Membrane protein biogenesis by the EMC

Recent cryo‐EM‐based models reveal how the ER membrane protein complex may accomplish insertion of protein transmembrane domains with limited hydrophobicity.

Insertion of strongly hydrophobic TMDs into the ER membrane is mediated by the Sec61 complex for co‐translational insertion and the GET complex for post‐translational insertion of tail‐anchors (Volkmar & Christianson, 2020). By contrast, the EMC inserts TMDs of limited hydrophobicity, frequently located at the N‐ or C‐termini of proteins, and is involved in biogenesis of multi‐spanning membrane proteins (Volkmar & Christianson, 2020).The EMC is highly conserved (Wideman, 2015). In vertebrates, ten subunits have been identified (EMC1‐10), two of which, EMC8 and EMC9, are homologous and the result of a vertebrate‐specific gene duplication (Wideman, 2015). In Saccharomyces cerevisiae, EMC8 has been lost (Wideman, 2015). Only EMC3 displays clear homology to other membrane protein insertases, the Oxa1 family (Wideman, 2015; Volkmar & Christianson, 2020). This family includes YidC, which inserts TMDs into the bacterial cytoplasmic membrane, usually in cooperation with the Sec61‐homologous SecYEG channel (Volkmar & Christianson, 2020). Their association, along with the SecDF ancillary complex, forms a holo‐translocon capable of protein secretion and TMD insertion, with striking similarities to the EMC complex (Martin et al, 2019).Recent work by Pleiner et al (2020) presented a 3.4 Å cryo‐EM structure of the human EMC purified via a GFP‐tag on EMC2 and incorporated into a phospholipid nanodisc. The complex is formed by nine proteins (EMC1‐8, EMC10) (Pleiner et al, 2020). EMC8 and EMC9 are structurally similar, and their association with EMC2 is mutually exclusive (O''Donnell et al, 2020). Of the 12 TMDs, nine constitute the pseudosymmetric central ordered core, with a basket‐shaped cytosolic vestibule formed primarily by alpha‐helices of the EMC3 and EMC6 TMDs and cytosolic EMC2 (Fig 1A; Pleiner et al, 2020). The L‐shaped lumenal domain of the EMC consists mostly of beta‐sheets (Fig 1A; Pleiner et al, 2020), flanked by a conspicuous and conserved amphipathic alpha‐helix of EMC1 sealing the vestibule at the interface between the membrane and the ER lumen, together with another smaller amphipathic helix contributed by EMC3 (Fig 1A; Pleiner et al, 2020). In the ER lumen, the two 8‐bladed propellers of EMC1 contact six of the eight other subunits and stabilize the entire complex (Fig 1A; Pleiner et al, 2020). Beta‐sandwiches of EMC7 and EMC10 are anchored to the EMC1 lumenal domain (Fig 1A; Pleiner et al, 2020). In the cytosol, the tetratricopeptide repeat (TPR) spiral of EMC2 forms a cup underneath the partially hydrophilic vestibule in the membrane between the TMDs of EMC3 and EMC6, bridging the cytosolic ends of TMDs of EMC1, 3 and 5 (Fig 1A; Pleiner et al, 2020). Cytosolic EMC8 is bound to the opposite face of EMC2 (Fig 1A).Open in a separate windowFigure 1Comparison of the structures of human and yeast EMC(A) Cryo‐EM 3D map of the human (emdb‐21929) and yeast (emdb‐21587) EMC, showing front and back views with individual subunits coloured. Membrane position, obtained from the OPM database, is shown by grey discs. (B) Close‐up view of the EMC cavity formed by EMC3 and EMC6. Left, shown in a hydrophobicity surface pattern. Right, surface representation overlapped with the TMDs of EMC3 and EMC6. EMC4, flexible and with a gate function at the substrate‐binding place, is shown in pink in the yeast representation. EMC4 is not visible at the atomic EMC human structure, although is observed as a weak density at the human model, accompanied by TMs of EMC7 and EMC10 (Pleiner et al, 2020). (C) The yeast EMC following > 5 µs of CG‐MD simulation. The protein is shown as surface and coloured as per Pleiner et al (2020). The computed densities of waters and phospholipid tails and phosphates are shown as blue, yellow and lime green densities, sliced to bisect the cavity for clarity. Right, inset of the EMC cavity. Methods: CG‐MD simulations were built using PDB 6WB9 in a solvated symmetric POPC/POPE/cholesterol membrane and run in the Martini forcefield as described in Martin et al (2019). 