15 min, respectively, before fixation and imaging. The peptides A-UBI (catalogue number 481418, Calbiochem) and NBD (480025, Calbiochem) were used at a final concentration of 20 mM on WT HDFs. Cells were incubated with aforementioned peptides for 2 h, followed by treatment with IL-1, before fixation.
Western blottings. Cells (10 6 ) were lysed in 200 ml of lysis buffer. After centrifugation at 10,000 g. Ten microlitres of supernatant corresponding to 10 5 cells were loaded onto a 12% SDS-PAGE, followed by western blotting using a monoclonal antibody against NEMO. The human nucleoside diphosphate kinase B was used as a loading control.
Sample preparation for SRLM. Glass coverslips were washed three times in optical grade acetone, methanol and deionized water. Coverslips were then sonicated in 0.1 M KOH for 45 min, rinsed and further sonicated in deionized water. Cells were resuspended in 100% ethanol and exposed to ultraviolet radiation. Cells were fixed with 3.7% stabilized formaldehyde for 10 min at room temperature and then permeabilized with 70% ethanol. After blocking with 1% BSA-Tween for 30 min, the samples were incubated with primary antibodies at 4°C overnight, washed three times with 1% BSA and incubated with secondary antibodies for 1 h at room temperature. To maximize the number of detected molecules, care was taken to minimize photobleaching: cells were embedded in oxygen scavenger buffer as previously described 25 , imaged on the same day of labelling and kept in the dark until imaged. Tetraspeck beads (200 nm diameter; Invitrogen) were mounted with the sample as fiduciary landmarks and later tracked for computational drift correction.
Image acquisition. Samples were imaged on a custom-built PALM/STORM microscope as described before 57 . For each coverslip, preselected fields of view were chosen typically containing one cell. We developed the automated acquisition sequence based on the open-source Micro-Manager microscopy acquisition software 58 . First, the field of view was centred to include as much of the cytosol as possible, to maximize the area acquired over the flatter cell membrane. Cells were then imaged by TIRF illumination with a 488 nm, 568 nm and/ or 647 nm laser. We then performed a SRLM image-stream acquisition for the 647 channel (635 nm laser-excitation at 1.7 kW cm À 2 , 662-690 nm emission) or the 568 channel (561 nm laser excitation at 2.4 kW cm À 2 , 589-625 nm emission), each composed of 20,000 images acquired at 33 ms intervals. Dual dSTORM images were generated by first performing an acquisition as described above in the 647 channel, followed by the 568 channel. Imaging parameters were set using Micro-Manager running on a desktop PC. Laser control was achieved with custom software 57 . Before particle detection and localization, fiduciary beads were identified and marked, typically 1-5. Sample drift was then calculated by tracking the group displacement of selected beads throughout the acquired image sequence. After particle detection and localization, the position of each particle was readjusted by subtracting the drift identified at the corresponding time point.
The diffraction limited z-stack acquisition was performed at 300 nm z-steps encompassing the size of the cells. This was performed to determine the proper localization and expression levels of the proteins of interest in the analysed cells.
Live-cell image acquisition. Imaging of NEMO-GFP-expressing cells was performed using an N-STORM inverted microscope (Nikon) in TIRF mode. A 488 nm excitation laser was angled through the back focal plane of an original magnification Â 100 TIRF objective (Nikon). A range of laser powers were tested for imaging the samples; as GFP intensity fluctuations are minimal regardless of laser power in the absence of a specialized imaging buffer, similar SOFI-based reconstructions were obtained at all powers tested. As a result, the live-cell movie displayed was obtained at 0.083 kW cm À 2 . Emitted signal from the excited GFP molecules was collected by an electron-multiplying charge-coupled device camera (iXon Ultra 897, Andor), with a pixel size of 160 nm.
Fixed-cell super-resolution image reconstruction. Final super-resolution reconstructions were then generated in Fiji 59 through the QuickPALM algorithm 57 by creating images with 20 nm pixel size and additively superimposing a Gaussian kernel of 20 nm full-width half-maximum to each particle.
Live-cell super-resolution image reconstruction. Final super-resolution reconstructions were generated in Fiji 59 through a custom-made SOFI-based algorithm using fourth-order SOFI and generating images with 40 nm pixel size. Superresolution Movie 1 analysed through the custom NanoJ-SRRF V1.0 ImageJ plugin is available upon request to the authors.
Quantitative analysis. Protein cluster segmentation was performed through a dedicated algorithm implemented in Jython, running within the Fiji software 59 .
Here the algorithm uses a Huang threshold method 60 to segment clustered particle detections, this approach showed optimal performance in the identification of NEMO particle clusters. Background detections (false detection) caused by noise fluctuations or motile unbound labelled antibodies were minimally present in the thresholded images. For each segmented particle cluster in the rendered image, the following properties were measured: particles density, area, circularity and diameter. We verified that cell illumination was considerably homogenous for the individual cell regions-of-interest, but slightly non-homogeneous for the full field of view due to the Gaussian profile of the laser illumination in our microscopy setup. In all graphs, n is the number of whole cells analysed per condition.
Statistical analyses. We used StatPlus to create histogram bin sets for the quantification of comparisons between cell types and treatments. The error bars are representative of s.e.m. Statistical analyses were performed using a two-tailed Student's t-test, to determine statistical significance that is expressed as *Po0.05. Data shown are that of biological replicates from various experiments performed multiple times.
Data availability. The data that support the findings of this study are available from the corresponding authors upon request.


