Defects in TRPM7 channel function deregulate thrombopoiesis through altered cellular Mg2+ homeostasis and cytoskeletal architecture

Stritt, Simon; Nurden, Paquita; Favier, Remi; Favier, Marie; Ferioli, Silvia; Gotru, Sanjeev K.; van Eeuwijk, Judith M M.; Schulze, Harald; Nurden, Alan T.; Lambert, Michele P.; Turro, Ernest; Burger-Stritt, Stephanie; Matsushita, Masayuki; Mittermeier, Lorenz; Ballerini, Paola; Zierler, Susanna; Laffan, Michael A.; Chubanov, Vladimir; Gudermann, Thomas; Nieswandt, Bernhard; Braun, Attila

doi:10.1038/ncomms11097

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Published: 29 March 2016

Defects in TRPM7 channel function deregulate thrombopoiesis through altered cellular Mg²⁺ homeostasis and cytoskeletal architecture

Simon Stritt ORCID: orcid.org/0000-0002-4299-4934^1,2,
Paquita Nurden^1,3,
Remi Favier^4,5,
Marie Favier^5,6,7,8,
Silvia Ferioli⁹,
Sanjeev K. Gotru^1,2,
Judith M M. van Eeuwijk^1,2,
Harald Schulze¹,
Alan T. Nurden³,
Michele P. Lambert^10,11,
Ernest Turro ORCID: orcid.org/0000-0002-1820-6563^12,13,14,15,
Stephanie Burger-Stritt¹⁶,
Masayuki Matsushita¹⁷,
Lorenz Mittermeier⁹,
Paola Ballerini⁴,
Susanna Zierler ORCID: orcid.org/0000-0002-4684-0385⁹,
Michael A. Laffan^18,19,
Vladimir Chubanov⁹,
Thomas Gudermann^9,20,21,
Bernhard Nieswandt^1,2 &
…
Attila Braun^1,2

Nature Communications volume 7, Article number: 11097 (2016) Cite this article

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Abstract

Mg²⁺ plays a vital role in platelet function, but despite implications for life-threatening conditions such as stroke or myocardial infarction, the mechanisms controlling [Mg²⁺]_i in megakaryocytes (MKs) and platelets are largely unknown. Transient receptor potential melastatin-like 7 channel (TRPM7) is a ubiquitous, constitutively active cation channel with a cytosolic α-kinase domain that is critical for embryonic development and cell survival. Here we report that impaired channel function of TRPM7 in MKs causes macrothrombocytopenia in mice (Trpm7^fl/fl-Pf4Cre) and likely in several members of a human pedigree that, in addition, suffer from atrial fibrillation. The defect in platelet biogenesis is mainly caused by cytoskeletal alterations resulting in impaired proplatelet formation by Trpm7^fl/fl-Pf4Cre MKs, which is rescued by Mg²⁺ supplementation or chemical inhibition of non-muscle myosin IIA heavy chain activity. Collectively, our findings reveal that TRPM7 dysfunction may cause macrothrombocytopenia in humans and mice.

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Introduction

Platelets are continuously produced from megakaryocytes (MKs) in the bone marrow by a cytoskeleton-driven process of which the molecular regulation is not fully understood. MKs extend long cytoplasmic protrusions into bone marrow sinusoids, where larger fragments, so-called preplatelets, are shed and further divide within the circulation to give rise to platelets (Supplementary Movie 1)^1,2,3.

Transient receptor potential melastatin-like 7 (TRPM7) channel and kinase domain, but not its kinase activity, are critical for embryonic development^4,5,6 and knockdown or cell-specific TRPM7 knockout approaches give rise to impaired cytoskeletal organization, cell migration, proliferation, polarization and survival. These defects could partially be explained by increased non-muscle myosin IIA heavy chain (NMMIIA)-mediated contractility of the actin cytoskeleton^{5,7,8,9,10,11,12,13}. Of note, among other substrates, the kinase domain of TRPM7 phosphorylates annexin I and NMMIIA, thus interfering with cell survival and cytoskeletal rearrangements^14,15. Interestingly, several variants of NMMIIA similarly altered the contractility of the actomyosin complex in MKs, thereby interfering with proplatelet formation in humans and mice¹⁶. During megakaryopoiesis, NMMIIA activity is suppressed by phosphorylation of its C-terminus, enabling MK polyploidisation and ultimately proplatelet formation¹⁷. However, for proper platelet fission and sizing, NMMIIA needs to be re-activated under shear stress in the circulation^16,18. Although both kinase and channel activity of TRPM7 have been proposed to regulate cytoskeletal dynamics, channel activity alone was sufficient to restore cell polarization, morphology and migration^10,13,19, suggesting a critical role of cations therein. Consequently, the differential role of TRPM7 channel versus kinase activity in the regulation of the cytoskeleton still remains unclear. Moreover, TRPM7 has been implicated as a key regulator of signal conductance in the murine heart by regulating the expression of different pacemaker channels, such as HCN4 (ref. 20). Although TRPM7-mediated cation influx has been detected in MKs (ref. 21), its role in thrombopoiesis has not been investigated to date.

Here we report that impaired channel function but not kinase activity of TRPM7 in MKs causes macrothrombocytopenia in Trpm7^fl/fl-Pf4Cre mice and likely in several members of a human pedigree, which, in addition, feature atrial fibrillation. The impaired proplatelet formation is associated with cytoskeletal alterations due to increased actomyosin contractility and can be rescued by either Mg²⁺ supplementation or chemical inhibition of NMMIIA activity. Collectively, our findings reveal TRPM7 dysfunction as a novel cause of macrothrombocytopenia in mice and potentially in humans too.

Results

Defects in TRPM7 cause macrothrombocytopenia

We identified TRPM7 as the key Mg²⁺ channel and magnesium transporter 1 (MagT1) to be expressed in murine platelets (Supplementary Fig. 1a) and generated MK- and platelet-specific TRPM7 knockout mice (Supplementary Fig. 1b,c). The absence of TRPM7 currents in patch clamp measurements confirmed the efficacy of the targeting strategy in primary bone marrow MKs (Supplementary Fig. 1d). Unexpectedly, these mice displayed a macrothrombocytopenia (Fig. 1a,b) with enlarged and spherical platelets, often containing large vacuoles as revealed by electron microscopy (Fig. 1c). In contrast, mice carrying a kinase-dead K1646R mutation in Trpm7 (ref. 6; Trpm7^KI) showed normal platelet counts, size and morphology, thus suggesting that the lack of TRPM7 channel function rather than its kinase activity accounts for the macrothrombocytopenia in the mutant mice (Supplementary Fig. 2a–e). In line with this notion, intracellular Mg²⁺ concentrations in Trpm7^fl/fl-Pf4Cre platelets, but not in Trpm7^KI platelets, were decreased (Fig. 1d; Supplementary Fig. 2f)⁶.

**Figure 1: TRPM7 deficiency causes mild macrothrombocytopenia in mice.**

Impaired proplatelet formation causes thrombocytopenia

A mildly reduced platelet lifespan in Trpm7^fl/fl-Pf4Cre mice (t½=43.6 h for wild type (WT) versus t½=35.7 h for Trpm7^fl/fl-Pf4Cre mice), however, was insufficient to explain the reduced platelet count and was not associated with altered platelet terminal galactose levels (Supplementary Fig. 3a,b). Immunostaining of whole-femora bone marrow sections (Fig. 2a) revealed an increased number of MKs in the mutant mice (6.3±0.3 for WT versus 13.3±1.6 for Trpm7^fl/fl-Pf4Cre mice; Fig. 2b). The MKs in Trpm7^fl/fl-Pf4Cre mice were also located further from bone marrow sinusoids than in controls (Fig. 2c) suggesting impaired migration of MK-precursors to bone marrow sinusoids. Although splenomegaly was not observed in Trpm7^fl/fl-Pf4Cre mice, we found an increased number of MKs in an expanded red pulp in the spleen and in line with the increased MK numbers, plasma thrombopoietin levels were decreased, thus further indicating deregulated megakaryopoiesis (Supplementary Fig. 4a–d).