3 µs unrestrained simulations were run, followed by 2.5 µs backbone restrained simulation for density calculation, done using VolMap in VMD (Humphrey et al, 1996).The 3.0 Å cryo‐EM structure of the yeast EMC presented by Bai and colleagues shows a very similar overall organization (Bai et al, 2020). Here, purification was via a 3xFLAG‐tag on EMC5, and the structure of the 8‐subunit complex (without EMC8/9) was visualized in detergent solution (Bai et al, 2020). The yeast complex has twelve TMDs like the human EMC, but unlike the human structure, EMC4 in yeast has three TMDs that are clearly visible (Bai et al, 2020). They are angled in the membrane pointing away from the complex at the cytosolic end (Fig 1A), and Bai et al (2020) propose that TMDs of EMC4, EMC3 and EMC6 form a substrate‐binding pocket similar to that of YidC. As in the human EMC, there are two amphipathic helices (EMC1 and EMC3) at the membrane/lumen interface (Fig 1A; Bai et al, 2020). In the ER lumen, yeast EMC1 only has one 8‐bladed beta‐propeller, to which the beta‐sandwiches of EMC7 and EMC10 are anchored (Fig 1A; Bai et al, 2020). In the cytosol, EMC2 bridges EMC3, 4 and 5, and its TPR repeats form a cup underneath the vestibule similar to human EMC2 (Fig 1A; Bai et al, 2020).The authors propose that insertion of a partially hydrophilic TMD by the yeast EMC is mechanistically similar to insertion by bacterial YidC (Bai et al, 2020). Yeast EMC is proposed to bind substrate between TMD2 of EMC3 and TMD2 of EMC4 in a pocket with polar and positively charged amino acids at either end and hydrophobic amino acids in the centre (Fig 1B; Bai et al, 2020). Much has been made of a conserved positive region within the EMC complex here, present in an equivalent position also in YidC (Kumazaki et al, 2014): It is claimed to be important for the incorporation of more‐hydrophilic TMDs and perhaps responsible for the “positive‐inside” orientation rule (von Heijne, 1992). Yeast and human EMC3 contain a specific R31 and R26 residue, respectively, conserved also in YidC and important for function of the EMC, as well as for YidC in Gram‐positive, but interestingly not Gram‐negative, bacteria (Chen et al, 2014; Pleiner et al, 2020; Bai et al, 2020). Another interesting feature, also conserved with YidC, is the flexibility of the TMDs flanking the substrate‐binding pocket, critical for EMC entry of substrates (Bai et al, 2020).In the human EMC, methionine residues in a cytosolic loop of EMC3 act as a substrate bait (Pleiner et al, 2020). Polar and charged residues within the substrate‐binding groove guide the lumenal domain across the membrane, facilitated by local membrane thinning (Pleiner et al, 2020; Fig 1B). The positive charges within the substrate‐binding site exclude signal peptides and enforce the “positive‐inside rule” (von Heijne, 1992; Pleiner et al, 2020). Flexible TMDs of EMC4, EMC7 and EMC10 forming a “lateral gate” of the substrate‐binding groove allow sampling of the bilayer by the substrate TMD (Pleiner et al, 2020). As the shortened TMDs of EMC3 and EMC6 cannot stably bind the substrate TMD, they favour its release into the bilayer (Pleiner et al, 2020). The EMC1 beta‐propeller(s) may recruit additional protein maturation factors in the ER lumen (Pleiner et al, 2020; Bai et al, 2020) or bind the Sec61 channel to allow cooperation between the two insertases (Bai et al, 2020).Arguably, the most interesting feature of the EMC complex is the location of a large interior cavity with distinctive hydrophilic character, which likely aids TMD insertion (Fig 1B). We ran a coarse‐grained molecular dynamics (CG‐MD) simulation of the yeast EMC structure, which highlights a profound perturbation of the phospholipid bilayer in the EMC interior cavity (Fig 1C). Here, a deep gorge forms in the cytoplasmic leaflet of the bilayer, allowing the cavity to become flooded with water (Fig 1C). Note the location of the lipid head groups here (lime green), which presumably define the site of amphipathic TMD insertion. The incursion of phospholipids into the centre of the EMC complex is a feature shared by the bacterial holo‐translocon (Martin et al, 2019) and perhaps by all membrane protein insertases. The shape and character of the EMC cavity presumably dictate its predisposition for less hydrophobic TMDs; it would be interesting to see whether the cavities of different insertases are similarly tailored to suit their substrates.     