extfieldsofstudy: Biology Medicine

s2fieldsofstudy: Biology Chemistry

text:
Ursolic acid enhances macrophage autophagy and attenuates atherogenesis[S]

Macrophage autophagy has been shown to be protective against atherosclerosis. We previously discovered that ursolic acid (UA) promoted cancer cell autophagy. In the present study, we aimed to examine whether UA enhances macrophage autophagy in the context of atherogenesis. Cell culture study showed that UA enhanced autophagy of macrophages by increasing the expression of Atg5 and Atg16l1, which led to altered macrophage function. UA reduced pro-interleukin (IL)-1β protein levels and mature IL-1β secretion in macrophages in response to lipopolysaccharide (LPS), without reducing IL-1β mRNA expression. Confocal microscopy showed that in LPS-treated macrophages, UA increased LC3 protein levels and LC3 appeared to colocalize with IL-1β. In cholesterol-loaded macrophages, UA increased cholesterol efflux to apoAI, although it did not alter mRNA or protein levels of ABCA1 and ABCG1. Electron microscopy showed that UA induced lipophagy in acetylated LDL-loaded macrophages, which may result in increased cholesterol ester hydrolysis in autophagolysosomes and presentation of free cholesterol to the cell membrane. In LDLR−/− mice fed a Western diet to induce atherogenesis, UA treatment significantly reduced atherosclerotic lesion size, accompanied by increased macrophage autophagy. In conclusion, the data suggest that UA promotes macrophage autophagy and, thereby, suppresses IL-1β secretion, promotes cholesterol efflux, and attenuates atherosclerosis in mice.

Cell culture and manipulation RAW264.7 cells were obtained from ATCC (Rockville, MD) and were maintained in DMEM with10% FBS. Thioglycollateelicited mouse peritoneal macrophages (pMs) were obtained as previously described (33) and cultured in DMEM with 10% FBS for 2 h, unattached cells were washed away, and the attached macrophages were then further cultured in appropriate medium. For LPS treatment, pMs were incubated for 6-24 h in the culture medium (DMEM with 0.25% FBS) with an addition of LPS and UA at the indicated concentrations. The cells were then washed with Dulbecco's PBS twice before being lysed for total RNA or protein extraction. Raw 264.7 cells and pMs were treated with UA (10 M) and TLR1/2 ligand Pam3CSK4 (500 ng/ml) or TLR4 ligand LPS (100 ng/ml) either alone or in combination, as indicated, for 6 h. In some experiments, cells were exposed to various autophagy modulators for 1 h prior to UA treatment, such as 500 nM Wortmannin to inhibit PI3K or 100 nM bafilomycin to inhibit degradation of LC3.

Mice
All mice were purchased from the Jackson Laboratory (Bar Harbor, ME) and housed in the University of South Carolina Animal Research Facility. All animal experiments were carried out in compliance with the National Institutes of Health guidelines and were approved by the Institutional Animal Care and Use Committee of the University of South Carolina. In the serum IL-1 measurement experiment, C57BL/6 mice were randomly grouped (four mice in the control group, five mice in the LPS group, and five mice in LPS plus UA group) and were injected intraperitoneally with either LPS (20 g/mouse) alone or together with UA (50 mg/kg), and blood was taken after 16 h for analysis of serum IL-1 by ELISA using kits from eBioscience (catalog number BMS6002INST; San Diego, CA). For atherosclerosis studies, female LDLR / mice were fed a Western diet and treated with UA for 11 weeks before they were euthanized for analysis. The Western diet contained 21% (wt/wt) anhydrous milkfat, 34% (wt/wt) sucrose, and a total of 0.2% (wt/wt) cholesterol (diet number TD.88137; Harlan Laboratories, Indianapolis, IN).

