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Guardians of the Genome: Iron–Sulfur Proteins in the Nucleus

Lorena Novoa-Aponte

Andres Leon-Torres

and

Caroline C. Philpott

Genetics and Metabolism Section, Liver Diseases Branch, National Institutes of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD 20892, USA

Authors to whom correspondence should be addressed.

Inorganics 2024, 12(12), 316; https://doi.org/10.3390/inorganics12120316

Submission received: 28 October 2024 / Revised: 22 November 2024 / Accepted: 25 November 2024 / Published: 6 December 2024

(This article belongs to the Special Issue Iron-Sulfur Clusters: Assembly and Biological Roles)

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Abstract

Iron–sulfur (Fe-S) clusters are essential cofactors found in many proteins in the mitochondria, cytosol, and nucleus of the cell. These versatile cofactors may undergo reversible oxidation–reduction reactions to enable electron transfers; they may be structural and confer stability to a folded protein; they may be regulatory and transduce an iron signal that alters the function or stability of a recipient protein. Of the nearly 70 proteins described in mammalian cells that bind Fe-S clusters, about half localize exclusively or partially to the nucleus, where they are required for DNA replication and repair, telomere maintenance, transcription, mitosis, and cell cycle control. Most nuclear Fe-S cluster proteins interact with DNA, including DNA polymerases, primase, helicases, and glycosylases. However, the specific roles of the clusters in the enzymatic activities of these proteins and their interplay with DNA remain a matter of debate. Defects in the metallation of nuclear Fe-S proteins cause genome instability and alter the regulation of cell division and proliferation, which are hallmarks of various genetic diseases and cancers. Here, we provide an inventory of the nuclear Fe-S cluster-binding proteins and discuss cluster types, binding sites, the process of cluster acquisition, and the potential roles of the cluster in the function of the proteins. However, many questions remain unresolved. We highlight critical gaps in our understanding of cluster delivery to nuclear client proteins, the potential for cluster repair, and the mechanistic roles that clusters play in these enzymes. Taken together, this review brings the focus to the nucleus of the human cell as a hotspot for Fe-S cluster proteins and aims to inspire new research on the roles of iron in DNA metabolism and the maintenance of genome integrity.

Keywords:

iron–sulfur cluster; nucleus; iron trafficking; DNA replication; DNA repair

Graphical Abstract

1. Introduction

Iron–sulfur (Fe-S) clusters are essential cofactors found in a wide variety of proteins across all domains of life. These clusters consist of iron and sulfur ions with different compositions, structures, redox states, and coordination ligands in proteins. The most frequently found clusters are the rhombic [2Fe-2S] and the cubic or cubane [4Fe-4S], but [3Fe-4S] and [4Fe-3S] clusters have also been described. The chemistry of these clusters makes them ideal for sensing redox signals, as they can accommodate different electronic configurations by accepting or donating unpaired electrons. Consequently, these clusters serve a wide spectrum of functions within proteins, including electron transfer, enzymatic catalysis, protein structural stability, cofactor transfer, oxygen and iron sensing, and redox stress [1]. These functions render Fe-S proteins crucial in multiple cellular processes, including energy production, metabolite transformation, DNA maintenance, gene expression regulation, protein synthesis, mitosis, and antiviral defense [2].

In the early stages of identifying and characterizing Fe-S clusters in the human proteome, the proteins discovered were primarily associated with mitochondrial and cytosolic functions, such as ferredoxins, aconitases, and respiratory complexes [3]. When the first Fe-S cluster bound to a helicase was reported in 2006, the authors stated that “Fe-S clusters are rare in nuclear proteins” [4]. Nearly two decades later, we see that this perspective has completely changed, with the nucleus now known to contain about 35 proteins that bind Fe-S clusters, turning the nuclear compartment into a hotspot for Fe-S-related processes. The inventory of nuclear Fe-S cluster-binding proteins described to date is listed in Table 1.

The link between iron metabolism and nucleic acid metabolism is becoming increasingly clear. This is evident in processes related to genome integrity, with Fe-S clusters found in many proteins involved in DNA replication and repair, as well as in transcription, with Fe-S clusters present in RNA polymerase, transcription factors, and RNA modification proteins. Although the precise role of these Fe-S clusters often remains elusive, their prevalence in proteins associated with nucleic acid transactions and the fact that they have not been replaced by less redox-active metals throughout evolution suggest that they play a crucial role in nucleic acid metabolism.