Mycobiota dysbiosis: a new nexus in intestinal tumorigenesis

While there is growing evidence that perturbation of the gut microbiota can result in a variety of pathologies including gut tumorigenesis, the influence of commensal fungi remains less clear. In this issue, Zhu et al (2021) show that mycobiota dysbiosis stimulates energy metabolism changes in subepithelial macrophages promoting colon cancer via enhancing innate lymphoid cell activity. These findings provide insights into a role of the gut flora in intestinal carcinogenesis and suggest opportunities for adjunctive antifungal or immunotherapeutic strategies to prevent colorectal cancer.Subject Categories: Cancer, Immunology, Metabolism

Recent work reports a role for the commensal gut flora in driving aberrant host immunity and malignant cytokine signaling.

There is growing evidence for an important role for the microbiota in influencing tumorigenesis (Helmink et al, 2019). It is now well documented that gut microbiota represents a highly diverse polymicrobial population of bacteria, fungi, viruses, and protozoa. Recent evidence highlights involvement of the bacterial component of the gut microbiota in protection or enhancement of colorectal tumorigenesis. In contrast, the importance of the mycobiota is less well understood although recently suggested to promote pancreatic oncogenesis and colitis‐associated colon cancer (CAC) (Wang et al, 2018; Aykut et al, 2019). Therefore, gut fungi may play a role in the development of other gastro‐intestinal cancer types, such as CRC. Notably, there is emerging evidence suggesting that mycobiota imbalance modulates immune cells and can trigger inflammatory bowel disease (IBD) (Richard & Sokol, 2019).Here, Zhu et al (2021) provide new insight into the association between mycobiota dysbiosis, immunomodulation, and tumorigenesis in the mouse gut (Fig 1).Open in a separate windowFigure 1Dectin‐3 deficiency induces fungal dysbiosis and tumorigenesis in mice by orchestrating immune cell metabolism and cytokine signalingIn the gut of wild‐type mice, the natural population of the commensal yeast Candida albicans is detected by the Dectin‐3 receptor located on the subepithelial macrophage cell surface. This recognition allows macrophages to maintain gut homeostasis by exerting an antifungal activity. In Dectin‐3‐deficient mice, the mycobiota becomes disrupted and aberrantly increased populations of C. albicans emerge. Elevated C. albicans load triggers increased glycolysis in macrophages and interleukin‐7 (IL‐7) secretion. Macrophage‐derived IL‐7 finally induces IL‐22 secretion by group‐3 innate lymphoid cells that in turn promote tumor cell proliferation in the gut epithelium.The current study (Zhu et al, 2021) is based on previous observations suggesting that human pathogenic fungi are recognized by the C‐type lectin receptor Dectin‐3. This led Zhu et al (2021) to test whether the mycobiota influenced gut tumor formation and is linked to immune recognition mediated by Dectin‐3. First, the authors demonstrated that mice lacking the Dectin‐3 receptor had increased colonic tumorigenesis in response to the azoxymethane (AOM) and dextran sodium sulfate (DSS). This was evident histologically in marked differences in tumor number, size, and burden in Dectin‐3‐deficient mice. Of note, immunohistochemical staining revealed that the lack of Dectin‐3 induced gut tumor formation by triggering epithelial cell proliferation rather than preventing cell apoptosis. In fact, first insight into the impact of microbes in CAC was suggested by the observation that co‐housed WT and Dectin‐3‐deficient mice displayed no difference in tumorigenesis. The pivotal role of the microbiota was then underlined in fecal transplantation experiments. Chemically induced germ‐free mice that received feces from Dectin‐3 tumor‐bearing mice displayed exacerbated tumor development compared to wild‐type controls. In addition, the fungal burden was specifically increased in tumor‐bearing Dectin‐3‐deficient animals. Deep profiling of the mycobiota alterations demonstrated an increase in a single yeast species, i.e., Candida albicans, that normally behaves as commensal in the gut (Papon et al, 2013; Wilson, 2019). Preliminary experiments suggested that the increased burden of C. albicans in Dectin‐3‐deficient tumor‐bearing mice is due to impaired antifungal killing by macrophages. Consistently, elevated C. albicans populations triggered glycolysis and inflammatory IL‐7 secretion from lamina propria macrophages, suggesting that Dectin‐3 deficiency‐induced fungal dysbiosis resulted in modulation of gut macrophage metabolism, promoting tumorigenesis. Exploring the molecular and cellular mechanisms that linked macrophage‐derived IL‐7 secretion and CRC development, Zhu et al (2021) showed in vitro that IL‐7 produced by subepithelial macrophages induced IL‐22 secretion by group‐3 innate lymphoid cells (ILC3s). In turn, up‐regulation of IL‐22 in Dectin‐3‐deficient mice contributed to the oncogenesis seen in these animals. Finally, a detailed analysis of tumor tissues collected from 172 patients with CRC showed correlation and poorer clinical outcome in patients with decreased expression of Dectin‐3, but increased expression of IL‐22 and mycobiota burden, although they did not directly link this to the presence of C. albicans in these patients.Overall, Zhu et al (2021) define a new cell paradigm linking mycobiota dysbiosis, macrophage energy metabolism, and innate lymphoid cell function to tumor development in the mouse gut. In this context, this study also sheds additional light on a new role of ILC3s, a recently described type of lymphoid effectors (Serafini et al, 2015). Indeed, ILC3s have been shown in the present article to act as cornerstone cells orchestrating cytokine‐regulated tumorigenesis in the gut. Beyond these pathophysiological considerations, the study opens up new opportunities for developing adjunctive antifungal or immunotherapeutic strategies for the prevention of high morbidity in CRC. Importantly, this enlightening article provides firm evidence that colonic C. albicans populations promote metabolic reprogramming in lamina propria macrophages and tumor cell formation. Metabolic reprogramming has been observed with other fungi, such as Aspergillus fumigatus, which induces metabolic rewiring of alveolar macrophages in the lung epithelium (Gonçalves et al, 2020). In line, the report by Zhu et al (2021) adds to previous work suggesting that mycobiota promotes pancreatic oncogenesis via activation of mannose‐binding lectins (Aykut et al, 2019). Mycobiota dysbiosis therefore stands out as an important new field of investigation in cancer research that is ripe for future exploration.     

USP26: a genetic risk factor for sperm X‐Y aneuploidy

Segregation of the largely non‐homologous X and Y sex chromosomes during male meiosis is not a trivial task, because their pairing, synapsis, and crossover formation are restricted to a tiny region of homology, the pseudoautosomal region. In humans, meiotic X‐Y missegregation can lead to 47, XXY offspring, also known as Klinefelter syndrome, but to what extent genetic factors predispose to paternal sex chromosome aneuploidy has remained elusive. In this issue, Liu et al (2021) provide evidence that deleterious mutations in the USP26 gene constitute one such factor.Subject Categories: Cell Cycle, Development & Differentiation, Molecular Biology of Disease

Analyses of Klinefelter syndrome patients and Usp26‐deficient mice have revealed a genetic influence on age‐dependent sex chromosome missegregation during male meiosis.

