Inflammation is a critical part of a healthy immune system, which protects us against harmful pathogens (such as bacteria or viruses) and works to restore damaged tissues. In the immune cells of our body, the inflammatory process can be activated through a group of inflammatory proteins that together are known as the NLRP3 inflammasome complex.

While inflammation is a powerful mechanism that protects the human body, persistent or uncontrolled inflammation can cause serious, long-term damage. The inappropriate activation of the NLRP3 inflammasome has been implicated in several diseases, including Alzheimer’s disease, heart disease, and diabetes. The NLRP3 inflammasome can be activated by different stimuli, including changes in cell volume and exposure to either molecules produced by damaged cells or toxins from bacteria. However, the precise mechanism through which the NLRP3 becomes activated in response to these stimuli was not clear.

The exit of chloride ions from immune cells is known to activate the NLRP3 inflammasome. Chloride ions exit the cell through proteins called anion channels, including volume-regulated anion channels (VRACs), which respond to changes in cell volume. Green et al. have found that, in immune cells from mice grown in the lab called macrophages, VRACs are the only chloride channels involved in activating the NLRP3 inflammasome when the cell’s volume changes. However, when the macrophages are exposed to molecules produced by damaged cells or toxins from bacteria, Green et al. discovered that other previously unidentified chloride channels are involved in activating the NLRP3 inflammasome.

These results suggest that it might be possible to develop drugs to prevent the activation of the NLRP3 inflammasome that selectively target specific sets of chloride channels depending on which stimuli are causing the inflammation. Such a selective approach would minimise the side effects associated with drugs that generically suppress all NLRP3 activity by directly binding to NLRP3 itself. Ultimately, this may help guide the development of new, targeted anti-inflammatory drugs that can help treat the symptoms of a variety of diseases in humans.

Inflammation is an important protective host-response to infection and injury, and yet is also detrimental during non-communicable diseases (Dinarello et al., 2012). Inflammasomes are at the heart of inflammatory responses. Inflammasomes are formed by a soluble pattern recognition receptor (PRR), in many cases the adaptor protein ASC (apoptosis-associated speck-like protein containing a CARD), and the protease caspase-1 (2). Inflammasomes form in macrophages in response to a specific stimulus to drive the activation of caspase-1, facilitating the processing of the cytokines pro-interleukin (IL)1β and pro-IL-18 to mature secreted forms, and the cleavage of gasdermin D to cause pyroptotic cell death (Evavold and Kagan, 2019). A number of different inflammasomes have been described, but potentially the inflammasome of greatest interest to non-communicable disease is formed by NLRP3 (NACHT, LRR and PYD domains-containing protein 3) (Mangan et al., 2018). The mechanisms of NLRP3 activation remain poorly understood.

