# Store-operated calcium entry dysfunction in CRAC channelopathy: Insights from a novel STIM1 mutation

Benedicte Alary, Pascal Cintas, Corentin Claude, Olivier Dellis, Corinne Thèze, Charles van Goethem, Mireille Cossée, Martin Krahn, Valérie Delague, Marc Bartoli

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## To cite this version:

Benedicte Alary, Pascal Cintas, Corentin Claude, Olivier Dellis, Corinne Thèze, et al., Store-operated calcium entry dysfunction in CRAC channelopathy: Insights from a novel STIM1 mutation. Clinical Immunology, 2024, 265, pp.110306. 10.1016/j.clim.2024.110306. hal-04639441

HAL Id: hal-04639441

https://hal.science/hal-04639441v1

Submitted on 14 Sep 2024

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# Store-operated calcium entry dysfunction in CRAC channelopathy: Insights from a novel STIM1 mutation

Benedicte Alary<sup>a</sup>, Pascal Cintas<sup>b</sup>, Corentin Claude<sup>c</sup>, Olivier Dellis<sup>c</sup>, Corinne Thèze<sup>d</sup>, Charles Van Goethem<sup>d</sup>, Mireille Cossée<sup>d,e</sup>, Martin Krahn<sup>a,f</sup>, Valérie Delague<sup>a</sup>, Marc Bartoli<sup>a,g,*</sup>

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## ARTICLE INFO

### Keywords:

STIM1
CRAC channelopathy
Immunodeficiency
Store-operated calcium entry
Splicing
Therapy

## ABSTRACT

Store-operated calcium entry (SOCE) plays a crucial role in maintaining cellular calcium homeostasis. This mechanism involves proteins, such as stromal interaction molecule 1 (STIM1) and ORAI1. Mutations in the genes encoding these proteins, especially STIM1, can lead to various diseases, including CRAC channelopathies associated with severe combined immunodeficiency. Herein, we describe a novel homozygous mutation, NM_003156 c.792-3C > G, in *STIM1* in a patient with a clinical profile of CRAC channelopathy, including immune system deficiencies and muscle weakness. Functional analyses revealed three distinct spliced forms in the patient cells: wild-type, exon 7 skipping, and intronic retention. Calcium influx analysis revealed impaired SOCE in the patient cells, indicating a loss of STIM1 function. We developed an antisense oligonucleotide treatment that improves *STIM1* splicing and highlighted its potential as a therapeutic approach. Our findings provide insights into the complex effects of *STIM1* mutations and shed light on the multifaceted clinical presentation of the patient.

## 1. Introduction

The store-operated calcium entry (SOCE) is a ubiquitous mechanism and is essential for maintaining calcium homeostasis [18]. SOCE allows the entry of extracellular calcium when the calcium stores in the reticulum are depleted. This mechanism is primarily mediated by two families of proteins, stromal interaction molecule (STIM) and calcium release-activated calcium channel protein (ORAI), which are calcium-released activated channels (CRACs). STIM proteins, which are calcium sensors generally located within the endoplasmic reticulum (ER) membrane, can detect decreased calcium concentrations ([(Ca²⁺)] in the reticulum through the EF-hand domains located in the N-terminal part of the proteins. A reduction in the amount of calcium activates STIM by inducing oligomerization and changes in the protein structure, which allows the cytosolic tail to activate ORAI, a calcium-permeable channel located in the plasma membrane. This calcium entry is then transported in the ER by sarcoplasmic (SR)/ER calcium ATPase 1 (SERCA1). Although there are several members of the STIM and ORAI families,

STIM1 and ORAI1 isoforms are the major SOCE-mediating proteins. STIM1 is more efficient than STIM2 in triggering calcium influx and only becomes active in response to significant depletion of intracellular calcium stores [9]. Splicing isoforms of STIM1 exist, some being muscle cell-specific, such as STIM1L, and others exhibiting an inhibitory effect on SOCE, such as STIM1A. The most highly and ubiquitously expressed STIM1 protein is a type I transmembrane protein comprising 685 amino acids. Similar to all other isoforms in the family, it comprises two EF-hand domains in the luminal portion, which can bind and detect calcium levels, and a sterile alpha motif (SAM), which promotes oligomerization of the protein upon activation. Three coiled-coil (CC) domains have been identified in the cytosol. CC1 expands to fill the gap between the reticulum and the plasma membrane during STIM1 activation. The CC2 and CC3 domains form the Stim Orai Activated Region (SOAR), which interacts directly with ORAI1 to open it.

Mutations in genes encoding CRAC channel proteins, especially ORAI1 and STIM1, can lead to specific diseases due to alterations in protein functions. Gain-of-function mutations associated with the

<footnote><sup>a</sup> Aix Marseille Univ, INSERM, MMG, U1251 Marseille, France</footnote>

<footnote><sup>b</sup> Centre de Référence Maladies Rares Neuromusculaire, CHU Toulouse, Toulouse, France</footnote>

<footnote><sup>c</sup> Université Paris-Saclay, INSERM, U1193 Orsay, France</footnote>

<footnote><sup>d</sup> Laboratoire de Génétique Moléculaire, CHU Montpellier, Montpellier, France</footnote>

<footnote><sup>e</sup> PhyMedExp (Physiologie et Médecine Expérimentale du Cœur et des Muscles), Université de Montpellier, Inserm U1046, CNRS UMR9214, Montpellier, France</footnote>

<footnote><sup>f</sup> Département de Génétique Médicale, Hôpital Timone Enfants, APHM, Marseille, France</footnote>

<footnote><sup>g</sup> CNRS, Marseille, France</footnote>

<footnote>* Corresponding author at: Aix Marseille Université, INSERM, MMG, U1251, 13385, Marseille Cedex 05, France.</footnote>
constitutive activation of SOCE are responsible for tubular aggregate myopathy (TAM) (MIM 160565) and Stormorken syndrome (MIM 185070). In contrast, loss-of-function mutations associated with insufficient SOCE are associated with CRAC channelopathies. CRAC channels are characterized by severe combined immunodeficiency (SCID) involving recurrent and chronic infections, autoimmunity, muscular hypotonia, ectodermal dysplasia, anhidrosis, and mydriasis (MIM 612783) [24].

