research papers
Structural and biochemical insights into Zn2+-bound EF-hand proteins, EFhd1 and EFhd2
aSchool of Life Sciences, Gwangju Institute of Science and Technology, Gwangju, Republic of Korea, bSteitz Center for Structural Biology, Gwangju Institute of Science and Technology, Gwangju, Republic of Korea, and cDepartment of Chemistry, Gwangju Institute of Science and Technology, Gwangju, Republic of Korea
*Correspondence e-mail: eom@gist.ac.kr
EF-hand proteins, which contain a Ca2+-binding EF-hand motif, are involved in regulating diverse cellular functions. Ca2+ binding induces conformational changes that modulate the activities of EF-hand proteins. Moreover, these proteins occasionally modify their activities by coordinating metals other than Ca2+, including Mg2+, Pb2+ and Zn2+, within their EF-hands. EFhd1 and EFhd2 are homologous EF-hand proteins with similar structures. Although separately localized within cells, both are actin-binding proteins that modulate F-actin rearrangement through Ca2+-independent actin-binding and Ca2+-dependent actin-bundling activity. Although Ca2+ is known to affect the activities of EFhd1 and EFhd2, it is not known whether their actin-related activities are affected by other metals. Here, the crystal structures of the EFhd1 and EFhd2 core domains coordinating Zn2+ ions within their EF-hands are reported. The presence of Zn2+ within EFhd1 and EFhd2 was confirmed by analyzing anomalous signals and the difference between anomalous signals using data collected at the peak positions as well as low-energy remote positions at the Zn K-edge. EFhd1 and EFhd2 were also found to exhibit Zn2+-independent actin-binding and Zn2+-dependent actin-bundling activity. This suggests the actin-related activities of EFhd1 and EFhd2 could be regulated by Zn2+ as well as Ca2+.
Keywords: EFhd1; EFhd2; crystal structure; EF-hands; actin-binding/bundling protein.
1. Introduction
Actin is distributed in the cytosol and within some ; Dos Remedios et al., 2003). These ABPs all contain domains related to actin binding or regulation. The calponin homology (CH) domain is one of the most common modules. It consists of six α-helices (helix I–VI) and forms a compact structure through a network of hydrophobic interactions (Yin et al., 2020; Bramham et al., 2002). The domain both mediates actin binding and serves a regulatory function (Gimona et al., 2002). Another actin-related domain is the formin homology 2 (FH2) domain, an independently folding region conserved in the formin homology family (Higgs & Peterson, 2005; Wallar & Alberts, 2003). The mostly α-helical FH2 domain forms a unique dimer tethered together at either end (Xu et al., 2004). It is able to influence actin dynamics and contribute to actin filament assembly and elongation (Higgs & Peterson, 2005).
and contributes to cell migration, division and trafficking as well as to the maintenance of the proper cell shape. All of these phenomena rely on F-actin, a filamentous polymer composed of G-actin monomers, which undergoes assembly, disassembly, severing, branching and bundling mediated by various actin-binding proteins (ABPs) during the course of its activity (Winder & Ayscough, 2005Several ABPs also contain EF-hand motifs (EF-hands), which are helix–loop–helix structures, mostly found in pairs, within the hydrophobic core of EF-hand domains (Day et al., 2002; Denessiouk et al., 2014). A number of EF-hand proteins undergo Ca2+-induced conformational changes that expose the hydrophobic surface of each EF-hand domain, whereas others are much less affected by Ca2+. EF-hands are required for transducing Ca2+ signals into metabolic or mechanical responses and also serve to buffer Ca2+ levels within cells (Nelson & Chazin, 1998). Among the EF-hand protein superfamily, EFhd1 and EFhd2 are classified as ABPs. EFhd1 is localized within mitochondria, where it acts as a Ca2+-sensor for mitoflash activation (Tominaga et al., 2006; Hou et al., 2016). It is related to neuronal differentiation and pro/pre B-cell development (Tominaga et al., 2006; Stein et al., 2017). Neuronal energy homeostasis and mitochondrial morphology are influenced by both EFhd1 and β-actin (Xie et al., 2018; Ulisse et al., 2020). As an ABP within mitochondria, EFhd1 may be involved in regulating mitochondrial morphology via its Ca2+-dependent β-actin-bundling activity (Mun et al., 2021).
EFhd2 is a cytoskeletal Ca2+-sensor protein localized in the cytosol (Purohit et al., 2014). It was first detected in human CD8+ lymphocytes, where it stabilizes actin filaments by regulating the accessibility of F-actin to cofilin, which depolymerizes the F-actin (Vuadens et al., 2004; Huh et al., 2013). EFhd2 also regulates cytokine expression and lamellipodial dynamics through modulation of actin dynamics (Ramesh et al., 2009; Kwon et al., 2013), and the Ca2+-dependent actin-bundling activity of EFhd2 contributes to cell migration and spreading (Kwon et al., 2013). In addition, EFhd2 may play an important role in several neurodegenerative diseases, as it is associated with tau in Alzheimer's disease and other neurological disorders (Ferrer-Acosta et al., 2013; Vega, 2016). Recently, it was found that EFhd2 is up-regulated in the cardiomyocytes during cardiac remodeling and repair (Giricz et al., 2020). Although the functions and cellular localization of EFhd1 and EFhd2 differ, the two proteins exhibit a high degree of sequence identity (65%) (Mun et al., 2021; Dutting et al., 2011; Park et al., 2016). The actin-bundling activity of EFhd2 requires Ca2+ for maintenance of the rigidity of EF-hands (Park et al., 2016), and EFhd1 likely requires Ca2+ for actin bundling for the same reason.