In this review, we discuss the current knowledge of nuclear Fe-S cluster proteins, examine accumulating evidence suggesting that Fe-S clusters may have additional and unexpected roles in cells, and highlight areas of research that may have practical significance but for which no definitive evidence has yet emerged.

Table 1. The inventory of Fe-S cluster-binding proteins that localize to the nucleus of mammalian cells. The localization of the proteins reported here is based on consolidating information from references (when available) and from three different databases: The Human Protein Atlas [5] (https://www.proteinatlas.org, accessed on 16 October 2024), The UniProt Knowledgebase [6] (https://www.uniprot.org, accessed on 16 October 2024), and COMPARTMENTS [7] (https://compartments.jensenlab.org, accessed on 16 October 2024). An extended version of the table, including the residues that coordinate the clusters, is available as in the Supplementary Materials. The asterisks (*) denote that Fe-S cluster ligands are shared between oligomeric partners from the CIA or ISC pathways. Question marks indicate unknown or debatable cluster-binding residues, cluster types, or stoichiometries. SAM: S-adenosyl-L-methionine.

Participates In	Name	Consolidated Localization	Fe-S Cluster Type
DNA replication/DNA repair	PRIM2	Nucleus	[4Fe-4S] (Cys)₄ [8,9]
	POLA1	Nucleus, Cytosol [10]	[4Fe-4S] (Cys)₄ [11]
	POLD1	Nucleus, Cytosol [12]	[4Fe-4S] (Cys)₄ [11]
	POLE	Nucleus	[4Fe-4S] (Cys)₄ [11,13]
	REV3L	Nucleus, Cytosol [14]	[4Fe-4S] (Cys)₄ [11]
	FANCJ	Nucleus, Cytosol [15,16]	[4Fe-4S] (Cys)₄ [4,17]
	DDX11	Nucleus [18]	[4Fe-4S] (Cys)₄ [19]
	RTEL1	Nucleus [16]	[4Fe-4S] (Cys)₄ [20]
	XPD	Nucleus, Cytosol [16,21]	[4Fe-4S] (Cys)₄ [4]
	DNA2	Nucleus, Mitochondria [22]	[4Fe-4S] (Cys)₄ [23,24]
	EXO5	Nucleus, Cytosol [25]	[4Fe-4S] (Cys)₄ [25]
	MUTYH	Nucleus, Mitochondria [26]	[4Fe-4S] (Cys)₄ [27,28]
	NTHL1	Nucleus, Mitochondria [29]	[4Fe-4S] (Cys)₄ [30,31]
RNA transactions	ELP3	Nucleus, Cytosol [32]	[4Fe-4S] (Cys)₃ SAM [33,34]
	RPC6	Nucleus [35]	[4Fe-4S] (Cys)₄ [36,37]
	CPSF4	Nucleus [38]	[2Fe-2S] (Cys)₃ (His)? [39,40]
	TYW1	Nucleus, Cytosol	[4Fe-4S] (Cys)₃ SAM [34,41]
	TYW1B	Nucleus	[4Fe-4S] (Cys)₃ SAM [34,41]
	CDK5RAP1	Nucleus, Mitochondria, Cytosol [42]	[4Fe-4S] (Cys)₄, [4Fe-4S] (Cys)₃ SAM [34,43]
Mitosis	KIF4A	Nucleus [44]	[4Fe-4S] (Cys)₄? [45]
Mitosis	KIF4B	Nucleus [44]	[4Fe-4S] (Cys)₄? [45]
Iron metabolism	NCOA4	Nucleus, Cytosol [46]	[3Fe-4S] (Cys)₄? [47,48]
Iron metabolism	FBXL5	Nucleus, Perinuclear region [49]	[2Fe-2S] (Cys)₄ [50]
Post-translational modifications	DPH1	Nucleus, Cytosol [51]	[4Fe-4S] (Cys)₃ SAM [52]
	DPH2	Nucleus, Cytosol	[4Fe-4S] (Cys)₃ SAM [52]
	ATE1	Nucleus, Cytosol [53]	[4Fe-4S] (Cys)₄ [54]
Respiration	SDHB	Mitochondria, Nucleus [55]	[2Fe-2S] (Cys)₄, [3Fe-4S] (Cys)₃, [4Fe-4S] (Cys)₄ [55]
Unknown	RFESD	Nucleus	[2Fe-2S] (Cys)₂ (His)₂
CIA	CIAPIN1	Nucleus, Mitochondria, Cytosol [56]	[2Fe-2S] (Cys)₄, [4Fe-4S] (Cys)₄? [57,58]
	BOLA2	Nucleus, Cytosol [59]	[2Fe-2S] * Ligands shared with GLRX3 [60,61]
	GLRX3	Nucleus, Cytosol [62]	[2Fe-2S] * Ligands shared with BOLA2 [60,61]
	NUBP2	Nucleus, Cytosol [63]	[4Fe-4S] * Ligands shared with NUBP1 [64,65]
ISC	NFU1	Nucleus, Mitochondria, Cytosol [66]	[4Fe-4S] * Ligands shared between dimers [66,67]
	NFS1	Nucleus, Mitochondria, Cytosol [68]	[2Fe-2S] * Ligands shared with ISCU [69,70]
	GLRX2	Nucleus, Mitochondria [71]	[2Fe-2S] * Ligands shared between dimers [72]