Most mutations that occur in *STIM1* are missense mutations that can lead to either a loss or gain of function [14]. In addition, insertions and deletions can lead to abnormal protein functions [17,20,23]. A single splice mutation has been described in *STIM1* [2]. This mutation affects a consensus splice acceptor site in *STIM1* before exon 8, resulting in the formation of several abnormal transcripts with partial retention of intron 7 or deletion of exon 8 but complete absence of the wild-type (WT) transcript. This mutation increases the risk of developing Kaposi sarcoma caused by human herpesvirus-8 [2].

Herein, we describe a new homozygous c.792-3C > G splicing mutation in *STIM1* (NM_003156) 1 bp upstream of the 3' acceptor splice site in *STIM1* intron 6 in a patient, born of consanguineous parents, and presenting a clinical profile of CRAC channelopathy without severe immunodeficiency. This novel mutation leads to abnormal splicing and production of non-functional proteins. As a therapeutic approach, we tested an antisense oligonucleotide (AON) to restore normal splicing in patient cells.

## 2. Materials and methods

### 2.1. Genetic analyses

Next-generation sequencing (NGS) was performed on the extracted DNA from patient cells using a customized probe targeting neuromuscular genes as previously described [11,27]. The pathogenicity of variants was assessed through a set of criteria reported in the literature [27] and guidelines from the American College of Medical Genetics and Genomics (ACMG) [22]. Prediction of splicing variants effect were scored with online bioinformatics tools including Human Splicing Finder (HSF, https://hsf.genomnis.com/login) and SpliceAI (https://spliceailookup.broadinstitute.org/) [3,10]. The frequency of each variant in the general population was then evaluated using the Genome AggregationV3 database (gnomAD, http://gnomad.broadinstitute.org/) while variants were identified using the Human Gene Mutation Database (HGMD®) [25].

The *STIM1* variant was confirmed by Sanger sequencing and reported according to Human Genome Variation Society recommendations (http://varnomen.hgvs.org).

### 2.2. Cell culture

Immortalized lymphoblastoid cells from a patient carrying a homozygous intronic mutation in *STIM1* were prepared from heparin blood samples transformed with Epstein-Barr virus (EBV) using standard procedures at the accredited Biological Resource Center, Biogénopôle, Timone Hospital of Marseille. The cell line was stored at the CRB TAC and belonged to a biological sample collection declared to the French Ministry of Health, and the use for research purposes was authorized by the French Ministry of Health.

The three-control immortalized lymphoblastoid cell lines were purchased from the Coriell Cell Repository.

Cells were cultured in Roswell Park Memorial Institute (RPMI) 1640 medium GlutaMAX™ supplemented with 10% heat-inactivated fetal bovine serum (FBS), 50 U/mL penicillin, and 50 µg/mL streptomycin (Gibco™).

### 2.3. Reverse transcription-polymerase chain reaction (RT-PCR) and sequencing

RNAs were extracted from controls and patient lymphoblastoid cells line using the PureLink™ RNA Mini Kit (Invitrogen™ Thermo Fisher) and reverse transcribed using a high-capacity cDNA reverse transcription kit (Thermo Fisher Scientific, Waltham, USA).

STIM1 transcripts were analyzed using PCR and Sanger sequencing with the following primers: forward primer in exon 5, 5'-ATACAGTGCTCTTTGGGCCT-3' and reverse primer in exon 9, 5'-GGAAGGTGCCAAAGAGTGTG-3'. After gel-cutting, the PCR bands were purified using the NucleoSpin™ Gel and PCR Clean-up Kit (Macherey-Nagel™) and then sequenced using GeneWize.

### 2.4. RNA sequencing (RNA-Seq)

RNA-Seq and bioinformatic analyses were performed at the Genomics and Bioinformatics facility (GBiM) located in the U 1251/Marseille Medical Genetics lab. mRNA sequencing (mRNA-Seq) was performed in triplicate on total RNA samples extracted from immortalized lymphoblastoid cells of the patient and controls. Before sequencing, the quality of the total RNA samples was assessed using an Agilent Bioanalyzer (Agilent Technologies, USA), and RNAs with RNA integrity numbers (RIN) of >8 was selected for sequencing and used for library preparation. For each sample, a library of stranded mRNA was prepared from 500 ng of total RNA after capture of RNA species with a poly(A) tail using the Kapa mRNA HyperPrep kit (Roche, Switzerland), according to the manufacturer's instructions. The quality and profile of individual libraries have been quantified and visualized using Qubit™ and the Agilent Bioanalyzer dsDNA High Sensibility Kit (Agilent Technologies, USA), respectively. Indexed libraries were pooled and sequenced (2*101 bp paired-end sequencing) using a NovaSeq6000 platform (Illumina, USA). The quality of sequencing reads was assessed using fastQC v0.11.5 (https://www.bioinformatics.babraham.ac.uk/project/s/fastqc/) software. Raw sequencing reads were mapped to the human reference genome GRCh38 (hg38) and sorted using STAR v2.7.2b software, and bam files were indexed using Sambamba v0.6.6 software. After mapping, the number of reads per feature (GENCODE v34 annotations) was determined using Stringtie v1.3.1c software [19].

The presence of splicing abnormalities at the *STIM1* locus was studied through visual inspection exon-exon junctions wing IGV v 2.16.1 software with splice junction tracks and Sashimi plots. For sashimi plots, low-count splicing events were filtered by setting the minimum junction coverage to five reads.