Notably, Ca2+ is not the only metal that binds to EF-hands. For instance, Pb2+ reportedly activates calmodulin (CaM) and calcium binding protein (CaBP) by binding with higher affinity than Ca2+ (Fullmer et al., 1985; Richardt et al., 1986; Hui & Vogel, 1998). Moreover, the of the Pb2+–CaM complex confirms that Pb2+ can substitute for Ca2+ within the EF-hands of CaM (Kursula & Majava, 2007). It is unknown, however, whether EF-hands in EFhd1 and/or EFhd2 bind Zn2+ or whether the Zn2+ binding affects their function, although we previously observed Zn2+-mediated multimerization of EFhd1 or EFhd2 via conserved residues at the crystal-packing interface of EFhd1. We previously reported EFhd1 and EFhd2 structures in the Ca2+-bound state (Mun et al., 2021; Park et al., 2016). Within the structure of EFhd1, however, we did not experimentally identify the metal ions, though both Ca2+ and Zn2+ were present under the crystallization conditions (Mun et al., 2021).
Here, we report the structures of the mouse EFhd1 and human EFhd2 core domains in the Zn2+-bound state (EFhd1Zn, residues 79–180; EFhd2Zn, residues 82–180). We demonstrate Zn2+ coordination within the EF-hands of EFhd1 and EFhd2 through analysis of anomalous signals. Lastly, we show that Zn2+ affects the actin-bundling activities of EFhd1 and EFhd2 but not their actin binding, which suggests the possibility that Zn2+ acts to regulate EFhd1 and EFhd2 in both physiological and pathophysiological processes.
2. Results
2.1. Structures of Ca2+- or Zn2+-bound EFhd1 and EFhd2
We previously reported the 2+-mediated multimerization of EFhd1 and EFhd2 (Mun et al., 2021). On the other hand, the effects of Zn2+ on the actin-binding/bundling activities of EFhd1 and EFhd2 remain unknown. Therefore, to assess the role of Zn2+ in actin binding/bundling and its effect on the structures of the two proteins, we performed structural and biochemical studies with Zn2+-bound EFhd1 and EFhd2. To obtain EFhd1Zn, we added a final concentration of 1 mM CaCl2 to the protein solutions and used a crystallization buffer containing 2.5 mM ZnSO4. To obtain EFhd2Zn, we added EGTA and EDTA to final molar concentrations 15 times higher (0.18 mM each) than that of the protein to the EFhd2 proteins to remove native ions and then dialyzed the proteins. Thereafter, ZnCl2 was added to the protein solution to a final concentration of 0.75 mM. In the case of EFhd1Ca, CaCl2 was added to the protein solution directly to a final concentration of 4 mM. We crystallized these proteins and determined the crystal structures of EFhd1Zn, EFhd2Zn and EFhd1Ca at resolutions of 1.72, 2.60 and 2.80 Å, respectively, using methods [Table 1, Fig. 1(a)]. EFhd1Zn, EFhd2Zn and EFhd1Ca comprised two EF-hands, a ligand mimic (LM) helix at the C-terminus, a PR region at the N-terminus and a C-terminal linker (Mun et al., 2021; Park et al., 2016).
of EFhd1 and suggested the occurrence of Zn
‡Rmerge = ∑h ∑i|I(h)i − 〈I(h)〉|/[∑h∑i I(h)i], where I(h) is the intensity of the reflection of h, ∑h is the sum over all reflections and ∑i is the sum over i measurements of reflection h. §Redundancy: we collected EFhd1Ca and EFhd2Zn datasets using 720 frames, and EFhd1Zn datasets using 360 frames due to the radiation decay. ¶Rwork = ∑hkl ||Fo|−|Fc||/(∑hkl|Fo|); 5% of the reflections were excluded for the Rfree calculation. |
σA-weighted mFo − DFc maps (Fo − Fc map) above 8σ were observed with these EFhd proteins, which suggests metal binding [Table 2, Figs. 1(b)–1(d)]. Considering our protein preparation protocols, we expected that the map originated from Ca2+ or Zn2+. To identify the coordinating metal, we calculated anomalous difference maps. Because Zn2+ and Ca2+ have different numbers of anomalous electrons at the wavelength the EFhd1 data were collected (Zn2+ f″ = 3.9 electrons, Ca2+ f″ = 0.9 electrons), the anomalous difference map should be weaker in the Ca2+-bound state than the Zn2+-bound state. In the case of EFhd1Ca, the peak heights in the anomalous difference map were 3.4 and 2.7σ for EF-hand 1 (EF1) and EF-hand 2 (EF2), respectively. EFhd1Zn had peak heights of 23.0 and 14.0σ in the anomalous difference maps for EF1 and EF2, respectively. This suggests the metal coordinated in EFhd1Ca was Ca2+ while that coordinated in EFhd1Zn was Zn2+ [Table 3, Figs. 1(c) and 1(f)].
‡Zn K-edge. §Near Zn K-edge. |
‡Zn K-edge. §Near Zn K-edge. |
At a wavelength near the Zn K-edge (λ = 1.2851 Å/9648 eV), the Ca2+ f″ and Zn2+ f″ were 0.9 and 0.5 electrons, respectively. For that reason, we cannot rule out the possibility that the anomalous signals for EFhd2Zn originated from Ca2+, because the wavelength for the EFhd2Zn data collection was near the Zn K-edge [Fig. 1(g)]. To identify the metal coordinated in EFhd2, we analyzed the difference between anomalous difference maps (ΔAno). We used a single EFhd2Zn crystal to collect datasets at the peak and remote positions of the Zn K-edge, which are termed EFhd2Zn(P) or EFhd2Zn(R), respectively [Table 4]. After calculation of the anomalous difference maps for each dataset, the anomalous difference map for EFhd2Zn(R) was subtracted from that for EFhd2Zn(P) to calculate the ΔAno map. The peak heights of the ΔAno map calculated from EFhd2Zn(P) and EFhd2Zn(R) were 11.1 and 9.3σ in EF1 and EF2, respectively [Figs. 1(h) and 1(i)]. This demonstrates that the metal coordinated in the EF-hands of EFhd2 was Zn2+.