**Figure 2: Cytoskeletal alterations account for the macrothrombocytopenia.**

In contrast to a previous report on a neuroblastoma cell line²², the formation of podosomes, F-actin rich structures that are thought to serve cell migration and proplatelet protrusion²³, was unaltered in Trpm7^fl/fl-Pf4Cre MKs (Supplementary Fig. 5a) suggesting that other defects must account for their more distant localization from bone marrow sinusoids (Fig. 2c). Interestingly, mutant MKs displayed an increased mean ploidy as compared with controls (16.9 N±1.6 N in controls versus 22.4 N±3.4 N for Trpm7^fl/fl-Pf4Cre mice; Fig. 2d) thus excluding impaired MK maturation as the cause of thrombocytopenia. Despite the increased ploidy in vivo, we found a decreased proplatelet formation for both foetal liver- (Fig. 2e) and bone marrow-derived (Supplementary Fig. 5b) Trpm7^fl/fl-Pf4Cre MKs in vitro. This was further confirmed in vivo by intravital two-photon microscopy of the bone marrow (0.88% min⁻¹±0.16% min⁻¹ of WT MKs formed proplatelets versus 0.23% min⁻¹±0.12% min⁻¹ of Trpm7^fl/fl-Pf4Cre MKs), which revealed that Trpm7^fl/fl-Pf4Cre MKs preferentially formed short and bulky proplatelet protrusions (0.85% min⁻¹±0.13% min⁻¹ of observed MKs in Trpm7^fl/fl-Pf4Cre mice versus 0.05% min⁻¹±0.04% min⁻¹ in WT mice) that remained attached to the cell body during the observation period (Fig. 2f,g; Supplementary Movies 1, 2, 3).

The actin and microtubule cytoskeleton are critical for proper proplatelet formation². We therefore analysed the cytoskeletal architecture of Trpm7^fl/fl-Pf4Cre MKs and found an increased content and aberrant organization of microtubules in proplatelet-forming, resting and spread MKs (Fig. 2h; Supplementary Fig. 6a–c). In support of this, extraction of the microtubule cytoskeleton of resting bone marrow-derived MKs by ultracentrifugation revealed an increased microtubule content in the Triton X100 insoluble pellet fraction (Supplementary Fig. 6d,e). Of note, increased microtubule stability is associated with post-translational modifications of α-tubulin that include detyrosination (Glu-tub) and acetylation (ac-tub) which could account for the increased number of microtubules in Trpm7^fl/fl-Pf4Cre MKs (ref. 24). Nonetheless, we found an increased prevalence of highly dynamic tyrosinated (Tyr) microtubules as evidenced by lower Glu-/Tyr- (2.5±0.3 for control versus 1.5±0.2 for Trpm7^fl/fl-Pf4Cre MKs) and ac-/Tyr-tubulin ratios (2.2±0.2 for control versus 1.6±0.2 for samples of Trpm7^fl/fl-Pf4Cre mice), suggesting that accelerated microtubule assembly/dynamics accounted for the observed alterations (Supplementary Fig. 6d–g). Moreover, in line with the critical role of microtubules in trafficking of intracellular cargo, electron microscopy revealed a non-homogeneous distribution of granules, tortuous membrane complexes and aberrantly sized proplatelets in mutant MKs (Fig. 2i; Supplementary Fig. 7). Furthermore, we found thick and densely packed proplatelets in bone marrow sinusoids with signs of apoptosis reflecting impaired proplatelet fragmentation and release of preplatelets from Trpm7^fl/fl-Pf4Cre MKs (Fig. 2f,j; Supplementary Movies 2 and 3). In line with the impaired proplatelet formation (Fig. 2e,g; Supplementary Fig. 5b and Supplementary Movies 2 and 3), the recovery of the platelet count after antibody-induced platelet depletion was slightly delayed in Trpm7^fl/fl-Pf4Cre mice (Supplementary Fig. 8). The pronounced increase above initial platelet counts might be attributed to the increased number of MKs in Trpm7^fl/fl-Pf4Cre mice and alternative platelet release mechanisms as recently shown by Nishimura et al.²⁵.

Altered NMMIIA activity impairs proplatelet formation

NMMIIA has been described as a downstream effector of TRPM7 kinase^14,22 and importantly, abnormal function of NMMIIA has been associated with impaired formation and fragmentation of proplatelets in humans and mice¹⁶. Moreover, Mg²⁺ was recently shown to modify NMMIIA activity by regulating ADP release and its affinity to actin filaments, thus further linking TPRM7 channel to NMMIIA function²⁶. Strikingly, analysis of NMMIIA localization in MKs on whole-bone-marrow sections in situ revealed a homogeneous distribution in the cell body of control (92.1±2.0% in WT versus 14.7±1.1% in Trpm7^fl/fl-Pf4Cre mice), while it predominated in the cell cortex of Trpm7^fl/fl-Pf4Cre MKs (0.4±0.8% in WT versus 48.1±3.6% in Trpm7^fl/fl-Pf4Cre mice; Fig. 3a,b). A similar distribution pattern was found in vitro for foetal liver-derived MKs where in addition a significant accumulation of NMMIIA in proplatelets of mature WT MKs was observed (Fig. 3c). Moreover, NMMIIA localization was also altered in Trpm7^fl/fl-Pf4Cre platelets. However, in contrast to MKs, NMMIIA predominated in the cell cortex of WT platelets reminiscent of the marginal band, whereas it was homogeneously distributed in platelets of Trpm7^fl/fl-Pf4Cre mice (Supplementary Fig. 9a–c). This observation is in line with a report on resting foreskin fibroblasts in which NMMIIA and α-tubulin staining overlapped extensively at the cell cortex under resting, non-contractile conditions. However, on induction of cell migration and activation of NMMIIA, the overlap of NMMIIA and α-tubulin staining decreased, allowing NMMIIA to exert its contractile effects on the actin cytoskeleton²⁷. To further analyse the cause of the absent NMMIIA staining and the altered localization to the cell cortex in Trpm7^fl/fl-Pf4Cre MKs in situ, we next allowed bone marrow-derived MKs to spread on a collagen type I-coated surface. Surprisingly, on spreading of Trpm7^fl/fl-Pf4Cre MKs (Fig. 3d,e) or platelets (Supplementary Fig. 10a–d) we found a rapid degradation of NMMIIA that could be rescued by pretreatment with the NMMIIA inhibitor blebbistatin or by Mg²⁺ supplementation (Fig. 3e–g; Supplementary Fig. 10e–h). Based on these findings we speculated that deregulated [Mg²⁺]_i in Trpm7^fl/fl-Pf4Cre cells may cause an increased activity of NMMIIA resulting in its rapid degradation on cell stimulation. In agreement with the increased co-localization of NMMIIA to actin filaments in resting Trpm7^fl/fl-Pf4Cre platelets (Supplementary Fig. 9), sedimentation of cross-linked actin filaments of resting platelets revealed an increased amount of NMMIIA in the pellet fraction (Supplementary Fig. 11a–c). On activation, NMMIIA efficiently cross-linked actin filaments and accumulated in the pellet fraction of WT platelets (Supplementary Fig. 11d–f). In sharp contrast, we found a marked reduction in NMMIIA on stimulation of Trpm7^fl/fl-Pf4Cre platelets (Supplementary Fig. 11d–f), similar to the findings in spread MKs (Fig. 3c) or platelets (Supplementary Fig. 10a–d). To exclude that the observed effects were due to the short observation period, bone marrow-derived MKs were cultured for 48 h in the presence of collagen types I or IV, two major components of the extracellular matrix in the bone marrow. Strikingly, we found a reduced content of NMMIIA in Trpm7^fl/fl-Pf4Cre MKs (reduction of 39.9±10.3% for collagen I and 76.9±14.5% for collagen IV) cultured in the presence of different collagens as compared with control cells (Fig. 3h; Supplementary Fig. 12a,b). Together these results suggested an increased NMMIIA activity under resting conditions and that NMMIIA undergoes a rapid degradation on stimulation of platelets and MKs from Trpm7^fl/fl-Pf4Cre mice thus allowing proplatelet formation and cell spreading.