### 2.5. Western blot analysis

Proteins were extracted from immortalized lymphoblastoid cell lines from controls and patient using radioimmunoprecipitation assay (RIPA) buffer (Life technologies) and a protease and phosphatase inhibitor cocktail (Life technologies). After a bicinchoninic acid (BCA) dosage, 20 µg of protein was loaded on each well of a 4–12% NuPAGE Bis-Tris gel for sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) (Life Technologies), using Chameleon Duo as a size marker. Proteins are transferred onto nitrocellulose membrane at 30 V overnight at 4 °C. The membranes were blocked using intercept blocking buffer (Li-Cor) for 30 min at room temperature (RT). Primary antibodies (anti-STIM1 N-terminal S6072 synthetic peptide corresponding to amino acids 61–74 of human *STIM1*, 1:500; anti-STIM1 C-terminal S6197 synthetic peptide corresponding to amino acids 657–683 of human *STIM1*, 1:500; and anti-GAPDH ab125247, 1:1000) were diluted in intercept blocking buffer supplemented with 0.1% Tween and incubated for 2 h at RT. After three washes in phosphate-buffered saline (PBS) + 0.1% Tween, membranes were incubated with secondary antibodies (IR Dye 680RD donkey anti-mouse and IR Dye 800CW donkey anti-rabbit, Li-Cor, 1:10,000) in the intercept blocking buffer for 45 min at RT.
The membranes were then washed thrice with PBS + 0.1% Tween and once with PBS. Proteins were visualized using the LI-COR system and quantified using ImageJ software, with GAPDH as a loading control.

## 2.6. Immunofluorescence

For immunofluorescence analysis, cells were centrifuged on slides using a Cytospin centrifuge (50,000 cells per slide) and fixed in 4% paraformaldehyde at RT for 10 min. The cells were permeabilized (0.5% Triton X-100 in PBS) for 2 min and then blocked with blocking buffer (0.1% Triton X-100, 5% goat serum, and 5% BSA in PBS) at RT for 1 h. Next, the cells were incubated with primary antibodies (Anti-STIM1 C-terminal S6197, 1:500) diluted in blocking buffer at RT for 2 h. After washing in PBS, the cells were incubated with the secondary antibody, donkey anti-rabbit 555 (1:700, ab150074), for 1 h at RT. After washing thrice with PBS, cells were mounted in ProLong™ Gold Antifade Mountant with 4',6-diamidino-2-phenylindole (DAPI, P36935). Cells were observed under a Confocal LSM 800 Airyscan microscope with a 63×/1.20 objective.

## 2.7. Cytosolic Ca²⁺ concentration

[Ca²⁺] cytosolic (cyt) was recorded using a ratiometric fluorescence technique as described previously [4,8]. Lymphoblastoid cell lines from the controls and from the patient were spun and resuspended in Ca²⁺-free Hepes buffered saline solution (HBSS; 135 mM NaCl, 5.9 mM KCl, 1.2 mM MgCl₂, 11.6 mM Hepes, and 11.5 mM glucose, adjusted to pH 7.3 with NaOH) and incubated in the dark with 4 µM Indo-1-AM for 45 min at room temperature with slow agitation. The cells were then centrifuged and resuspended in dye-free HBSS. For the experiments, 0.5–1 × 10⁶ cells were suspended in 2 mL Ca²⁺-free HBSS in a quartz cuvette and inserted into a spectrofluorophotometer (RF-1501 Shimadzu Corporation, Kyoto, Japan) connected to a computer (Dell Computer Corp., Montpellier, France). A temperature of 37 °C was maintained by circulating water in a thermostatic bath. Ultraviolet light at 360 nm was used to excite Indo-1, and emissions at 405 and 480 nm were recorded. The background and autofluorescence of the cell suspensions were subtracted from the recordings. The maximum dye fluorescence (Rmax) was obtained by adding 1 µM ionomycin to the cell suspension in the presence of 10 mM CaCl₂. Minimum fluorescence was determined following the depletion of external Ca²⁺ with 5 mM ethylene glycol tetraacetic acid (EGTA). [Ca²⁺]cyt was calculated according to the equation [Ca²⁺]cyt = Kd (R-Rmin)/(Rmax-R)F, where Kd is the apparent dissociation constant of the Ca²⁺-dye (230 nM for Indo-1), R is the ratio of fluorescence at 405 and 480 nm for Indo-1, and F is the ratio of fluorescence values recorded at 480 nm in the absence and presence of 10 mM CaCl₂ [8].

To visualize the SOCE as a result of activation of ORAI1 channels by STIM1 proteins, we performed a classical two phases protocol: 1) 1 mM EGTA was added to the cells to remove the traces of Ca²⁺ ions. Next, cells were treated with 1 µM thapsigargin (TG), a well-known inhibitor of ER SERCA pumps, inducing a passive depletion of ER Ca²⁺ ions, allowing the oligomerization of STIM1 proteins and the opening of ORAI1 channels. After complete removal of cytosolic Ca²⁺ ions (400 s), 1 mM external CaCl₂ was added to visualize the massive [Ca²⁺]cyt due to SOCE. The experiments were repeated ten times with two immortalized cell lines in parallel. The difference in calcium entry was evaluated using Student's t-test. Differences with a p-value <0.05 were considered statistically significant.

## 2.8. Exercise test

An electrophysiological exercise test was performed to explore muscle channelopathy as described by Fournier and colleagues [7]. We performed an exercise test consisting of performing an effort for 10 s, followed by recording the Compound Muscle Action Potential (CMAP)

every 10 s for 1 min by electrical stimulation of the ulnar nerve at the level of the wrist and recording on the adductor of the 5th finger. This test was repeated thrice. We then performed a long exercise test after a 5-min adduction effort of the 5th finger and a similar recording of the CMAP on the adductor of the 5th finger. Recordings were performed every minute for the first 5 min then every 5 min for up to 45 min.

## 2.9. Treatment of cells with antisense oligonucleotide

AON were designed by selecting complementary sequences at the end of intron 6 containing the alternative splice acceptor site. AON was ordered from Eurogentec (Angers, France) with the following sequence: CAGGGAGCUGGAGGAGG. AON was 2'-O-methylated on the first two and last two bases and were phosphorothioate-modified on all bases except the last base. These chemical modifications increase stability and improve affinity for the target sequence [1].

Cells were plated in a six-well plate on the day of treatment at a rate of 1 million cells in 800 µL of RPMI 1640 medium (Gibco™) supplemented with 1× L-Glutamine (Gibco™). AON were transfected using Oligofectamine transfection reagent (Thermo Fisher Scientific, USA). Each well contained 2 µL of Oligofectamine, 20 µL of 20 µM AON or PBS and 178 µL of Opti-MEM (Gibco™). After 4 h of transfection, 1 mL of RPMI 1640 medium supplemented with 20% heat-inactivated FBS, 100 U/mL penicillin, 100 µg/mL streptomycin, and 1× L-Glutamine are added to each well. After 24 or 48 h of treatment, the cells were counted, rinsed in PBS after 5 min of centrifugation at 1000 g and used for labeling or extraction.