Within the crystal structures of EFhd1Ca, EFhd1Zn and EFhd2Zn, the anomalous difference maps were observed near the α4 of each protein, which is situated at the crystal-packing interface [Figs. 1(e)–1(g)]. Analyzing the metal coordination geometry using CheckMyMetal and the MetalPDB server, we expected that those peaks originated from Zn2+ (Zheng et al., 2017; Putignano et al., 2018). With EFhd1, the peak height in the anomalous difference map for the metal coordinated at the crystal-packing interface was 23.0σ in the case of EFhd1Ca. The metals coordinated at the crystal-packing interface of EFhd1Zn had peak heights of 65.0 or 17.2σ in the anomalous difference map [Table 3, Figs. 1(e) and 1(f)]. In the case of EFhd2Zn, the peak height of the ΔAno map was 9.6σ at the crystal-packing interface [Figs. 1(h) and 1(i)]. Considering both the anomalous signals and the metal coordination geometry, we deemed the metal at the interface to be Zn2+. Collectively, the two EF-hands of EFhd1 and EFhd2 are able to coordinate Zn2+ as well as Ca2+, and Zn2+ could also be coordinated at the crystal-packing interface.
2.2. Comparison of the overall structures of Ca2+- and Zn2+-bound EFhd1 and EFhd2
The overall structures of EFhd1Zn and EFhd2Zn superimposed well onto each other [the root mean square deviation (RMSD) of EFhd1Zn and EFhd2Zn was 0.381 Å for 80 Cα atoms; Fig. 2(a)]. Moreover, when we superimposed EFhd1Zn on EFhd1Ca and EFhd2Zn on EFhd2Ca (PDB entry 5i2l; Park et al., 2016) to analyze the structural differences between the Zn2+- and Ca2+-bound states, we found that they too superimposed well [RMSD for EFhd1Zn and EFhd1Ca was 0.102 Å for 98 Cα atoms; RMSD for EFhd2Zn and EFhd2Ca was 0.224 Å for 87 Cα atoms; Figs. 2(b) and 2(c)] (Park et al., 2016). Earlier findings suggest that EFhd1 and EFhd2 maintain open conformations irrespective of the presence of Ca2+ (Mun et al., 2021; Park et al., 2016). EFhd1Zn and EFhd2Zn also maintained open conformations similar to those of EFhd1Ca and EFhd2Ca. Collectively, these results show that the overall structure of EFhd1Zn is similar to that of EFhd2Zn, and the binding of Zn2+ has little effect on the conformations of the EFhd1 and EFhd2 core domains.
2.3. Structural comparison of the EF-hands in the Zn2+-bound EFhd1 and EFhd2 core domains
Consensus residues for Ca2+ or Mg2+ coordination within EF-hands are at positions 1(X), 3(Y), 5(Z), 7(−Y), 9(−X) and 12(−Z). Conventionally, one water molecule participates in metal coordination at position 9(−X). These residues coordinate Ca2+ through seven ligands with pentagonal bipyramidal geometry and coordinate Mg2+ through six ligands with octahedral geometry (Lewit-Bentley & Réty, 2000; Grabarek, 2006; Nelson & Chazin, 1998). In EFhd1Zn and EFhd2Zn, both EF-hands coordinated Zn2+. Zn1 and Zn2 in both EFhd1Zn and EFhd2Zn were each coordinated by seven oxygen atoms. In addition, two water molecules coordinated Zn1 at positions 3(Y) and 9(−X). The Gly at position 3(Y) (G106 in EFhd1, G107 in EFhd2) and the Asp at position 9(−X) (D112 in EFhd1, D113 in EFhd2) are not used to coordinate Zn1 [Figs. 3(a) and 3(c)]. Therefore, the geometry of the Zn1 coordination formed a distorted pentagonal bipyramid in the two proteins. Only one water coordinated Zn2 at position 9(−X) (S148 in EFhd1, S149 in EFhd2), which is consistent with the conventional one water coordination at that position [Figs. 3(b) and 3(d)]. As a result, the geometry of the Zn2 coordination formed the typical pentagonal bipyramid. To compare their EF-hands, we superimposed the structure EF1 or EF2 in EFhd1Zn on EFhd2Zn and found that both superimposed well [RMSD for EF1 in EFhd1Zn and EFhd2Zn was 0.300 Å for 32 Cα atoms; RMSD for EF2 in EFhd1Zn and EFhd2Zn was 0.456 Å for 36 Cα atoms; Figs. 3(e) and 3(f)]. In addition, measurement of the average distance between Zn2+ and its coordinating ligands in the EF-hands revealed that the overall average of Zn2+ coordination distances in EFhd1Zn and EFhd2Zn are similar to the average Zn2+–oxygen distance (2.3 ± 0.5 Å) (Table S1 of the supporting information) (Ireland & Martin, 2019).
2.4. Structural comparison of the EF-hands of the Zn2+- and Ca2+-bound states of EFhd1 or EFhd2
We next compared the structures of the EF-hands in the Ca2+- and Zn2+-bound states. When we superimposed the EF-hands of EFhd1Ca and EFhd1Zn and those of EFhd2Ca and EFhd2Zn, they both superimposed well (RMSD for EF1 in EFhd1Ca and EFhd1Zn = 0.100 Å for 38 Cα atoms; RMSD for EF2 in EFhd1Ca and EFhd1Zn = 0.088 Å for 35 Cα atoms; RMSD for EF1 in EFhd2Ca and EFhd2Zn = 0.157 Å for 30 Cα atoms; and RMSD for EF2 in EFhd2Ca and EFhd2Zn = 0.253 Å for 36 Cα atoms). In addition, these proteins showed similar geometries in the Ca2+- and Zn2+-bound states. In both proteins EF1 and EF2 formed the distorted or typical pentagonal bipyramid for Ca2+ and Zn2+ coordination [Figs. 4(a)–4(d)]. The side-chain topologies of the EF-hand loop were similar between EFhd1Zn and EFhd1Ca and between EFhd2Zn and EFhd2Ca. This suggests the metal coordination geometries of Ca2+ and Zn2+ are similar within EFhd proteins.