The NLRP3 inflammasome is activated through several routes which have been termed the canonical, non-canonical, and the alternative pathways (Mangan et al., 2018). Activation of the canonical NLRP3 pathway, which has received greatest attention thus far, typically requires two stimuli; an initial priming step involving a pathogen associated molecular pattern (PAMP), typically bacterial endotoxin (lipopolysaccharide, LPS), to induce expression of pro-IL-1β and NLRP3, and a second activation step usually involving a damage associated molecular pattern (DAMP), such as adenosine triphosphate (ATP) (Mariathasan et al., 2006). In 1996, Perregaux and colleagues discovered that hypotonic shock was effective at inducing the release of mature IL-1β when applied to LPS-treated human monocytes and suggested the importance of a volume-regulated response (Perregaux et al., 1996). It was later discovered that hypotonicity induced release of IL-1β via activation of the NLRP3 inflammasome (Compan et al., 2012), and that this was linked to the regulatory volume decrease (RVD), which is a regulated reduction in cell volume in response to hypo-osmotic-induced cell swelling, and was inhibited by the chloride (Cl^-) channel blocker NPPB (5-nitro-(3-phenylpropylamino)benzoic acid) (Compan et al., 2012). The RVD is regulated by the Cl^- channel VRAC (volume regulated anion channel). The molecular composition of the VRAC channel was established to consist of an essential LRRC8A sub-unit in combination with other (B-E) LRRC8 sub-units (Qiu et al., 2014; Voss et al., 2014). We recently reported that fenamate NSAIDs could inhibit the canonical NLRP3 inflammasome by blocking a Cl^- channel, which we suggested could be VRAC (Daniels et al., 2016). We also further characterised the importance of Cl^- flux in the regulation of NLRP3, showing that Cl^- efflux facilitated NLRP3-dependent ASC oligomerisation (Green et al., 2018). Given the poor specificity of many Cl^- channel inhibitors, we set out to systematically determine the importance of VRAC and the RVD to NLRP3 inflammasome activation. Given that hypertonic buffers inhibit ATP and other NLRP3 activating stimuli-induced IL-1β release from macrophages (Perregaux et al., 1996; Compan et al., 2012), we hypothesised that cell swelling, and the RVD, would be central to all NLRP3 activating stimuli. Using pharmacological and genetic approaches, we discovered that VRAC exclusively regulated RVD-dependent NLRP3 activation in response to hypotonicity, and not NLRP3 activation in response to other canonical stimuli. Thus, we provide genetic evidence for the importance of Cl^- in regulating NLRP3 via the VRAC dependence of the hypotonicity response, and suggest the presence of additional Cl^- sensing mechanisms regulating NLRP3 in response to DAMPs.

Following publication of the cryo-electron microscopy (cryo-EM) structure of VRAC (Deneka et al., 2018) with inhibitor 4-(2-butyl-6,7-dichloro-2-cyclopentyl-indan-1-on-5-yl)oxobutyric acid (DCPIB) (Figure 1A; Kern et al., 2019), we were able to investigate the interaction of established VRAC inhibitors with the channel using molecular modelling. DCPIB was first computationally redocked using Molecular Operating Environment (MOE 2015.08, Chemical Computing Group, Canada) into the homohexameric VRAC structure (PDB code 6NZW, resolution 3.2 Å) (Kern et al., 2019). The resulting pose produced a reasonable overlay with the cryo-EM conformation of DCPIB, giving a root-mean-square deviation in atomic position of 2.6 Å (Figure 1B,C). DCPIB exhibits an ionic interaction of its carboxylate group with the cationic side-chain of one of the Arg103 residues comprising the electropositive selectivity filter of VRAC (Kern et al., 2019). Known VRAC inhibitors (Daniels et al., 2016; Hélix et al., 2003; Droogmans et al., 1999) possessing carboxylic acid groups (flufenamic acid (FFA)), mefenamic acid (MFA) and N-((4-methoxy)−2-naphthyl)−5-nitroanthranilic acid (MONNA), the tetrazole moiety (NS3728) and sulfonic acid groups (4-sulfonic calix[6]arene) were then docked into the DCPIB site of VRAC. The most favourably bound poses of these ligands were similarly found to block the pore in a cork-in-bottle manner (Kern et al., 2019) at the selectivity filter; the ligands’ ionised acidic groups formed strong electrostatic interactions with Arg103 (Figure 1D–H). Tamoxifen, a basic inhibitor of VRAC was also docked into the cryo-EM structure (Figure 1I). Accordingly, tamoxifen docked with its cationic tertiary amino group remote to the Arg103 side-chains (Figure 1l and Figure 1—figure supplement 1); these side-chains instead formed cation-π interactions with the phenyl group of tamoxifen (Figure 1l). The interaction of the tamoxifen pose was computed as having a calculated ligand-binding affinity of −6.5 kcal mol⁻¹ via the molecular mechanics/generalised Born volume integration (MM/GBVI) method (Labute, 2008; Galli et al., 2014). The binding energies of the anionic ligands were also predicted as favourable, ranging from −5.1 (DCPIB) to −5.8 kcal mol⁻¹ (MONNA). This range excludes the larger 4-sulfonic calix[6]arene (calixarene), which gave a binding energy of −9.9 kcal mol⁻¹; we note that the GBVI implicit solvent model may be underestimating the high desolvation cost of this polyanionic ligand and therefore overestimating the magnitude of the corresponding binding energy of this compound.