MTT assay
The pMs were cultured in phenol red-free medium in 24well plates. Cytotoxicity of UA was determined using an MTT cell viability assay kit from ATCC Bioproducts TM (Manassas, VA) following the manufacturer's instructions. The 96-well microplates were read using a SpectraMax M5 microplate reader (Molecular Devices, Sunnyvale, CA); absorbance at 570 nm was measured.
Lipid-laden macrophages play a crucial role in the initiation of atherogenesis, and inflammatory responses of macrophages are a driving force for atherosclerotic progression and plaque growth (17)(18)(19)(20)(21). Unbalanced macrophage foam cell apoptosis and efferocytosis lead to necrotic core formation in advanced plaques and plaque instability (22). Recently, limited but convincing studies have shown that macrophage autophagy is atheroprotective by substantially altering all these aspects, including the enhancement of cholesterol efflux, suppression of inflammation, and improvement of efferocytosis (11,15,23). Evidence showing that atherosclerosis progression is accompanied by increasing defects in macrophage autophagy provides further enthusiasm regarding the possibility that enhancement of macrophage autophagy may be an effective way to reduce or reverse atherosclerosis. However, currently known autophagy inducers, including mTOR inhibitors and toll-like receptor (TLR)7 ligands, are less than ideal therapeutics in the context of atherosclerosis, due to hyperlipidemic (24) and pro-inflammatory effects, respectively (25). Therefore, the search for new macrophage autophagy stimuli is urgent.
Ursolic acid (UA), a natural pentacyclic triterpene carboxylic acid found in various plants, is present in the human diet and possesses a wide range of biological functions, such as anti-oxidation, anti-inflammation (26), anti-cancer (27), and anti-hyperlipidemia (28). Some studies have reported that UA reduces obesity, glucose intolerance, and fatty liver (29,30). These beneficial properties are of apparent value for the treatment of atherosclerosis, although the underlying molecular mechanisms are largely unknown. Indeed, limited animal studies have shown the potential anti-atherogenic effects of UA (31). Our recent finding that UA promotes cancer cell autophagy (32) prompted us to determine whether this autophagy-enhancing function also holds true in macrophages. To this end, we performed both in vitro and in vivo studies and found that UA enhances macrophage autophagy, suppresses lipopolysaccharide (LPS)-induced IL-1 production and secretion, and promotes cholesterol efflux from acetylated (Ac)LDLloaded macrophages. Furthermore, we found that UA attenuated atherogenesis in LDLR / mice fed a Western diet. Our results suggest that UA may be developed as a new anti-atherogenic agent by virtue of its ability in enhancing macrophage autophagy in an atherogenic environment. and permeabilized for 10 min with Triton X-100 (0.1% in PBS). All subsequent steps were conducted at room temperature. The cells were blocked with 0.01 M glycine/PBS three times, 30 min each time, and then with 5% BSA in PBS for 1 h and stained with primary antibody (rabbit anti-LC3 or goat anti-IL-1 at 1:200 in blocking buffer) for 1 h. After washing three times with PBS, the cells were stained with Alexa Fluor 555-conjugated goat anti-rabbit IgG (1:500 in PBS) and with Alexa Fluor 488-conjugated donkey anti-goat IgG (1:500 in PBS) for 1 h. The cells were stained with 4′,6diamidino-2-phenylindole (5 g/ml) for 5 min and then washed with PBS. The coverslips were mounted onto glass slides with fluorescent mounting medium (Dabco) and analyzed on an Olympus FV1000 laser scanning confocal microscope (Tokyo, Japan).

Transmission electron microscopy
Cells were cultured on Permanox four chamber slides for different lengths of time. Cells were fixed with 1-2% glutaraldehyde for 30 min and then rinsed with PBS three times; they were then incubated in 1% osmium tetroxide with 1.5% potassium ferricyanide for 30 min and then rinsed with water. The cells were dehydrated by incubating them in 70% ethanol for 15 min twice, then 95% ethanol for 15 min twice, and finally 100% ethanol for 20 min. The cells were then incubated in ethanol and acetonitrile (1:1) for 10 min, then with acetonitrile for 10 min twice; next they were incubated in PolyBed 812 and acetonitrile (1:2) for 1 h, then with PolyBed 812 and acetonitrile (2:1) for 1 h, and within pure PolyBed 812 overnight. Cells were embedded in mold 48 h at 60°C, which was cut and examined by transmission electron microscopy; 50 cells in each group were examined.

Cellular cholesterol analysis
The intracellular total cholesterol and FC were measured by an intracellular cholesterol quantitation kit (Sigma-Aldrich) according to the manufacturer's instructions. Briefly, macrophages were trypsinized with trypsin-EDTA followed by centrifugation. The cell pellets were resuspended in a mix of chloroform:isoporpanol:IGEPAL CA-630 (7:11:0.1). The samples were centrifuged at 13,000 g for 10 min to remove insoluble material. The organic phase was transferred to new tubes and air dried at 50°C for 30 min, followed by blowing the samples with nitrogen to remove organic solvent. The lipid was dissolved with 200 l cholesterol assay buffer; 50 l of the reaction mix was added to each standard and sample well in a 96-well plate and incubated for 60 min at 37°C in the dark. The absorbance at OD 570 nm was measured with a Spectra Max M5 microplate reader (Molecular Devices).

Plasma lipid analysis
Blood was obtained from mice following an overnight fast and the plasma was obtained by centrifugation of the blood at 4,000 g for 20 min. Plasma cholesterol and triglyceride levels were measured by enzymatic colorimetric assays with cholesterol reagent and triglycerides GPO reagent kits (Raichem, SanDiego, CA). Plasma cholesterol lipoprotein profiles were determined using a fast-performance LC system (AKTA purifier, GE Healthcare Biosciences, Pittsburgh, PA) equipped with a Superose 6 10/300 GL column (GE Healthcare). Pooled mouse plasma (100 l) was loaded onto the column and eluted at a constant flow rate of 0.5 ml/min with 1 mM sodium EDTA and 0.15 M NaCI. Fractions of 0.5 ml were collected and cholesterol concentration from each fraction was measured.