2.5. Structural comparison of the EF-hands in EFhd1Zn and EFhd2Zn with other Zn2+-bound proteins
EFhd proteins were able to coordinate Zn2+ as well as Ca2+ within their EF-hands. Similarly, several other proteins, including Tse3 and calmodulin (CaM), also coordinated Zn2+ within their EF-hand or EF-hand-like motif. In the case of Tse3, an EF-hand-like motif was able to coordinate Ca2+ or Zn2+, and the average coordination distance for Zn2+ is 2.5 Å, which is slightly longer than the average Zn2+–oxygen distance (2.3 ± 0.5 Å; Table S1) (Lu et al., 2014). When we superimposed the EF-hand-like motifs of the Ca2+- and Zn2+-bound Tse3 structures [Tse3Ca (PDB entry 4n80; Lu et al., 2014), Tse3Zn (PDB entry 4n7s; Lu et al., 2014)], they were superimposed well, with an RMSD of 0.125 Å for 31 Cα atoms. Moreover, those EF-hand-like motifs had similar metal coordination geometries: a pentagonal bipyramid with seven ligands, including one water molecule [Fig. 4(e)]. This shows that the structure and metal coordination geometry of the EF-hand-like motif of Tse3 are comparable to those of EF2 of EFhd proteins but different from those of EF1.
CaM also coordinates both Zn2+ and Ca2+ within EF-hands [CaMZn (PDB entry 4hex; Kumar et al., 2013), CaMCa (PDB entry 1cll; Chattopadhyaya et al., 1992)]. Within the structure of CaMZn, which comprises two chains (chains A and B) within the there are three Zn2+-bound EF-hands: EF4 in chain B and EF2 and EF3 in chain A. In the case of EF4, Ca2+ or Zn2+ is coordinated by seven ligands forming a general pentagonal bipyramid [Fig. 4(f)]. In EF2, seven oxygen atoms, including a water oxygen, coordinated Ca2+ (Ca1) with typical pentagonal bipyramidal geometry. Zn2+ (Zn1) was coordinated within EF2 of CaMZn by six oxygen atoms. Because there is no water in the Zn1 coordination, it formed a distorted pentagonal bipyramidal geometry with a vacancy at position 9(−X) [Fig. 4(g)]. In EF3 of CaMCa, Ca2 coordination and geometry were the same as those in EF2 and formed the typical pentagonal bipyramid. Zn2 coordination and geometry in EF3 were also the same as those in EF2 and formed a distorted pentagonal bipyramid with the absence of a water at position 9(−X) [Fig. 4(h)]. The average metal coordination distances differed slightly between Ca2+ and Zn2+. The average coordination distances with Ca2+ in CaM EF4, EF2 and EF3 were 2.4, 2.3 and 2.4 Å, respectively; the average coordination distances with Zn2+ in these two EF-hands were 2.3 Å for EF4 and 2.4 Å for both EF2 and EF3 (Table S1). In CaMZn, all of the average coordination distances for Zn2+ were around 2.3 ± 0.5 Å, the average Zn2+–oxygen distance. When we superimposed EF2 of CaMZn and CaMCa or EF3 of CaMZn and CaMCa, the topologies of the alpha helices in CaMZn and CaMCa differed slightly in both cases [Figs. 4(g) and 4(h)]. Thus, there were no significant structural differences between the Ca2+- and Zn2+-bound EF-hands in EFhd proteins or EF4 of CaM. For EF2 and EF3 of CaM, however, differences between the Ca2+- and Zn2+-bound EF-hands were detected.
ALG-2 belongs to the penta-EF-hand (PEF) protein family and contains three metal binding EF-hands (Jia et al., 2001). When we superimposed the EF-hands of Ca2+-bound and Zn2+-bound ALG-2 [ALG-2Ca (PDB entry 2zn9; Suzuki et al., 2008) and ALG-2Zn (PDB entry 2zn8; Suzuki et al., 2008)], they superimposed well [RMSD for EF1 = 0.265 Å for 31 Cα atoms; RMSD for EF3 = 0.292 Å for 35 Cα atoms; Figs. 4(i) and 4(j)]. Ca1 was coordinated by six oxygen atoms, including a water oxygen and excluding the S40 oxygen, and assumed a pentagonal bipyramidal geometry. In ALG-2Zn, Zn1 was coordinated by the six oxygen atoms in EF1 with the distorted pentagonal bipyramidal geometry [Fig. 4(i)]. Ca2 was also coordinated by six oxygen atoms in EF3 and formed a distorted pentagonal bipyramid. No water molecule was observed. On rare occasions, two Zn2+ ions, Zn2 and Zn3, simultaneously bound to EF3 in ALG-2Zn. Zn2 and Zn3 were coordinated by six and three oxygen atoms, forming a trigonal prism and distorted tetrahedron, respectively. Asp105 which participated in the Ca2+ coordination bound to Zn2 and Zn3 as a bidentate ligand, as did Glu114, and Asp111, which did not participate in Ca2+ coordination, bound to Zn3 [Fig. 4(j)]. The average coordination distances for Ca2+ in the two EF-hands were 2.4 and 2.5 Å, respectively; the average coordination distances for Zn2+ in EF1 and EF3 were the same as those for Ca2+, which are slightly longer than the average Zn2+–oxygen distance (Table S1). Therefore, the structures of the EF-hands in ALG-2Zn were similar to those in ALG-2Ca, but the metal coordination of ALG-2Zn showed diverse geometries unlike those of ALG-2Ca. These results show that the EF-hand structures and metal coordination of ALG-2Ca are similar to those of EFhd1Ca and EFhd2Ca, but those of ALG-2Zn differ from those of EFhd1Zn and EFhd2Zn.