Multilayered mechanisms have evolved to ensure successful X‐Y recombination, as a prerequisite for subsequent normal chromosome segregation. These include a distinct chromatin structure as well as specialized proteins on the pseudoautosomal region (Kauppi et al, 2011; Acquaviva et al, 2020). Even so, X‐Y recombination fails fairly often, especially in the face of even modest meiotic perturbations. It is perhaps not surprising then that X‐Y aneuploidy—but not autosomal aneuploidy—in sperm increases with age (Lowe et al, 2001; Arnedo et al, 2006), as does the risk of fathering sons with Klinefelter syndrome (De Souza & Morris, 2010).Klinefelter syndrome is one of the most common aneuploidies in liveborn individuals (Thomas & Hassold, 2003). While most human trisomies result from errors in maternal chromosome segregation, this is not the case for Klinefelter syndrome, where the extra X chromosome is equally likely to be of maternal or paternal origin (Thomas & Hassold, 2003; Arnedo et al, 2006). Little is known about genetic factors in humans that predispose to paternal XY aneuploidy, i.e., that increase the risk of fathering Klinefelter syndrome offspring. The general notion has been that paternally derived Klinefelter syndrome arises stochastically. However, fathers of Klinefelter syndrome patients have elevated rates of XY aneuploid sperm (Lowe et al, 2001; Arnedo et al, 2006), implying a persistent defect in spermatogenesis in these individuals rather than a one‐off meiotic error.To identify possible genetic factors contributing to Klinefelter syndrome risk, Liu et al (2021) performed whole‐exome sequencing in a discovery cohort of > 100 Klinefelter syndrome patients, followed by targeted sequencing in a much larger cohort of patients and controls, as well as Klinefelter syndrome family trios. The authors homed in on a mutational cluster (“mutated haplotype”) in ubiquitin‐specific protease 26 (USP26), a testis‐expressed gene located on the X chromosome. Effects of this gene’s loss of function (Usp26‐deficient mice) on spermatogenesis have recently been independently reported by several laboratories and ranged from no detectable fertility phenotype (Felipe‐Medina et al, 2019) to subfertility/sterility associated with both meiotic and spermiogenic defects (Sakai et al, 2019; Tian et al, 2019). With their Klinefelter syndrome cohort findings, Liu et al (2021) also turned to Usp26 null mice, paying particular attention to X‐Y chromosome behavior and—unlike earlier mouse studies—including older mice in their analyses. They found that Usp26‐deficient animals often failed to achieve stable pairing and synapsis of X‐Y chromosomes in spermatocytes, produced XY aneuploid sperm at an abnormally high frequency, and sometimes also sired XXY offspring. Importantly, these phenotypes only occurred at an advanced age: XY aneuploidy was seen in six‐month‐old, but not two‐month‐old Usp26‐deficient males. Moreover, levels of spindle assembly checkpoint (SAC) proteins also reduced in six‐month‐old males. Thus, in older Usp26 null mice, the combination of less efficient X‐Y pairing and less stringent SAC‐mediated surveillance of faithful chromosome segregation allows for sperm aneuploidy, providing another example of SAC leakiness in males (see Lane & Kauppi, 2019 for discussion).Liu et al’s analyses shed some light on what molecular mechanisms may be responsible for the reduced efficiency of X‐Y pairing and synapsis in Usp26‐deficient spermatocytes. USP26 codes for a deubiquitinating enzyme that has several substrates in the testis. Because USP26 prevents degradation of these substrates, their levels should be downregulated in Usp26 null testes. Liu et al (2021) show that USP26 interacts with TEX11, a protein required for stable pairing and normal segregation of the X and Y chromosomes in mouse meiosis (Adelman & Petrini, 2008). USP26 can de‐ubiquitinate TEX11 in vitro, and in Usp26 null testes, TEX11 was almost undetectable. It is worth noting that USP26 has several other known substrates, including the androgen receptor (AR), and therefore, USP26 disruption likely contributes to compromised spermatogenesis via multiple mechanisms. For example, AR signaling‐dependent hormone levels are misregulated in Usp26 null mice (Tian et al, 2019).The sex chromosome phenotypes observed in Usp26 null mice predict that men with USP26 mutations may be fertile, but producing XY aneuploid sperm at an abnormally high frequency, and that spermatogenic defects should increase with age (Fig 1). These predictions were testable, because the mutated USP26 haplotype, present in 13% of Klinefelter syndrome patients, was reasonably common also in fertile men (7–10%). Indeed, sperm XY aneuploidy was substantially higher in fertile men with the mutated USP26 haplotype than in those without USP26 mutations. Some mutation carriers produced > 4% aneuploid sperm. Moreover, age‐dependent oligospermia was also found associated with the mutated USP26 haplotype.Open in a separate windowFigure 1Mutated USP26 as genetic risk factor for age‐dependent X‐Y defects in spermatogenesisMouse genetics demonstrate that deleterious USP26 mutations lead to less‐efficient X‐Y pairing and recombination with advancing age. Concomitant decrease of spindle assembly checkpoint (SAC) protein levels leads to less‐efficient elimination of metaphase I spermatocytes that contain misaligned X and Y chromosomes. This allows for the formation of XY aneuploid sperm in older individuals and subsequently increased age‐dependent risk for fathering Klinefelter syndrome (KS) offspring, two correlates also observed in human USP26 mutation carriers. At the same time, oligospermia/subfertility also increases with advanced age in both Usp26‐deficient mice and USP26 mutation‐carrying men, tempering Klinefelter syndrome offspring risk but also decreasing fecundity.As indicated by its prevalence in the normal control population, the USP26 mutated haplotype is not selected against in the human population. With > 95% of sperm in USP26 mutation carriers having normal haploid chromosomal composition, the risk of producing (infertile) Klinefelter syndrome offspring remains modest, likely explaining why USP26 mutant alleles are not eliminated. Given that full Usp26 disruption barely affects fertility of male mice during their prime reproductive age (Felipe‐Medina et al, 2019; Tian et al, 2019; Liu et al, 2021), there is little reason to assume strong negative selection against USP26 variants in humans. USP26 as the first‐ever genetic risk factor predisposing to sperm X‐Y aneuploidy and paternal origin Klinefelter syndrome offspring in humans, as uncovered by Liu et al, may be just one of many. 90% of Liu et al’s Klinefelter syndrome cases were not associated with USP26 mutations. But even in the age of genomics, discovery of Klinefelter syndrome risk factors is not straightforward, since most sperm of risk mutation carriers will not be XY aneuploid and thus not give rise to Klinefelter syndrome offspring. In addition, as Usp26 null mice demonstrate, both genetic and non‐genetic modifiers impact on penetrance of the XY aneuploidy phenotype: Spermatogenesis in the absence of Usp26 was impaired in the DBA/2 but not the C57BL/6 mouse strain background (Sakai et al, 2019), and in older mice, there was substantial inter‐individual variation in the severity of the X‐Y defect (Liu et al, 2021). In human cohorts, genetic and non‐genetic modifiers are expected to blur the picture even more.Future identification of sex chromosome aneuploidy risk factors has human health implications beyond Klinefelter syndrome. Firstly, XXY incidence is not only relevant for Klinefelter syndrome livebirths—it also contributes to stillbirths and spontaneous abortions, at a 4‐fold higher rate than to livebirths (Thomas & Hassold, 2003). Secondly, persistent meiotic X‐Y defects can, over time, result in oligospermia and even infertility. Since the mean age of first‐time fathers is steadily rising and currently well over 30 years in many Western countries, age‐dependent spermatogenic defects will be of ever‐increasing clinical relevance.     