2. Metallation of Nuclear Fe-S Cluster Proteins

The assembly of Fe-S clusters in mammalian cells involves a complex interplay between two machineries: the mitochondrial Iron–Sulfur Cluster (ISC) pathway and the Cytosolic Iron–Sulfur Assembly (CIA) pathway. This organized, multistep process relies on the functions of multiple proteins located inside and outside the mitochondria and consists of common underlying mechanisms of cluster synthesis, trafficking, and insertion into the final recipient proteins. Cofactor maturation of nuclear [4Fe-4S] proteins depends on the function of both the ISC and CIA pathways, as evidenced by the defects in the metallation of nuclear Fe-S proteins when depleting either ISC or CIA components [11,65,73,74,75,76,77,78,79]. Excellent recent reviews have focused on the known aspects of the de novo assembly of clusters inside and outside the mitochondria [2,80,81]. Although numerous studies on yeast and mammalian cells indicate that mitochondria provide the raw materials for the de novo assembly of clusters, studies on mammalian cells suggest that the de novo assembly of Fe-S clusters also occurs in the cytosol via dual localization of some ISC components in the mitochondria and the cytosol [66,68,82,83,84]. However, the relative contribution of each pathway to the assembly of clusters in mammalian cells remains unclear.

Similarly, the source of the iron used for the assembly of clusters has not been determined. In the cytosol, the iron chaperone poly C-binding protein 1 (PCBP1) binds iron from the kinetically labile, exchangeable iron pool (called the labile iron pool) and delivers it to ferritin and non-heme iron enzymes [85,86,87,88]. The co-chaperone complex composed of PCBP1 and BOLA2, formed via a bridging Fe ligand, serves as an intermediate for the assembly of [2Fe-2S] clusters on the chaperone complex formed by BOLA2 and the monothiol glutaredoxin GLRX3 [89]. In mitochondria, however, the source of iron for cluster assembly is still an open question due to the lack of known iron chaperones residing in the mitochondrial compartment. Although some early studies suggested that frataxin could act as the mitochondrial iron donor for the ISC [90,91,92,93], mounting evidence does not support an iron chaperone function for frataxin, but instead indicates its role as an allosteric activator of the persulfide transfer process during the initial steps of Fe-S cluster assembly [81,94,95]. Consequently, the source of iron for mitochondrial ISC-mediated cluster synthesis remains unknown.

A fully assembled Fe-S cluster may be transferred to a recipient apoprotein that requires the cluster for function. This associative transfer occurs via protein–protein interactions, which protect the labile cofactor from chemical attack during delivery and prevent the cofactor from engaging in harmful redox chemistry. More than one system may be involved in the transfer of clusters to recipient apoproteins. The insertion of Fe-S clusters into apoproteins is facilitated by chaperone proteins such as HSC20 [82,84] or co-chaperone systems of the CIA-targeting complex (CTC), composed of three proteins: CIAO1, MMS19, and FAM96B/CIAO2B [74,77,78,96,97]. A summary of the Fe-S cluster trafficking proteins that have been shown to interact with nuclear Fe-S proteins is included in Supplementary Table S1. To assist the incorporation of Fe-S clusters into target proteins, HSC20 recognizes target proteins containing a short-peptide (LYR) motif [82]. In contrast, the CTC guides the Fe-S clusters from the CIA to client apoproteins containing a conserved [LIM]-[DES]-[WF] C-terminal tripeptide motif, called the CTC recognition motif, which facilitates the identification of about a quarter of the known CTC clients or their adaptors [98]. However, this C-terminal signature is only present in a handful of the known Fe-S cluster-binding proteins (Table S1). It is still unclear how apo-client recognition occurs for the remaining 75% of the Fe-S cluster proteins that do not contain the described CTC recognition motif. Alternative recognition motifs and new adaptor proteins mediating cluster transfer from CTC to apoproteins are likely yet to be discovered.

