SUMMARY
We have isolated a mouse strain with a single missense mutation in the gene encoding MLKL, the essential effector of necroptotic cell death. The resulting substitution lies within the two-helix ‘brace’ and confers constitutive, RIPK3 independent, killing activity to MLKL. Mice homozygous for MlklD139V develop lethal inflammation within days of birth, implicating the salivary glands and pericardium as hotspots for necroptosis and inflammatory infiltration. The normal development of MlklD139V homozygotes until birth, and the absence of any overt phenotype in heterozygotes provides important in vivo precedent for the capacity of cells to clear activated MLKL. These observations offer an important insight into the potential disease-modulating roles of three common human MLKL polymorphisms that encode amino acid substitutions within or adjacent to the brace region. Compound heterozygosity of these variants is found at up to 12-fold the expected frequency in patients that suffer from a pediatric autoinflammatory disease, CRMO.
INTRODUCTION
Necroptosis is a form of programmed cell death associated with the production of pro-inflammatory cytokines, destruction of biological membranes and the release of intracellular Damage Associated Molecular Patterns (DAMPs) (Newton and Manning, 2016). Necroptosis depends on the activation of pseudokinase Mixed Lineage Kinase domain-Like (MLKL) by Receptor Interacting Protein Kinase 3 (RIPK3) (Murphy et al., 2013; Sun et al., 2012; Zhao et al., 2012). RIPK3-mediated phosphorylation of MLKL triggers a conformational change that facilitates the translocation to, and eventual irreversible disruption of, cellular membranes. While the precise biophysical mechanism of membrane disruption is still a matter of debate, it is consistently associated with the formation of an MLKL oligomer and the direct association of the four-helix bundle domain (4HB) of MLKL with membranes (Cai et al., 2014; Chen et al., 2014; Dondelinger et al., 2014; Hildebrand et al., 2014). In mouse cells, the expression of the murine MLKL 4HB domain alone (residues 1-125), 4HB plus brace helix (1-180), or the expression of phosphomimetic or other single site pseudokinase domain (PsKD) mutants is sufficient to induce membrane translocation, oligomerization and membrane destruction (Hildebrand et al., 2014; Murphy et al., 2013). While capable of disrupting synthetic liposomes when produced recombinantly, similarly truncated and equivalent single site (PsKD) mutant forms of human MLKL do not robustly induce membrane associated oligomerization and cell death without forced dimerization (Petrie et al., 2018; Quarato et al., 2016; Tanzer et al., 2016). Furthermore, both mouse and human MLKL mutants have been reported that have the capacity to form membrane associated oligomers, but fail to cause irreversible membrane disruption and cell death (Hildebrand et al., 2014; Petrie et al., 2018). Recent studies have revealed that necroptosis downstream of MLKL phosphorylation and membrane association can be modulated by processes that utilize the Endosomal Sorting Complex Required for Transport (ESCRT) family of proteins. One model proposes a role for ESCRT in limiting necroptosis via plasma membrane excision and repair (Gong et al., 2017) while other models limit plasma membrane disruption by ESCRT-mediated endosomal trafficking and the release of MLKL in endosomes (Yoon et al., 2017) or the shedding of phosphorylated MLKL in extracellular vesicles (Zargarian et al., 2017).
In mice, the absence of MLKL does not appear to have obvious deleterious developmental or homeostatic effects (Murphy et al., 2013; Wu et al., 2013). However, genetic deletion of Fadd, Casp8 or Ripk1, leads to inappropriate activation of MLKL and ensuing necroptosis during embryogenesis and is incompatible with life beyond embryonic day (E)10.5, E10.5 and 1-3 days post-natally, respectively (Dillon et al., 2014; Kaiser et al., 2014; Kelliher et al., 1998; Rickard et al., 2014b; Varfolomeev et al., 1998; Yeh et al., 1998; Zhang et al., 2011). Exploring the precise physiological consequences of inappropriate MLKL activation in these scenarios is complicated by the fact that FADD, Caspase-8 and RIPK1 also play important roles in cellular processes other than modulation of MLKL-induced necroptotic cell death (Alvarez-Diaz et al., 2016; Kaiser et al., 2011; Kang et al., 2004; Newton et al., 1998; Oberst et al., 2011; Rickard et al., 2014b).
Aberrant levels of MLKL-dependent cell death contribute to disease in several genetic and experimental mouse models (Anderton et al., 2017; Dannappel et al., 2014; Hockendorf et al., 2016; Newton et al., 2016; Rickard et al., 2014a; Rickard et al., 2014b). In humans, MLKL mRNA and protein levels are positively correlated with survival of patients with pancreatic adenocarcinoma, cervical-, gastric-, ovarian- and colon-cancers (reviewed by (Lalaoui and Brumatti, 2017)). Interestingly, high levels of phosphorylated MLKL are associated with reduced survival in esophageal and colon cancer patients (Liu et al., 2016b). Two missense MLKL somatic mutations identified in human cancer tissue have been found to confer a reduction in necroptotic function in cell-based assays (Murphy et al., 2013; Petrie et al., 2018). One recent study reported a significant enrichment of an ultra rare MLKL stop-gain gene variant p.Q48X in Hong Kong Chinese patients suffering from a form of Alzheimer’s disease (Wang et al., 2018) however more common germline MLKL gene variants are only weakly associated with human disease in GWAS databases. In two recent studies, lethal immunodeficiency, arthritis and intestinal inflammation was reported in patients homozygous for ultra rare-loss of function RIPK1 mutations (Cuchet-Lourenco et al., 2018; Li et al., 2019), however to date, MLKL gene variants have not been directly implicated in any severe Mendelian forms of human disease.
We have identified a single base pair germline mutation of mouse Mlkl that encodes a missense alteration to the MLKL mouse brace region and confers constitutive activation independent of upstream necroptotic stimuli. Given this mutant Mlkl allele is subject to the same developmental and environmental controls on gene expression as wildtype Mlkl, the postnatal lethality in these mice provides novel insight into the physiological and pathological consequences of dysregulated necroptosis. In parallel these findings inform the potential functional significance of three common human MLKL polymorphisms that encode non-conservative amino acid substitutions within, or in close proximity to, the brace helix that is mutated in the MlklD139V mouse.