Consequently, these suggested that the Ca2+ coordination geometry within EF-hands consistently formed a pentagonal bipyramid, whereas the Zn2+ coordination geometry assumed a variety of forms.
2.6. Zn2+-mediated actin-binding/bundling activities of EFhd1 and EFhd2
EFhd1 and EFhd2 both exhibit Ca2+-independent actin-binding and Ca2+-dependent actin-bundling activity (Mun et al., 2021; Huh et al., 2013; Kwon et al., 2013). Because the EF-hands of EFhd1 and EFhd2 are situated within the actin-binding site, we hypothesized that Zn2+ may affect the actin-binding and/or bundling activities of EFhd1 and EFhd2 (Kwon et al., 2013). To determine whether EFhd1 and/or EFhd2 bind F-actin in the presence of Zn2+, we performed in vitro high-speed co-sedimentation assays using α- and β-actin with full-length EFhd1 and EFhd2 [Figs. 5(a) and 5(b)]. As previously reported, EFhd1 and EFhd2 bound α- and β-actin independently of Ca2+ (Mun et al., 2021). As reflected by the percentage bound, the binding affinity of β-actin for EFhd1 was higher than for EFhd2 (β-actin EGTA EFhd1: 34 ± 7%, β-actin EGTA EFhd2: 23 ± 2%, β-actin Ca2+ EFhd1: 32 ± 4% and β-actin Ca2+ EFhd2: 21 ± 1%), whereas the binding affinity of α-actin was similar for EFhd1 and EFhd2 (α-actin EGTA EFhd1: 23 ± 4%, α -actin EGTA EFhd2: 25 ± 3%, α-actin Ca2+ EFhd1: 25 ± 4% and α-actin Ca2+ EFhd2: 22 ± 2%) [Fig. 5(c)]. In the presence of Zn2+, EFhd1 and EFhd2 bound to F-actin and showed similar binding affinities for β- and α-actin (β-actin 20 µM Zn2+ EFhd1: 29 ± 3%, β-actin 20 µM Zn2+ EFhd2: 26 ± 1%, α-actin 20 µM Zn2+ EFhd1: 24 ± 4%, and α-actin 20 µM Zn2+ EFhd2: 22 ± 4%) [Fig. 5(c)]. Thus, both EFhd1 and EFhd2 bind actin in the presence of Ca2+, Zn2+ or EGTA. Consequently, actin binding is both Ca2+- and Zn2+-independent.
Lastly, we used 2+ affects the actin-bundling activities of EFhd1 and EFhd2. Because the subcellular localization of α-actin is in the cytosol and β-actin is in mitochondria, we separately analyzed the actin-bundling activities of EFhd1 and EFhd2 with β-actin and α-actin [Figs. 5(d) and 5(e)] (Xie et al., 2018; Storch et al., 2007; Reyes et al., 2011). In the electron micrographs, we observed F-actin bundles in the presence of Ca2+ or Zn2+, but not in the presence of EGTA. We therefore conclude that EFhd1 and EFhd2 are capable of mediating Ca2+-dependent and Zn2+-dependent actin bundling.
with negative staining to assess whether Zn3. Discussion
The Ca2+-binding proteins EFhd1 and EFhd2 regulate Ca2+-dependent F-actin bundling (Mun et al., 2021; Kwon et al., 2013; Park et al., 2016). However, cells contain a variety of metals, and it is unknown whether metals other than Ca2+ also affect EFhd1 and EFhd2 function. In the present study, we determined the crystal structures of EFhd1 and EFhd2 coordinating Zn2+ within their EF-hands and at the crystal-packing interface. In addition, we determined that EFhd1 and EFhd2 bind actin independently of Ca2+ or Zn2+ and also exhibit Ca2+-dependent and Zn2+-dependent actin-bundling activity.
Smaller than Ca2+ (Zn: r = 0.74 Å versus Ca: r = 0.99 Å), Zn2+ contributes to diverse physiological functions (Kambe et al., 2015; Allouche et al., 1999). When Zn2+ interacts with proteins, Cys, His, Asp and Glu are frequently involved in its coordination (Vahrenkamp, 2007; Laitaoja et al., 2013). Moreover, its lack of effects makes Zn2+ suitable for different coordination numbers and binding geometries in different biological settings (Laitaoja et al., 2013). Through analysis of EF-hand structures, which are able to coordinate Ca2+ or Zn2+, it was found that the Zn2+ can be coordinated through more diverse metal coordination geometries than Ca2+ (Grabarek, 2006, Kumar et al., 2013; Lu et al., 2014; Suzuki et al., 2008). Through anomalous signal analysis, we demonstrated that EFhd1 and EFhd2 are able to coordinate not only Ca2+ but also Zn2+ within their EF-hands. For the EFhd proteins, the binding of Ca2+ or Zn2+ had little effect on the conformations of EFhd1 or EFhd2, which probably explains why they are able to mediate actin bundling in the presence of Zn2+. In an earlier study, revealed the presence of bundled actin filaments in ZnO-treated cells (Garcia-Hevia et al., 2016). Because EFhd2 is localized in the cytosol, Zn2+-dependent actin bundling mediated by EFhd2 may have contributed to the bundled actin filaments detected in that study.