**Figure 3: Altered localization and accelerated degradation of NMMIIA in MKs.**

Deregulated [Mg²⁺]_i perturbs NMMIIA activity

We hypothesized that deregulated Mg²⁺ homeostasis²⁶ and increased NMMIIA activity may account for the impaired proplatelet formation in Trpm7^fl/fl-Pf4Cre MKs (refs 16, 26). In support of this, either inhibition of NMMIIA activity or Mg²⁺ supplementation could almost fully restore proplatelet formation of Trpm7^fl/fl-Pf4Cre MKs in vitro (Fig. 4a,b). Of note, while pretreatment of foetal liver- (Fig. 4c) or bone marrow-derived MKs (Supplementary Fig. 13) with the Ca²⁺ chelators 1,2-bis(2-aminophenoxy) ethane-N,N,N′,N′-tetraacetic acid tetrakis(acetoxymethyl ester) (BAPTA-AM) or ethylene glycol tetraacetic acid (EGTA) did not exert gross effects; non-specific chelation of Ca²⁺ and Mg²⁺ with ethylenediaminetetraacetic acid (EDTA) significantly reduced proplatelet formation by MKs, suggesting that Mg²⁺ is critical for this process^28,29. According to a previous report⁸, Mg²⁺ supplementation should restore [Mg²⁺]_i in mutant MKs, which cannot be fully achieved through an upregulation of MagT1 expression under normal culture conditions (Fig. 4d,e).

**Figure 4: Altered NMMIIA regulation accounts for the impaired proplatelet formation.**

Inhibition of NMMIIA restores cytoskeletal architecture

Besides increasing cortical tension through relaxation of the underlying cytoskeleton^17,30, blebbistatin treatment reduced the prevalence and stability of microtubules in proplatelet protrusions, as evidenced by decreased Glu-/Tyr- (1.1±0.1 for WT and 1.3±0.1 for samples of Trpm7^fl/fl-Pf4Cre mice) and ac-/Tyr-tubulin ratios (1.3±0.1 for WT and 1.2±0.2 for samples of Trpm7^fl/fl-Pf4Cre mice; Supplementary Fig. 14). Immunostaining (Fig. 5a–c) and electron microscopy (Fig. 5d) revealed an increased number of aberrantly organized microtubules in Trpm7^fl/fl-Pf4Cre platelets as compared with controls, similar to findings in mutant MKs. This correlated with an increased presence of highly dynamic Tyr-tubulin (151.4±3.7% of controls; Fig. 5e–g), leading to accelerated and uncontrolled microtubule polymerization in mutant cells (Fig. 5a,h). Stable microtubules in Trpm7^fl/fl-Pf4Cre platelets displayed a similar pattern of post-translational modifications of α-tubulin (104.2±14.2% of control for Glu-, and 97.4±9.4% of control for ac-tubulin) and were efficiently disassembled on cold challenge, thus excluding enhanced stability as cause for the increased microtubule content (Fig. 5e–h). Interestingly, pretreatment of WT platelets with EDTA, but not EGTA, BAPTA-AM or MgCl₂, mimicked these cytoskeletal alterations (Fig. 6a–c), changes that could also be reverted by blebbistatin (Fig. 6d–f), thus further supporting the notion that reduced [Mg²⁺]_i alters the subcellular localization and activity of NMMIIA resulting in cytoskeletal disorganization. In agreement with this, we observed an increased content of filamentous actin in resting Trpm7^fl/fl-Pf4Cre platelets (Supplementary Fig. 15a,b) and a decreased polymerization of filamentous actin on platelet activation (Fig. 6g; Supplementary Fig. 15b,c). Moreover, spread Trpm7^fl/fl-Pf4Cre platelets displayed an increased surface area (Fig. 6h; Supplementary Fig. 16) most likely reflecting the activation-dependent rapid degradation of NMMIIA and consequently the loss of coherent cytoplasmic contractile forces normally generated by activated NMMIIA (refs 31, 32).

**Figure 5: Altered cytoskeletal organization in *Trpm7*^fl/fl-Pf4Cre platelets.**

**Figure 6: Deregulated [Mg²⁺]_i accounts for the altered cytoskeletal organization.**

Variants in TRPM7 may cause macrothrombocytopenia

We next hypothesized that some DNA variants of extreme low-frequency affecting TRPM7 channel function might also cause macrothrombocytopenia in humans. Examining the results of genome sequencing of 702 cases with bleeding and platelet disorders of unknown genetic basis in the BRIDGE database of the NIHR BioResource—Rare Diseases revealed three cases with a coding variant unobserved in nearly 81,000 control DNA samples: UCN 0012 with p.C721G (c.2161T>G), UCN 0025 with p.R902C (c.2704C>T) variant, and UCN 0110 with p.G1353D (c.4058G>A; Supplementary Table 1). Two of these three index cases (p.C721G and p.R902C) showed low-platelet counts and macrothrombocytopenia. Interestingly, the variant of index patient UCN 0110 (p.G1353D (c.4058G>A)) was located close to the α-kinase domain and the absence of macrothrombocytopenia is in agreement with the results on the Trpm7^KI mice (Supplementary Table 1 and Supplementary Fig. 2). Unfortunately, the family with the p.R902C variant was unavailable for further studies.

Further studies were focussed on the index case UCN 0012 with a p.C721G (c.2161T>G) variant and pedigree members. Sanger sequencing showed that the p.C721G variant was present in two further pedigree members with macrothrombocytopenia, but was absent in one asymptomatic pedigree member, indicating segregation with the TRPM7 genotype (Fig. 7a, Supplementary Table 2 and Supplementary Fig. 17). For the fourth patient (pedigree member 2), now deceased, macrothrombocytopenia was detected during her life (Fig. 7a). All other blood cell parameters and platelet function were normal for all p.C721G patients. Strikingly, paroxysmal atrial fibrillation was also present for the index case (pedigree member 5) and her mother (pedigree member 2).

**Figure 7: A human genetic variant in *TRPM7* causes macrothrombocytopenia.**

Similarly to Trpm7^fl/fl-Pf4Cre mouse platelets, we found a reduced content of Mg²⁺ and an increased concentration of Ca²⁺ in platelets from all tested patients with the p.C721G substitution as compared with healthy controls (Fig. 7b). Patch clamp studies on HEK293 cells confirmed that the p.C721G variant reduced TRPM7 channel activity by 85±4% as compared with WT controls (Fig. 7c) despite being localized to the cell membrane (Supplementary Fig. 18). Likewise, in vitro studies on the p.R902C variant revealed, although less pronounced, a reduced TRPM7 channel activity by 39±6% (Supplementary Fig. 19). Platelets from all tested patients displayed an increased size, with a spherical shape and contained numerous vacuoles. Moreover, while platelets from controls displayed the typical discoid shape and microtubules organized into coils, the marginal band, platelets from the p.C721G patients showed an aberrant distribution of granules and an increased number and anarchic organization of microtubules (Fig. 7d; Supplementary Fig. 20). The abnormal cytoskeletal organization could be confirmed by immunostaining on resting p.C721G platelets (Fig. 7e,f; Supplementary Figs 21 and 22) and similarly to the Trpm7^fl/fl-Pf4Cre platelets, which was not associated with increased microtubule stability (Fig. 7g; Supplementary Fig. 21).

Altered regulation of NMMIIA in p.C721G platelets

Since the ultrastructure of human platelets from individuals carrying the p.C721G variant strongly resembled that of Trpm7^fl/fl-Pf4Cre mouse platelets, we next analysed the distribution and stability of NMMIIA. Whereas in control platelets NMMIIA localized to the marginal band, it was homogeneously distributed throughout the cytoplasm of platelets from the patients (Fig. 8a,b; Supplementary Fig. 22). In line with the observations on Trpm7^fl/fl-Pf4Cre mouse platelets, spread platelets from the patients showed an increased surface area, a strong decrease in NMMIIA and an increased content of microtubules (Fig. 8c,d; Supplementary Figs 23 and 24). As for the mouse model, blebbistatin prevented loss of NMMIIA on spreading (Fig. 8d,e; Supplementary Fig. 25) and rescued the cytoskeletal organization of resting platelets after cold challenge (Fig. 8f,g; Supplementary Fig. 26). As mentioned above, patients 2 and 5 suffered from atrial fibrillation which might be associated with alterations in [Mg²⁺]_i (ref. 33). Furthermore, lack of TRPM7 has been associated with altered expression of channels, such as HCN4 encoding the pacemaker current in the conduction system of the heart that has also previously been linked to atrial fibrillation^20,34.