RESULTS
Generation of a constitutively active form of MLKL
An ENU mutagenesis screen was performed to identify mutations that ameliorate thrombocytopenia in Mpl−/− mice (Kauppi et al., 2008). A G1 founder, designated Plt15, had a modestly elevated platelet count of 189×106/mL compared to the mean for Mpl−/− animals (113±57×106/mL) and yielded 19 Mpl−/− progeny. Ten of these mice had platelet counts over 200×106/mL, consistent with segregation of a dominantly acting mutation (Fig. 1A). Linkage analysis and sequencing (see Experimental Procedures) identified an A to T transversion in Mlkl that was heterozygous in all mice with an elevated platelet count (Fig. 1B). The MlklPlt15 mutation results in a non-conservative aspartic acid-to-valine substitution at position 139. In the full length mMLKL structure D139 forms a salt bridge with an arginine residue at position 30 (α2 helix) of the MLKL four-helix bundle (4HB) domain (Murphy et al, 2013) (Fig. 1C). This salt bridge represents one of a series of electrostatic interactions between residues in helix α2 of the MLKL 4HB domain and the two-helix ‘brace’ region. D139 of mouse MLKL is conserved in all MLKL orthologues in vertebrata reported to date (Fig. 1D). We have shown that the exogenous expression of the 4HB domain of murine MLKL alone is sufficient to kill mouse fibroblasts whereas exogenous expression of full length MLKL does not, indicating that this ‘electrostatic zipper’ may play an important role in suppressing the killing activity of the MLKL 4HB (Hildebrand et al., 2014). To determine if MLKLD139V exhibited altered ability to induce necroptotic cell death relative to MLKLWt, we stably expressed these full length proteins under the control of a doxycycline-inducible promoter in immortalized mouse dermal fibroblasts (MDF) isolated from Wt, Mlkl−/−, Ripk3−/− or Ripk3−/−;Casp8−/− mice. While expressed at comparable levels, MLKLD139V induced markedly more death than MLKLWt, on each of the genetic backgrounds tested (Fig. 1E-F, Supp. Fig. 1A). This indicates that MLKLD139V is a constitutively active form of MLKL, capable of inducing necroptotic cell death independent of upstream signaling and phosphorylation by its activator RIPK3. Consistent with this interpretation, exogenous expression of MLKLD139V in Ripk3−/−;Casp8−/− MDFs was sufficient to induce the organelle swelling and plasma membrane rupture characteristic of TNF induced necroptosis when examined by Transmission Electron Microscopy (Fig. 1G).
Constitutively active mouse MLKL causes a lethal perinatal inflammatory syndrome
To define the phenotypic consequences of constitutively active MLKL in the absence of any confounding effects resulting from Mpl-deficiency, all subsequent studies were performed on a Mpl+/+ background. Homozygous MlklD139V/D139V pups were born at expected Mendelian frequencies (Supp. Table I) and were ostensibly normal macroscopically and histologically at E19.5 (Supp. Fig. 2A-D). However, by 3 days of age, although outwardly indistinguishable from littermates (Fig. 2A), they exhibited reduced body weight (Supp. Fig. 2B) and failed to thrive, with a maximum observed lifespan of 6 days under conventional clean housing conditions. Like MlklWt/D139V mice, Mlklnull/D139V compound heterozygotes were present at the expected frequency at P21 and developed normally to adulthood (Supp. Table II). Thus, the constitutive activity of MLKLD139V was not affected by the presence of normal MLKL protein suggesting it is the absolute allelic dose of MlklD139V that determines perinatal lethality. To confirm that the phenotype of the ENU derived MlklD139V mice was due to the MlklD139V missense mutation, we independently generated MlklD139V mice using CRISPR-Cas9 genomic editing. Homozygote CRISPR-MlklD139V/D139V mice also died soon after birth (Supp. Table III).
Hematoxylin-Eosin stained-sections from both P2 and P3 MlklD139V/D139V pups revealed multifocal acute inflammation characterized by neutrophilic infiltration, dilated blood vessels and edema (Fig. 2B) in the dermis and subcutis of the head and neck. These inflammatory features were not observed in MlklWt/Wt or MlklWt/D139V littermates, nor in Mlkl−/− mice of the same age (Supp. Fig. 2I). Cells of hematopoietic origin, revealed by immunohistochemical staining for CD45, were sparsely distributed throughout the lower head and neck and confined predominantly to a clearly delineated developing lymph node in MlklWt/Wt and MlklWt/D139V littermates (Fig. 2C). In contrast, CD45+ cells were more numerous and distributed throughout the cutis, subcutis and salivary glands of MlklD139V/D139V pups (Fig. 2C). A mixture of diffuse and focal inflammatory infiltration was also observed within the mediastinum and pericardial space of all P2/P3 MlklD139V/D139V pups examined, as was a marked paucity of thymic cortical lymphocytes (Fig. 2D, Supp. Fig 2E), phenotypes not evident in E19.5 embryos (Supp. Fig. 2D). Apart from small foci of hepatocyte and enterocyte loss/necrosis evident in the livers and small intestines of some MlklD139V/D139V pups examined (data not shown), no other lesions were observed by histopathology. Consistent with this inflammatory phenotype significantly elevated levels of several pro-inflammatory cytokines and chemokines were evident in the plasma of both E19.5 and P3 MlklD139V/D139V pups (Fig. 2E, F). Blood glucose levels were normal (Supp. Fig. 2 F, G).
Hematopoetic defects in MlklD139V mice
Although blood cell numbers were unchanged in MlklD139V/D139V pups at E19.5 relative to MlklWt/Wt and MlklWt/D139V littermates, by P3 significant deficits were evident in total white blood cell count, lymphocyte and platelet numbers (Fig. 3A-C, Supp. Fig. 3A). Similarly, the numbers of hematopoietic stem and progenitor cells were present at normal proportions in fetal livers of E18.5 MlklD139V/D139V pups, although increased levels of intracellular ROS were uniformly evident (Fig. 3D-E, Supp. Fig. 3B). By P2, deficits in CD150+CD48+ and CD150+CD48− populations were present (Fig. 3F), accompanied by increased AnnexinV binding (which indicates either phosphatidyl serine exposure or plasma membrane rupture) in all lineages (Fig. 3G). In adult MlklWt/D139V mice, numbers of hematopoietic stem and progenitor cells were unaffected (Fig. 3H); however, upon myelosuppressive irradiation, recovery of hematopoietic cell numbers was delayed and characterized by increased expression of ROS and Annexin V (Supp. Fig. 3C, D). When challenged with the cytotoxic drug 5-fluorouracil (5-FU), blood cell recovery in MlklWt/D139V mice was similarly delayed (Fig. 3I). In competitive transplants in which test MlklWt/D139V or MlklWt/Wt marrow was co-injected with wild type competitor marrow in 10:1 excess, as expected, MlklWt/Wt marrow contributed to 90% of recipient blood cells 8 weeks after transplantation and maintained that level of contribution for 6 months (Fig. 3J). In contrast, MlklWt/D139V marrow performed poorly, contributing to 25% and 51% of recipient blood cells at these times (Fig. 3J). Similarly, while wild type fetal liver cells contributed to the vast majority of blood cells in irradiated recipients up to 6 months after transplantation, cells from MlklD139V/D139V embryos failed to compete effectively during this period (Fig. 3K). Heterozygote MlklWt/D139V fetal liver cells contributed poorly in the first month following the graft but recovered to contribute more after six months (Fig. 3K). Thus, while tolerated under steady-state conditions, heterozygosity of MlklD139V is deleterious under conditions of hematopoietic stress. Bone marrow-derived HSCs from MlklWt/D139V adults and fetal liver-derived HSCs from MlklWt/D139V and MlklD139V/D139V pups also formed fewer and smaller colonies in the spleens of lethally irradiated recipient mice after 8 days (Supp. Fig. 3E).