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We then tested the ability of six of the compounds described in Figure 1 (DCPIB, calixarene (Calix), tamoxifen, FFA, MONNA, NS3728) to inhibit hypotonicity-induced VRAC-dependent Cl^- flux using the iodide (I^-) quenching of halide-sensitive YFP H148Q/I152L (Galietta et al., 2001) in live HeLa cells (Figure 2A). In this model, I^- enters the cell through open Cl^- channels to induce quenching of a mutant EYFP. In response to hypotonic shock to induce VRAC opening, YFP fluorescence was immediately quenched, which was significantly inhibited by tamoxifen (10 µM), MONNA (50 µM), DCPIB (10 µM), FFA (100 µM), and NS3728 (10 µM), but not calixarene (100 µM) (Figure 2A,B). VRAC also regulates RVD in response to cell swelling (Qiu et al., 2014; Voss et al., 2014). We measured the RVD by measuring the change in cellular fluorescence in calcein-loaded primary mouse bone-marrow-derived macrophages (BMDMs) in response to hypotonicity. Hypotonicity caused a rapid increase in cell volume which declined over time, characteristic of an RVD response (Figure 2C). Similar to the quenching assay, RVD was also significantly inhibited in the presence of tamoxifen, MONNA, DCPIB, FFA and NS3728, but not calixarene (Figure 2C,D). These data suggest that all the molecules in our panel, except calixarene, are bona-fide VRAC inhibitors at the concentrations tested.

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We tested whether the panel of VRAC inhibitors characterised above could block NLRP3 inflammasome activation and release of IL-1β in response to DAMP stimulation. Primary BMDMs were primed with LPS (1 µg mL⁻¹, 4 hr), before activation of NLRP3 by ATP (5 mM, 2 hr). Inhibitors were given at the same dose they inhibited VRAC above in HeLa cells 15 min before the addition of ATP and were then present for the duration of the experiment. Of the panel of verified VRAC inhibitors, only MONNA, FFA and NS3728 consistently inhibited ATP-induced IL-1β release (Figure 3A). At the dose used in this assay, DCPIB did not consistently inhibit ATP-induced IL-1β release (Figure 3A), but at higher concentrations did inhibit NLRP3 activation (Figure 3—figure supplement 1). Likewise, pyroptosis, as measured by LDH release, was significantly reduced by MONNA and FFA, and was unaffected by tamoxifen or calixarene (Figure 3B). The inhibitors that blocked ATP-induced IL-1β release also inhibited ASC oligomerisation, caspase-1 activation, and gasdermin D cleavage (Figure 3C). These data show that some very effective VRAC inhibitors failed to inhibit activation of the NLRP3 inflammasome and release of IL-1β, suggesting that VRAC may not be the molecular target of these molecules inhibiting the inflammasome.

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ATP-induced NLRP3 activation is dependent upon K⁺ efflux, whereas NLRP3 activation by treatment with the imidazoquinoline compound imiquimod is K⁺ efflux-independent (Groß et al., 2016). We therefore sought to test if VRAC inhibitors that were effective at blocking ATP-induced inflammasome activation were specific to K⁺ efflux-sensitive mechanisms. FFA (100 µM) and NS3728 (10 µM) were effective at blocking IL-1β release after treatment with the K⁺ ionophore nigericin (10 µM, 2 hr) (Figure 3D), but were unable to block imiquimod (75 µM, 2 hr)-induced IL-1β release (Figure 3E). Similarly, ASC oligomerisation, caspase-1 activation and gasdermin D cleavage induced by nigericin were sensitive to FFA and NS3728 pre-treatment (Figure 3F). However, ASC oligomerisation, caspase-1 activation and gasdermin D cleavage induced by imiquimod were not affected by FFA and NS3728 pre-treatment (Figure 3F). Increased extracellular KCl (25 mM) was sufficient to block nigericin-induced activation, but not imiquimod, demonstrating the K⁺ dependency of nigericin (Figure 3F). These data suggest that these Cl^- channel inhibiting compounds exclusively target the K⁺-dependent canonical pathway of NLRP3 activation.