Restraining CDK1–cyclin B activation: PP2A on the cUSP(7)

USP7 inhibitors are gaining momentum as a therapeutic strategy to stabilize p53 through their ability to induce MDM2 degradation. However, these inhibitors come with an unexpected p53‐independent toxicity, via an unknown mechanism. In this issue of The EMBO Journal, Galarreta et al report how inhibition of USP7 leads to re‐distribution of PP2A from cytoplasm to nucleus and an increase of deleterious CDK1‐dependent phosphorylation throughout the cell cycle, revealing a new regulatory mechanism for the progression of S‐phase cells toward mitosis to maintain genomic integrity.Subject Categories: Cell Cycle, Post-translational Modifications, Proteolysis & Proteomics

Recent work reveals untimely activation of mitotic cyclin‐dependent kinase as a molecular basis for p53‐independent cell toxicity of USP7 deubiquitinase inhibitors.

The G2‐M transition in the eukaryotic cell cycle is a critical point to ensure that cells with damaged DNA are unable to enter the mitotic phase. This checkpoint is highly regulated by a number of kinases, including ATR, ATM and WEE1, and ends upon activation of the CDK1–cyclin B1 kinase complex (Visconti et al, 2016). Since premature activation of CDK1–cyclin B1 causes replication fork collapse, DNA damage, apoptosis, and mitotic catastrophe (Szmyd et al, 2019 and references therein), restricting CDK1–cyclin B1 activity prior to mitosis is key to maintaining genomic integrity.A body of recent work has suggested that the deubiquitinase USP7 is a master regulator of genomic integrity; it is required for DNA replication in numerous ways, including indirect regulation of cyclin A2 during the S‐phase, origin firing, and replication fork progression. USP7 also regulates mitotic entry by stabilizing PLK1, another kinase which is highly active in the M phase and ensures proper alignment of chromatids prior to segregation. Notably, USP7 inhibitors have become an attractive cancer therapeutic strategy based on their ability to trigger degradation of MDM2, and thereby stabilize p53 (Valles et al, 2020). However, there is growing evidence of USP7 inhibitor‐related toxicity that is not mediated through p53 (Lecona et al, 2016; Agathanggelou et al, 2017), indicating that USP7 inhibitors impact other cellular processes. Therefore, Galarreta et al (2021) investigated the potential functional relationship between USP7 and CDK1, given the role of both factors in regulating the cell cycle.Through a series of in vitro experiments, the authors confirmed that five USP7 inhibitors induce premature mitotic kinase activity, including increased MPM2 signal (indicative of mitosis‐specific phosphorylation events) and phosphorylation of histone H3 Ser10 (H3S10P) in all cells, regardless of where they are in the cell cycle. To determine whether USP7 affects CDK1 during the cell cycle, Galarreta et al (2021) demonstrate that cell lines treated with USP7 inhibitors exhibit reduced levels of inhibitory Tyr‐15 phosphorylation on CDK1 and increased cyclin B1 presence in the nucleus, suggesting activation of the CDK1–cyclin B1 complex. Furthermore, treatment with the CDK1 inhibitor RO3306 rescues the USP7 inhibitor‐dependent increase of mitotic activity.These observations suggest that CDK1 has the potential to catalyze mitosis‐specific phosphorylation irrespective of cell cycle phase and that cells rely on USP7‐specific deubiquitination to suppress or reverse premature CDK1 activity. Surprisingly, despite the nuclear localization of cyclin B and decrease in inhibitory CDK1 Tyr‐15 phosphorylation, USP7 inhibitors failed to drive cells into mitosis. How might this be? Nuclear localization of cyclin B normally occurs just minutes before the onset of mitosis and nuclear envelope breakdown (Santos et al, 2012), yet the nucleus remains intact following USP7 inhibition. Moreover, the decrease in Tyr‐15 phosphorylation suggests the ATR‐ and WEE1‐dependent G2/M checkpoint is inactivated by USP7 inhibition. Do these data hint at the presence of an additional, unknown regulatory mechanism controlling mitotic entry independent of the G2/M checkpoint and nuclear localization of the CDK1–cyclin B complex?To determine whether CDK1 is the driver of USP7 inhibitor toxicity, Galarreta et al exposed cells to CDK1 inhibitors and observed a reduction in apoptosis. Furthermore, CDK1 inhibitors promote cell survival in cells treated with three structurally unrelated USP7 inhibitors. Finally, CDC25A‐deficient mouse embryonic stem cells, which constitutively express low levels of CDK1, resist USP7 inhibition. Together, these data suggest that the USP7 inhibitor‐dependent toxicity is the result of CDK1‐mediated cell death. The authors note that the phosphatase PP2A is an antagonist for CDK1 in addition to being a candidate USP7 substrate (Lecona et al, 2016; Wlodarchak & Xing, 2016), and thus, they turned their attention to elucidating the connection between USP7 and PP2A. Combining biochemical and immunofluorescence