Homozygous MlklD139V fibroblasts are less sensitive to necroptotic stimuli and have low levels of MLKL protein
To examine if the constitutive activity of exogenously expressed MLKLD139V results in an enhanced propensity for necroptosis in cells that express MLKLD139V under the control of its endogenous promoter, we immortalized MDFs from MlklWt/Wt, MlklWt/D139V and MlklD139V/D139V littermates and from Mlkl−/− E19.5 pups. As expected, we observed no significant difference in the sensitivity of these cells to an apoptotic stimulus such as TNF plus Smac mimetic (Fig. 4A). However we observed a significant and consistent decrease in sensitivity to TNF induced necroptosis using three different pan-caspase inhibitors Q-VD-OPh, Z-VAD-fmk and IDUN-6556 in a MlklD139V dose dependent manner (Fig. 4A). While MDFs isolated from MlklD139V/D139V homozygotes were up to 60% less sensitive to TNF-induced necroptosis compared to MlklWt/Wt MDFs, they were not completely resistant like Mlkl−/− MDFs (Fig. 4A).
Surprisingly, while there were no obvious differences in the levels of MLKLWT and MLKLD139V protein following inducible exogenous expression (Fig. 1F), MLKL was virtually undetectable by Western blot in MlklD139V/D139V cells (Fig. 4B). There was, however, no significant reduction in Mlkl transcript levels in these cells suggesting that this reduction was post-transcriptionally regulated (Supp. Fig. 4A). The reduction in MLKLD139V protein levels was also evident in whole body protein lysates prepared from E14 embryos (Supp. Fig. 4B). Lysates from E14 embryos also clearly show that MlklWt/D139V heterozygotes have intermediate levels of MLKL, reflecting the intermediate sensitivity of MlklWt/D139V MDFs to necroptotic stimuli (Fig. 4A).
MLKLD139V and RIPK3-phosphorylated wildtype MLKL is turned over in a proteasome and lysosome dependent manner
Measuring the half-life of endogenously expressed MLKLD139V is not possible using conventional ‘pulse chase’ methods because this mutant protein induces necroptotic cell death, so we capitalized on our previous observation that an N-terminally FLAG-tagged MLKL 4HB forms a high molecular weight membrane-associated complex just like the untagged form, but, unlike the untagged version, does not kill cells (Hildebrand et al., 2014). Consistent with this observation, N-FLAG full-length mouse MLKL was phosphorylated by RIPK3 following stimulation with TSI, and formed a high molecular weight membrane associated complex, but did not induce cell death when inducibly expressed in Mlkl−/− MDFs (data not shown).
Using this system we were able to measure the half-life of MLKL by inducing N-FLAG-MLKLWT or N-FLAG-MLKLD139V expression in Mlkl−/− MDFs for 15 hours in doxycycline then washing and culturing them in the absence of doxycycline for a further 2-24 hours. In the absence of a stimulus (UT), the levels of N-FLAG-MLKLWT remained consistent over the 24-hour period (Fig. 4C), indicating that wild type MLKL is a stable protein in MDFs. However, when these cells were treated with a necroptotic stimulus (TSI) the levels of wild type MLKL rapidly declined even though these cells were unable to undergo a necroptotic cell death. This indicates that RIPK3 induced phosphorylation, oligomerization or translocation to the membrane induces turnover of MLKL in a cell death independent manner. Consistent with the fact that untagged MLKLD139V behaves as an auto-activated form of MLKL (Fig. 1E), the half-life of N-FLAG-MLKLD139V(4-6 hours) was similar to the WT version stimulated with TSI (Fig. 4C). Thus, the absence of endogenously expressed MLKLD139V in E14 embryo lysates and cultured fibroblasts can be attributed to the reduced post-translational stability of this mutant auto-activated form of the protein.
To determine which cellular mechanism(s) are required for the clearance of activated MLKL, we included a series of proteasome, lysosome and specific protease inhibitors during the ‘chase’ period after doxycycline was withdrawn (schematic in Fig. 4D). The doses of these inhibitors were carefully titrated to minimize apoptotic cell death during the assay (Supp. Fig. 4C). Nevertheless, even at the very low doses used, the proteasome inhibitor PS341 reduced the clearance of TSI stimulated N-FLAG-MLKLWT (Fig. 4D). This protection was particularly evident when specifically probing for phospho(p)-MLKL. Chloroquine, Bafilomycin and NH4Cl also partially protected against p-MLKL clearance (Fig. 4D). These agents have multifaceted actions, but interfere with the processes of lysosomal acidification and/or the fusion of autophagosomes/endosomes with lysosomes and thus prevent protein degradation by lysosomal proteases. Loss of total N-FLAG-MLKLD139V was also prevented by PS341, however it was not possible to probe for p-MLKL as this activated form of MLKL is not phosphorylated in this assay due to the absence of TSI stimulation (Fig. 4E).
The reduced half-life of activated MLKL supports recent findings by others that mechanisms exist for the clearance of activated forms of MLKL (Gong et al., 2017; Yoon et al., 2017; Zargarian et al., 2017). Based on these findings we hypothesized that this MLKL-clearance mechanism limits the capacity of MLKLD139V to kill MlklD139V hetero and homozygote cells in culture and in vivo by maintaining protein levels below a critical threshold. To test whether this protective mechanism could be overwhelmed, we incubated MDFs with agents that have been shown to induce Mlkl expression (TNF, interferons (IFN) β and γ) (Rodriguez et al., 2016; Rusinova et al., 2013; Tanzer et al., 2017; Thapa et al., 2013), or inhibit its turnover (proteasome and lysosome inhibitors). MLKLD139V protein in untreated MlklD139V/D139V MDFs was undetectable by Western blot but became faintly detectable following stimulation with such stimuli (Fig. 4B & Supp. Fig. 4D). This correlates with moderate but statistically significant increases in cell death (particularly when compared with the lack of sensitivity to conventional necroptotic stimuli (Fig. 4A)), when exposed to IFNβ alone and in combination with proteasome or lysosome inhibitors (Fig. 4F). An allele-dose dependent sensitivity is also evident in primary MDFs (Supp. Fig. 4E). Together, these experiments provide evidence for the existence of steady-state MLKL surveillance and turn-over mechanisms that suppress cell death by lowering the abundance of activated MLKL below a killer threshold – both at the cellular and whole animal level.