Atherosclerosis analysis
Hearts and aortas were dissected from mice and processed as described previously (35). The base of the heart containing the

Macrophage cholesterol efflux
The cholesterol efflux assay was processed as previously described (34). Attached mouse macrophages in 24-well plates were cultured in DMEM for 24 h followed by cholesterol-loading with 100 g/ml AcLDL incorporated with 3 H-cholesterol for 3 days. Then efflux medium (either FBS-free DMEM or DMEM containing 20 g/ml human apoAI or 50 g/ml human HDL) with the absence or presence of 10 M UA was added to initiate cholesterol efflux for 24 h. 3 H radioactivity in efflux medium and cells was counted using a Beckman LS6500 liquid scintillation counter (Beckman Coulter, Indianapolis, IN). Cholesterol efflux rate was calculated as the percentage of the radioactivity counts in the medium. 3 H-labeled cholesterol was purchased from Perkin-Elmer (Waltham, MA). AcLDL and human HDL were from Biomedical Technologies (Stoughton, MA).

Western blot
Protein samples were loaded onto SDS-PAGE gels for electrophoresis. The size-separated proteins were then transferred to nitrocellulose membranes (Amersham Biosciences, Piscataway, NJ). The indicated primary antibodies and HRP-conjugated secondary antibodies were used to detect target proteins. Signal was detected using an ECL kit (Bio-Rad). For stripping, membranes were submerged for 30 min at 55-60°C in a buffer containing 100 mM 2-mercaptoethanol, 2% (v/v) SDS and 62.5 mM Tris-HCl (pH 6.7), with agitation, and then were washed three times with PBST.

Measurement of cytokines in cell culture medium or serum
Cytokine protein concentrations in cell culture medium or serum were measured by ELISA, using kits from eBioscience (San Diego, CA) following the manufacturer's instructions. The macrophage-conditioned culture medium was diluted by 30-to 180fold for the measurement. The 96-well microplates were read using a SpectraMax M5 microplate reader.

Immunofluorescence staining
Macrophages were seeded in 8-well chamber slides. Cells were fixed with 4% paraformaldehyde for 3 min at room temperature showed that UA at 10 M was 2-fold more effective in increasing LC3I/II expression than rapamycin at 100 nM (Fig. 1B).
The autophagy-enhancing effect of UA was also evident as observed by electron microscopy. As shown in Fig. 1C, UA-treated pMs had numerous double-membrane structures that were characteristic of nascent autophagosomes, autophagosomes, and autolysosomes, while much fewer of these structures were detected in control cells. Autophagosomes and nascent autophagosomes could not be clearly distinguished in the micrographs. The structure indicated as "autophagosomes" could also be nascent autophagosomes. In autolysosomes, the segregated organelles lose their distinct shape and are usually replaced by electrondense materials. The diameter of typical autophagosomes or autolysosomes is 0.5-2 m. These electron microscopy data have confirmed that UA induces autophagy in pMs.
To examine the mechanisms by which UA enhances macrophage autophagy, qPCR was performed and showed that in both mouse pMs and Raw264.7 cells, UA at 10 M significantly increased the expression of autophagyrelated proteins Atg5 and Atg16L1, but not that of Atg4c and Beclin-1 (BECN1) (Fig. 1D, E). UA did not change LC3 mRNA expression in macrophages either (data not shown).