# 3. Results

## 3.1. Clinical description

The patient was born to consanguineous parents (Fig. 1A) and was 42 years old at the time of this study. The patient was born at full term without any neonatal distress. She presented with a delay in walking acquisition associated with difficulties in sucking during the early years. During childhood, she presented with a progressive worsening of the muscular deficit of the pelvic girdle and difficulty climbing stairs. From the age of 15 years, she developed fluctuations of weakness that occurred as a myasthenic crisis that could last several weeks without rhabdomyolysis. Electrophysiological and immunological tests for neuromuscular junction abnormalities were negative. She underwent three muscle biopsies, which revealed irregularities in fiber size. The deficit progressed with the loss of walking ability the age of 25 years and was associated with a distal deficit in flexion and extension of the hands and feet. Several tests, including an exercise test, have been performed in the absence of a diagnosis to detect possible channelopathy. A short exercise test did not demonstrate any modification of CMAP post-exercise. However, after a long exercise period, there was an immediate post-exercise reduction in CMAP of approximately 18%, reaching 34% at after 2 min. Gradually, the CMAP amplitude returned to the pre-exercise value after 15 min (data not shown).

This patient has the particularity of having presented numerous pathologies mostly carcinologic and auto-immune, such as Hodgkin's lymphoma at the age of 15 years and recently a breast carcinoma; Crohn's disease, sclerosing cholangitis, thrombocytopenic purpura and primary hyperparathyroidism. The patient presented with amelogenesis imperfecta.

## 3.2. Identification of a novel mutation in STIM1

Next-generation sequencing (NGS) analysis of the exons and exon-intron junctions of 185 genes involved in neuromuscular disorders [11,27] revealed a homozygous variant of *STIM1* (NM_003156), c.792-3C > G (Fig. 1B), localized near the acceptor splice site of *STIM1* exon 7. This variant is rare in the general population (gnomAD V3:0.0007%).
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Identification of a novel STIM1 mutation. (A) Family pedigree with the index case indicated by an arrow. (B) Electropherogram showing the identified STIM1 mutation c.792-3C > G (NM_003156). (C) Schematic representation of the STIM1 gene with the newly identified mutation in red. The different splicing possibilities and associated proteins are shown: WT splicing and WT protein (solid line connection). Utilization of a cryptic splice site leading to the formation of a transcript with 21 extra nucleotides and the insertion of 7 supplementary amino acids (dot-dash line connection) in the CC1 domain of the protein. Exon 7 skipping leading to a truncated protein p.Leu265Phefs*4 (dotted line connection). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>**Fig. 2.** Effect of the mutation at the transcriptional level. (A) RT-PCR analysis of STIM1 using mRNA from the patient's (Patient) and a control's (Ctrl 3543) immortalized lymphoblastoid cell line as well as a negative control without mRNA (–Ctrl). (B) Chromatogram obtained after sanger sequencing of the different RT-PCR bands. (C) Sashimi plot generated by IGV centered on exon 6, 7 and 8 of STIM1 after RNA sequencing data from the patient and the control (Ctrl 3543). Alignments in exons are represented as read densities while splice junction reads are drawn as arcs connecting pairs of exons, with the number of reads aligning to the junction indicated.</figcaption></figure>
This homozygous variant was confirmed using Sanger sequencing (Fig. 1B). No other candidate variants were identified in the 185 analyzed genes.

We used the *Human Splicing Finder* (HSF) and *Splice AI* tools to predict the effect of this mutation on *STIM1* splicing [3,10]. HSF predicted skipping of exon 7, and Splice AI predicted a loss of the acceptor with a score of 0.69, as well as a gain of a new acceptor site with a score of 0.5 (Fig. 1C). This new cryptic splice site is located 21 nucleotides before the canonical splice site. Thus, the mutation is likely to lead to a transcript with retention of the last 21 nucleotides from intron 6, and subsequently, the mutation is located at position -3 in this intron.

### 3.3. Effect of the mutation at the transcriptional level

To determine if this variant, located prior the 3' splice site of exon 7, has an effect on splicing, we performed RT-PCR using RNA extracted from the patient and control immortalized lymphoblastoid cell lines. The primers used were located in exons 5 and 9, and the flanking exons were likely affected. After RT-PCR, we loaded the PCR products onto an agarose gel and observed a single band (C1) at the expected size of 597 bp in the control, corresponding to the normal splicing of exons 5 to 9 (Fig. 2A). In contrast, there were three different fragments at 618, 597, and 419 pb in the patient sample, likely matching the transcripts with the retention of the last 21 nucleotides of intron 6 (P1), the WT transcript (P2), and the transcript with exon 7 skipping (P3), respectively.

Sequencing of these fragments using Sanger sequencing (Fig. 2B) confirmed that i) the highest band (P1) matched the retention of the last 21 nucleotides from intron 6, clearly showing the C > G variation found in patient cells, ii) the lowest band (P3) matched a transcript skipping whole exon 7, whereas iii) the middle band (P2) displayed the WT transcript (C1).

To determine the proportion of each transcript, we performed RNA-Seq of the total RNA extracted from the patient and control cells, which confirmed the presence of three different spliced transcripts (Fig. 2C), visible on the sashimi plot. The exons 6, 7 and 8 are seen as read densities while splice junction reads are drawn as arcs connecting pairs of exons, with the number of reads aligning to the junction indicated [12]. In summary, the homozygous c.792-3C > G splicing variant found in *STIM1* (NM_003156) led to the production of three different types of splicing events in this region. First, we observe a WT splicing, identical to that in controls (solid line in Fig. 1C). Then there is splicing by skipping exon 7 resulted in a reading frame shift with the appearance of a premature stop codon, which was predicted to produce a putative truncated protein, p.Leu265Phefs\*4 (dotted line in Fig. 1C). Finally, there is also splicing with retention of the 3' section of intron 6, causing the in-frame addition of 21 nucleotides with the following sequence: CTCCCTGCATTGCCCCCCCAG, as observed in the zoomed Sashimi plot (Fig. 2C). If a protein is produced from this transcript, it includes the insertion of the following seven amino acids into the CC1 domain: SSLHCPP (dot-dash line Fig. 1C).