Interestingly, genetic deletion of Tnfr1, Myd88 and Ifnar did not provide any extension to the lifespan of MlklD139V homozygote pups (Table I), indicating that the removal of any one of these routes to NF-κB- and interferon-mediated gene upregulation is not sufficient to protect against a double allelic dose of MlklD139V. Similarly, combined genetic deletion of Casp8 and Ripk3 did not rescue or extend the life of MlklD139V/D139V mice, indicating that post-natal death is not mediated by bystander extrinsic apoptotic cell death that may occur secondary to initial waves of MLKLD139V-mediated necroptosis and associated inflammatory cytokine release (Table I). To test whether the death of MlklD139V/D139V neonates was mediated by activation of the inflammasome we also crossed this line with the Caspase 1/11 null mouse strain (Kuida et al., 1995; Li et al., 1995). This did not enhance the lifespan of MlklD139V/D139V pups (Table I).
Three of the four most frequent missense gene variants in human MLKL encode amino acid substitutions within or immediately adjacent to the brace region
Given the severe inflammatory phenotype of murine MlklD139V/D139V neonates and the significant defects in stress hematopoiesis observed in murine MlklWt/D139V adults, we explored the prevalence of brace region variation in human MLKL. Examination of the gnomAD database (Lek et al., 2016), which contains human MLKL exome or genome sequence data from a total of over 141,456 individuals revealed that the second and third highest frequency human MLKL missense coding variants; rs34515646 (R146Q) and rs35589326 (S132P), alter the same brace helix (Table II, Fig. 5A). The 4th most common human MLKL polymorphism, rs144526386 (G202*V) is a missense polymorphism identified exclusively in the context of a shorter splice isoform of MLKL (*) named ‘MLKL2’ (Arnez et al., 2015) (Table II, Fig. 5B). The full length canonical transcript of MLKL encodes a 471 amino acid protein, while alternatively spliced MLKL2 encodes an isoform of MLKL that is 263 amino acids long and is missing a large portion of the pseudokinase domain which functions to repress the killing potential of the 4HB domain (Cai et al., 2014; Chen et al., 2014; Dondelinger et al., 2014; Hildebrand et al., 2014) and recruit co-effectors like RIPK3 and HSP90 ((Jacobsen et al., 2016; Petrie et al., 2018). Glycine202* is encoded by an extension to exon 9 that is unique to the MLKL2 splice isoform (Fig. 5A, B).
While the amino acid substitution MLKLR146Q is classified as ‘tolerated’ and ‘benign’ by SIFT/POLYPHEN 2 algorithms (Adzhubei et al., 2013; Sim et al., 2012) (Supp. Table IV), R146 of human MLKL shows NMR chemical shift perturbations in the presence of the negatively charged phospholipids IP3 and IP6, indicating a possible role in membrane association and disruption (Dovey et al., 2018; Quarato et al., 2016). Ser-132 lies at the intersection of a dynamic disordered loop and the first structured residue of the conserved brace helix 1 (Fig. 5A) (Murphy et al., 2013; Petrie et al., 2018; Su et al., 2014). A Serine-to-Proline substitution at this position is predicted to significantly impact the conformation of the immediately adjacent W133 (brace helix) and in turn, the closely situated W109 (4 helix bundle) (Supp. Fig. 5A). When mapped to a model of MLKL splice-isoform 2 (Arnez et al., 2015) Glycine 202* is predicted to be on an isoform 2-specific helix and to form an interface along with S132 and R146 of brace helix 1. While the precise structural consequence of these three brace polymorphisms is unknown, modelling of human MLKL predicts that disruption in the brace region favours adoption of an activated conformation (Petrie et al, 2018). Consistent with this prediction, the murine equivalent of the human S132P variant, mMLKLS131P, formed high molecular weight membrane-associated complexes and killed MDFs in the absence of a necroptotic stimulus (Fig. 5 C, D) when expressed at close to endogenous levels (Supp. Fig. 5B).
MLKL brace helix variants appear in trans at a higher frequency in a cohort of CRMO patients than in healthy controls
To investigate if human MLKL brace region polymorphisms play a role in human autoinflammatory disease we examined their frequency in cohorts suffering from Ankylosing Spondylitis (AS), chronic recurrent multifocal osteomyelitis (CRMO), Guillain Barre Syndrome (GBS) and Synovitis, Acne, Pustulosis, Hyperostosis and Osteitis (SAPHO) Syndrome. The individual minor allele frequencies of R146Q, S132P and G*202V are not enriched in these disease cohorts relative to healthy controls when population distribution is accounted for (Supp. Tables IV and V). However these alleles occur in trans (making ‘compound heterozygotes’ – schematic in Fig. 5E) in 3 out of 128 CRMO patients. This is 29 times the frequency that these combinations are observed in healthy NIH 1000 genomes samples (where there are only 2 compound heterozygotes for these polymorphisms out of 2504 healthy individuals sequenced), or at 10-12 times the frequency when only European CRMO patients and two separate healthy European control populations were compared (Table III).
DISCUSSION
In contrast to apoptosis, necroptosis is widely held to be an inflammatory form of cell death. However, definitive evidence for this proposition has yet to emerge. Because MLKL is activated by inflammatory stimuli such as TNF it is very difficult to separate cause from effect. The identification of an auto-activating mutant of MLKL (MlklD139V) in mice has allowed us to explore the consequences of inappropriate necroptosis in the absence of such confounding factors. Furthermore it has led to significant insights into developmental processes sensitive to MLKL activation and into physiological mechanisms that exist to neutralize activated MLKL. These turnover mechanisms critically control cell fate, determining whether auto-active MLKLD139V is present at a sufficient level to promote cell death.