UA suppresses IL-1 production in macrophages
It has been reported that UA has anti-inflammatory effects on macrophages (26). To investigate the effects of UA on pro-inflammatory cytokines such as IL-1, IL-6, and TNF, pMs were treated with LPS (100 ng/ml) and/or UA (10 M) for 6 h, and the expression of the abovementioned cytokines was examined using qPCR. As shown in Fig. 2A and supplementary Fig. 1A, LPS significantly increased the mRNA expression of IL-1, IL-6, and TNF; UA only slightly (without statistical significance) suppressed LPS-induced expression of these cytokines. However, Western blot analysis showed that UA substantially reduced the pro-IL-1 protein level in both LPS-treated pMs and Raw264.7 cells (Fig. 2B).
We next examined whether UA suppresses mature IL-1 secretion. Mouse pMs were treated with LPS or Pam3CSK4 with or without UA for 16 h and the conditioned medium was collected for ELISA measurement of IL-1 protein levels. The result showed that LPS and Pam3CSK4 stimulated the secretion of IL-1, whereas UA significantly inhibited the secretion of IL-1 (Fig. 2C). However, UA did not significantly reduce the secretion of IL-6 or TNF in LPS-treated mouse pMs (supplementary Fig. 1B).
To determine whether UA suppresses IL-1 production in vivo, we injected C57BL/6 mice intraperitoneally with either PBS (control), LPS alone, or LPS together with UA (50 mg/kg). Sixteen hours after injection, blood samples were taken for analysis of serum IL-1 concentration by ELISA. The results showed that LPS significantly increased serum IL-1 levels and UA significantly suppressed the induction (Fig. 2D). We developed a method to measure mouse serum UA concentration by coupling HPLC aortic root was embedded in OCT medium and frozen at 20°C. The aorta (from the ascending region to the abdominal region) was fixed in 10% neutral buffered formalin at room temperature overnight and then kept in PBS before removing the adventitia. For analysis of atherosclerosis in the aortic root, 10 m serial sections were collected. Atherosclerotic lesion areas of 10 serial sections were visualized with hematoxylin and eosin staining (hematoxylin: GeneTex, Inc., ready-to-use, catalog number GTX73341; eosin: Sigma-Aldrich, eosin B, catalog number 861006-10G), and lipid-stained lesions were visualized by Oil Red O (ORO) staining and quantified by Image-Pro Plus 6.0. Rat anti-mouse moma2 (Abcam) was used for immunostaining for macrophage analysis on serial sections of the aortic root, and biotinylated goat anti-rabbit IgG (Vector Laboratories, Burlingame, CA; catalog number BA-1000) was used as the secondary antibody. Immunoreactivity was amplified using the Vectastain ABC kit (Vector Laboratories, catalog number PK6100), and signal was enhanced by peroxidase enhancer (GeneTex, Irvine, CA; GTX82979) and finally reacted with the substrate from AEC Chromogen/FRP substrate kit (GeneTex; GTX82977). For collagen staining of the lesions, the 10 m-thick aortic root sections were stained by Movat's pentachrome using a Movat pentachrome stain kit from Scytek Laboratory (catalog number MPS-2; Logan, UT) following the manufacturer's instructions, and visualized and analyzed by a Nikon NiU research microscope equipped with a NIS-Elements software package and a high resolution CMOS color camera as well as Image-Pro Plus 6.0. The en face aortas were stained with ORO and the plaque areas were analyzed by ImageJ.

Statistical analysis
All data were summarized as mean ± SEM. To compare two groups on a continuous response variable (e.g., gene expression levels and atherosclerosis size), we used a t-test. To compare multiple groups on a continuous response variable, we used a oneway ANOVA followed by Bonferroni post assay to compare the selected two groups. Normality and homoscedasticity were tested to ensure that parametric testing was appropriate. P  0.05 was considered statistically significant. GraphPad Prism 5 software was used to carry out all statistical analysis.

UA enhances macrophage autophagy
The structure of UA was confirmed by NMR; the purity was determined by mass spectroscopy and HPLC to be >98% (data not shown). Its cytotoxicity was measured by MTT assay. UA was dissolved in DMSO as a stock solution of 10 mM and then diluted into appropriate media. Mouse pMs and Raw264.7 cells were treated with 0-40 M UA for up to 5 days. Data showed that at concentrations up to 10 M, UA did not cause any apparent cytotoxicity to both cell types, but it led to cell death at >30 M (data not shown). Therefore, we used UA at 10 M or lower for all the following experiments. To examine whether UA increases macrophage autophagy, we treated mouse pMs with UA and Wortmannin (an autophagy inhibitor). UAenhanced autophagy was evidenced by increased LC3-I/II proteins, whereas Wortmannin completely blocked UAinduced LC3 increase (Fig. 1A). Next, using Raw264.7 cells, we confirmed UA's autophagy-inducing activity. Data The overlap between IL-1 and LC3 signals indicates that autophagy induced by UA may sequester pro-IL-1 in autophagic vesicles.