It is broadly assumed that Fe-S clusters are inserted into client apoproteins during protein synthesis and folding in the cytosol and remain bound during the lifetime of the Fe-S cluster protein. This seems necessary for proteins for which the cluster is buried deeply within the tertiary structure of the protein. For example, xanthine oxidase (XDH) contains two Fe-S clusters buried in the interior of the protein. The free exchange of these clusters seems unlikely due to the steric hindrance inherent in the structure of the folded protein (Figure 1A). However, evidence is lacking that shows that Fe-S metallation steps must occur exclusively during folding in the cytosol and before the nuclear protein is translocated to the nucleus. An additional hypothesis points to the possibility of cluster transfer after full translation and folding of the apoproteins. Two interesting observations may shed light in this regard: the evidence of Fe-S cluster transfer via adaptor-mediated protein–protein interactions [98], and the presence of multiple CIA components in the nucleus of the cell.

First, emerging evidence suggests that a subset of Fe-S proteins require an adaptor protein for their recognition by the CTC [98,99]. This is the case for the DNA primase subunit 2 (PRIM2) with its adaptor PRIM1, the elongator subunit ELP3 with its adaptor ELP4, and the ribosome recycling factor ABCE1 with its adaptor LTO1 [98,99]. These three [4Fe-4S] cluster proteins receive clusters from the CTC but do not contain the C-terminal [LIM]-[DES]-[WF] CTC recognition motif in their sequences. Instead, the CTC recognition motif is present in the C-terminus of the corresponding adaptor proteins [98]. This suggests that the transfer of the metalloclusters through adaptor-mediated protein–protein interactions should occur once the proteins have been synthesized and folded and the client–adaptor complexes have been formed. Supporting this idea, the PRIM2/PRIM1 complex appears to form prior to Fe-S cluster incorporation into PRIM2 [100]. Both PRIM2 and PRIM1, while in complex, directly interact with the CIA-targeting complex for cluster transfer [101]. This may only be possible for proteins harboring readily accessible, solvent-exposed cluster-binding sites. Figure 1B–D show that the three proteins, ABCE1, PRIM2, and ELP3, that use an adaptor protein for CTC recruitment have, in fact, solvent-exposed clusters. Thus, it is probable that the CTC donates cofactors to fully or partially folded proteins with accessible cluster-binding sites, while proteins with buried clusters may utilize a co-translational system that delivers clusters during protein folding. Further studies are needed to clarify how this parallel process occurs.

Figure 1. A comparison of solvent-exposed and buried Fe-S clusters in proteins. The structures (left) and close-ups of the Fe-S binding domains (square, right). (A) X-ray structure of human xanthine oxidase (XDH) showing the two [2Fe-2S] clusters deeply buried within the protein structure (PDB 2CKJ (https://www.rcsb.org/structure/2CKJ, accessed on 30 September 2024)). (B) Cryo-EM structure of the human ribosome recycling factor ABCE1 showing the two [4Fe-4S] clusters situated in solvent-exposed loops at the N-terminal domain (PDB 7A09 (https://www.rcsb.org/structure/7A09, accessed on 1 October 2024)). The clusters in ABCE1 are prone to degradation [102]. Each cluster in ABCE1 is coordinated by three nearby Cys residues and one distant Cys residue in the primary sequence, suggesting a structural role for these clusters [103]. (C) Cryo-EM structure of human DNA primase subunits, PRIM1 (green) and PRIM2 (blue) (PDB 8QJ7 (https://www.rcsb.org/structure/8QJ7, accessed on 2 October 2024)). The [4Fe-4S] cluster bound to PRIM2 locates at the junction of the N-terminal and C-terminal domains close to the protein surface. PRIM1 is proposed to serve as an adaptor for the recognition of PRIM2 by the CTC to facilitate its metallation. (D) Cryo-EM structure of the yeast elongator complex proteins ELP3 (green) and ELP4 (yellow) (PDB 8ASV (https://www.rcsb.org/structure/8ASV, accessed on 2 October 2024)). The radical SAM enzyme ELP3 has a [4Fe-4S](Cys)₃ cluster, which is exposed to solvent via two channels [103]. ELP4 is proposed to serve as an adaptor for the recognition of ELP3 by the CTC to facilitate its metallation.