While MLKL phosphorylation might serve as an immuno-histochemical marker for necroptosis ordinarily, in the MlklD139V mice it is not possible to pinpoint exactly which cell type/s undergo necroptosis. Nevertheless, the presence of high levels of circulating pro-inflammatory cytokines in MlklD139V/D139V pups at E19.5 relative to MlklWt/Wt and MlklWt/D139V littermates suggests that necroptosis and ensuing inflammation occurs in the sterile in utero environment. This is not enough to overtly retard prenatal development or affect hematopoietic cell populations other than by moderately reducing circulating platelet levels. However, upon birth and/or exposure to the outside environment the capacity of homozygous MlklD139V/D139V pups to suppress MLKLD139 Vactivity appears overwhelmed and they die within days of birth. This is clearly a dose-dependent effect because both MlklD139V/Wt and MlklD139V/null heterozygous mice are viable. We therefore speculate that transcriptional upregulation of MlklD139V overwhelms the turnover and/or membrane repair mechanisms that counteract MLKL activation (Gong et al., 2017; Yoon et al., 2017). Post-natal death cannot be prevented by combined deficiencies in Ripk3 and Casp8 or indeed deficiency of any other inflammatory gene that we tested, including Tnfr1, Myd88 or Ifnar. This further supports the idea that excessive MLKL-induced necroptosis can generate an inflammatory response in the absence of other inflammatory mediators. Difficulty with suckling due to inflammatory infiltration of the head and neck and resulting failure to thrive is one possible explanation for the lethality. However, the narrow window of mortality for MlklD139V/D139V pups and marked pericardial immune infiltration make heart failure another potential cause of sudden neonatal death.
One of the most unexpected findings from our study is the physiological importance of endogenous mechanisms that limit the ability of activated MLKL to kill cells. While others have recently shown that an ESCRT dependent repair mechanism can help protect membranes from limited MLKL damage it was not feasible to demonstrate the physiological relevance of this finding (Gong et al., 2017; Yoon et al., 2017). Our data suggest both proteasomal and lysosomal mechanisms also exist to dispose of activated MLKL. While proteasomal degradation is usually considered to be cytoplasmic and completely separate from lysosomal degradation, it was notable that low doses of either the proteasome inhibitor, PS341, or chloroquine (that inhibits lysosome acidification) limited p-MLKL degradation to very similar extents. This creates the possibility that these mechanisms or the previously described ESCRT mechanism intersect. Finally, the ability of these mechanisms to hold heterozygous levels of active MLKL in check without deleterious consequences in vivo supports the idea that direct inhibition of activated MLKL may be an effective means to therapeutically prevent unwanted necroptotic cell death.
The MlklD139V brace mutant mouse strain may be a useful model to study the role of necroptosis in human health and disease. According to current allele frequencies in gnomAD, up to 8% of individuals globally are heterozygous for missense MLKL gene variants within the brace-coding region. This percentage of people with brace variants indicates that this region is highly tolerant to missense mutation (Fig. 5F, Supp. Fig. 5C). High tolerance to missense variation in a coding sequence is often used to filter out potential pathogenic variants in human genetic studies because it indicates that such variations are likely to be functionally neutral (Traynelis et al., 2017). However, the first brace helix is both highly evolutionarily conserved yet also tolerant of missense mutations in the human population (Fig. 1D, Fig. 5F,G). Furthermore in vivo and in vitro data show that amino acid substitutions in the brace region have profound effects on MLKL function (Davies et al., 2018; Quarato et al., 2016). Therefore, overlayed with structural, biochemical, cell and animal-based evidence of function, it is tempting to speculate that these human MLKL brace region variants have accumulated not simply by chance, but through positive evolutionary selection. While defective emergency hematopoiesis is likely to be subject to negative evolutionary selection, MlklD139V mouse-derived HSCs are only defective following chemo- or radio-ablation. Given that these forms of HSC depletion are unlikely to have been a significant selective force during human evolution, we speculate that these human brace polymorphisms have achieved high frequencies in the human population because they have conferred a selective advantage to infectious disease. Evidence for positive selection has been found for over 300 immune-related gene loci and many of these have been found to be associated with the incidence of autoimmune and autoinflammatory disease in modern humans (Gutierrez-Arcelus et al., 2016; Ramos et al., 2015). Many of these variants have also been mechanistically linked to defense against a particular pathogen (Karlsson et al., 2014; Ramos et al., 2015). While increased numbers and examination of independent cohorts will be required to confirm the statistical enrichment of human MLKL brace variants occurring in trans in CRMO, this patient cohort offers a tantalizing first clue into their potential as modifiers of complex/polygenic inflammatory disease.
EXPERIMENTAL PROCEDURES
Mice
All mice were backcrossed to C57BL/6 mice for >10 generations or generated on a C57BL/6J background. Mlkl−/-, Tnfr1−/−, Myd88−/−, IFNAR1−/−, Ripk3−/−, Casp8−/− and Casp1/Casp11−/− mice were generated as described (Adachi et al., 1998; Beisner et al., 2005; Hwang et al., 1995; Kuida et al., 1995; Li et al., 1995; Murphy et al., 2013; Newton et al., 2004; Peschon et al., 1998). Mice designated as E19.5 were obtained by Caesarean section from mothers that received progesterone injections at E17.5 and E18.5. An independent mouse strain that carried the D139V mutation in the Mlkl gene (MLKLD139V CRISPR) was generated using CRISPR/Cas9 as previously described (Wang et al., 2013). Briefly, one sgRNA of the sequence GGAAGATCGACAGGATGCAG (10ng/μl), an oligo donor of the sequence ATTGGAATACCGTTTCAGATGTCAGCCAGCCAGCATCCTGGCAGCAGGAAGATCGA CAGGTTGCAGAAGAAGACGGgtgagtctcccaaagactgggaaagagtaggccagggttgggggtagggtgg (10ng/uL) and Cas9 mRNA (5ng/μL) were injected into the cytosol of C57BL/6J zygotes. Mice were sequenced across the mutated region to confirm incorporation of the altered codon and analysis was performed after at least 2 back-crosses to C57BL/6. The relevant Animal Ethics Committee approved all experiments.