Interestingly, RNA-seq data showed that, in the patient sample, reads were found throughout intron 6, whereas they were not found in the control as shown in Fig. 2C.

### 3.4. Effect of the mutation at the protein level

To determine whether these three types of transcripts led to protein production, we performed immunoblotting and immunocytochemistry.

The expected molecular weights of the proteins obtained from the WT, intronic-retention, and exon-skipping transcripts were 77.4, 78.1, and 30.9 kDa, respectively. The WT (77.4 kDa) band migrates to a relative molecular weight of 90 kDa.

Using an antibody targeting the N-terminal region of STIM1, which theoretically recognizes all three different proteins, we obtained a single band of the same size in all tested individuals (patient and three control cell lines). Our experimental conditions were unable to differentiate WT

proteins from proteins with the potential insertion of an additional seven amino acids because the expected sizes were too close. However, the patient cells showed a significant decrease of approximately 33% in the amount of STIM1 compared to the controls (Fig. 3A).

Additionally, the absence of bands at approximately 30 kDa in the patient's sample revealed that the exon 7 skipped transcript did not lead to a stable protein (Fig. 3A).

Similar to the results obtained with the N-terminal antibody, a significant decrease of 33% in STIM1 protein was observed in the patient using the C-terminal antibody (Fig. 3B).

The N-terminal antibody used for immunocytochemical analyses did not reveal any staining both in control and patient cells. Whereas the STIM1 C-terminal antibody staining is present in the control cells but was hardly visible in the patient's immortalized lymphoblastoid cell lines (Fig. 3C). However, a protein was present as observed by immunoblotting with the same antibody under denaturing conditions (Fig. 3B).

### 3.5. Functional abnormalities

Because STIM1 expression levels were approximately 67% in the patient cells, we further investigated whether the produced proteins were functional. We performed an experiment evaluated the ability of STIM1 to activate ORAI1 channels, resulting in SOCE, upon calcium storage depletion (Fig. 4A). Thapsigargin (TG), a well-known inhibitor of the ER Ca²⁺ ATPase SERCA, was used to induce Ca²⁺ release from the ER and activate STIM1. As shown in Fig. 4, in the absence of extracellular Ca²⁺ ions, TG induced a similar transient increase in [Ca²⁺]cyt in both the patient and control cells as estimated by the area under the curve (Fig. 4B: 37171 ± 1348 vs. 39,590 ± 4580 arbitrary unit for control and patient cells respectively, n = 10). As there is no extracellular calcium and the ER cannot fill up in the presence of TG, this [Ca²⁺]cyt is only due to Ca²⁺ ions released by the ER. This showed that the amount of Ca²⁺ in the ER was similar in control and patient cells. After peaking in the cytosol, calcium ions released by the ER are progressively extracted from the cells by the plasma membrane Ca²⁺ ATPases (PMCA), inducing the slow decrease of [Ca²⁺]cyt.

In the presence of TG, Ca²⁺ ions stored in the ER lumen are released, inducing STIM1 oligomerization and the interaction of STIM1 with Orai1 channels, allowing their opening and SOCE. The addition of 1 mM extracellular Ca²⁺ ions induced a large and rapid [Ca²⁺]cyt increase in the control lymphoblastoid cell line (Fig. 4C: [Ca²⁺]cyt peak of 1654 ± 100 nM, n = 10) owing to the massive entry of Ca²⁺ ions through ORAI1 channels. However, in patient cells, after Ca²⁺ addition, the peak of [Ca²⁺]cyt was significantly reduced by ≈ 95% (Fig. 4C:136 ± 14 nM, n = 10, p < 0.001), meaning there is a huge decrease in calcium entry.

These results indicate that the STIM1-ORAI1 pathway is completely impaired by mutations in *STIM1* in the patient cells.

### 3.6. Therapeutic approach

To restore normal STIM1 splicing, we designed an antisense oligonucleotide (AON), which specifically masks the cryptic acceptor site, as shown in Fig. 5A. The AON penetrates the nucleus and hybridizes complementarily with a pre-messenger RNA sequence. With this method, we aimed to improve the recognition of the WT 3' splice site, thereby increasing the proportion of WT transcripts. After 24 h of treatment without AON, STIM1 C-terminal staining was barely visible. In contrast, after treatment with AON, STIM1 staining was observed in almost all the patient cells (Fig. 5B). Thus, by masking the cryptic donor splice site, we restored to a staining of STIM1, comparable to that in the control (Fig. 3C).

## 4. Discussion

In this study, we identified a novel homozygous mutation, c.792-3C
<figure><img src="image_7.png" /><figcaption>**Fig. 3.** Protein analysis. Western blot analysis on immortalized cell lysates from three different controls (Ctrl 735, 647 and 3543) and the patient (Patient) for STIM1 N-terminus (A) and STIM1 C-terminus (B). Data are mean ± SEM for *n* = 3. Statistical analysis was performed with Mann-Whitney test (*: p-value <0.05). (C) Immunofluorescence staining of STIM1 (using C-terminal antibody S6197, red) and the nucleus (DAPI, blue) in immortalized control (Ctrl 3543) and patient (Patient) lymphoblastoid cell line. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Functional test. (A) Cytosolic Ca²⁺ concentration ([Ca²⁺]cyt) measurement in immortalized cell line from the patient (red) and control (black) using Indo-1 fluorescence. Cells were treated with thapsigargin (1 µM, black arrow) to induce Ca²⁺ release from endoplasmic reticulum and opening of the SOCE channels. Then, 1 mM CaCl₂ was added, allowing Ca²⁺ entry through the SOCE channels. Curves are mean ± SEM (grey) from 10 repeats. (B) ER calcium release in patient and control cells are represented by the area under the curve after thapsigargin treatment. (C) Quantification of the Store Operated Calcium Entry through ORAI1 channel. Statistical analyses were performed with a t-test of Student (***: p-value <0.01). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Therapeutic approach. (A) Design of an AON (in green) masking the cryptic acceptor splice site. (B) Immunofluorescence staining of STIM1 (C-terminal antibody S6197, in red) and the nucleus (DAPI, in blue) in immortalized patient lymphoblastoid cell line treated with or without AON for 24 h. No STIM1 signal was apparent in patient cells without AON treatment, while STIM1 signal is observed in almost all patient cells after AON treatment. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