**Figure 8: Altered NMMIIA activity accounts for the aberrant cytoskeletal organization.**

Discussion

In the present study, we provide evidence that defects in TRPM7 channel function cause macrothrombocytopenia in mice and likely in humans too. These results demonstrate a critical role of TRPM7-mediated Mg²⁺ influx in regulating NMMIIA activity and cytoskeletal rearrangements during thrombopoiesis. In support of this, it has been shown that Mg²⁺ inhibits NMMIIA activity by reducing the ADP release rate and its affinity for actin²⁶. Furthermore, using Trpm7^KI mice (Supplementary Fig. 2), we convincingly show that these effects occur independently of TRPM7 α-kinase activity. However, it is too early to exclude that the kinase domain also acts as a molecular hub orchestrating yet unknown signalling events, and thereby contributes to the development of macrothrombocytopenia.

The observed loss of NMMIIA in Trpm7^fl/fl-Pf4Cre bone marrow MKs in situ and in vitro on spreading could represent a physiological process to overcome the inhibitory effects of increased NMMIIA activity enabling proplatelet formation (Fig. 3a,b,d–h; Supplementary Figs 10–12). In support of this hypothesis, NMMIIA degradation was most pronounced on culture of bone marrow-derived MKs on collagen type IV, which predominates in the vascular niche where proplatelet formation occurs (Fig. 3a,b,h; Supplementary Fig. 12). Furthermore, using different antibodies and experimental approaches, we provide several independent lines of evidence that the observed loss of NMMIIA on activation or spreading of TRPM7-deficient cells represents protein degradation. However, it is important to note that we cannot entirely exclude epitope masking due to modifications on NMMIIA (Figs 3 and 8; Supplementary Figs 10–12 and 23–25).

It has recently been reported that plasma Thpo levels are also regulated via Ashwell–Morell receptor-dependent clearance of desialylated platelets, which in turn triggers expression of Thpo in hepatocytes and thus regulates platelet production³⁵. However, it is important to note that in agreement with the moderately reduced platelet lifespan, we did not observe differences in platelet terminal galactose levels suggesting that the decreased plasma Thpo levels are reciprocally associated to the increased number of MKs in the bone marrow and spleen (Fig. 2a,b; Supplementary Figs 3 and 4)³⁶. Furthermore, the marked increase in platelet counts after immuno-depletion might result from an alternative platelet release mechanism as recently described by Nishimura et al.²⁵.

The here-described human disorder (Fig. 7; Supplementary Tables 1 and 2) is clearly distinguishable from MYH9-related platelet disease, which can be associated with hearing loss, cataracts or renal failure³⁷. However, patients with variants in TRPM7 and MYH9 both display macrothrombocytopenia with more spherical platelets and partially, increased actomyosin contractility (Figs 7 and 8; Supplementary Figs 21–26)¹⁶. The different clinical outcomes and symptoms of patients with different genetic variants of TRPM7 are most likely due to the location and functional consequences of the mutations (Supplementary Table 1). While both the p.C721G and the p.R902C variant are located in the channel domain and reduce channel activity (Fig. 7c and Supplementary Fig. 19), the p.G1353D variant is not located within the channel domain which may likely explain the distinct phenotype and absence of macrothrombocytopenia in the index patient UCN 0025. Similar genotype–phenotype-relationships have been observed for patients suffering from MYH9-related disorder^38,39, Wiskott–Aldrich syndrome^2,40 or filaminopathy A⁴¹. Although we could not observe a dominant negative effect of the p.C721G variant on TRPM7 channel function in vitro, we speculate that the defect might be masked by the high-expression levels of TRPM7 in the used system or diminished channel activity may not represent the only mechanistic explanation for the observed phenotypes. Furthermore, Trpm7^+/− mice did not display macrothrombocytopenia suggesting that gene haploinsufficiency does not account for the observed disease (Supplementary Fig. 27). However, the impaired channel activity and the striking phenotypic similarities in platelets from patients and Trpm7^fl/fl-Pf4Cre mice strongly suggest that the variants in TRPM7 account for the observed macrothrombocytopenia.

Our study provides for the first time several independent lines of evidence that proper regulation of Mg²⁺ homeostasis in MKs by TRPM7 plays a critical role in thrombopoiesis and platelet sizing, both in humans and mice. Collectively, our results highlight the clinical need to carefully control platelet count and size in patients with deregulated Mg²⁺ homeostasis. Ultimately, our findings suggest, after careful consideration and assessment of adverse events, that Mg²⁺ supplementation may be used as a potential treatment of patients with increased activity of NMMIIA to manage thrombocytopenia. Despite the possibility of severe potential side effects in patients with impaired kidney function, Mg²⁺ supplementation is considered as a relatively safe therapeutic intervention^42,43,44. Nevertheless, further studies in animal models and patients are required to assess the efficacy and safety of Mg²⁺ supplementation in the management of certain cases of thrombocytopenia.

Finally, the fact that two of the patients in our study also suffered from paroxysmal atrial fibrillation promotes TRPM7 as a novel candidate interfering with conductance in cardiac cells.

Methods

Semi-quantitative reverse transcription PCR

Total platelet RNA was isolated after lysis using TRIzol reagent (15596018, Invitrogen) and tissue RNA was gained using the Qiagen RNeasy kit. To generate cDNA 1 μg RNA was reverse transcribed with the SuperScript II reverse transcriptase (18064014, Invitrogen) according to the manufacturer’s instructions. β-actin transcripts were determined as control. Primers used for the amplification of Mg²⁺ transporters and channels are listed in Supplementary Table 3.

Mice

Conditional Trpm7-deficient mice were generated by intercrossing Trpm7^fl/fl mice (exon 17 flanked by loxP sites) with mice carrying the Cre-recombinase under the platelet factor 4 (Pf4) promoter⁴⁵ (Supplementary Fig. 1b). Trpm7^fl/fl mice were described earlier⁴, Pf4-Cre mice were kindly provided by Dr Radek Skoda and Trpm7^KI mice were generously provided by Dr Masayuki Matsushita⁶. All mice used in experiments were 12- to 16-week-old and sex-matched, if not stated otherwise. For experiments on MKs, only male animals were used. All animal experiments were approved by the district government of Lower Franconia (Bezirksregierung Unterfranken). Trpm7^fl/fl-Pf4Cre mice were genotyped via PCR on genomic DNA extracted from ear biopsies using the following primer pair: Trpm7KO-F: 5′-gaggtactggcaattgtgagc-3′ and Trpm7KO-R: 5′-accacaaaatctctgccctct-3′ yielding a 1,200-bp product for the floxed, 1000, bp product for the WT and a 400-bp fragment for the recombined allele (Supplementary Fig. 1c). For all experiments, the respective Trpm7^fl/fl (for Trpm7^fl/fl-Pf4Cre mice) or Trpm7^WT/WT (for Trpm7^KI/KI mice) littermate controls were used. All mice were derived from the following breeding strategies for Trpm7^fl/fl X Trpm7^fl/fl-Pf4Cre, yielding 50% Trpm7^fl/fl WT and 50% Trpm7^fl/fl-Pf4Cre mice or Trpm7^WT/KI X Trpm7^WT/KI resulting in 25% Trpm7^WT/WT, 50% Trpm7^WT/KI and 25% Trpm7^KI/KI mice, respectively.