Linkage analysis
We mapped the chromosomal location of the Plt15 mutation by mating affected mice to 129/Sv Mpl−/− mice to produce N2 (backcross) and F2 (intercross) generations. A genome wide scan using 20 N2 mice with the highest platelet counts (287±74×106/ml, compared with 133±75×106/ml for the overall population, Fig. 1A) localized the mutation to a region of chromosome 8 between D8Mit242 and D8Mit139 and linkage to this region was then refined. Analysis of the F2 population revealed a significant reduction in the frequency of mice homozygous for C57BL/6 alleles in this interval (e.g. D8Mit200 3/81 F2 mice homozygous C57BL/6, p=2.2×10−5 χ2-test), suggesting the Plt15 mutation results in recessive lethality. The refined 2.01 Mb interval contained 31 annotated genes, only five of which appeared to be expressed both in the hematopoietic system and during embryogenesis (http://biogps.gnf.org/): Dead box proteins 19a and 19b (Ddx19a and Ddx19b), Ring finger and WD repeat domain 3 (Rfwd3), Mixed lineage kinase domain like (Mlkl), and WD40 repeat domain 59 (Wdr59). Sequencing identified a single mutation, an A to T transversion in Mlkl that was heterozygous in all mice with an elevated platelet count.
Reagents
Antibodies
Rat-anti mRIPK3 and rat anti-mMLKL 8F6 (selected for affinity to residues 1-30 of mouse MLKL) and rat anti-MLKL 3H1 (MLKL brace region) were produced in-house. Anti-Pro Caspase 8 (#4927) and GAPDH (#2113) were purchased from Cell Signaling Technology. Anti-mouse P-MLKL (ab196436) and anti-Actin (ab5694) were purchased from Abcam. Anti-VDAC (AB10527) was purchased from Millipore. FC-hTNF was produced in house and used at a final concentration of 100ng/mL. Recombinant mouse IFNγ and β were purchased from R&D Systems (Minneapolis, MN, USA) Q-VD-OPh and ZVAD were purchased from MP Biomedicals (Seven Hills, NSW, Australia). Smac mimetic also known as Compound A, and the caspase inhibitor IDN-6556 were a gift from TetraLogic (Malvern, PA, USA). Propidium iodide, doxycycline, and bafilomycin were purchased from Sigma-Aldrich (Castle Hill, NSW, Australia).
Cell line generation and culture
Primary mouse dermal fibroblasts were prepared from skin taken from the head and body of E19.5 pups delivered by C-section or from the tails of adult mice as described (Etemadi et al., 2013). Primary MDFs were immortalized by stable lentiviral transduction with SV40 large T antigen. Immortalized MDFs were stably transduced with exogenous mouse and human MLKL cloned into the pFTRE 3G vector, which was generated by Toru Okamoto, and allows doxycycline-inducible expression as described (Murphy et al., 2013). Cells were maintained in culture as previously described (Tanzer et al., 2017).
Cell death assays
Cell death assays were performed as described previously using 5 × 104 MDFs per well in 24 well tissue culture plates (Murphy et al., 2013). Doxycycline (20 ng/mL) was added together with death stimuli. Fc-hTNF was produced in house and used at 100ng/mL, Compound A Smac mimetic and IDN6556 were used at 500 nM and 5 μM respectively. ZVAD and QVD-OpH were used at 25 μM and 10 μM respectively. Mouse and human interferons gamma and beta were used at 30 ng/mL, PS341 and MG132 at 2 nM and 200 nM respectively and Bafilomycin at 300 nM.
MLKL turn-over assays
5 × 104 MDFs per well were plated in 24 well tissue culture plates and allowed to settle. Doxycycline (20 ng/mL) +/-TNF, Smac Mimetic and IDN6556 was added. After 15 hr, ‘no dox’ and ‘0’ wells were harvested. Media was removed from remaining wells and cells were washed with PBS and fresh media containing IDN6556 was re-added. Wells were then harvested 2, 4, 6, 8 and 24 hours from this point. Cells were harvested by direct lysis in reducing SDS-PAGE lysis buffer.
MLKL protection assays
5 × 104 MDFs per well were plated in 24 well tissue culture plates and allowed to settle. Doxycycline (20 ng/mL) was added. After 18 hrs, ‘no dox’ and ‘T0’ samples were harvested. Media was removed and cells washed before addition of fresh media containing TSI or IDN alone for 3 hrs. Cells were washed again and media restored with IDN6556 alone (UT), or IDN6556 + inhibitor (MG132 (200 nM), PS341 (10-40 nM), Choloroquin (50 μM), Bafilomycin (300 nM), Ca-074 Me (20 μM), TLCK (100 μM) and AEBSF (100 μM)) for a further 21 hours. Cells were harvested by direct lysis in reducing SDS-PAGE lysis buffer.
Transmission Electron Microscopy
Murine dermal fibroblasts prepared from mice of the indicated genotypes were untreated or stimulated with the indicated agents for the indicated hours. Then, cells were fixed with 2% glutaraldehyde in 0.1 M phosphate buffer, pH 7.4, postfixed with 2% OsO4, dehydrated in ethanol, and embedded in Epok 812 (Okenshoji Co.). Ultrathin sections were cut with an ultramicrotome (ultracut N or UC6: Leica), stained with uranyl acetate and lead citrate, and examined with a JEOL JEM-1400 electron microscope. The viability of a portion of these cells was determined by measuring LDH release as described previously (Murai et al., 2018).
Mouse histopathology
Caesarian-sectioned E19.5 and Day P2/3 pups were euthanized by decapitation and fixed in 10% buffered formalin. 5 μm coronal sections were taken at 200 μm intervals for the full thickness of the head, 5 μm sagittal sections were taken at 300 μm intervals for the full thickness of the body. A thorough examination of these sections was performed by histopathologists Aira Nuguid and Tina Cardamome at the Australian Phenomics Network, Melbourne. Findings were confirmed by Veterinary Pathologist Prof. John W. Finney, SA Pathology, Adelaide and clinical Pathologist Prof. Catriona McLean, Alfred Hospital, Melbourne.
Measurement of relative thymic cortical thickness
Representative images of thymus sections were analysed to determine relative cortical thickness using ImageJ. Briefly, medullary areas were identified on the basis of H and E staining and removed from the larger thymus structure using the Image J Image Calculator function to isolate the cortical region. The thickness of the cortical region, defined by the radius of the largest disk that can fit at a pixel position, was determined using the Local Thickness plugin in ImageJ (http://www.optinav.info/Local_Thickness.htm).
Immunohistochemistry
Following terminal blood collection, P0 and P3 pups were fixed for at least 24 hours in 10% buffered formalin and paraffin embedded before microtomy. Immunohistochemical detection of Cleaved caspase 3 (Cell Signaling Technology #9661) and CD45 (BD) was performed as described previously (Rickard et al., 2014b).