UA promotes cholesterol efflux from lipid-loaded macrophages
Autophagy has been shown to enhance cholesterol efflux to apoAI (12)(13)(14). To examine whether UA promotes macrophage cholesterol efflux by enhancing autophagy, we used mouse pMs to perform experiments using previously established protocol (35). We found that UA (10 M) significantly increased cholesterol efflux to apoAI, but not to HDL (Fig. 3A). An autophagic flux inhibitor, bafilomycin A1, completely diminished the cholesterol efflux enhancement by UA (Fig. 3B). qPCR and Western blots showed that UA did not increase the mRNA expression and protein levels of ABCA1 and ABCG1 (Fig. 3C, D), indicating that the cholesterol efflux enhancing effects of UA are not caused by ABCA1 or ABCG1 induction.
To search for evidence of autophagic vacuoles in lipidloaded macrophages and, if present, whether these were associated with lipid droplets (LDs), electron microscopic analysis was performed. The results clearly showed that UA increased lipophagy in AcLDL-loaded macrophages (Fig. 3E). LDs were easily identifiable as round lightdensity structures in AcLDL-loaded cells [ Fig. 3E(b), white and mass spectroscopy; we found, after injection of UA at 50 mg/kg, that serum UA concentration peaked at 12 M at 2 h post injection and dropped to below 3 M at 24 h post injection (data not shown).
A previous study demonstrated that autophagy inhibits inflammasome activation and IL-1 secretion in response to endotoxin (9). IL-1 is produced by activated macrophages as a proprotein (pro-IL-1), which is proteolytically processed to its active form by caspase 1. Inflammasome activation results in increased availability of caspase 1 and, therefore, accelerated process of pro-IL-1. Interestingly, we found that UA did not change the expression of caspase 1 (data not shown). Other studies have shown that, in macrophages, autophagy may also regulate IL-1 secretion by encapsulating and degrading pro-IL-1 in autolysosomes (10). To examine whether UA promotes autophagic sequestration and degradation of pro-IL-1, we incubated Raw264.7 cells with UA and LPS, and detected the intracellular location of IL-1 and LC3 by confocal microscopy. As shown in Fig. 2E, when Raw264.7 cells were treated with LPS for 24 h, IL-1 staining in the cytosol was significantly greater compared with control; UA-treated cells had dramatically more autophagosomes (LC3 staining) than control or LPS-treated cells. When the cells were treated with LPS and UA, IL-1 staining became weaker and appeared to be colocalized with LC3. could reduce atherosclerosis in mouse models. Female LDLR / mice at an age of 8 weeks were randomly divided into two groups (9-10 mice per group) and fed a Western diet for 11 weeks. In the DMSO group, the mice were injected intraperitoneally with vehicle (0.1% DMSO in 1 ml PBS) every other day; while in the UA group, the mice were injected with UA (50 mg/kg in 1 ml PBS with 0.1% DMSO) every other day. Food intake and body weight were measured weekly. We noticed that, while the food intake was not different between the two groups (supplementary Fig. 2A), the body weights of the UA group mice were lower than the vehicle group in the first 3 weeks after the initiation of Western diet and UA treatment, but they caught up in the following weeks; at the endpoint, body weights of the two groups were not significantly different (supplementary Fig. 2B). The mice were euthanized and comprehensive analyses were performed to compare the two groups. We did not find any significant differences between the two groups of mice in total serum cholesterol (supplementary Fig. 2C), triglycerides (supplementary Fig. 2D), and fasting blood glucose (supplementary Fig. 2E), solid arrow] and UA reduced the number of LDs and increased the frequency of LDs with areas of high density and asymmetrically localized multi-membrane structures, indicating that autophagosomes occur on the surface of the LDs [Fig. 3E(c), black solid arrow].
To examine whether UA increases cholesterol ester (CE) hydrolysis through enhancing autophagy, we measured total cholesterol and FC in mouse pMs loaded with 50 g/ml AcLDL in the absence or presence of 10 M UA in serum-free DMEM for 24 h; and the CE and FC/CE ratio were calculated. We found that, while UA did not significantly reduce total cholesterol loading to the macrophages, it significantly increased the FC/CE ratio (Fig. 3F), indicating that UA, indeed, increased CE hydrolysis in macrophages.

UA attenuates atherogenesis in hypercholesterolemic mice
Because UA was shown to suppress macrophage IL-1 production and secretion and enhance macrophage cholesterol efflux, we next examined whether UA treatment significance (Fig. 4C). However, the collagen content (percentage) in the lesions, as determined by Movat's pentachrome staining, was significantly larger in the UA group (22.9 ± 2.0%) than in the vehicle group (14.7 ± 2.6%) (Fig.  4D). Furthermore, the atherosclerotic lesion area in en face aortas was significantly smaller in the UA group than in the vehicle group (supplementary Fig. 3B, C).
We also stained IL-1 and LC-3 in the aortic root slides to examine whether UA induced autophagy and reduced IL-1 in atherosclerotic lesions. The autophagy levels (reflected by LC-3 staining) in atherosclerotic lesions were increased in the UA group compared with the control group, especially in the macrophage-rich lesions. Meanwhile, the positivity of IL-1 was significantly decreased in the atherosclerotic lesions of the UA group compared with the control group. More interestingly, in the lesions of the UA group, there was significantly more IL-1 and LC-3 overlapping (Fig. 5). Our results suggested that UA treatment has increased autophagy in the atherosclerotic lesion, as well as pro-IL-1 degradation in the autophagosome.