Many Cl^- channel inhibiting drugs are known to inhibit multiple Cl^- channels, and we established that very effective VRAC inhibitors (tamoxifen and DCPIB) had negligible effect on NLRP3 activation at VRAC inhibiting concentrations. Thus, to conclusively determine the role of VRAC in NLRP3 inflammasome activation we generated a macrophage specific LRRC8A knockout (KO) mouse using CRISPR/Cas9 (Figure 4A). The generation of a macrophage-specific LRRC8A KO was required as whole animal LRRC8A KO mice do not survive beyond 4 weeks and have retarded growth (Kumar et al., 2014). Lrrc8a^fl/fl mice were bred with mice constitutively expressing Cre under the Cx3cr1 promoter, as previously shown to be expressed in monocyte and macrophage populations (Yona et al., 2013). This generated mice with the genotype Lrrc8a^fl/fl:Cx3cr1^cre (KO) with littermates Lrrc8a^fl/fl:Cx3cr1^WT (WT). Cell lysates were prepared from BMDMs and peritoneal macrophages isolated from WT and KO mice and were western blotted for LRRC8A confirming that Lrrc8a KO cells were knocked out for LRRC8A (Figure 4B). Functional loss of LRRC8A was confirmed using the calcein RVD assay described above. BMDMs were subjected to a hypotonic shock and changes in calcein fluorescence measured over time. In WT cells, there was a characteristic RVD (Figure 4C). However, in Lrrc8a KO cells there was complete loss of the RVD response (Figure 4C,D). The absence of RVD was also strikingly evident by observation of the cells by phase contrast microscopy (Figure 4E, Videos 1 and 2). Treatment of Lrrc8a KO BMDMs with DCPIB and subsequent hypotonic shock resulted in a further dysregulation of cell volume compared to WT cells or KO cells treated with hypotonic shock alone, suggesting that additional Cl^- channels act to constrain cell volume in the absence of a functional RVD (Figure 4—figure supplement 1). These data confirm functional KO of the VRAC channel in macrophages.

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We next used the Lrrc8a KO macrophages to test the hypothesis that VRAC and the RVD were important for NLRP3 inflammasome activation and IL-1β release in response to DAMP stimulation. WT BMDMs and Lrrc8a KO BMDMs were primed with LPS (1 µg mL⁻¹, 4 hr) and then treated with the NLRP3 inflammasome activators ATP (5 mM, 2 hr), nigericin (10 µM, 2 hr), silica (300 µg mL⁻¹, 2 hr), or imiquimod (75 µM, 2 hr). Knocking out LRRC8A had no effect on the release of IL-1β (Figure 5A) or cell death (Figure 5B). We then used western blotting to determine ASC oligomerisation and caspase-1 activation. In response to the NLRP3 inflammasome activators nigericin, ATP, and imiquimod, there was no effect of LRRC8A KO on ASC oligomerisation or caspase-1 activation (Figure 5C). Furthermore, IL-1β release in response to ATP or nigericin was still inhibited by the VRAC inhibitors flufenamic acid (FFA, 100 µM), and NS3728 (10 µM) in the Lrrc8a KO BMDMs, confirming that these inhibitors are inhibiting NLRP3 inflammasome activation by a VRAC-independent mechanism (Figure 5D). Flufenamic acid and NS3728 also inhibited ASC oligomerisation and caspase-1 activation as determined by western blot in the Lrrc8a KO BMDMs to the same extent as in the WT (Figure 5E). We then used a murine peritonitis model described previously (Daniels et al., 2016) to investigate the role of LRRC8A in vivo. First, we tested if the VRAC inhibitor NS3728 was effective at blocking NLRP3 in vivo. Wild-type C57BL6/J mice were injected intraperitoneally with NS3728 (50 mg kg⁻¹), the NLRP3 inhibitor MCC950 (50 mg kg⁻¹), or vehicle control, at the same time as LPS (1 µg, 4 hr). NLRP3 was then further activated by intraperitoneal injection of ATP (100 mM, 500 µL, 15 min) and IL-1β release was measured by ELISA of the peritoneal lavage (Figure 5F) and plasma (Figure 5G). Addition of ATP induced a significant increase in the release of IL-1β into peritoneal lavage and this was inhibited by MCC950 and NS3728, indicating an NLRP3-dependent response. IL-6 levels were unaltered in both peritoneal lavage and plasma by addition of NS3728 (Figure 5—figure supplement 1A,B). These data show that NS3728 was able to inhibit NLRP3 in vivo. To determine the role of VRAC in this model, we repeated this experiment in our macrophage Lrrc8a KO mice and their littermate controls. Macrophage Lrrc8a KO mice exhibited normal proportions of myeloid cells in the peritoneum as assessed by flow cytometry (Figure 5—figure supplement 1C–G). Loss of macrophage LRRC8A had no effect on the IL-1β levels in the peritoneal lavage in response to LPS and ATP (Figure 5H), or in the plasma (Figure 5I). Moreover, similar to our in vitro findings, NS3728 was still effective at inhibiting this response in the absence of LRRC8A (Figure 5H,I). These data suggested that VRAC was dispensable for NLRP3 activation by DAMP stimulation, and that the VRAC inhibitors are effective at inhibiting NLRP3 in the absence of VRAC, suggesting the presence of another target.