studies, Galarreta et al (2021) demonstrate that USP7 interacts with two subunits of PP2A, and this interaction increases in response to USP7 inhibition. Inhibiting USP7 furthermore triggers PP2A re‐localization from the cytoplasm to the nucleus and increases the phosphorylation levels of PP2A substrates, such as AKT and PRC1. DT‐061, a chemical activator of PP2A, reduces CDK1 phosphorylation events, suggesting that PP2A deregulation is a key mediator of USP7 inhibitor‐related toxicity. Using phosphoproteomics to analyze cells treated with a USP7 inhibitor or PP2A‐inhibiting okadaic acid, the authors reveal that both treatments share a significant number of altered phosphorylated targets—especially those related to mitosis, the cell cycle, and epitopes with a CDK‐dependent motif. Thus, the effects of USP7 inhibitors on CDK1 appear to be mediated through PP2A localization to the nucleus.These unexpected findings raise several questions that potentially impact the current view of cell cycle regulation. For example, how does USP7 regulate PP2A localization and is this important for reversing CDK1‐dependent phosphorylation of mitotic substrates prior to mitosis? Does PP2A accumulation in the nucleus explain the failure of USP7‐inhibited cells to enter mitosis despite cyclin B1 nuclear localization? A role for ubiquitin signaling as a regulator of CDK1 in interphase cells has not been reported, and accordingly, new investigations will be needed to unravel the mechanisms by which USP7 controls PP2A localization.Another important question that arises is whether or not CDK1 has sufficient basal activity to phosphorylate numerous mitotic proteins independent of cell cycle phase. The observation that USP7 and PP2A act to prevent the improper accumulation of CDK1‐dependent phosphorylation even in G1 phase cells suggests this to be the case. Alternatively, USP7 activity may be required to prevent abnormal pairing of CDK1 with a cyclin that is ubiquitously expressed across the cell cycle. If so, more research will be needed to uncover how ubiquitin signaling ensures CDK1 only pairs with cyclin A and cyclin B once they accumulate later in the cell cycle.Interestingly, USP7 inhibition also causes a rapid loss in DNA synthesis of S‐phase cells, prompting the authors to perform a time course experiment to decipher the order of events following treatment (i.e., does CDK1 activation precede or follow termination of DNA replication?). High‐throughput microscopy and flow cytometry analysis reveal an immediate reduction of DNA replication, an increase of CDK1 activity, and elevated DNA damage before a detectable increase in H3S10P. Long‐term exposure of USP7 inhibitors leads to DNA damage restricted only to cells with corresponding high levels of H3S10P and MPM2. Overall, these results illustrate how inhibition of USP7 activates CDK1, disrupting DNA replication and inducing DNA damage (Fig 1).Open in a separate windowFigure 1USP7 regulates CDK1In untreated cells, CDK1 is suppressed by USP7 and PP2A, and CDK1‐cyclin B is only active during the G2/M transition. In response to treatment, USP7 facilitates PP2A localization to the nucleus. This allows CDK1 to initiate premature mitotic activity throughout the cell cycle, resulting in increased DNA damage and cellular toxicity.The finding that USP7 inhibitors caused a rapid shutdown of DNA replication brings to mind the recent findings by several groups, that CDK1 activation occurs concomitantly with the S/G2 transition and that premature CDK1 activation in S‐phase terminates replication (Akopyan et al, 2014; Lemmens et al, 2018; Saldivar et al, 2018; Deng et al, 2019; Branigan et al, 2021). According to these studies, coupling of CDK1 activation to the S/G2 transition is regulated by ATR‐CHK1 signaling, a pathway activated by DNA replication to restrain CDK1 through Tyr‐15 phosphorylation. Galarreta et al''s observation that USP7 inhibition overrides ATR‐CHK1 (i.e., Tyr‐15 phosphorylation) highlights the fundamental importance of ubiquitin signaling, and potentially PP2A localization, for ensuring proper S‐to‐M progression and genome maintenance. Ultimately, the mechanistic details of Galarreta et al''s observations remain to be elucidated, and undoubtedly, their findings will inspire future investigations. Moreover, their discovery may lead to a new strategy targeting CDK1 to mitigate unwanted toxicities in the clinic.     

Glycolysis regulates Hedgehog signalling via the plasma membrane potential

The authors regret omitting the citation of a bioRxiv preprint study by preprint: Emmons‐Bell et al (2020), who independently discovered the role of ion channel‐dependent membrane depolarization for Smo membrane accumulation in the fly wing disc. This study used a different methodological approach and did not describe the mechanism of how membrane potential affects hedgehog signaling. The reference is herewith added.     

Molecular basis of cross‐species ACE2 interactions with SARS‐CoV‐2‐like viruses of pangolin origin