DISCUSSION
In this study, we demonstrate that a phytonutrient, UA, enhances macrophage autophagy, suppresses macrophage as well as overall fast-performance LC lipoprotein profile (supplementary Fig. 2F, G). ELISA showed that IL-1 protein levels were reduced in the serum of the UA group compared with the control group, while the serum levels of IL-5 were not different between the two groups of mice (Fig. 4A). Interestingly, serum IL-6 and TNF levels were also significantly reduced in UA-treated mice (supplementary Fig. 2H). The reduction in these cytokines may be secondary to IL-1 reduction and the resulting reduced chronic systemic inflammation in mice.
The UA group of mice developed significantly less atherosclerosis compared with the vehicle group. As visualized by hematoxylin and eosin staining, mean lesion area in the aortic root was 3.72 ± 0.27 and 6.01 ± 0.58 × 10 5 m 2 in the UA group and the vehicle group, respectively, reflecting a 38% reduction in the UA group (supplementary Fig. 3A). We further stained the aortic root lesions for neutral lipid with ORO. The results showed that the lipid staining area in the lesions of the UA group (3.91 ± 0.31 × 10 5 m 2 ) was significantly smaller than that in the vehicle group (5.79 ± 0.22 × 10 5 m 2 ), with a 32% reduction (Fig. 4B). We stained lesional macrophages using a mouse macrophage-specific antibody, MOMA-2, and found that the UA group displayed a slightly lower percentage of macrophage area in aortic root atherosclerotic lesions compared with the vehicle group (63.4 ± 2.6% vs. 70.2 ± 2.7%), but without statistical Values are presented as mean ± SEM relative to DMSO control. *P < 0.05. B: The effect of UA on macrophage cholesterol efflux to apoAI was diminished by bafilomycin A1 (Baf). *P < 0.05. C, D: UA did not affect ABCA1 and ABCG1 mRNA expression (C) and protein levels (D) in cholesterol-loaded pMs. pMs were loaded with AcLDL, with or without UA, ABCA1, and ABCG1 mRNA and protein levels were determined by qPCR or Western blot. Values are presented as mean ± SEM relative to vehicle control. E. Transmission electron microscopy images of pMs: control (a); AcLDL (b); AcLDL+UA (c). White arrows, LDs; black arrows, LDs with areas of high density and asymmetrically localized multi-membrane structures; N, nucleus. Scale bar: 2 m. F: Total cellular cholesterol and FC/CE ratio were measured for mouse pMs loaded with AcLDL for 24 h in the absence or presence of UA.
inhibitors at levels sufficient for autophagy induction in macrophages causes systemic immunosuppression. The characterization of UA as a macrophage autophagy inducer in this study may add this natural compound as a new compound for clinical development toward a new anti-atherogenic agent. We show that UA enhances macrophage autophagy by increasing mRNA expression of Atg5 and Atg16l1 and protein levels of LC3I and LC3II; electron microscopy confirmed that UA increases autophagosome formation in macrophages. However, the more detailed molecular mechanisms warrant further investigation; for example, what targets UA directly binds to in the cell or on the cell membrane, and how UA modulates Atg5 and Atg16l1 expression, but not several other autophagy-related genes.
Macrophages contribute to atherogenesis by serving as inflammatory machinery and as a lipid deposit tank. Autophagy has been shown to reduce macrophage inflammation and foam cell formation by various mechanisms. In the arm of inflammation in the arterial intima, oxidized lipids and other inflammatory stimuli provoke macrophages to produce a variety of inflammatory cytokines, including IL-1, IL-6, and TNF-, among which IL-1 plays a dominant role in the inflammatory cascade in the plaques. In this study, we show that UA significantly reduced intracellular pro-IL-1 levels and mature IL-1 secretion in LPS-treated macrophages, although it did not reduce IL-1 mRNA expression. Increasing evidence suggests that autophagy is necessary to keep down the endogenous sources of inflammasome and IL-1 activation (36). Macrophages derived from Atg16l1-deficient mice produced higher levels of IL-1 (9). Additional studies using cells of human origin demonstrated that the inhibition of autophagy led to the upregulated production of IL-1 (37,38). To examine how UA regulates IL-1 production in macrophages, we used immunofluorescence confocal microscopy to observe colocalization of IL-1 and LC3 in Raw 264.7 cells after treatment by TLR ligands combined with UA. The overlap between IL-1 and LC3 indicated that autophagy induced by UA brought pro-IL-1 into the lumen of autophagic vacuoles for degradation, thereby reducing pro-IL-1 in the cells, which was usually processed by caspase 1 to generate mature IL-1 for secretion; although UA-induced autophagy did not suppress active caspase 1 generation through inflammasomes.
Oxidized or other modified LDL, unlike normal LDL, is readily taken up by the macrophage through scavenger receptors SR-A or CD36, resulting in the formation of macrophage foam cells. Accumulation of lipid-loaded macrophage foam cells is a central feature in the formation of atherosclerotic plaques (39,40). Maintaining macrophage cholesterol homeostasis is essential for the prevention and treatment of atherosclerosis. In this study, we provide evidence that UA enhanced apoAImediated cholesterol efflux from macrophage foam cells through autophagy. Our data show that UA enhanced apoAI-mediated cholesterol efflux, which can be blocked by the autophagic flux inhibitor, bafilomycin A1. IL-1 secretion, and promotes cholesterol efflux from cholesterol-loaded macrophages. Furthermore, we show that administration of UA attenuates atherogenesis in hyperlipidemic mouse models. This represents the first report that a natural compound suppresses atherosclerosis through a mechanism that is associated with macrophage autophagy enhancement.