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RVD in response to hypo-osmotic-induced cell swelling is documented as an inducer of NLRP3 inflammasome activation and IL-1β release (Perregaux et al., 1996; Compan et al., 2012). Since Lrrc8a KO BMDMs could no longer control their volume in response to hypotonic shock, we tested whether NLRP3 inflammasome activation by hypotonicity was altered. LPS-primed (1 µg mL⁻¹, 4 hr) BMDMs were incubated in a hypotonic solution (4 hr) which caused IL-1β and LDH release from WT cells, and which was significantly inhibited in Lrrc8a KO BMDMs (Figure 6A,B). There was no difference in IL-1β release or cell death between ATP-stimulated WT and KO BMDMs (Figure 6A,B). Caspase-1 cleavage and IL-1β processing induced by hypotonicity were also completely inhibited in the absence of LRRC8A (Figure 6C), indicating the response was completely dependent on both NLRP3 and VRAC. Moreover, hypotonicity-induced ASC oligomerisation was also dependent on VRAC (Figure 6D). These data show that in response to hypo-osmotic stress, VRAC was essential for NLRP3 inflammasome activation.

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Pharmacological and biochemical evidence supporting an important role of Cl^- ions in the activation of the NLRP3 inflammasome has been provided by various studies over the years (Perregaux et al., 1996; Compan et al., 2012; Daniels et al., 2016; Green et al., 2018; Verhoef et al., 2005), although conclusive genetic evidence has been lacking. The promiscuous nature of many Cl^- channel inhibiting drugs, and an unresolved molecular identity of major Cl^- channels, have prevented the emergence of conclusive genetic proof. However, the discovery that the Cl^- channel regulating the RVD (VRAC) was composed of LRRC8 sub-units, and that LRRC8A was essential for channel activity, offered us the opportunity to investigate the direct importance of VRAC in the regulation of NLRP3. Hypotonicity induces cell swelling which is corrected by the VRAC-dependent RVD (Qiu et al., 2014; Voss et al., 2014). The RVD was previously linked to NLRP3 activation (Compan et al., 2012). Thus by knocking out LRRC8A, and thus VRAC, we would discover that VRAC was essential for RVD-induced NLRP3 inflammasome activation, providing strong evidence for the direct requirement of a Cl^- channel in NLRP3 inflammasome activation. Given that there are a number of inflammatory conditions and models that can be targeted by administration of a hyper-tonic solution (e.g. Theobaldo et al., 2012; Schreibman et al., 2018; Shields et al., 2003; Petroni et al., 2015), it is possible that the VRAC-dependent regulation of NLRP3 in response to hypotonicity could represent a therapeutic target.