Correction to: The EMBO Journal (2021) 40: e107786. DOI 10.15252/embj.2021107786 | Published online 8 June 2021The authors would like to add three references to the paper: Starr et al and Zahradník et al also reported that the Q498H or Q498R mutation has enhanced binding affinity to ACE2; and Liu et al reported on the binding of bat coronavirus to ACE2.Starr et al and Zahradník et al have now been cited in the Discussion section, and the following sentence has been corrected from:“According to our data, the SARS‐CoV‐2 RBD with Q498H increases the binding strength to hACE2 by 5‐fold, suggesting the Q498H mutant is more ready to interact with human receptor than the wildtype and highlighting the necessity for more strict control of virus and virus‐infected animals”.to“Here, according to our data and two recently published papers, the SARS‐CoV‐2 RBD with Q498H or Q498R increases the binding strength to hACE2 (Starr et al, 2020; Zahradník et al, 2021), suggesting the mutant with Q498H or Q498R is more ready to interact with human receptor than the wild type and highlighting the necessity for more strict control of virus and virus‐infected animals”.The Liu et al citation has been added to the following sentence:“In another paper published by our group recently, RaTG13 RBD was found to bind to hACE2 with much lower binding affinity than SARS‐CoV‐2 though RaTG13 displays the highest whole‐genome sequence identity (96.2%) with the SARS‐CoV‐2 (Liu et al, 2021)”.Additionally, the authors have added the GISAID accession IDs to the sequence names of the SARS‐CoV‐2 in two human samples (Discussion section). To make identification unambiguous, the sequence names have been updated from “SA‐lsf‐27 and SA‐lsf‐37” to “GISAID accession ID: EPI_ISL_672581 and EPI_ISL_672589”.Lastly, the authors declare in the Materials and Methods section that all experiments employed SARS‐CoV‐2 pseudovirus in cultured cells. These experiments were performed in a BSL‐2‐level laboratory and approved by Science and Technology Conditions Platform Office, Institute of Microbiology, Chinese Academy of Sciences.These changes are herewith incorporated into the paper.     

The hSSB1 orthologue Obfc2b is essential for skeletogenesis but dispensable for the DNA damage response in vivo

We recently became aware of panel duplications in Supplementary Figures S6 and S7, due to pasting errors of similar flow cytometry images during figure preparation. This concerned the first two panels in the top row of Suppl. Fig S6A; second and third panel in the bottom row of Suppl. Fig S7B; and third and fourth panel in the bottom row of Suppl. Fig S7C.Furthermore, we noted a typographical error in Suppl. Fig S7B (top row, sixth plot), where the indicated percentage was wrongly given as 1.4%, instead of 1.1%. These errors did not change the results or the interpretation of the data. We deeply apologize to the scientific community for any confusion these errors may have caused. The updated appendix is published with this corrigendum.The original FlowJo analysis plots related to the affected figures are published as source data with this corrigendum. Please note that initial labelling of the experiments in these files referred to the official gene name Obfc2b informally as hSSB1, and Obfc2a‐shRNAs as ‘sh1’ and sh4’.Open in a separate windowFigure S6AOriginalOpen in a separate windowFigure S6ACorrectedOpen in a separate windowFigure S7BOriginalOpen in a separate windowFigure S7BCorrectedOpen in a separate windowFigure S7COriginalOpen in a separate windowFigure S7CCorrected     

Structural basis of transport and inhibition of the Plasmodium falciparum transporter PfFNT

In “Structural basis of transport and inhibition of the Plasmodium falciparum transporter PfFNT” by Lyu et al (2021), the authors depict the inhibitor MMV007839 in its hemiketal form in Fig 3A and F, Fig 4C, and Appendix Figs S10A, B and S13. We note that Golldack et al (2017) reported that the linear vinylogous acid tautomer of MMV007839 constitutes the binding and inhibitory entity of PfFNT. The authors are currently obtaining higher resolution cryo‐EM structural data of MMV007839‐bound PfFNT to ascertain which of the interconvertible isoforms is bound and the paper will be updated accordingly.     

Syncytia formation by SARS‐CoV‐2‐infected cells

In the supporting information of the article, the authors noticed that there was an error in Movie EV1. The right panel (SARS‐CoV‐2 + IFITM1) showed the same PI channel data (red) as the middle panel (SARS‐CoV‐2). This mistake occurred during the assembly of the merged movie file and does not change the interpretation of the data. A corrected version of the movie is herewith updated.     

Experts in science communication: A shift from neutral encyclopedia to equal participant in dialogue

Even if the predominant model of science communication with the public is now based on dialogue, many experts still adhere to the outdated deficit model of informing the public. Subject Categories: Genetics, Gene Therapy & Genetic Disease, S&S: History & Philosophy of Science, S&S: Ethics

During the past decades, public communication of science has undergone profound changes: from policy‐driven to policy‐informing, from promoting science to interpreting science, and from dissemination to interaction (Burgess, 2014). These shifts in communication paradigms have an impact on what is expected from scientists who engage in public communication: they should be seen as fellow citizens rather than experts whose task is to increase scientific literacy of the lay public. Many scientists engage in science communication, because they see this as their responsibility toward society (Loroño‐Leturiondo & Davies, 2018). Yet, a significant proportion of researchers still “view public engagement as an activity of talking to rather than with the public” (Hamlyn et al, 2015). The highly criticized “deficit model” that sees the role of experts as educating the public to mitigate skepticism still persists (Simis et al, 2016; Suldovsky, 2016).Indeed, a survey we conducted among experts in training seems to corroborate the persistence of the deficit model even among younger scientists. Based on these results and our own experience with organizing public dialogues about human germline gene editing (Box 1), we discuss the implications of this outdated science communication model and an alternative model of public engagement, that aims to align science with the needs and values of the public.Box 1

The DNA‐dialogue project

The Dutch DNA‐dialogue project invited citizens to discuss and form opinions about human germline gene editing. During 2019 and 2020, this project organized twenty‐seven dialogues with professionals, such as embryologists and midwives, and various lay audiences. Different scenarios of a world in 2039 (https://www.rathenau.nl/en/making‐perfect‐lives/discussing‐modification‐heritable‐dna‐embryos) served as the starting point. Participants expressed their initial reactions to these scenarios with emotion‐cards and thereby explored the values they themselves and other participants deemed important as they elaborated further. Starting each dialogue in this way provides a context that enables everyone to participate in dialogue about complex topics such as human germline gene editing and demonstrates that scientific knowledge should not be a prerequisite to participate.An important example of “different” relevant knowledge surfaced during a dialogue with children between 8 and 12 years in the Sophia Children’s Hospital in Rotterdam (Fig 1). Most adults in the DNA‐dialogues accepted human germline gene modification for severe genetic diseases, as they wished the best possible care and outcome for their children. The children at Sophia, however, stated that they would find it terrible if their parents had altered something about them before they had been born; their parents would not even have known them. Some children went so far to say they would no longer be themselves without their genetic condition, and that their condition had also given them experiences they would rather not have missed.Open in a separate windowFigure 1 Children participating in a DNA‐dialogue meeting. Photographed by Levien Willemse.     