Platelet preparation

Mice were bled under isoflurane anaesthesia. Blood was collected in heparin (20 U ml⁻¹, Ratiopharm) and centrifuged twice for 6 min at 300 g. Platelet-rich plasma (PRP) was supplemented with 2 μl ml⁻¹ of apyrase (0.02 U ml⁻¹; A6410, Sigma-Aldrich) and 5 μl ml⁻¹ prostacyclin I2 (PGI₂) (0.1 μg ml⁻¹; P6188, Sigma-Aldrich) and platelets were pelleted by centrifugation for 5 min at 800 g, washed twice with Tyrodes-N-2-hydroxyethyl-piperazine-N′2-ethanesulfonic acid (HEPES) buffer (134 mM NaCl, 0.34 mM Na₂HPO₄, 2.9 mM KCl, 12 mM NaHCO₃, 5 mM HEPES, 5 mM glucose, 0.35% bovine serum albumin (BSA), pH 7.4) containing 2 μl ml⁻¹ apyrase and 5 μl ml⁻¹ PGI₂.

Blood samples of patients were obtained after informed consent in accordance with the Declaration of Helsinki. Ethical approval was obtained from Inserm RBM 04–14 for the project ‘Network on the inherited diseases of platelet function and platelet production’ and by the Ethics Committee of the University Hospital Würzburg. Fresh blood samples of patients and healthy volunteers were collected in 1/10 volume of acid-citrate-dextrose and centrifuged for 10 min at 200 g. PRP was collected, supplemented with 2 μl of apyrase (0.02 U ml⁻¹, Sigma-Aldrich) and 5 μl PGI₂ (0.1 μg ml⁻¹, Sigma-Aldrich) per ml PRP. Before adherence to poly-L-lysine (PLL)-coated slides or spreading on different matrices, platelets were pelleted by centrifugation for 10 min at 800 g and washed twice with Tyrodes-HEPES buffer containing 2 μl ml⁻¹ apyrase and 5 μl ml⁻¹ PGI₂. The samples were allowed to rest for 30 min at 37 °C prior to being used in experiments.

Flow cytometry

Diluted blood (1:20) was incubated for 15 min at room temperature (RT) with fluorophore-conjugated antibodies (2 μg ml⁻¹) directed against platelet surface glycoproteins. Platelet count and size (FSC) were assessed using a FACSCalibur (BD Biosciences) flow cytometer or with an automated blood cell analyser (Sysmex KX-21N).

Transmission electron microscopy of platelets

To analyse platelet ultrastructure, washed platelets were fixed with 2.5% glutaraldehyde (16,210, Electron Microscopy Sciences) in 50 mM cacodylate buffer (12,201, pH 7.2; AppliChem). After embedding in epon 812 (14,900, Electron Microscopy Sciences), ultrathin sections were generated and stained with 2% uranyl acetate (22,400, Electron Microscopy Sciences) and lead citrate (17,800, Electron Microscopy Sciences). Samples were visualized with an EM900 transmission electron microscope (Carl Zeiss).

Immunostaining on resting or spread platelets

Coverslips were either coated with PLL (Sigma-Aldrich), fibrinogen (100 μg ml⁻¹; F4883, Sigma-Aldrich) or collagen-related peptide (CRP) (ref. 46; 6 μg ml⁻¹) overnight at 4 °C. For spreading on fibrinogen, platelets were stimulated with 0.01 U ml⁻¹ thrombin (10602400001, Roche) and allowed to spread for the indicated time. If indicated, platelets were pre-incubated with toxins interfering with cytoskeletal dynamics such as colchicine (10 μM; A4082, AppliChem) before the spreading assay. After the indicated time points, platelets were either fixed with 4% paraformaldehyde (PFA) in PBS and analysed or processed for immunofluorescence staining of the cytoskeleton. Therefore, platelets were fixed and permeabilized in PHEM buffer (60 mM piperazine-N,N-bis-2- ethanesulfonic acid, 25 mM HEPES), 10 mM EGTA, 2 mM MgCl₂, pH 6.9) supplemented with 4% PFA and 1% IGEPAL CA-630, stained with anti-α-tubulin-Alexa F488 (3.33 μg ml⁻¹, 322588 (B-5-1-2), Invitrogen), anti-NMMIIA (10 μg ml⁻¹, #3403, polyclonal, CellSignaling) and phalloidin-Atto647N (170 nM, 65906, Fluka) and mounted with Fluoroshield (F6182, Sigma-Aldrich). All used fluorophore-conjugated secondary antibodies were purchased from Invitrogen. Samples were visualized using a Leica TCS SP5 confocal microscope (Leica Microsystems).

Inductively coupled plasma mass spectrometry (ICP-MS)

The cation content of 4 × 10⁷ platelets was analysed by ICP-MS on platelets from PRP. ICP-MS analysis was performed by ALS Scandinavia AB (Lulea, Sweden).

Determination of platelet lifespan

The clearance of platelets from the circulation was determined by the retro-orbital injection of 5 μg DyLight 488-labelled anti-GPIX antibody^2,47 in PBS into male mice. The percentage of labelled platelets was determined by daily blood withdrawal and subsequent analysis by flow cytometry using fluorophore-conjugated platelet-specific antibodies².

Immunofluorescence staining on whole-femora cryosections

Femora of male mice were isolated, fixed with 4% paraformaldehyde (A3813, AppliChem) and 5 mM sucrose (S0389, Sigma-Aldrich), transferred into 10% sucrose in PBS and dehydrated using a graded sucrose series. Subsequently the samples were embedded in Cryo-Gel (39475237, Leica Biosystems) and shock frozen in liquid nitrogen. Frozen samples were stored at −80 °C. Cryosections (7-μm-thick) were generated using the CryoJane tape transfer system (Leica Biosystems) and probed with Alexa488-conjugated anti-glycoprotein (GP) Ib (ref. 48) antibodies (7A9, 1.33 μg ml⁻¹), to specifically label platelets and MKs, and Alexa647-conjugated anti-CD105 antibodies (3.33 μg ml⁻¹, 120402 (MJ7/18), Biolegend) to stain the endothelium. Nuclei were stained using 4′,6-diamidino-2-phenylindole (1 μg ml⁻¹, D1306, Invitrogen). Samples were visualized with a Leica TCS SP5 confocal microscope (Leica Microsystems).

Histology

Sections (3-μm-thick) of formalin-fixed paraffin embedded spleens and femora of male mice were prepared deparaffinized and stained with haematoxylin (MHS32, Sigma-Aldrich) and eosin (318906, Sigma-Aldrich). MK number, morphology and localization were analysed with an inverted Leica DMI 4000 B microscope.

In vitro differentiation and cultivation of bone marrow MKs

Haematopoietic stem cells were isolated from male mouse bone marrow single-cell suspensions using a magnetic bead-based negative depletion kit (anti-rat-IgG Dynabeads, Invitrogen) in combination with rat-anti-mouse antibodies directed against CD45R/B220, TER-119, CD3, Ly-6G/C and CD11b (each 0.5 μg per 10⁷ cells, 103216 (RA3-6B2), 116214 (TER-119), 100208 (17A2), 108414 (RB6-8C5) and 101214 (M1/70), Biolegend). Experiments were performed according to the manufacturer’s protocol. Cells were cultured in MK-medium supplemented with 50 μg ml⁻¹ recombinant hirudin (Schering) at 37 °C, 5% CO₂ for 3 days, before MK enrichment using a two step BSA density gradient. On day 4, the percentage of proplatelet-forming MKs was determined using a light microscope (Zeiss).

MK spreading

In vitro cultivated bone marrow MKs of male mice were allowed to adhere and spread at 37 °C and 5% CO₂ on coverslips coated with fibrillar collagen type I (50 μg ml⁻¹; Nycomed), fibrinogen (100 μg ml⁻¹; F4883, Sigma-Aldrich) or CRP (6 μg ml⁻¹) for the indicated time. MK spreading was stopped by fixation and permeabilization of the cells using PHEM buffer supplemented with 4% PFA and 1% IGEPAL CA-630.