Cytokine quantification
All Plasma was stored at −80°C prior to cytokine analyses. Cytokines were measured by Bioplex Pro mouse cytokine 23-plex assay (Bio- Rad #M60009RDPD) according to manufacturer’s instructions. When samples were designated ‘<OOR’ (below reference range) for a particular cytokine, they were assigned the lowest value recorded for that cohort (as opposed to complete exclusion or inclusion as ‘zero’ which would artificially inflate or conflate group averages respectively). Values are plotted as fold change relative to the mean value for the Wt/Wt samples, and p values were calculated in Microsoft Excel using a 2 tailed TTEST, assuming unequal variance. Data is only shown for cytokines that displayed statistically significant differences between genotypes at either of or both day E19.5 and day P3.
Hematological Analysis
Blood was collected from P0 and P3 pups into EDTA coated tubes using heparinized glass capillary tubes from the neck cavity immediately after decapitation. After centrifugation at 500G for 5 min, 5-15 μL of plasma was carefully removed and this volume was replaced with PBS. Blood cells were resuspended and diluted between 8-20 fold in DPBS for automated blood cell quantification using an ADVIA 2120 hematological analyzer within 6 hours of harvest. Blood was collected from adult mice retro-orbitally into tubes containing EDTA and analyzed using an ADVIA120 automated hematological analyzer (Bayer).
Transplantation Studies
Donor bone marrow or fetal liver cells were injected intravenously into recipient C57BL/6-CD45Ly5.1/Ly5.2 mice following 11Gy of gamma-irradiation split over two equal doses. Recipient mice received neomycin (2 mg/mL) in the drinking water for 4 weeks. Long term capacity of stem cells was assessed by flow cytometric analysis of donor contribution to recipient mouse peripheral blood and/or hematological organs up to 6 months following engraftment. Recovery from cytotoxic insult was assessed by automated peripheral blood analysis at regular times following treatment of mice with 150 mg/kg 5-fluorouracil (5-FU).
Flow Cytometry
To analyze the contribution of donor and competitor cells in transplanted recipients, blood cells were incubated with a combination of the following antibodies: Ly5.1-PE, Ly5.2-FITC, Ly5.2-biotin or Ly5.2 PerCPCy5.5 (antibodies from Becton Dickenson, Ca). If necessary, cells were incubated with a streptavidin PECy5.5 (BD), mixed with propidium iodide (Sigma) and analysed on a LSRI (BD Biosciences) flow cytometer. To analyse the stem- and progenitor cell compartment, bone marrow cells were incubated with biotinylated or Alexa700 conjugated antibodies against the lineage markers CD2, CD3, CD4, CD8, B220, CD19, Gr-1 and Ter-119.
For stem and progenitor cell detection antibodies against cKit, Sca-1, CD48, AnnexinV, CD105, FcγRII/III or CD135 in different combinations (see antibody list for details). Finally FluoroGold (AAT Bioquest Cat#17514) was added for dead cell detection. Cells were then analysed on LSRII or Fortessa1 (BD Biosciences) flow cytometers.
Reactive Oxygen Species (ROS) detection
ROS was detected by using Chloromethyl-H2DCFDA dye according to the manufacturer’s instructions (Invitrogen Cat#C6827). In brief, bone marrow cells were loaded with 1μM Chloromethyl-H2DCFDA for 30 minutes at 37°C. Loading buffer was then removed, and cells were placed into 37°C StemPro-34 serum free medium (ThermoFisher Cat#10639011) for a 15 minute chase period. After incubation cells were placed on ice and stained with surface antibodies suitable for FACS analysis. Cells were analysed using a LSRII flow cytometer (Becton Dickinson).
Quantitative PCR
RNA was prepared using Trizol (Invitrogen) according to the manufacturer’s instructions and 10μg was used for first strand cDNA synthesis using SuperScript II (Life Technologies). ∼0.5 μg of cDNA was then used in a TaqMan PCR reaction with Universal PCR mastermix and murine Mlkl (Mm1244222_n1) and GAPDH (Mm99999915_m1) Taqman probes (ThermoFisher) on an ABI 7900 Fast Real-Time PCR instrument (Applied Biosystems). Mlkl expression relative to GAPDH control was determined using SDS version 2.3 program (Applied Biosystems) and expressed as ΔCT values.
Statistics (Mouse and cell-based assays)
Please consult figure legends for description of error bars used. All P values were calculated in Microsoft Excel or Prism using an unpaired, two tailed t-test, assuming unequal variance. * p ≤ 0.05, ** p ≤ 0.01, ***p ≤ 0.005
Whole Exome Sequencing
DNA from CRMO probands and their family members (when available) was purified from saliva or blood and prepared for whole exome sequencing (WES). The samples underwent WES at several different times, enriched using the Agilent SureSelect Human All Exon V4, V5 or V6+UTR (Agilent Technologies) before sequencing at either Otogenetics, Inc (Atlanta, GA), Beckman Coulter Genomics (Danvers, MA), or at the University of Iowa Genomics Core (Iowa City, IA). The fastq files were quality-checked and processed to vcf format as described previously (Cox et al., 2017). Variants for all samples were called together using GATK’s Haplotype Caller (McKenna et al., 2010) and were recalibrated and hard-filtered in GATK as described previously (Cox et al., 2017). Variants were annotated with minor allele frequencies (MAFs) from 1000 genomes (Genomes Project et al., 2015), ExAC and gnomAD (Lek et al., 2016) and with information regarding the effect of each variant using SNPSift/SNPEff (Cingolani et al., 2012). The databases used for annotation were dbNSFP2.9 (Liu et al., 2016a) (for MAFs) and GRCh37.75 for protein effect prediction.
Ancestry Determination
Ancestry was determined for each CRMO proband using the LASER software package (Wang et al., 2014). A vcf file including ten probands at a time was uploaded to the LASER server and the TRACE analysis was selected using the Worldwide panel. For probands with indeterminate ancestry using the Worldwide panel, the European and Asian panels were used. Principal component values for each proband were plotted using R Statistical Software and the code provided in the LASER package.