Several recent studies have convincingly shown that macrophage autophagy plays a protective role against atherogenesis in mouse models. However, using macrophage autophagy induction as an anti-atherosclerosis strategy is a distant goal because none of the known autophagy inducers or enhancers have been shown to be beneficial without reasonable side effects in preclinical studies. For example, systemic administration of mTOR Fig. 4. UA attenuates atherosclerosis in LDLR-deficient mice fed a Western diet. The mice were fed a Western diet for 11 weeks and administered vehicle or UA as described. The aortic roots were embedded in OCT medium and frozen at 20°C immediately after being cut away from the aorta. For analysis of atherosclerosis, 10 micron-thick sections were collected in the region of the proximal aorta, starting from the end of the aortic sinus. A: UA significantly reduced serum IL-1 levels, but did not significantly alter serum IL-5 levels in the mice. B: The lipid content in lesions was determined by ORO staining and quantified by Image-Pro Plus 6.0. The quantitative analysis and representative images are shown. C: The areas of macrophages in lesions were determined by immunostaining for MOMA-2 (a macrophage marker). D: Trichrome staining for quantification of the lesion collagen content. Quantitative analysis and representative images are shown. Scale bar: 250 m.
UA has anti-atherogenic effects on female LDLR / mice induced to be diabetic and fed a high fat diet supplemented with UA for 11 weeks. The mechanism was suggested to be monocyte recruitment suppression. Our results are in agreement with these findings because the absolute MOMA-2-stained area in the plaques of our LDLR / mice was reduced, although the area percentage reduction did not reach statistical significance. On the other hand, Messner et al. (43) reported a UA dosedependent increase in atherosclerosis in Western dietfed male apoE / mice receiving UA in drinking water for 24 weeks. The mechanisms were proposed to be endothelial cell death induction and serum IL-5 reduction caused by UA consumption. In our study, we did not examine the endothelial proliferation and apoptosis in UA-treated mice, but did not observe a significant decrease of serum IL-5 levels in LDLR / mice. As correctly pointed out by Tannock (44), all these studies, including ours, were designed differently, including the animal model, the UA administration route, the dose, and the duration, and the mechanistic studies evaluated different cell types. In our study, we administered UA using peritoneal injection, avoiding the potential metabolism of UA in the gut and first pass effects in the liver. The plasma UA concentration determined in mice was 3-12 M within 24 h after a single injection, which is close to the concentrations we used in the in vitro macrophage experiments, indicating that the macrophage autophagyenhancing mechanism identified in in vitro experiments can at least partially explain the anti-atherogenic effects. Indeed, we also directly observed enhanced autophagy in the plaques of UA-treated mice. Nevertheless, more studies are needed to confirm the effects of UA on atherosclerosis and to more comprehensively examine the mechanisms.
In conclusion, our data suggest that UA enhances macrophage autophagy and thus reduces pro-IL-1 processing into mature IL-1, and promotes macrophage cholesterol efflux (as depicted in Fig. 6). Both of these effects are antiatherogenic, contributing to the attenuated atherogenesis in LDLR / mice fed a Western diet. Our study provides evidence that UA might be an optimal anti-atherogenic therapy by virtue of its macrophage autophagy-enhancing activity.
By doing so, UA did not increase ABCA1 and ABCG1 expression; instead, UA increased lipophagy in macrophages, evidenced by the fusion of autophagosomes with LDs in the electron microscopic images. Our finding that enhanced autophagy is associated with improved cholesterol efflux from macrophages is consistent with previous reports (13,14). It is worth noting that our electron microscopic images show that UA increased the frequency of LDs with areas of high density and asymmetrically localized multi-membrane structures. For larger LDs, autophagosomes only engulfed a discrete part of the LDs, which then pinched off as membrane vesicles enriched in LC3 (autolipophagosomes), in this way autophagosomes occur on the surface of these large organelles. Such partial engulfment of organelles by autophagy has been described for the nucleus and endoplasmic reticulum (41,42).
We further demonstrate that the beneficial effects of UA on macrophage inflammatory response and cholesterol homeostasis translate well into atherosclerosis reduction in mouse models. We show that peritoneal injection of UA at 50 mg/kg once every other day in LDLR / mice fed a Western diet for 11 weeks significantly reduced atherosclerotic plaque formation. Detailed composition analysis for LDLR / mice showed that UA reduced the overall lesion size, the lipid containing area, and the macrophage content, while it increased the collagen content, suggesting that UA treatment results in smaller and more stable lesions in mice. Further immunohistochemical analysis revealed that UA treatment reduced IL-1 levels, but increased LC3 levels, in atherosclerotic plaque, and that UA significantly increased the colocalization of IL-1 and LC3 in the lesions, indicating that UA, indeed, reduced IL-1 concentration in plaque through enhancing autophagy and IL-1 sequestration and degradation in the autophagic pathway. ELISA data further confirmed the reduction of IL-1 in serum. These in vivo data demonstrate that the macrophage autophagy-inducing effects indeed confer UA with anti-atherogenic capacity.
There are two previous studies that investigated the effects of UA on atherosclerosis using mouse models that obtained opposite results. Ullevig et al. (31) showed that  UA enhances autophagy in macrophages by increasing expression of autophagy-related proteins Atg5 and Atg16l1. Autophagosome sequesters pro-IL-1 and leads to pro-IL-1 degradation in lysosomes, diverting it from being processed by caspase 1 into mature IL-1 for secretion. Autophagosomes also interact with LDs, encircling some lipids to form autolipophagosomes, and then fuse with the lysosome to form an autolysosome, which processes the containing lipids and presents FC to the cell membrane for ABCA1mediated cholesterol efflux.