Immunofluorescence staining of cultured or spread MKs

To visualize the cytoskeleton, the cultured MKs were spun onto glass slides (Shandon Cytospin 4, Thermo Scientific), fixed and permeabilized in PHEM buffer supplemented with 4% PFA and 1% IGEPAL CA-630 and blocked with 1% BSA in PBS. F-actin was stained using phalloidin-Atto647N (170 nM, 65906, Fluka) and tubulin was stained with an anti-α-tubulin-Alexa F488 (3.33 μg ml⁻¹, 322588 (B-5-1-2), Invitrogen) antibody. WASp (10 μg μl⁻¹, #4860, CellSignaling) or NMMIIA (10 μg ml⁻¹, #3403, polyclonal, CellSignaling) staining served as podosome marker. Nuclei were stained using 4′,6-diamidino-2-phenylindole (1 μg μl⁻¹, D1306, Invitrogen) before mounting of samples with Fluoroshield (F6182, Sigma-Aldrich). Visualization was performed with a Leica TCS SP5 confocal microscope (Leica Microsystems).

Determination of MK ploidy

To determine bone marrow MK ploidy, both femora of male mice were isolated and the bone marrow was flushed and homogenized. Non-specific binding sites of the 5D7 antibody were blocked by incubation of the cell suspension with 0.02 μg μl⁻¹ anti-FcγR antibody (553142 (2.4G2), BD Pharmingen). Afterwards, MKs were stained using a fluorescein isothiocyanate-conjugated anti-CD41 antibody (10 μg ml⁻¹, 5D7). Finally, cells were fixed, permeabilized and DNA was stained using 50 μg ml⁻¹ propidium iodide (P3561, Invitrogen) staining solution with 100 μg ml⁻¹ RNaseA (EN0202, Fermentas) in PBS. Analysis was performed by flow cytometry and FlowJo software (Tree Star Inc.).

Plasma thrombopoietin levels

Plasma Thpo levels were determined using the Mouse Thrombopoietin Quantikine ELISA Kit (DY488, R&D Systems). Briefly, plasma was collected, diluted (1:5 in Reagent Diluent) and immediately applied as duplicates onto the anti-Thpo-IgG coated 96-well plate and incubated for 2 h at room temperature. The plate was washed five times and incubated for 2 h with 100 μl of horseradish peroxidase-conjugated anti-mouse Thpo antibodies. After another five washing steps, tetramethylbenzidine solution was added and incubated for 30 min at room temperature. The reaction was aborted by the addition of 100 μl of diluted hydrochloric acid. Optical densities of the samples were determined using a Multiskan Ascent (96/384) plate reader (MTX Lab Systems) at 450 nm. Wavelength correction was performed at 570 nm.

In vitro differentiation of foetal liver-derived MKs

The livers of 13.5–14.5-day-old female and male mouse foetuses were isolated from time-mated female mice and single-cell suspensions were prepared in MK-medium (IMDM medium containing 10% FCS, 1% penicillin/streptomycin (31980097, Gibco) and 50 ng ml⁻¹ recombinant Thpo (ref. 49). The homogenized foetal liver cells were cultured for 72 h at 37 °C and 5% CO₂ and mature MKs were enriched on day three of culturing using a BSA density gradient (3% and 1.5% BSA in PBS; A7030, Sigma-Aldrich). If indicated the cells were treated with 25 μM blebbistatin (B0560, Sigma-Aldrich) or maintained in culture medium supplemented with 10 mM MgCl₂. On day 4, the percentage of proplatelet-forming MKs was determined by counting the total number of MKs as well as the number of proplatelet-forming MKs under a light microscope (Zeiss).

Two-photon intravital microscopy of the bone marrow

Male and female mice were anaesthetized and a 1-cm incision was made along the midline to expose the frontoparietal skull while carefully avoiding damage to the bone tissue. The mouse was placed on a customized metal stage equipped with a stereotactic holder to immobilize its head^2,3. Bone marrow vasculature was visualized by injection of tetramethylrhodamine dextran (8 μg per g body weight, 2 MDa, Molecular Probes). Platelets and MKs were antibody-stained (0.6 μg per g body weight anti-GPIX-Alexa Fluor 488). Images were acquired with a fluorescence microscope equipped with a × 20 water objective with a numerical aperture of 0.95 and a TriM Scope II multiphoton system (LaVision BioTec), controlled by ImSpector Pro-V380 software (LaVision BioTec). Emission was detected with HQ535/50-nm and ET605/70-nm filters. A tuneable broad-band Ti:Sa laser (Chameleon, Coherent) was used at 760 nm to capture Alexa Fluor 488 and rhodamine dextran fluorescence. ImageJ software (NIH) was used to generate movies.

Transmission electron microscopy of bone marrow MKs

For transmission electron microscopy of MKs, bone marrow was flushed from 12- to 16-week-old male mice using Karnovsky fixative (2% PFA, 2.5% glutaraldehyde in 0.1 M cacodylate buffer) and incubated overnight at 4 °C. Subsequently, fatty components of the samples were fixed with 2% osmium tetroxide in 50 mM sodium cacodylate (pH 7.2), stained with 0.5% aqueous uranyl acetate, dehydrated with a graded ethanol series and embedded in Epon 812. Ultrathin sections were stained with 2% uranyl acetate (in 100% ethanol) followed by lead citrate. Images were taken on a EM900 transmission electron microscope (Zeiss).

Platelet depletion

Circulating platelets were depleted in mice by injection of 20 μg per 30 g body weight anti-GPIbα-antibodies (Emfret, Eibelstadt, Germany) and platelet counts were monitored by flow cytometry for 10 days.

Immunoblot

Denatured resting or convulxin-stimulated (0.5 μg ml⁻¹) platelets or untreated and MgCl₂ supplemented MKs were lysed and separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS–PAGE) and blotted onto polyvinylidene difluoride membranes. NMMIIA (1 μg ml⁻¹, #3403, polyclonal, CellSignaling; 1 μg ml⁻¹, MP3791; ECM Biosciences), Gapdh (1 μg ml⁻¹, G5262, Sigma-Aldrich) or MagT1 (10 μg ml⁻¹, AP5056a, Abgent) were probed with the respective antibodies and detected using horseradish peroxidase-conjugated secondary antibodies (0.33 μg ml⁻¹) and enhanced chemiluminescence solution (JM-K820-500, MoBiTec). Images were recorded using a MultiImage II FC Light Cabinet (Alpha Innotech cooperation) device. Uncropped immunoblotting images are shown in Supplementary Figs 28 and 29.

Extraction and sedimentation of the cytoskeleton

Resting or activated (0.1 U ml⁻¹ thrombin (10602400001, Roche) platelets (5.7 × 10⁷) or MKs (5 × 10⁶) were lysed in PHEM buffer containing 1% Triton X100, 6 μM taxol (A4667, AppliChem) for microtubule sedimentation or 2 μM phalloidin (A1488, AppliChem) for F/G-actin sedimentation and protease inhibitors (P8340, Sigma-Aldrich). Polymerized and soluble fractions were separated by centrifugation for 30 min at 100.000 g (microtubules) or 16.000 g (F/G-actin), respectively, and 37 °C in a TLA-100 rotor (Beckman Coulter). Total platelet lysates, soluble supernatants (S) and insoluble pellets (P) were supplemented with SDS–PAGE buffer containing 5% β-mercaptoethanol (M6250, Sigma-Aldrich). Samples were separated by SDS–PAGE, blotted onto polyvinylidene difluoride membranes and probed with anti-Tyr-tubulin (0.5 μg ml⁻¹, MAB1864 (YL1/2), Milipore), anti-acetylated tubulin (1 μg ml⁻¹, sc-23950 (6-11B-1), Santa Cruz Biotechnology Inc.), anti-detyrosinated-tubulin (1 μg ml⁻¹, AB3201, Milipore), NMMIIA (1 μg ml⁻¹, #3403, polyclonal, CellSignaling; 1 μg ml⁻¹, MP3791; ECM Biosciences) or β-actin (1 μg ml⁻¹, SAB5500001 (SP124), Milipore) antibodies.