> G, within the *STIM1* gene in a patient from a consanguineous family. This patient presented with a spectrum of pathologies, mostly autoimmune and carcinogenic, such as Crohn's disease, sclerosing cholangitis, thrombocytopenic purpura, primary hyperparathyroidism, Hodgkin's lymphoma, and breast carcinoma. She also had muscle weakness and amelogenesis imperfecta. This specific intronic mutation induces complex splicing. This discovery sheds light on the potential implications of altered STIM1 expression and its multifaceted clinical manifestations.

In patient cells, three distinct STIM1 transcripts were present in roughly equal proportions, each accounting for approximately 33% of the total transcripts. Notably, skipping exon 7 and introducing a premature stop codon did not yield a truncated protein. Instead, it is likely targeted for degradation through nonsense-mediated decay (NMD). This suggests the presence of a cellular quality control system that identifies and eliminates potentially mutated STIM1 transcripts.

In the RNA-Seq data, reads were identified throughout intron 6 only in the patient sample. The mutation, which reduces the strength of the splice site, leads not only to skipping of exon 7 or retention of the 3' part of intron 6, but also to more complex forms such as a form with the total retention of intron 6. Byun et al. showed that the only splicing mutation reported in *STIM1* also results in varying degrees of intronic retention of the affected intron [2].

Using western blotting, we showed that approximately 67% of the proteins appeared at the expected size of STIM1 full-length in the patient compared to controls. However, notably, this method cannot distinguish between WT proteins and those with the insertion of seven amino acids, because of partial intron 6 retention. The proteins observed in the patient cells corresponded to 33% of the WT transcripts and 33% of the transcripts with intronic retention. This proportion was consistent with the proportions identified using RNA-Seq, and visible on Sashimi Plot, in which these transcripts were equally abundant.

Although 67% of the proteins were detected by western blotting under denaturing conditions, immunochemistry failed to detect any proteins in the patient's cells. This can be explained by the fact that the transcript with 21 bp intron 6 retention has a preserved reading frame and therefore may produce a protein with seven additional amino acids. This insertion appears after amino acid 263 within the cytosolic section of the protein in the C-terminal extension of helix α1 found in coiled-coil domain 1. Thus, this molecular alteration highlights a change in protein

structure that may block detection using standard immunocytochemical techniques.

In addition, Lupas et al. analyzed the frequency of amino acids in coiled-coil domains and observed that prolines are rarely encountered and should not be positioned consecutively within this structure [15]. However, in the case of intronic retention, two consecutive proline residues are inserted into the CC1 domain, leading to an aberrant tertiary structure. This structural anomaly could potentially destabilize the cytosolic portion of the protein and mask the epitope binding site of the C-terminal antibody, as verified by immunocytochemistry. In contrast, during immunoblot, the proteins are denatured, losing their tertiary structure and thereby exposing the antibody-binding site.

The CC1α1 domain maintains STIM1 in an inactive state with a clamped conformation by interaction with the CC3 domain. In particular, residues 258-261 are crucial for this interaction, and mutations in this region lead to constitutive activation of the protein, which is constantly in an extended conformation [5,16]. In contrast, mutations in the loop between α1 and α2, which begins at amino acid 272, reinforce the closed conformation of the protein. It has been shown that weakening CC1 α1-α2 interhelical interactions promotes the resting state of STIM1 and decreases the protein activation kinetics [21].

The insertion of these seven amino acids appears in the middle of critical regions involved in the regulation of STIM1 activation. This insertion may enhance the closed conformation of STIM1, resulting in a loss of protein function that is manifested in biochemical tests.

In the present case, despite the autosomal recessive nature of the disease, splicing defects did not preclude the presence of the WT form. However, the presence of this variant did not mitigate the onset of symptoms. This phenomenon can be explained in two ways: either the WT variant is insufficient, resulting in haploinsufficiency, or the mutated variant exerts a dominant-negative effect, thereby hindering the WT variant function. In scenarios involving protein oligomerization, mutated monomers with dominant-negative traits can disrupt and impair the functionality of the entire complex [26]. Although RNA-Seq indicated a proportion of WT transcripts, the absence of detectable proteins using immunocytochemistry may be attributed to the potential of abnormal proteins to induce misconfiguration of STIM1 dimers during their resting state. Consequently, these altered dimers may evade detection under standard physiological conditions, particularly when using a C-terminal antibody.

This hypothesis of a dominant-negative effect is supported by functional tests at both the cellular and clinical levels. At the cellular level, comparisons between patient and control groups revealed identical reticular calcium storage levels. Following brief effort, the patient muscle mobilized only the reticular calcium, and no decrease in CMAP was observed (Fig. Sup. 1 A). However, divergence emerged as the patient's cells displayed an inability to activate SOCE. This deficiency was highlighted by the absence of calcium influx following the depletion of intracellular stores induced by TG. Indeed, the patient's cells show only a small influx of calcium following depletion of intracellular store likely due to $Ca^{2+}$ ion leakage through channels other than ORAI1. Typically, during intense exercise, calcium stores are rapidly depleted and SOCE replenishes these stores. After intense exercise, the patient showed a significant reduction in CMAP post-exercise, indicating an inability to effectively restore the calcium stores that are essential for proper muscle function. Interestingly, patient muscular capacity was restored 15 min after post-exercise, suggesting that an alternative mechanism facilitated the replenishment of calcium stores (Fig. Sup. 1B). This process may explain the absence of a severe form of combined immunodeficiency typically associated with STIM1 loss-of-function mutations [6]. The complex nature of the cellular responses requires further exploration to fully understand the physiological implications of this mutation.