MLKL variant quantification
1000 Genomes
Vcf files from 1000 genomes were annotated and filtered as described previously (Cox, 2018). Values for MLKL variants rs35589326 (S132P), rs34515646 (R146Q), and rs144526386 (G202V) as well as all MLKL coding variants were queried and tabulated for allele and genotype count for participants of all ancestry (n=2504), and for those of European ancestry (n=503). Compound heterozygous variants were evident due to the phasing of all variants in the 1000 genomes dataset. CRMO: Allele and genotype counts for all MLKL coding variants were tabulated in probands of European ancestry (n=101) and for all probands (n=128). Compound heterozygous variants were identified using parental sequence data. AS: DNA from all subjects in AS cohort were genotyped using the Illumina CoreExome chip following standard protocols at the Australian Translational Genomics Centre, Princess Alexandra Hospital, Brisbane. Bead intensity data was processed and normalized for each sample and genotypes called using the Illumina Genome Studio software. All the samples listed in the table have been passed quality control process. GB: Genotyping was performed in an ISO15189-accredited clinical genomics facility, Australian Translational Genomics Centre (ATGC), Queensland University of Technology. All samples were genotyped by Illumina HumanOmniExpress (OmniExpress) BeadChip (Blum et al., 2018). QUT controls: A collection of healthy control data of verified European ancestry from various cohort studies, complied by the Translational Genomics Group, QUT and typed on an Illumina CoreExome microarray. Includes data from the The UK Household Longitudinal Study, led by the Institute for Social and Economic Research at the University of Essex and funded by the Economic and Social Research Council. The survey was conducted by NatCen and the genome-wide scan data were analysed and deposited by the Wellcome Trust Sanger Institute. Information on how to access the data can be found on the understanding Society website https://www.understandingsociety.ac.uk/.
Statistical Analysis (Human data)
Statistical comparisons were performed at the level of allele frequency or the level of compound heterozygote sample frequency using either a Fisher’s exact test or a Chi-Squared test with Yates correction as specified under each table. Compound heterozygous variants were quantified and compared at the individual rather than the allelic level, where individuals with and without qualifying variants were compared at the allelic level.
Web resources
gnomAD – https://gnomad.broadinstitute.org/ http://asia.ensembl.org
OrthoDB – https://www.orthodb.org
CADD – https://cadd.gs.washington.edu/
Clustal Omega – https://www.ebi.ac.uk/Tools/msa/clustalo/
WEBLOGO – https://weblogo.berkeley.edu/logo.cgi
Missense Tolerance Ratio (MTR) Gene Viewer – http://biosig.unimelb.edu.au/mtr-viewer
UK biobank – https://www.ukbiobank.ac.uk
AUTHOR CONTRIBUTIONS
Conceptualization: JMH, MK, JMM, WSA, JS.
Methodology: JMH, JMM, WA, JS, HA, JR
Investigation: JMH, MK, IJM, ZL, AC, SM, EJP, MAS, MCT, SNY, CH, SEG, JC, MDS, PG, ECJ, KR, AT, JS, TSS, JGZ, CCA, GMT, ECH, TAW, DS, CAG, JC, AH, NS, SKS, DC, DM, MSH, CGV, CM, MB, SLM, JMM
Resources: JMH, JGZ, VA, RML, AGB, BWD, MAFS, NV, DH, MK, WZ, KW, NV, JT, SB, JR, CGV, PM, MAB, BTK, PJF, JMM, WSA, JS
Writing-Original Draft: JMH, WA, JS
Writing – Review and Editing: JMH, MK, EJP, PAC, PM, PJF, SLM, JMM, WSA, JS Supervision-JMH, MP, PJF, HN, JMM, WA, JS
Funding Acquisition-JMH, JMM, WA, JS
ACKNOWLEDGEMENTS
We thank all the following people for their technical assistance; Jiami Han, Cynthia Liu, Jasmine McManus, Janelle Lochland (WEHI). Aira Nuguid and Tina Cardamone (APN histopathology – The University of Melbourne). Thomas Boudier (WEHI Centre for Dynamic Imaging). The WEHI Histology Service, WEHI Antibody Facility and WEHI Bioservices. Y. Uchiyama and S. Kakuta who advised the interpretation of the results of TEM. Victoria Jackson and Annette Jacobsen for important insight and discussion. The generation of MlklD139V mice by CRISPR/Cas9 was performed by Andrew Kueh and Marco Herold (WEHI MAGEC laboratory) supported by the Australian Phenomics Network (APN) and the Australian Government through the National Collaborative Research Infrastructure Strategy (NCRIS) program.
This work is supported by; Project grant (1105023) and Fellowships (0541951 and 1142669) from the Australian National Health and Medical Research Council (NHMRC) to JMH. Project grant (1105023) and Fellowship (1107149) from the NHMRC to JS. Program grant (1113577) and Fellowship (1058344) from the NHMRC (WSA). JMM-Project grant (1124735) and Fellowship (1105754) from the NHMRC (JMM). NIH training grants T32GM008629 and T32GM082729-01 (AJC). R01AR059703 from the National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) at the National Institutes of Health (PJF and AGB), the Marjorie K. Lamb Professorship PJF. Program grant (1113577) and Fellowship (1058344) from the NHMRC (WSA). Grants-in-Aid from Scientific Research (B) 17H04069 (to HN) from Japan Society for the Promotion of Science (JSPS), and Scientific Research on Innovative areas 26110003 (to HN), the Japan Agency for Medical Research and Development (AMED) through AMED-CREST with a grant number JP18gm1210002 (to HN), and Private University Research Branding project (to HN) from a MEXT (Ministry of Education, Culture, Sports, Science and Technology). Victorian International Research Scholarship (Z. Liu and MCT). Australia Postgraduate Award (CAD). SLM acknowledges funding from NHMRC grants (1144282,1142354 and 1099262), The Sylvia and Charles Viertel Foundation, HHMI-Wellcome International Research Scholarship and Glaxosmithkline. Fellowship from the Lorenzo and Pamela Galli Charitable Trust (ECJ). NHMRC grants 1107425 and 1045549 and The Sylvia & Charles Viertel Senior Medical Research Fellowship (MP). DBA was supported by the Jack Brockhoff Foundation (JBF 4186, 2016) and NHMRC Fellowship (APP1072476). Supported in part by the Victorian Government’s OIS Program. NHMRC IRIISS and Victorian Government Operational Infrastructure Support schemes. NHMRC Project and Targeted Research grants 1006769, 512672 and 512381 to MFS.
MAB acknowledges the Department of Industry, Innovation, Science, Research and Tertiary Education Collaborative Research Network and Diabetes Australia for their support. IJM was supported by the Victorian Cancer Agency, and by generous support from the Felton Bequest.
We gratefully acknowledge the contribution of genotype data by Dr Yorgi Mavros (University of Sydney), Professor Nick Martin (QIMR), Professor Jim Rosenbaum (Oregon Health and Science University), and Professor Maxime Breban and the Groupe Française d’Etude Génétique des Spondylarthrites (GFEGS). We are grateful to Professor BP Wordsworth of the University of Oxford, UK for access to genotype data on ankylosing spondylitis cases collected in studies funded, in part, by Arthritis Research UK (grants 19536 and 18797), by the Wellcome Trust (grant 076113) and by the Oxford Comprehensive Biomedical Research Centre ankylosing spondylitis chronic disease cohort (theme A91202).