In resting cells, the cytosolic [Ca2+] is ∼100 nM, whereas the cytosolic [Zn2+] is tightly controlled in the picomolar to low nanomolar range (Kambe et al., 2015; Esteras & Abramov, 2020; Patergnani et al., 2020). In addition, the EF-hands of EFhd2 exhibit high Ca2+-binding affinities (Kd of EF1 = 96 ± 15 nM, Kd of EF2 = 70 ± 1 nM), implying that EFhd2 is likely to have Ca2+ bound within resting cells. The binding affinity of EFhd1 for Ca2+ has not been previously reported. We therefore measured the affinity of Ca2+ for EFhd1 using ITC (Kd = 22.3 ± 0.2 nM; Fig. S2 of the supporting information). The mitochondrial [Ca2+] is similar to that in the cytosol, whereas [Zn2+] is in the picomolar range under resting conditions (Kambe et al., 2015; Esteras & Abramov, 2020; Patergnani et al., 2020). Thus, EFhd1 may also mainly coordinate Ca2+ in resting cells due to the higher [Ca2+] than [Zn2+]. On the other hand, under conditions of Zn2+ overload, the mitochondrial [Zn2+] can reach the submicromolar range (Sensi et al., 2003), and Zn2+-mediated multimerization of EFhd1 may occur, as we previously reported (Mun et al., 2021).
EFhd2 is a novel amyloid protein that forms filaments using its coiled-coil region to self-oligomerize. EFhd2 and tau granules have been observed in fractions obtained from Alzheimer disease (AD) brains, suggesting a novel amyloid protein may form nucleation centers to induce the formation of tau aggregates (Ferrer-Acosta et al., 2013). We previously suggested that Zn2+ mediates multimerization of EFhd1 and EFhd2 through protein aggregation. In addition, we confirmed that Zn2+ mediates crystal-packing interactions between EFhd2 molecules, which raises the possibility of the involvement of Zn2+-mediated multimerization in AD. Consistent with that idea, Zn2+ is reported to be significantly elevated in the AD neuropil (Lovell et al., 1998). We therefore suggest that Zn2+ may be the seed for self-oligomerization of the novel amyloid protein EFhd2.
In the present study, we determined the crystal structures of EFhd1 and EFhd2 in the Zn2+-bound state, without the coiled-coil region. We also found that Zn2+-bound full-length EFhd1 and EFhd2 bind actin and mediate actin bundling. However, understanding the coiled-coil regions of EFhd1 and EFhd2 is important, given its association with self-oligomerization and actin-bundling activity (Kwon et al., 2013; Ferrer-Acosta et al., 2013). We therefore anticipate that structural studies of full-length EFhd1 and EFhd2 alone and in complex with actin will be useful for achieving a fuller understanding of the biological functions of these two proteins.
4. Materials and methods
4.1. Plasmid
Mouse EFhd1 ΔNTD (residues 69−240) and the human EFhd2 core domain (residues 70−184) were amplified from full-length mouse EFhd1 (residues 1−240) and human EFhd2 (residues 1−240), respectively, using (PCR). The amplified EFhd1 ΔNTD was cloned into a modified pET28a vector (Novagen) containing an N-terminal His6 tag and a tobacco etch virus (TEV) protease cleavage site (Glu–Asn–Leu–Tyr–Phe–Gln/Gly). The amplified EFhd2 core domain was cloned into a modified pET41a vector containing glutathione S-transferase (GST) with a TEV protease cleavage site. Full-length EFhd1 was cloned into a modified pET28a vector (Novagen) with an N-terminal His6-TEV tag. Full-length EFhd2 was cloned into a modified pET28a vector carrying an N-terminal His6 tag.
4.2. Protein expression and purification of EFhd1 ΔNTD (residues 69−240)
Protein expression and purification of mouse EFhd1 ΔNTD were performed as reported previously (Mun et al., 2021). The target protein was finally purified through a HiLoad 16/60 Superdex 75 gel-filtration column (GE Healthcare Life Sciences) pre-equilibrated with the final buffer [20 mM HEPES-NaOH (pH 7.5), 150 mM NaCl, 0.4 mM PMSF and 14.3 mM β-ME]. The purified protein was concentrated using a 10 K centrifugal filter (Millipore) and stored at −80°C. During purification, the presence of EFhd1 protein was confirmed using SDS–PAGE, and protein degradation was observed following incubation with TEV protease.
4.3. Protein expression and purification of EFhd2 core domain (residues 70−184)
Overall expression of the EFhd2 core domain was similar to that of EFhd1 ΔNTD. Cells transformed with the EFhd2 core domain were harvested by centrifugation, and the cell pellet was suspended in a lysis buffer [50 mM HEPES-NaOH (pH 7.5), 300 mM NaCl, 0.4 mM PMSF and 14.3 mM β-ME], lysed by sonication and centrifuged at 14 000g for 50 min at 4°C. The supernatant was then subjected to GST-bind agarose (Elpis) After washing with the lysis buffer, the target protein was eluted with lysis buffer supplemented with 30 mM glutathione, and the eluted protein was incubated with TEV protease overnight at 4°C to cleave the N-terminal GST-TEV tag. The target protein was further purified through a HiLoad 16/60 Superdex 75 gel-filtration column (GE Healthcare Life Sciences) pre-equilibrated with the final buffer [20 mM HEPES-NaOH (pH 7.5), 150 mM NaCl]. To obtain Zn2+-bound EFhd2 protein, the purified protein was treated with a 15-fold excess of EGTA and EDTA for 30 min at 4°C to remove pre-bound metal ions. The protein was then dialyzed in the final buffer for 24 h at 4°C, changing the buffer every 8 h. The dialyzed protein was concentrated using a 10 K centrifugal filter (Millipore) to 9.4 mg ml−1 and treated with 0.75 mM ZnCl2. The resultant Zn2+-bound protein was stored at −80°C.
