Mediterranean fever gene (MEFV) is located on chromosome 16 (16p13.3) and comprises 10 exons with over 370 variants identified to date (1). MEFV encodes for a 781 amino acid protein called pyrin. Pyrin interacts with microtubules and serves as one of the potential initiators of rapid innate immune responses through alterations in cytoplasmic homeostasis, incited by harmful stimuli, termed “homeostasis-altering molecular processes” (HAMPs) (2). Upon activation, pyrin oligomerizes with other cellular proteins, forming a macromolecular complex called ‘pyrin inflammasome’ which subsequently activates caspase 1. This, in turn, allows the release of pro-inflammatory interleukins (IL) from their inactive precursors and induces pyroptosis via the gasdermin D pathway (3).

Some mutations in the MEFV gene cause Familial Mediterranean fever (FMF), which stands as the most common hereditary autoinflammatory disease globally but also pathological phenotypes without FMF diagnosis criteria in many patients with increased inflammatory levels. FMF is mainly characterized by recurrent, transient episodes of peritonitis, pleuritis, arthritis, rash, with accompanying fever (4, 5). Among the prevalent pathogenic mutations are M694V (c.2080A > G), M694I (c.2082G > A), V726A (c.2177T > C), and M680I (c.2040G > C and c.2040G > A), all of which are localized in exon 10. These variants collectively represent nearly 75% of all FMF cases (6). Colchicine remains the most common FMF treatment exhibiting a complete response rate of 64% and a partial response rate of 31% over the long term. Nevertheless, approximately 5–10% of patients exhibit poor responsiveness to colchicine, requiring an additional therapeutical intervention (7). Among these alternatives are IL-1α/β antagonists, such as anakinra or canakinumab, which have demonstrated significant efficacy in reducing the frequency of attacks and enhancing quality of life, accompanied by favorable safety profiles (8). However, their use is restrained by financial constraints and diminished efficacy over prolonged treatment periods (9).

A common feature of mutations in the MEFV gene is neutrophil overactivation. Studies observed that unstimulated neutrophils from patients with the M694V mutation exhibited spontaneous secretion of higher levels of S100A12, IL-18, IL -1β and caspase 1 compared to healthy controls (10, 11). Furthermore, heterozygous, compound heterozygous, or homozygous M694V-positive patients displayed elevated serum levels of S100A12 and IL-18 during both inactive and subclinical phases, in contrast to individuals lacking the M694V mutation. Additionally, Sharma D. et al. (12) discovered a novel regulatory mechanism in an FMF mouse model involving a feedback loop for the TNF/pyrin inflammasome axis, wherein TNF signaling and inflammasome activation synergistically amplify the inflammatory response.

While pyrin inflammasome activation is known to correlate with IL-1β maturation and release (13), and thereby capable of activating caspase 1, it remains unclear whether the pyrin inflammasome directly participates in the IL-18 processing (14). Therefore, the question arises: Why is there a passive increase in IL-18 levels in the serum of FMF patients compared to healthy controls? Recent studies suggest that NLRP1 could be the major inflammasome responsible for regulating IL-18 (15). The objective of this work is to study the potential implications of NLRP1 in MEFV mutation patient-derived peripheral blood mononuclear cells (PBMCs) and monocyte cell model (THP1), aiming to shed light on new molecular mechanism in the FMF disease and explore potential novel treatments.

We started with blood samples from different previously described MEFV mutation patients (Table S1) and healthy controls. Patients 1 and 2 showed a new insertion mutation in the coding region of MEFV gene, the p.Glu128_Asn130dup classified as likely pathogenic by INFEVERS and only previously described in two patients by Logan et al (15). Our initial analysis was focused on measuring the levels of IL-1β and IL-18 (Total IL-18, IL-18BP and free IL-18) in serum. Our findings revealed an increased concentration of total IL-18, IL-18BP and free IL-18 (Fig. 1B-1D) in the serum compared to the control group, but no significant in the case of IL-1β (Fig. 1A). This corroborates findings from previous studies cited in the introduction. Despite the short-lived nature of inflammatory attacks in FMF, these elevated IL-18 levels may account for the observed peaks in immune reactivity. Moreover, this could elucidate the effectiveness of interleukin antagonists such as anakinra.

To evaluate the reactivity after stress induction in MEFV mutated cells, PBMCs were isolated from patients and controls blood samples and stimulated with LPS treatment. Our first approach for assessing the inflammatory response of PBMCs involved lactate dehydrogenase (LDH) quantification, a marker of pyroptosis (Fig. 2A). Results showed that PBMCs derived from MEFV patients are more sensitive to LPS compared to controls. Because this could be associated to the cytokine releases, we measured both IL-18 and IL-1β in the PBMCs medium. Again, in basal conditions, IL-18 was increased and even more so after LPS stimulation (Fig. 2B) compared with IL-1β which remained at low levels in basal conditions and only was increased after LPS stimulation (Fig. 2C). This phenomenon could explain the fever peaks observed in MEFV patients following bacterial or viral infections.