However, VRAC was only essential for RVD-induced NLRP3 activation and was not involved in the NLRP3 response to DAMP stimulation. The fact that our VRAC channel-inhibiting drugs block DAMP-induced NLRP3 activation suggests that additional Cl^- channels (or alternative targets) are involved in coordinating NLRP3 responses to other stimuli. Chloride intracellular channel proteins (CLICs 1–6) form anion channels and regulate a variety of cellular processes (Littler et al., 2010; Argenzio and Moolenaar, 2016). Localisation of CLIC1 and 4 to membrane fractions in macrophages is increased by LPS stimulation, and RNAi knockdown of both CLIC1 and 4 impaired LPS and ATP-induced IL-1β release from macrophages (Domingo-Fernández et al., 2017). In addition to CLICs 1 and 4, CLIC5 is also implicated in NLRP3-dependent IL-1β release (Tang et al., 2017). Knockdown of CLICs 1, 4, and 5 inhibits NLRP3 inflammasome activation in response to the soluble agonists ATP and nigericin, and also the particulate DAMP monosodium urate crystals (Tang et al., 2017). Thus, it appears that multiple Cl^- channels encode diverse signals arising from DAMP stimulation, or from altered cellular homeostasis, to trigger NLRP3 inflammasome activation. Importantly our data suggest that Cl^- channels are only important to NLRP3 activation dependent upon K⁺ efflux, highlighting the further potential for selective pathway modulation and therapeutic development. The relationship between K⁺ and Cl^- efflux requires further investigation, and whether, in RVD, Cl^- efflux is a pre-requisite for K⁺ efflux.

Although activation of VRAC is best understood under hypotonic conditions, VRAC activation has also been reported to occur in response to a variety of stimuli (Osei-Owusu et al., 2018). It is also possible that VRAC may regulate NLRP3 inflammasome activation in response to stimuli other than hypotonicity that were not tested here. For example, sphingosine-1-phosphate (S1P) activates VRAC in mouse macrophages (Burow et al., 2015), and we previously reported that sphingosine, and S1P, could activate NLRP3 (Luheshi et al., 2012). VRAC is also thought to be important for cell death induced by apoptosis inducing drugs (Sørensen et al., 2016; Planells-Cases et al., 2015), and we also previously reported that apoptosis inducing drugs could activate the NLRP3 inflammasome in activated macrophages (England et al., 2014) suggesting another potential area of relevance. While VRAC is directly permeable to Cl^- it is also possible that additional Cl^- channels are involved in the hypotonicity-induced swelling response. For example, VRAC activation caused by swelling can activate other Cl^- channels, notably anoctamin 1 (e.g. Liu et al., 2019; Benedetto et al., 2016). Furthermore, whilst our research suggests an importance of Cl^-, VRAC is also permeable to small molecules including cGAMP (Zhou et al., 2020) and ATP (Dunn et al., 2020), highlighting additional ways through which VRAC could contribute to inflammation.

Inhibiting the NLRP3 inflammasome has become an area of intense research interest due to the multiple indications of its role in disease (Mangan et al., 2018). The inhibitor MCC950 is now thought to bind directly to NLRP3 to cause inhibition (Coll et al., 2019; Tapia-Abellán et al., 2019), although it has also been reported to inhibit Cl^- flux from macrophages treated with nigericin (Jiang et al., 2017), and was found to bind directly to CLIC1 (Laliberte et al., 2003), so it is possible that some of its inhibitory activity may be attributable to an effect on Cl^-. We found that Cl^- channel inhibition blocked IL-1β release in a NLRP3-dependent model of peritonitis, and previously reported protective effects of the fenamate NSAIDs in rodent models of Alzheimer’s disease that we attributed to an effect on Cl^- channel inhibition (Daniels et al., 2016). Thus, it is possible that targeting Cl^- channels offers an additional route to inhibit NLRP3-dependent inflammation in disease.