Cold-induced microtubule disassembly

Microtubules were depolymerized by incubation of platelets at 4 °C; reassembly was allowed by subsequent rewarming at 37 °C. Samples maintained at 4 °C and 37 °C as well as rewarmed platelets were fixed and permeabilized in PHEM buffer supplemented with 4% PFA and 1% IGEPAL CA-630 and allowed to adhere to a PLL-coated coverslips. Samples were immunostained using an anti-β₁-tubulin (2.5 μg ml⁻¹, T4026, clone TUB 2.1, Sigma-Aldrich) or anti-α-tubulin Alexa F488 antibody (3.33 μg ml⁻¹, 322588 (B-5-1-2), Invitrogen) and visualized using a Leica TCS SP5 confocal microscope (Leica Microsystems).

Actin polymerization

Washed platelets were incubated with a DyLight-649–labelled anti-GPIX antibody derivative (20 μg ml⁻¹⁾. Subsequently, platelets were either left unstimulated or were stimulated with the indicated agonists for 2 min. Platelets were fixed with 0.55 volume of 10% paraformaldehyde in PHEM buffer and treated with 0.1 volume 1% Triton X100. Subsequently, platelets were stained with 10 μM phalloidin-fluorescein isothiocyanate (P5282, Sigma-Aldrich) for 30 min and analysed on a FACSCalibur.

Recruitment of patients

The index case and pedigree members were enrolled to the French ‘Network on the inherited diseases of platelet function and platelet production’ (INSERM RBM 01–14). Further subjects with bleeding and platelet disorders (BPD) and with unrelated rare disorders (control data) were enrolled to the BRIDGE-BPD project of the NIHR BioResource—Rare Diseases study (UK REC 13/EE/0325). Here, gene variants were retained in an analysis set if they occur at a frequency of <1:10,000 in the exome aggregation consortium database based on over 60,000 individuals (http://exac.broadinstitute.org/). All cases provided informed written consent at enrolment.

The BRIDGE bleeding and platelet disorders collection

We searched for cases with coding variants of extreme low frequency in the TPRM7 gene by analysing data from 702 index cases with BPDs of unknown genetic basis recruited to the BRIDGE study of the NIHR BioResource—Rare Diseases. Gene variants were identified in genome sequencing data by comparison with reference human genome build GRCh37 and consequences were predicted using the Ensembl version 75 TPRM7 transcript ENST00000313478. Variants predicted to alter the amino acid sequence of the protein were retained in the analysis set if they occurred at frequencies of <1 in 10,000 out of 67,000 individuals of the Exome Aggregation Consortium (ExAC) database, <1 in 1,000 in the 10,000 subjects of UK10K database and <1 in 100 in 3,453 subjects with unrelated rare disorders or unaffected pedigree members recruited to other branches of the NIHR BioResource. We selected cases with extreme low-frequency variants and on the basis of clinical and laboratory characteristics, which were coded using Human Phenotype Ontology (HPO) terms⁵⁰ retrieved from the BRIDGE-BPD study database. We then focused our further exploration on the cases with extremely rare variants that were unobserved in the nearly 81,000 control DNA samples. The first step was to contact the families to explain the necessity to control these results by performing co-segregation studies. One family met these requirements and additional family members were examined in Paris at a French Centre for inherited platelet diseases.

Co-segregation studies

Genomic DNA from peripheral blood mononuclear cells was extracted using a commercial DNA purification kit (Qiagen) and exon 17 of TRPM7 was amplified by polymerase chain reaction using the respective primer pairs: TRPM7_Ex17_forw: 5′-ggagaatgtgctctggattc-3′ and TRPM7_Ex17_rev: 5′-gccaatcatccatcttgctc-3′, expected product size 606 bp. PCR products were purified with the help of QIAquick PCR purification kit (Qiagen) and processed for Sanger sequencing.

Platelet function testing

Platelet function testing of human blood was performed by light transmission turbidometric aggregometry. To this end, PRP (2.5 × 10⁵ platelet per μl) was isolated and stimulated with different concentrations of agonists (ADP, 5 and 10 mM; collagen I, 1 and 5 mg ml⁻¹; epinephrine, 5 mM; arachidonic acid, 1 mM; ristocetin, 0.6 and 1.5 mg ml⁻¹; and TRAP (thrombin receptor activating peptide), 20 mM). Light transmission was followed over time with PPP set as 100% light transmission.

Transient expression of WT and p.C721G TRPM7 variants

The TRPM7 p.C721G and p.R902C variants were generated by site-directed mutagenesis on the pcDNA3.1-TRPM7 expression construct⁵¹ using the QuickChange II XL site-directed mutagenesis kit (200517, Agilent Technologies) with the following primer pairs: Trpm7_C721G_for: 5′-ctggagtaattcaaccggcctcaagttagcgtttc-3′ and Trpm7_C721G_rev: 5′-gaaactgctaacttgaggccggttgaattactccag-3′. The mutation was confirmed by sequencing. For transient expression of WT TRPM7 and the p.C721G variant, human embryonic kidney (HEK) 293 cells were maintained at 37 °C and 5% CO₂ in Earle’s minimal essential medium supplemented with 10% foetal bovine serum, 100 μg ml⁻¹ streptomycin and 100 U ml⁻¹ penicillin (Invitrogen). Cells were transiently co-transfected with eukaryotic expression vectors encoding WT TRPM7 or the p.C721G and p.R902C variant with 100 ng of an enhanced green fluorescent protein (EGFP) reporter construct (pcDNA3.1) using Lipofectamine2000 (Invitrogen) according to the manufacturer’s instructions and processed for patch clamp measurements or confocal microscopy.

Electrophysiology

Patch clamp experiments were performed at a whole-cell configuration. Currents were elicited by a ramp protocol from −100 to +100 mV over 50 ms acquired at 0.5 Hz and a holding potential of 0 mV. Inward currents were extracted at −80 mV, outward currents at +80 mV and plotted versus time. Data were normalized to cell size as pA pF⁻¹. Capacitance was measured using the automated capacitance cancellation function of the EPC10 (HEKA). Nominally Mg²⁺-free extracellular solution contained 140 mM NaCl, 3 mM CaCl₂, 2.8 mM KCl, 0 mM MgCl₂, 10 mM HEPES-NaOH, 11 mM glucose (pH 7.2, 300 mOsm). Intracellular solution contained 120 mM Cs-glutamate, 8 mM NaCl, 1 mM MgCl₂, 10 mM HEPES, 10 mM BAPTA, 5 mM EDTA (pH 7.2, 300 mOsm).

Image analysis

All images of an experiment were acquired with the same laser power, scan speed, detector settings, processed equally and analysed blinded. Microtubule surface area was determined with the help of Fiji⁵² by thresholding the α-tubulin staining of platelets and MKs using the same settings for both groups. NMMIIA distribution in resting platelets was analysed on three-dimensional surface plots and profile plots with the help of Fiji⁵². Means of the first and last maxima and the mean between the first and last minima of the histograms were determined and the NMMIIA_max to NMMIIA_min was calculated and depicted. NMMIIA distribution in spread MKs was analysed by grouping and counting the different distribution patterns. The surface of spread platelets was measured by thresholding the F-actin staining. To analyse the distribution/content of NMMIIA in spread platelets the area of the NMMIIA staining was correlated to the total cell size (F-actin) staining.

Data analysis

The presented results are mean±s.d. from at least three independent experiments per group, if not otherwise stated. Data distribution was analysed using the Shapiro–Wilk test and differences between control and knockout mice were statistically analysed using Student’s t-test or Wilcoxon–Mann–Whitney test, respectively. P-values<0.05 were considered as statistically significant *P<0.05; **P<0.01; ***P<0.001. Results with a P value>0.05 were considered as not significant (NS).

Additional information

How to cite this article: Stritt, S. et al. Defects in TRPM7 channel function deregulate thrombopoiesis through altered cellular Mg²⁺ homeostasis and cytoskeletal architecture. Nat. Commun. 7:11097 doi: 10.1038/ncomms11097 (2016).