However, because NLRP1 inflammasome has been shown to trigger IL-18 release (15, 16), we evaluated the effect of NLRP1 siRNA in LPS treated PBMCs from patients. NLRP1 siRNA were previously tested on THP1 cells (Supplementary Fig. 1). Interestingly, LPS-induced IL-1β and IL-18 release and LDH levels were reduced under NLRP1 siRNA conditions, highlighting the relevance of NLRP1 in this disease.

The release of both interleukins was hampered after NLRP1 silencing, corroborating new findings about the linked regulation of several inflammasomes during the same pro-inflammatory stimuli (17, 18). To confirm these results, we measured protein expression through western blotting. Every inflammatory marker was upregulated in MEFV cells, including pyrin, NLRP1, caspase 1 and active IL-1β protein levels, which were again reduced after NLRP1 silencing (Fig. 2D). Because these findings were observed in PBMCs from MEFV variant patients but these had not FMF diagnostic criteria, we extended our experiment to 5 more patients with more prevalent mutations. For this, we select patients 5 and 6 who have the homozygote (son) and heterozygote (mother) E148Q mutation and the dual P369S/R408Q mutations which have also been associated to the FMF diseases in many patients. Patient 7 has homozygote M694V mutation, and patients 8 and 9 who have the dual P369S/R408Q mutations. These new patients also showed increased serum levels of total IL-18, IL-18BP and free IL-18 compared to healthy controls (Fig. 1C-D), LPS-induced IL-1β and IL-18 release and increased LDH levels which were reduced under NLRP1 siRNA conditions (Fig. 3A-C), and upregulated inflammatory markers which also were reduced by NLRP1 siRNA (Fig. 3D).

NLRP3 has also been associated to the inflammation in FMF patients who have shown increased NLRP3 protein expression (19, 20). In our study, we also observe increased NLRP3 in PBMCs from patients (Fig. 2D). To elucidate the responsible of the high IL-18 levels in FMF patients, we also treated the patient's cells with MCC950, a specific NLRP3 inhibitor (19) for 4 hours. While IL-18 levels were not reduced, LDH release was mildly decreased, suggesting that although NLRP3 is present, it may play a secondary role in this disease (Supplementary Fig. 2A and B).

As a cellular model for FMF disease or MEFV gain-of-function mutations, we treated THP-1 cells with Toxin A from Clostridium difficile (TcdA) with or without LPS priming, including a THP-1 CRISPR/Cas9 NLRP1 KO strain (Supplementary Fig. 3). TcdA is commonly used as a pyrin inflammasome activator (20, 21). After treatment, cells showed an increased LDH release with no significant difference with the prior priming step (Fig. 4A). Pyrin and both NLRP1 and NLRP3 proteins showed increased expression even without LPS and in combination TcdA. The presence of NLRP1 and NLRP3 after TcdA exposure seems logical, as both inflammasomes are associated with various bacterial infections, including Bacillus anthracis toxins (22). TcdA treatment increased IL-18 release by fivefold; however, in NLRP1 KO cells, there was no significant difference compared to the negative control (Fig. 4B). These results support the idea that NLRP1 is necessary for IL-18 release during pyrin inflammasome activation. It appears that in the case of the MEFV mutation, pyrin may play a leading role. However, NLRP1 could be essential for its full activation in an upstream pathway or might function as a compensatory mechanism. In fact, NLRP1 siRNA also reduced pyrin protein expression in control and patients PBMCs despite the LPS treatment.

MEFV mutations relies on increased pyrin expression and activity as previously described. Our results showed that focusing on only one inflammasome for long term treatments could be not the best choice. Most inflammasomes exhibit overlapping functions due to their critical roles in combating pathogens or addressing cellular dysfunction. Some are specialized in detecting specific bacterial strains, like the NLRP12 inflammasome targeting Yersinia pestis (23), while others, such as NLRC3 (24) and presumably NLRP14 (25), regulate the activity of other inflammasomes. This diverse array of complexes renders the inflammasome system remarkably versatile, capable of adapting to various combinations of stressors. The connection between pyrin and NLRP1, or even more inflammasomes, could explain the lack of effectivity in some MEFV patients. NLRP1 may gain increased significance over time due to an imbalance in the inflammatory process. Unfortunately, there are currently no effective inhibitors for the NLRP1 inflammasome. From a therapeutical point of view, identifying a potential NLRP1 inflammasome inhibitor could open new windows for treatment, not only for FMF but also for many other inflammatory diseases. The development of novel inflammasome inhibitors, specifically targeting NLRP1, could improve therapeutic outcomes.