Restoring the natural splicing of STIM1 can improve the 'function of the protein and, thus, the phenotype observed in the patient. To achieve this objective, we designed an AON to reduce the proportion of transcripts with intronic sequence retention, while increasing the abundance
of WT transcripts. After treatment, STIM1 labeling was restored based on the immunocytochemistry analysis, indicating the effect of treatment. However, additional studies are required to confirm whether the presence of STIM1 is restored, as well as its functional activity and capacity to rescue the phenotype. These findings highlighted the viability of this treatment and its potential as a therapeutic avenue for addressing the conditions of the patient.

Treatments using AONs are currently being used for other pathologies, particularly Duchenne muscular dystrophy, for which several AONs are being tested in clinical trials [13]. Our study can be considered a proof of concept for the development of therapeutic AONs from the perspective of treatment for this patient.

## Funding

This work was supported by AFM-Telethon grant under the strategic program Mothard.

## CRediT authorship contribution statement

**Benedicte Alary:** Writing – review & editing; Writing – original draft, Methodology, Investigation, Formal analysis, Conceptualization.
**Pascal Cintas:** Writing – review & editing, Methodology, Investigation.
**Corentin Claude:** Investigation. **Olivier Dellis:** Writing – review & editing, Methodology, Investigation, Formal analysis. **Corinne Thêze:** Investigation. **Charles Van Goethem:** Investigation. **Mireille Cossée:** Writing – review & editing, Methodology, Investigation. **Martin Krahn:** Writing – review & editing, Funding acquisition. **Valérie Delague:** Writing – review & editing, Methodology, Funding acquisition. **Marc Bartoli:** Writing – review & editing, Writing – original draft, Supervision, Project administration, Methodology, Funding acquisition, Conceptualization.

## Declaration of competing interest

None.

## Data availability

The data that support the findings of this study are available on request from the corresponding author.

## Acknowledgements

We sincerely thank the patient. We appreciate the assistance of the staff of the GBIM facility for RNA-seq analysis. We sincerely thanks Dr. Claude Villard for his comment on mass spectrometry. We would like also to thank Dr. Anne Plauzolles for critical reading of the manuscript.

## Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.clim.2024.110306.

## References

[1] A. Aartsma-Rus, Overview on AON design, in: A. Aartsma-Rus (Ed.), Exon Skipping, Methods in Molecular Biology, Humana Press, Totowa, NJ, 2012, pp. 117–129, https://doi.org/10.1007/978-1-61779-767-5_8.

[2] M. Byun, A. Abhyankar, V. Lelarge, S. Plancoulaine, A. Palanduz, L. Telhan, B. Boisson, C. Picard, S. Dewell, C. Zhao, E. Jouanguy, S. Feske, L. Abel, J.-L. Casanova, Whole-exome sequencing-based discovery of STIM1 deficiency in a child with fatal classic Kaposi sarcoma, J. Exp. Med. 207 (2010) 2307–2312, https://doi.org/10.1084/jem.20101597.

[3] F.-O. Desmet, D. Hamroun, M. Lalande, G. Collod-Béroud, M. Claustres, C. Béroud, Human splicing finder: an online bioinformatics tool to predict splicing signals, Nucleic Acids Res. 37 (2009) e67, https://doi.org/10.1093/nar/gkp215.

[4] A. Djillani, O. Nüße, O. Dellis, Characterization of novel store-operated calcium entry effectors. Biochim. Biophys. Acta BBA - Mol. Cell Res. 1843 (2014) 2341–2347, https://doi.org/10.1016/j.bbamcr.2014.03.012.

[5] M. Fahrner, M. Muik, R. Schindl, C. Butorac, P. Stathopoulos, L. Zheng, I. Jardin, M. Ikura, C. Romanin, A coiled-coil clamp controls both conformation and clustering of stromal interaction molecule 1 (STIM1), J. Biol. Chem. 289 (2014) 33231–33244, https://doi.org/10.1074/jbc.M114.610022.

[6] S. Feske, CRAC channels and disease – from human CRAC channelopathies and animal models to novel drugs, Cell Calcium 80 (2019) 112–116, https://doi.org/10.1016/j.ceca.2019.03.004.

[7] E. Fournier, M. Arzel, D. Sternberg, S. Vicart, P. Laforet, B. Eymard, J. Willer, N. Tabti, B. Fontaine, Electromyography guides toward subgroups of mutations in muscle channelopathies, Ann. Neurol. 56 (2004) 650–661, https://doi.org/10.1002/ana.20241.

[8] G. Grynkiewicz, M. Poenie, R.Y. Tsien, A new generation of Ca2+ indicators with greatly improved fluorescence properties, J. Biol. Chem. 260 (1985) 3440–3450.

[9] P.G. Hogan, A. Rao, Store-operated calcium entry: mechanisms and modulation, Biochem. Biophys. Res. Commun. 460 (2015) 40–49, https://doi.org/10.1016/j.bbcrr.2015.02.110.

[10] K. Jaganathan, S. Kyriazopoulou Panagiotopoulou, J.F. McRae, S.F. Darbandi, D. Knowles, Y.I. Li, J.A. Kosmicki, J. Arbelaez, W. Cui, G.B. Schwartz, E.D. Chow, E. Kanterakis, H. Gao, A. Kia, S. Batzoglou, S.J. Sanders, K.K.-H. Farh, Predicting splicing from primary sequence with deep learning, Cell 176 (2019) 535–548.e24, https://doi.org/10.1016/j.cell.2018.12.015.

[11] R. Juntas Morales, A. Perrin, G. Solé, D. Lacourt, H. Pegeot, U. Walther-Louvier, P. Cintas, C. Cances, C. Espil, C. Theze, R. Zenagui, K. Yauy, E. Cosset, D. Renard, V. Rigau, A. Maues De Paula, E. Uro-Coste, M.-C. Arne-Bes, M.-L. Martin Négrier, N. Leboucq, B. Acket, E. Malfatti, V. Biancalana, C. Metay, P. Richard, J. Rendu, F. Rivier, M. Koenig, M. Cossée, An integrated clinical-biological approach to identify Interindividual variability and atypical phenotype-genotype correlations in myopathies: experience on a cohort of 156 families, Genes 12 (2021) 1199, https://doi.org/10.3390/genes12081199.