In summary, we have reported that hypotonicity-induced NLRP3 inflammasome activation depends exclusively on the Cl^- channel VRAC, and that different Cl^- sensing and regulating systems coordinate the activation of NLRP3 in response to DAMPs. This opens the possibility of discrete Cl^- regulating mechanisms conferring selectivity and information about the nature of the NLRP3 activating stimulus. Thus, this investigation has opened the door to further studies on Cl^- regulation of NLRP3 and identified the possibility of selective therapeutic intervention strategies informed by the nature of the disease or DAMP stimulus, potentially minimising complications of immunosuppression caused by a blanket NLRP3 inhibition.

All data generated or analysed during this study are included in the manuscript and supporting files. Source data are provided for graphs summarised where individual data points are not already provided in the figure, i.e Figure 2A, 2C and 4C.

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Author details

1. Jack P Green

   1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
   2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

   Contribution

   Data curation, Formal analysis, Investigation, Visualization, Methodology, Project administration, Writing - review and editing

   Contributed equally with

   Tessa Swanton

   For correspondence

   Jack.green@manchester.ac.uk

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1108-3422

2. Tessa Swanton

   1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
   2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

   Contribution

   Data curation, Formal analysis, Investigation, Visualization, Methodology, Writing - review and editing

   Contributed equally with

   Jack P Green

   Competing interests

   No competing interests declared

3. Lucy V Morris

   1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
   2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

   Contribution

   Formal analysis, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

4. Lina Y El-Sharkawy

   Division of Pharmacy and Optometry, School of Health Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, The University of Manchester, Manchester, United Kingdom

   Contribution

   Formal analysis, Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

5. James Cook

   1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
   2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

   Contribution

   Investigation, Writing - review and editing

   Competing interests

   No competing interests declared

6. Shi Yu

   1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
   2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

   Contribution

   Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0651-4769

7. James Beswick

   Division of Pharmacy and Optometry, School of Health Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, The University of Manchester, Manchester, United Kingdom

   Contribution

   Resources, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

8. Antony D Adamson

   Genome Editing Unit Core Facility, Faculty of Biology, Medicine and Health, University of Manchester, Manchester, United Kingdom

   Contribution

   Resources, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

9. Neil E Humphreys

   1. Genome Editing Unit Core Facility, Faculty of Biology, Medicine and Health, University of Manchester, Manchester, United Kingdom
   2. EMBL-ROME, Epigenetics and Neurobiology Unit, Adriano Buzzati-Traverso Campus, Monterotondo, Italy

   Contribution

   Resources, Investigation, Methodology, Writing - review and editing

   Competing interests

   No competing interests declared

10. Richard Bryce

    Division of Pharmacy and Optometry, School of Health Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, The University of Manchester, Manchester, United Kingdom

    Contribution

    Formal analysis, Supervision, Investigation, Methodology, Writing - review and editing

    Competing interests

    No competing interests declared

11. Sally Freeman

    Division of Pharmacy and Optometry, School of Health Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, The University of Manchester, Manchester, United Kingdom

    Contribution

    Resources, Formal analysis, Supervision, Methodology, Writing - review and editing

    Competing interests

    No competing interests declared

12. Catherine Lawrence

    1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
    2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

    Contribution

    Supervision, Funding acquisition, Methodology, Project administration, Writing - review and editing

    Competing interests

    No competing interests declared

13. David Brough

    1. Division of Neuroscience and Experimental Psychology, School of Biological Sciences, Faculty of Biology, Medicine and Health, Manchester Academic Health Science Centre, University of Manchester, Manchester, United Kingdom
    2. Lydia Becker Institute of Immunology and Inflammation, University of Manchester, Manchester, United Kingdom

    Contribution

    Conceptualization, Resources, Supervision, Funding acquisition, Writing - original draft, Project administration, Writing - review and editing

    For correspondence

    david.brough@manchester.ac.uk

    Competing interests

    No competing interests declared

    "This ORCID iD identifies the author of this article:" 0000-0002-2250-2381

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