Despite the importance and novelty of our findings, we acknowledge several limitations in our study. First, many of our results are based on blood samples from patients. More significant findings might emerge if we were able to study the immune environment and inflammation in other tissues. Second, validating our results in larger cohorts is highly desirable, and we are actively working to expand the repository of these valuable samples. Third, studying inflammasome components and their activation during fever episodes would provide critical insights. Therefore, further research in this area is essential.

Biological samples

In this study, we used primary cells obtained from blood samples from healthy donors and people with MEFV mutations. Peripheral blood mononuclear cells (PBMCs) were obtained from the Hospital Puerta del Mar in Cádiz, Spain. Study protocols were approved by the corresponding Ethical Committees. All subjects recruited for this study were adults who provided written informed consent. Information on renal disease parameters from affected patients is summarized in Table S1. Gender and age are also indicated in Table S1 unless not available. PBMCs and serum were obtained from patient blood using Cytiva Ficoll-Paque™ PLUS (Thermo Fisher, Waltham, MA, USA, 11768538).

Cell culture

Commercial THP-1 cell line was purchased from ATCC (Manassas, VI, USA, TIB202).

10⁵ PMBC/well and 5*10⁵ THP-1/well cells were seeded in 6 wells plates containing 2ml of RPMI growth medium supplemented with 10% FBS and 1% antibiotics (Thermo Fisher, Waltham, MA, USA, 11548876), and kept at 37ºC in a CO2 5% incubator. Cells were pelleted by centrifugation at 1000g 5’. Medium was refreshed prior treatment. LPS (Sigma, Saint Louis, MO, USA, L4391) was used at 1ug/ml for 4 hours. TcdA (RyD Systems, Minneapolis, MI, USA, 8619-GT-020) was used for 100ng/ml for 3 hours.

CRISPR/Cas9 NLRP1 KO

10⁶ THP1 cells were resuspended in 2ml RPMI medium containing 1:100 ViralEntry Transduction Enhancer (Applied Biological Materials, Richmond, Canada, G515). Then, 1ug/ml of NLRP1 sgRNA CRISPR/Cas9 All-in-One Lentivirus set (Applied Biological Materials, Richmond, Canada, 31906111) was added and kept for 10’ at room temperature followed spinoculation by centrifugation at 900g for 20’. Then, cells were splitted and cultured in a 6 wells plate containing 1:100 ViralEntry Transduction Enhancer. After 3 days, growth medium was changed and replaced with selection medium containing 0,5 µg/ml of puromycin (Santa Cruz Biotechnologies, Dallas, TX, USA, sc-108071). Selection was kept for another 3 days and the culture was split in T25 culture flasks, then gene expression analysis was performed prior study experiments.

Gene expression

Expression of NLRP1 was analysed by SYBR Green (Takara, Kusatsu, Japan, RR420W) quantitative PCR. cDNA was obtained from extracted mRNA using the iScript cDNA KIT (Biorad, Hercules, CA, USA). The qPCR was performed in a CFX96 Connect Real-Time PCR Detection System (Biorad, Hercules, CA, USA). Actin expression was used as reference gene. 2-ΔΔCT method was used as a relative quantification strategy for data analysis. This method is a convenient way to calculate relative gene expression levels between different samples in that it directly uses the threshold cycles (CTs) generated by the qPCR system for calculation.

Protein electrophoresis

For SDS-PAGE, proteins were extracted using RIPA buffer (Thermo Fisher, Waltham, MA, USA, 89901). Pelleted cells were resuspended in RIPA buffer and kept in ice for 30’, then they were sonicated for 10’’. Next samples were centrifugated at 12.000g for 2’. Protein quantification was done using the Lowry method using DC Protein Assay (Biorad, Hercules, CA, USA, 5000116). Protein concentration was 20ug/ul, preserved in LDS NuPAGE buffer (Thermo Fisher, Waltham, MA, USA, NP0008) and heatshocked at 95ºC for 2’. Gel electrophoresis was performed using 4–20% Mini-PROTEAN® TGX Stain-Free™ Protein Gels (Biorad, Hercules, CA, USA) at 200V in Tris-Glycine-SDS buffer (Biorad, Hercules, CA, USA, 1610772) for 45’.

Western Blotting

Protein transfer was made using a TurboTransfer (Biorad, Hercules, CA, USA) at 25V for 10’. After transferring the proteins to 0.45 uM nitrocellulose membranes (Biorad, Hercules, CA, USA), these were incubated for 1 hour in BSA 5% in PBS-Tween20 0.05% and then overnight at 4ºC with primary antibodies at 1:1000. Then washed twice with PBS-Tween20 and incubated with the corresponding secondary 1:5000 antibody for 1h at RT. Protein loading was checked using reference protein (Tubulin). Stripping was not used.