4.4. Protein expression and purification of full-length EFhd1 and EFhd2
To investigate their actin-binding and bundling activities, we purified full-length EFhd1 and EFhd2. The protein expression and purification of EFhd1 and EFhd2 were performed as previously reported (Mun et al., 2021). The two proteins were finally purified through a HiLoad 16/60 Superdex 75 gel-filtration column (GE Healthcare Life Sciences) pre-equilibrated with the final buffer containing 20 mM HEPES-NaOH (pH 7.5), 150 mM NaCl, 0.8 mM PMSF and 5 mM DTT. The purified protein was then concentrated using a 10 K centrifugal filter (Millipore) and stored at −80°C.
During purification, the presence of full-length EFhd1 and EFhd2 proteins was confirmed using SDS–PAGE.
4.5. Crystallization of the Ca2+- and Zn2+-bound EFhd1 core domain and Zn2+-bound EFhd2 core domain
We initially attempted to crystallize Ca2+- and Zn2+-bound EFhd1 ΔNTD (residues 69−240). Purified EFhd1 ΔNTD was incubated for at least 20 min on ice after the addition of 4 mM CaCl2 or 1 mM CaCl2 to 20.0 mg ml−1 and 11.2 mg ml−1 protein. Thereafter, 1 mM CaCl2 containing the protein was screened using the sitting-drop vapor-diffusion method in a 96-well sitting drop `IQ' plate (SPT Labtech). We found that EFhd1 ΔNTD was degraded, and the core domain (residues 79−180) was crystallized. The EFhd1 core domain formed rod-shaped crystals after 1 week in reservoir solution containing 80 mM HEPES–NaOH (pH 7.0), 2 mM ZnSO4 and 25%(v/v) Jeffamine ED-2003 (Molecular Dimensions). Additional refinements of the crystallization conditions were performed using the sitting-drop vapor-diffusion method. Drops were prepared by mixing 1 µl of 1 mM CaCl2 containing the protein and 1 µl of reservoir solution or 3 µl of 4 mM CaCl2 containing the protein and 1 µl of reservoir solution. In the former mixing solution, the Zn2+-bound EFhd1 core domain (EFhd1Zn) crystals were obtained using reservoir solution containing 0.1 M Tris–HCl (pH 8.5), 2.5 mM ZnSO4 and 25%(w/v) Jeffamine ED-2001 (Hampton Research). In the latter mixing solution, crystals of Ca2+-bound EFhd1 core domain (EFhd1Ca) were obtained using reservoir solution containing 0.1 M Tris–HCl (pH 8.5), 0.4 mM ZnSO4 and 25% Jeffamine ED-2001 (Hampton Research). For data collection, crystals were cryoprotected by transferring them to mother liquor containing 30%(v/v) glycerol and flash freezing in liquid nitrogen.
To obtain crystals of Zn2+-bound EFhd2 core domain (EFhd2Zn), we performed an initial screening using the sitting-drop vapor-diffusion method in a 96-well sitting drop `IQ' plate (SPT Labtech). The EFhd2 core domain formed cubic crystals after 1 week in reservoir solution containing 0.1 M HEPES-NaOH (pH 7.5), 25%(w/v) PEG 8000. Additional refinements of the crystallization conditions were performed using the sitting-drop vapor-diffusion method with drops prepared by mixing 1 µl of protein and 1 µl of reservoir solution. Zn2+-bound EFhd2 core domain crystals were obtained using the same reservoir solution used for the initial screening. For data collection, crystals were cryoprotected by transferring them to mother liquor containing 20% glycerol and 1 mM ZnCl2 and flash freezing in liquid nitrogen.
4.6. X-ray data collection, and refinement
X-ray diffraction data for EFhd1Ca, EFhd1Zn and EFhd2Zn were collected at 100 K using synchrotron X-ray sources on beamline 5C at the Pohang Accelerator Laboratory (PAL, South Korea). Ultimately, we collected the best resolution diffraction data for EFhd1Ca, EFhd1Zn and EFhd2Zn at 2.80, 1.72 and 2.60 Å resolution, respectively. Diffraction data were collected at wavelengths corresponding to the peak position of the Zn K-edge (λ = 1.2826 Å/9669 eV) or near the Zn K-edge (λ = 1.2851 Å/9648 eV). The crystals belonged to the P212121 (a = 44.3, b = 47.9 and c = 63.4 Å for EFhd1Ca; a = 44.2, b = 47.5 and c = 63.7 Å for EFhd1Zn; and a = b = c = 92.8 Å for EFhd2Zn; with α = β = γ = 90°). The diffraction data for EFhd1Ca and EFhd1Zn were indexed, processed and scaled using the HKL2000 suite (Otwinowski & Minor, 1997). The diffraction data for EFhd2Zn were indexed and integrated using DIALS/xia2 in CCP4i2, and data merging and scaling were performed using AIMLESS from CCP4 (Winter et al., 2018; Evans & Murshudov, 2013; Winn et al., 2011; Winter, 2010; Potterton et al., 2018). was carried out using Phaser-MR in the Phenix program suite, using the structures of the EFhd1 (PDB entry 7clt; Mun et al., 2021) and EFhd2 core domains (PDB entry 5i2l) as the templates (Mun et al., 2021; Park et al., 2016; McCoy et al., 2007; Liebschner et al., 2019). Additional model building was performed using the program Coot (Emsley & Cowtan, 2004). Iterative was performed with phenix.refine (Liebschner et al., 2019; Afonine et al., 2012). The N- and C-terminals of EFhd1Ca, EFhd1Zn and EFhd2Zn were partially disordered. Details of the data collection and are provided in Table 1.