[12] Y. Katz, E.T. Wang, J. Silterra, S. Schwartz, B. Wong, H. Thorvaldsdóttir, J. T. Robinson, J.P. Mesirov, E.M. Airoldi, C.B. Burge, Quantitative visualization of alternative exon expression from RNA-seq data, Bioinformatics 31 (2015) 2400–2402, https://doi.org/10.1093/bioinformatics/btv034.

[13] T. Koo, M.J. Wood, Clinical trials using antisense oligonucleotides in Duchenne muscular dystrophy, Hum. Gene Ther. 24 (2013) 479–488, https://doi.org/10.1089/hum.2012.234.

[14] R.S. Lacruz, S. Feske, Diseases caused by mutations in ORAI1 and STIM1, Ann. N. Y. Acad. Sci. 1356 (2015) 45–79, https://doi.org/10.1111/nyas.12938.

[15] A. Lupas, M. Van Dyke, J. Stock, Predicting coiled coils from protein sequences, Science 252 (1991) 1162–1164, https://doi.org/10.1126/science.252.5009.1162.

[16] G. Ma, M. Wei, L. He, C. Liu, B. Wu, S.L. Zhang, J. Jing, X. Liang, A. Senes, P. Tan, S. Li, A. Sun, Y. Bi, L. Zhong, H. Si, Y. Shen, M. Li, M.-S. Lee, W. Zhou, J. Wang, Y. Wang, Y. Zhou, Inside-out Ca2+ signalling prompted by STIM1 conformational switch, Nat. Commun. 6 (2015) 7826, https://doi.org/10.1038/ncomms8826.

[17] H. Okuma, F. Saito, J. Mitsui, Y. Hara, Y. Hatanaka, M. Ikeda, T. Shimizu, K. Matsumura, J. Shimizu, S. Tsuji, M. Sonoo, Tubular aggregate myopathy caused by a novel mutation in the cytoplasmic domain of STIM1, Neurol. Genet. 2 (2016) e50, https://doi.org/10.1212/NXG.0000000000000050.

[18] H.L. Ong, K.P. Subedi, G.-Y. Son, X. Liu, I.S. Ambudkar, Tuning store-operated calcium entry to modulate Ca2+-dependent physiological processes, Biochim. Biophys. Acta BBA-Mol. Cell Res. 1866 (2019) 1037–1045, https://doi.org/10.1016/j.bbamcr.2018.11.018.

[19] M. Pertea, G.M. Pertea, C.M. Antonescu, T.-C. Chang, J.T. Mendell, S.L. Salzberg, StringTie enables improved reconstruction of a transcriptome from RNA-seq reads, Nat. Biotechnol. 33 (2015) 290–295, https://doi.org/10.1038/nbt.3122.

[20] C. Picard, C.-A. McCarl, A. Papolos, S. Khalil, K. Lüthy, C. Hivroz, F. LeDeist, F. Rieux-Laucat, G. Rechavi, A. Rao, A. Fischer, S. Feske, *STIM1* mutation associated with a syndrome of immunodeficiency and autoimmunity, N. Engl. J. Med. 360 (2009) 1971–1980, https://doi.org/10.1056/NEJMoa0900082.

[21] P. Rathner, M. Fahrner, L. Cerofolini, H. Grabmayr, F. Horvath, H. Kroath, M. Gupta, E. Ravera, M. Fragai, M. Bechmann, T. Renger, C. Luchinat, C. Romanin, N. Müller, Interhelical interactions within the STIM1 CC1 domain modulate CRAC channel activation, Nat. Chem. Biol. 17 (2021) 196–204, https://doi.org/10.1038/s41589-020-00672-8.

[22] S. Richards, N. Aziz, S. Bale, D. Bick, S. Das, J. Gastier-Foster, W.W. Grody, M. Hegde, E. Lyon, E. Spector, K. Voelkerding, H.L. Rehm, Standards and guidelines for the interpretation of sequence variants: a joint consensus recommendation of the American College of Medical Genetics and Genomics and the Association for Molecular Pathology, Genet. Med. 17 (2015) 405–424, https://doi.org/10.1038/gim.2015.30.

[23] A. Salvi, C. Skrypnyk, N. Da Silva, J.A. Urtizberea, M. Bakhiet, C. Robert, N. Lévy, A. Megarbané, V. Delague, M. Bartoli, A novel bi-allelic loss-of-function mutation in *STIM1* expands the phenotype of STIM1-related diseases, Clin. Genet. 100 (2021) 84–89, https://doi.org/10.1111/cge.13959.

[24] R. Silva-Rojas, J. Laporte, J. Böhm, STIM1/ORAI1 loss-of-function and gain-of-function mutations inversely impact on SOCE and calcium homeostasis and cause multi-systemic Mirror diseases, Front. Physiol. 11 (2020) 604941, https://doi.org/10.3389/fphys.2020.604941.

[25] P.D. Stenson, M. Mort, E.V. Ball, M. Chapman, K. Evans, L. Azevedo, M. Hayden, S. Heywood, D.S. Millar, A.D. Phillips, D.N. Cooper, The human gene mutation database (HGMD®): optimizing its use in a clinical diagnostic or research setting,
Hum. Genet. 139 (2020) 1197–1207, https://doi.org/10.1007/s00439-020-02199-3.

[26] R.A. Veitia, Exploring the molecular etiology of dominant-negative mutations, Plant Cell 19 (2008) 3843–3851, https://doi.org/10.1105/tpc.107.055053.

[27] R. Zenagui, D. Lacourt, H. Pegeot, K. Yauy, R. Juntas Morales, C. Theze, F. Rivier, C. Cances, G. Sole, D. Renard, U. Walther-Louvier, X. Ferrer-Monasterio, C. Espil, M.-C. Arné-Bes, P. Cintas, E. Uro-Coste, M.-L. Martin Negrier, V. Rigau, E. Bieth, C. Goizet, M. Claustres, M. Koenig, M. Cossée, A reliable targeted next-generation sequencing strategy for diagnosis of myopathies and muscular dystrophies, especially for the Giant titin and Nebulin genes, J. Mol. Diagn. 20 (2018) 533–549, https://doi.org/10.1016/j.jmoldx.2018.04.001.