Interleukin measurement

ELISA Kits for IL-18 (EH0011), IL-18BP (EH5043) and IL-1β (EH0185) were purchased from FineTest (Boulder, CO, USA). The assays were performed using patient blood serum or cell culture medium according to the manufacturer's protocol. Briefly, 300 µL of serum or medium was collected and centrifuged at 10.000g for 5’ for each condition. Then, 100 µL of undiluted sample was added to each well and incubated for 90’ at 37°C. The plate was washed twice, followed by the addition of 100 µL of biotin-labeled antibody working solution to each well, incubated for 60 minutes at 37°C, and washed three times. Next, 100 µL of HRP-Streptavidin conjugate solution was added for 30’ at 37°C, followed by five washes. Subsequently, 90 µL of 3,3′,5,5′-Tetramethylbenzidine (TMB) substrate was added and incubated for 15’ at 37°C. Finally, 50 µL of stop solution was added, and the absorbance was measured at 450 nm.

Free IL-18 was calculated using the IL-18BP concentration and total IL-18 concentration, in accordance with the law of mass action (26), with a 1:1 stoichiometry in the complex of IL-18 and IL-18BP and complex dissociation constant (KD) of 0.4 nM.

LDH assay

The LDH/Lactate Dehydrogenase Assay Kit (Colorimetric) (Abcam, Cambridge, UK, ab102526) was used according to the manufacturer's protocol. Briefly, 200 µL of medium was collected and centrifuged at 10,000 g for 5 minutes. Then, 50 µL of undiluted sample was added to each well, mixed with 48 µL of LDH assay buffer and 2 µL of LDH substrate mix solution, and incubated for 30’ at 37°C. Absorbance was measured at 450 nm.

Silencer RNA assay

Silencer RNA Assay was performed using Lipofectamine® RNAiMAX Reagent (Thermo Fisher, Waltham, MA, USA, 13778-030) following the manufacturer's protocol. Briefly, 5 × 10⁵ cells per well were seeded in a 6-well plate with 2 mL of optimized medium. The siRNA-lipid complex solution was prepared by mixing the Lipofectamine solution (150 µL RPMI without FBS or antibiotics, plus 9 µL Lipofectamine) and the siRNA solution (150 µL RPMI without FBS or antibiotics, plus 3 µL siRNA at 10 µM) and incubating for 5 minutes at room temperature. Then, 250 µL of the siRNA-lipid complex was added to each well. Cells were incubated for 72 hours before treatment and/or analysis.

Reagents

Bovine Serum Albumim (A7030), Triton X-100 (X100-100ML) and PMA (P8139) were purchased from MERCK (Darmstadt, Germany). The following reagents were obtained from Thermo Fisher Scientific (Waltham, MA, USA): RPMI 1640 medium (31870074) and silencer select siRNA NLRP1 (4392420 id: s22521). Primary antibodies: anti ASC antibody (Abcam, Cambridge, UK, ab283684), anti-Caspase 1 antibody (Cell Signaling, Danvers, MA, USA, 3866), anti NLRP1 antibody (Abclonal, Woburn, MA, USA, A16212), anti NLRP3 antibody (Abclonal, Woburn, MA, USA, A5652), anti β actin antibody (Abcam, Cambridge, UK, ab6276), anti interleukin 1β antibody (Cell Signaling, Danvers, MA, USA, 12703) and, anti- Pyrin/MEFV antibody (Cell Signaling, Danvers, MA, USA, 40649).

Statistics

Non parametric student t-test (Mann-Whitney) was used to compare data between 2 groups. The Mann-Whitney U test does not rely on the assumption of normal distribution in the data. It is particularly useful when dealing with ordinal or interval data that may not meet the normality assumption required by parametric tests. It can be applied to a wide range of data types, including ordinal, interval, and some ratio data. The test is robust against outliers and skewed data, making it a good choice when dealing with datasets that may have extreme values. The Mann-Whitney U test is sensitive to differences in both central tendency and shape of the distribution, making it versatile for various types of comparisons. The test assumes that the observations in one group are independent of the observations in the other group. If this assumption is violated, the results may be invalid. It assumes that the shapes of the distributions in the two groups are similar. If the shapes are substantially different, the Mann-Whitney U test may not provide an accurate assessment of differences. Finally, the test may be less accurate when there are ties in the data (i.e., multiple observations with the same value). Some adjustments or corrections may be necessary in such cases.

All results are expressed as mean ± SD of 3 independent experiments and a p-value < 0.05 was considered as statistically significant. Level of significance is denoted by asterisks *P < 0.05, **P < 0.01.