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Acknowledgements

We thank the patients and healthy volunteers for donating blood samples; Jonas Müller, Stefanie Hartmann and Birgit Midloch for excellent technical assistance. We are thankful to Prof David E. Clapham for providing the Trpm7^fl/fl mice. We acknowledge Prof Willem Ouwehand, the NIHR BioResource—Rare Diseases and the associated BRIDGE-BPD consortium for performing whole-exome sequencing of the index cases. We thank the microscopy platform of the Bioimaging Centre (Rudolf Virchow Centre) for providing technical infrastructure and support. This work was supported by the Deutsche Forschungsgemeinschaft (SFB 688 to H.S., B.N. and A.B.). S.S. and J.M.M.v.E. were supported by a grant of the German Excellence Initiative to the Graduate School of Life Sciences, University of Würzburg. V.C., S.Z. and T.G. were supported by the Deutsche Forschungsgemeinschaft, TRR 152. Research in the Ouwehand laboratory was supported by programme grants from the European Commission, NIHR and the BHF under numbers RP-PG-0310-1002 and RG/09/12/28096 and the laboratory also receives funding from NHS Blood and Transplant. M.A.L. was supported by the Imperial College London Biomedical Research Centre.

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Authors and Affiliations

Chair of Experimental Biomedicine, University Hospital, University of Würzburg, Josef-Schneider-Strasse 2, 97078 Würzburg, Germany,
Simon Stritt, Paquita Nurden, Sanjeev K. Gotru, Judith M M. van Eeuwijk, Harald Schulze, Bernhard Nieswandt & Attila Braun
Rudolf Virchow Centre, University of Würzburg, Josef-Schneider-Strasse 2, 97078 Würzburg, Germany,
Simon Stritt, Sanjeev K. Gotru, Judith M M. van Eeuwijk, Bernhard Nieswandt & Attila Braun
Institut Hospitalo-Universitaire LIRYC, Plateforme Technologique d'Innovation Biomédicale, Hôpital Xavier Arnozan, Avenue du Haut Lévêque, 33604 Pessac, France,
Paquita Nurden & Alan T. Nurden
Assistance Publique—Hôpitaux de Paris, Haematological Laboratory, Armand Trousseau Children Hospital, 26 Avenue du Docteur Arnold-Netter, 75012 Paris, France,
Remi Favier & Paola Ballerini
Inserm U1170, Gustave Roussy, University Paris Sud, 114 Rue Edouard Vaillant, 94805 Villejuif, France,
Remi Favier & Marie Favier
Inra, UMR_INRA 1260, 27 Boulevard Jean Moulin, 13385 Marseille, France,
Marie Favier
Aix Marseille Université, 58 Boulevard Charles Livon, 13284 Marseille, France,
Marie Favier
Inserm UMR_S 1062, 27 Boulevard Jean Moulin, 13385 Marseille, France,
Marie Favier
Walther-Straub-Institute for Pharmacology and Toxicology, Ludwig-Maximilians University Munich, Goethestraße 33, 80336 Munich, Germany,
Silvia Ferioli, Lorenz Mittermeier, Susanna Zierler, Vladimir Chubanov & Thomas Gudermann
Division of Haematology, Children’s Hospital of Philadelphia, 3401 Civic Center Blvd, Philadelphia, Pennsylvania 19104, USA,
Michele P. Lambert
Department of Paediatrics, Perelman School of Medicine at the University of Pennsylvania, 3400 Civic Center Blvd, Philadelphia, Pennsylvania 19104, USA,
Michele P. Lambert
Department of Haematology, University of Cambridge, Cambridge Biomedical Campus, Francis Crick Ave, Cambridge CB2 0S, UK,
Ernest Turro
NHS Blood and Transplant, Cambridge Biomedical Campus, Francis Crick Ave, Cambridge CB2 0S, UK,
Ernest Turro
Medical Research Council Biostatistics Unit, Cambridge Institute of Public Health, Cambridge Biomedical Campus, Robinson Way, Cambridge CB2 0SR, UK,
Ernest Turro
NIHR BioResource—Rare Diseases, Cambridge University Hospitals, Cambridge Biomedical Campus, Hills Rd, Cambridge CB2 0QQ, UK,
Ernest Turro
Department of Internal Medicine I, Endocrinology and Diabetes Unit, University Hospital of Würzburg, Oberdürrbacher Strasse 6, 97080 Würzburg, Germany,
Stephanie Burger-Stritt
Department of Molecular & Cellular Physiology, Graduate School of Medicine, University of the Ryukyus, 207 Uehara, Okinawa 903-0215, Japan,
Masayuki Matsushita
Centre for Haematology, Hammersmith Campus, Imperial College Academic Health Sciences Centre, Imperial College London, Du Cane Road, London SW7 2AZ, UK,
Michael A. Laffan
Imperial College Healthcare NHS Trust, Du Cane Road, London, SW7 2AZ, UK
Michael A. Laffan
Comprehensive Pneumology Center Munich (CPC-M), German Center for Lung Research, Max-Lebsche-Platz 31, 81377 Munich, Germany,
Thomas Gudermann
DZHK (German Centre for Cardiovascular Research), Munich Heart Alliance, Lazarettstraße 36, 80636 Munich, Germany,
Thomas Gudermann

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Simon Stritt
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Paquita Nurden
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Remi Favier
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Marie Favier
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Silvia Ferioli
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Judith M M. van Eeuwijk
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Harald Schulze
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Alan T. Nurden
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Michele P. Lambert
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Ernest Turro
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Stephanie Burger-Stritt
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Masayuki Matsushita
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Lorenz Mittermeier
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Paola Ballerini
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Susanna Zierler
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Michael A. Laffan
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Vladimir Chubanov
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Thomas Gudermann
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Bernhard Nieswandt
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Contributions

S.S. designed the research, performed the experiments, analysed the data and wrote the manuscript. P.N. and R.F. performed the experiments, analysed the data and contributed to the writing of the manuscript. M.F., S.F., S.K.G. and J.M.M.v.E. performed the experiments and analysed the data. L.M. helped with the breeding of mice. H.S. and A.T.N. helped with genetic analyses and commented on the manuscript. M.P.L. collected the phenotype data. E.T. performed genetic analyses. S.B.-S. provided ethics support and helped with the blood withdrawals. M.M. provided Trpm7^KI mice. P.B. analysed the data. S.Z. and V.C. performed the experiments, analysed the data and commented on the manuscript. M.A.L. co-established and co-directed the BRIDGE-BPD consortium and directed the genetic analysis by E.T. T.G. analysed the data and commented on the manuscript. B.N. analysed the data and wrote the manuscript. A.B. supervised the research, performed the experiments, analysed the data and wrote the manuscript.

Corresponding authors

Correspondence to Bernhard Nieswandt or Attila Braun.

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Competing interests

The authors declare no competing financial interests.

Supplementary information

Supplementary Information

Supplementary Figures 1-29, Supplementary Tables 1-3 and Supplementary Note (PDF 6801 kb)

Supplementary Movie 1

Representative intravital two-photon microscopy time-lapse movie of bone marrow MKs in the skull of a control mouse, showing proplatelet formation. Scale bar, 50 μm. (MOV 5992 kb)

Supplementary Movie 2

Representative intravital two-photon microscopy time-lapse movie of bone marrow MKs in the skull of a Trpm^{7fl/fl-Pf4Cre} mouse with mature MKs attempting to form proplatelets but fail to do so. Scale bar, 50 μm. (MOV 8076 kb)

Supplementary Movie 3

Representative intravital two-photon microscopy time-lapse movie of bone marrow MKs in the skull of a Trpm^{7fl/fl-Pf4Cre} mouse during proplatelet formation (arrow) showing short and bulky proplatelet protrusion that remains attached to the MK body. Scale bar, 50 μm. (MOV 3512 kb)

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Stritt, S., Nurden, P., Favier, R. et al. Defects in TRPM7 channel function deregulate thrombopoiesis through altered cellular Mg²⁺ homeostasis and cytoskeletal architecture. Nat Commun 7, 11097 (2016). https://doi.org/10.1038/ncomms11097

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Received: 20 August 2015
Accepted: 18 February 2016
Published: 29 March 2016
DOI: https://doi.org/10.1038/ncomms11097

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