Structural basis of transport and inhibition of the Plasmodium falciparum transporter PfFNT

Subject Categories: Membranes & Trafficking, Microbiology, Virology & Host Pathogen Interaction, Structural Biology

We recently reported the first structures of the Plasmodium falciparum transporter PfFNT, both in the absence and presence of the inhibitor MMV007839 (Lyu et al, 2021). These structures indicated that PfFNT assembles as a pentamer. The bound MMV007839 was found in the middle of the elongated channel formed by each PfFNT protomer, adjacent to residue G107. MMV007839 exists in two tautomeric forms and can adopt either a cyclic hemiketal‐like structure or a linear vinylogous acid conformation (Fig ​(Fig3A).3A). Unfortunately, these two tautomeric forms could not be clearly distinguished based on the existing cryo‐EM data at 2.78 Å resolution. The bound MMV007839 inhibitor was reported as the cyclic hemiketal‐like form in the structure in Figs ​Figs3A3A and ​andF,F, and ​and4C,4C, Appendix Figs S10A and B, and S13 and in the online synopsis image.Open in a separate windowFigure 3Cryo‐EM structure of the PfFNT‐MMV007839 complex

1. Chemical structure of MMV007839. The compound can either be in cyclic hemiketal‐like or linear vinylogous acid tautomeric forms.
2. Cryo‐EM density map of pentameric PfFNT viewed from the parasite’s cytoplasm. Densities of the five bound MMV007839 within the pentamer are colored red. The five protomers of pentameric PfFNT are colored yellow, slate, orange, purple, and gray.
3. Ribbon diagram of the 2.18‐Å resolution structure of pentameric PfFNT‐MMV007839 viewed from the parasite’s cytoplasm. The five protomers of pentameric PfFNT are colored yellow, slate, orange, purple, and gray.
4. Ribbon diagram of pentameric PfFNT‐MMV007839 viewed from the extracellular side of the parasite. The five protomers of pentameric PfFNT are colored yellow, slate, orange, purple, and gray.
5. Ribbon diagram of pentameric PfFNT‐MMV007839 viewed from the parasite’s membrane plane. The five protomers of pentameric PfFNT are colored yellow, slate, orange, purple, and gray. Densities of the five bound MMV007839 are depicted as red meshes.
6. The MMV007839‐binding site of PfFNT. The bound MMV007839 is colored green. Density of the bound MMV007839 is depicted as black mesh. Residues involved in forming the inhibitor binding site are colored yellow. The hydrogen bonds are highlighted with black dotted lines.

Open in a separate windowFigure 4Structure of the central channel in the PfFNT‐MMV007839 protomer

• CA cartoon of the central channel formed within a PfFNT protomer. The channel contains one constriction site in this conformational state. Residues forming the constriction and the K35‐D103‐N108 and K177‐E229‐N234 triads are illustrated as sticks. Residues F94, I97, and L104, which form the first constriction site in the apo‐PfFNT structure, are also included in the figure.

Eric Beitz alerted us to the findings reported by his group that the linear vinylogous acid tautomer of MMV007839 constitutes the binding and inhibitory entity of PfFNT (Golldack et al, 2017).     

The breakthrough paradox: How focusing on one form of innovation jeopardizes the advancement of science

Research needs a balance of risk‐taking in “breakthrough projects” and gradual progress. For building a sustainable knowledge base, it is indispensable to provide support for both. Subject Categories: Careers, Economics, Law & Politics, Science Policy & Publishing

Science is about venturing into the unknown to find unexpected insights and establish new knowledge. Increasingly, academic institutions and funding agencies such as the European Research Council (ERC) explicitly encourage and support scientists to foster risky and hopefully ground‐breaking research. Such incentives are important and have been greatly appreciated by the scientific community. However, the success of the ERC has had its downsides, as other actors in the funding ecosystem have adopted the ERC’s focus on “breakthrough science” and respective notions of scientific excellence. We argue that these tendencies are concerning since disruptive breakthrough innovation is not the only form of innovation in research. While continuous, gradual innovation is often taken for granted, it could become endangered in a research and funding ecosystem that places ever higher value on breakthrough science. This is problematic since, paradoxically, breakthrough potential in science builds on gradual innovation. If the value of gradual innovation is not better recognized, the potential for breakthrough innovation may well be stifled.

While continuous, gradual innovation is often taken for granted, it could become endangered in a research and funding ecosystem that places ever higher value on breakthrough science.

Concerns that the hypercompetitive dynamics of the current scientific system may impede rather than spur innovative research have been voiced for many years (Alberts et al, 2014). As performance indicators continue to play a central role for promotions and grants, researchers are under pressure to publish extensively, quickly, and preferably in high‐ranking journals (Burrows, 2012). These dynamics increase the risk of mental health issues among scientists (Jaremka et al, 2020), dis‐incentivise relevant and important work (Benedictus et al, 2016), decrease the quality of scientific papers (Sarewitz, 2016) and induce conservative and short‐term thinking rather than risk‐taking and original thinking required for scientific innovation (Alberts et al, 2014; Fochler et al, 2016). Against this background, strong incentives for fostering innovative and daring research are indispensable.