4.7. Anomalous X-ray diffraction data collection at peak and remote positions of the Zn K-edge
To confirm Zn2+-binding by the EF-hands of EFhd2Zn, we collected diffraction data at wavelengths corresponding to the peak (λ = 1.2823 Å/9669 eV) and low-energy remote positions of the Zn K-edge (λ = 1.2917 Å/9599 eV). The diffraction data for EFhd2Zn [data title: EFhd2Zn(P), EFhd2Zn(R)] were indexed, processed and scaled using the HKL2000 suite (Otwinowski & Minor, 1997). was carried out with Phaser-MR in the Phenix program suite, using the structures of the EFhd2 core domain (PDB entry 5i2l) as the template (McCoy et al., 2007; Liebschner et al., 2019). We created difference between anomalous difference maps (ΔAno) using the program Coot (Emsley & Cowtan, 2004). Details of the data collection and are provided in Table 4.
|
4.8. Structural analysis
All structural figures were generated using PyMOL (version 1.8.6.0; Schrödinger LLC). The σA-weighted mFo − DFc, anomalous difference and ΔAno maps were converted to the CCP4 format using phenix.maps tools (Liebschner et al., 2019; Pražnikar et al., 2009) and were visualized in PyMOL.
4.9. Measurement of the Ca2+ binding affinity of wild-type EFhd1 using ITC
To assess the Ca2+ binding affinity of the EFhd1 core domain, we purified EFhd1 (69–200), which is more stable than EFhd1 ΔNTD or the full-length protein. For protein expression and purification of EFhd1 (69–200), the His6–TEV tag cleavage and gel-filtration steps were same as those used for EFhd1 ΔNTD, except the final buffer contained 20 mM HEPES–NaOH (pH 7.5) and 150 mM NaCl. After purification, EFhd1 (69−200) was treated with a tenfold excess of EGTA and EDTA for 30 min at 4°C to remove pre-bound metal ions. The protein was then dialyzed for 24 h at 4°C in buffer containing 50 mM Tris–HCl (pH 8.5) and 20 mM NaCl. The dialyzed EFhd1 (69–200) was again treated for 30 min at 4°C with a tenfold excess of EGTA and EDTA, after which the protein was again dialyzed for 24 h using the same dialysis buffer, which was refreshed every 8 h. The dialyzed protein was concentrated to 20 µM, and the ligand solution (0.3 mM CaCl2) was prepared in the same buffer. Each EFhd1 (69–200) sample was titrated with 30 injections of ligand (6 µl) in a VP-ITC calorimeter (MicroCal). All measurements were carried out at 20°C, and binding isotherm analysis and fitting were conducted using the Origin software supplied with the calorimeter.
4.10. In vitro actin-binding assay
Actin co-sedimentation assays were performed as previously described (Mun et al., 2021; Kwon et al., 2013). In brief, non-muscle actin (85% β-actin and 15% γ-actin) derived from human platelets and muscle actin (α-actin) derived from rabbit skeletal muscle (Cytoskeleton Inc.) were mixed in G-buffer [0.2 mM CaCl2, 5 mM Tris–HCl (pH 8.0)] to produce actin stock and were polymerized in an actin polymerization buffer [100 mM KCl, 2 mM MgCl2, 0.5 mM ATP, 0.2 mM Tris–HCl (pH 8.0)] for 1 h at 24°C. Solutions (50 µl) containing polymerized actin (8 µM) were incubated with EFhd1 (12 µM) or EFhd2 (12 µM) for 30 min at 24°C in the presence of 1 mM EGTA, 1 mM CaCl2, or 20 µM ZnCl2. Actin filaments with each protein were pelleted by centrifugation at 100 000g for 2 h at 24°C (for the actin-binding assay). Equal amounts of pellet and supernatant were resolved with SDS–PAGE, and the protein bands were visualized by Coomassie Blue staining. The percentage of each protein in the pellet was quantified with densitometry using ImageJ version 1.53k, and a percentage of pellet histogram was plotted using the OriginPro software (version 9.1; OriginLab Corporation, Northampton, MA, USA; Schneider et al., 2012).
4.11. Negative-staining imaging
Muscle and non-muscle actin (Cytoskeleton Inc.) were polymerized in F-actin buffer containing 100 mM KCl, 2 mM MgCl2, 0.5 mM ATP and 0.2 mM Tris–HCl at pH 8.0. Mixtures (50 µl) of F-actin (4 µM) and full-length EFhd1 (6 µM) or EFhd2 (6 µM) in the presence of 1 mM EGTA, 20 µM CaCl2 or 20 µM ZnCl2 were allowed to react for 1 h. For grid preparation, 2 µl of reaction mixture were loaded onto C-flat holey gold grids (CF-1.2/1.3-4Au-50) and blotted with filter paper to remove excess sample. The sample-loaded grid was then stained in a solution of 1%(w/v) uranyl acetate. The grids were immersed in the stain solution for 20 min, blotted with filter paper to remove excess stain and air-dried. The samples were imaged using an FEI Tecnai G2 F30 S-Twin transmission electron microscope operated at 300 kV.
Supporting information
Supporting figures and table. DOI: https://doi.org/10.1107/S2052252523001501/jt5066sup1.pdf
Acknowledgements
We thank the staff at beamlines BL-5C, 7A and 11C of the Pohang Accelerator Laboratory (Pohang, Republic of Korea) for their kind help with data collection. SHE and SAM planned and designed the experiments. SAM and JYK performed gene cloning and expression. SAM carried out purification, crystallization, in vitro actin co-sedimentation assay and analysis. JP performed data collection, data processing and TP, MJ and JY performed data collection and investigation. SHE and SAM wrote the manuscript. All authors contributed to the article and approved the submitted version. The authors declare that they have no conflict of interest.
crystal titration experiments,Funding information
This work was supported by the National Research Foundation (NRF) of the Korean government (grant no. NRF-2021R1A2C1006267) and the `GIST Research Institute (GRI) IIBR' grant funded by Gwangju Institute of Science and Technology in 2022.
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