Although solvent-exposed clusters are inherently more susceptible to oxidative damage [104,105,106,107] and may require replacement or repair to maintain their function, proteins with exposed clusters may also exchange cofactors more readily. This lability could confer to these proteins the capacity to sense and respond to environmental cues. For example, the variable metallation status of the iron-responsive protein 1 (IRP1/ACO1) [108], the nuclear receptor coactivator 4 (NCOA4) [109], or the F-box and leucine-rich repeat protein 5 (FBXL5) [50,110] determines the biological activity of these three proteins. They are central to the regulation of intracellular iron metabolism, and Fe-S cluster binding allows for coordination of the intracellular iron supply with their cellular activities. These three proteins also harbor solvent-exposed clusters. However, it remains to be demonstrated whether other Fe-S proteins, including nuclear Fe-S enzymes with solvent-exposed cluster-binding sites, may exist as a pool of fully folded proteins that incorporate the cluster only when required to respond to cellular needs. Interestingly, although most of the proteins involved in the assembly and delivery of clusters are present in the cytosol, the nucleus of the cell also houses many of them including MMS19 [21,111], CIAPIN1 [56], CIAO1 [111], CIAO2B [21], GLRX3 [62], NFU1 [66], NFS1 [68], NUBP2 [63], GLRX2 [71], and PCBP1 [5]. Whether these proteins assist in the in situ transfer of iron or Fe-S clusters to client proteins in the nucleus remains elusive.

3. Fe-S Cluster Proteins Involved in Nuclear DNA Transactions

Defects in Fe-S cluster biogenesis result in DNA damage and genome instability. This is due to the requirement of Fe-S clusters for the function of multiple enzymes involved in DNA transactions [76,77,78,112,113,114,115]. In fact, the fastest growing category of Fe-S binding proteins consists of enzymes that participate in nucleic acid metabolism, including DNA polymerases, primases, glycosylases, helicases, nucleases, transcription factors, RNA polymerases, and RNA-modifying enzymes.

3.1. Fe-S Clusters and DNA Replication

DNA replication in eukaryotic cells relies entirely on several Fe-S cluster-binding proteins, whose function and cluster-binding properties are summarized below. Fe-S clusters may link the initiation of DNA replication to intracellular iron availability through licensing factors that ensure replication occurs only when the cell’s iron levels are sufficient to meet the high demand for this essential cofactor during DNA synthesis.

3.1.1. DNA Polymerases

The human genome encodes at least 14 DNA-dependent DNA polymerases [116]. Among these, DNA polymerases Pol α, δ, and ε, all located in the nucleus, are considered the replicative DNA polymerases. These multi-subunit complexes are classified as class B DNA polymerases and play crucial roles in precise nuclear DNA duplication [116]. Pol α initiates DNA replication at origins by synthesizing an RNA-DNA hybrid primer of 20–30 nucleotides. This primer is further extended by the highly processive Pol δ and ε on the lagging and leading strands of DNA, respectively [116] (Figure 2A). The catalytic subunits of all eukaryotic replicative DNA polymerases (POLA1, POLD1, and POLE) harbor a [4Fe-4S] cluster that is essential for replisome stability [11,117,118]. The fourth and final member of the class B DNA polymerase family, the error-prone DNA polymerase Pol ζ (zeta) [116], also binds a [4Fe-4S] cluster using a Cys-rich motif at the C-terminal domain of its catalytic subunit, REV3L [11]. Mutations that prevent cluster binding to Pol ζ abrogate its activity in UV mutagenesis repair [119]. Fe-S cluster binding has not been described for other classes of DNA polymerases [11]. Interestingly, although all class B DNA polymerases bind the cubane [4Fe-4S] cluster via a classical 4 Cys arrangement, the site of cluster binding varies. The cysteine motifs CysA and CysB, located at the C-terminus, are conserved among the four members of the family. CysB serves as the cluster-binding site for Pol δ and Pol ζ [119]. In contrast, the catalytic domain of Pol ε binds the [4Fe-4S] cluster at an additional Cys-rich motif in the N-terminus, known as CysX, which is unique to Pol ε (C651, C654, C663, and C747) [13,117]. This positions the cluster in an exposed area, away from the DNA template, at the base of the processivity domain (P-domain), near the junction of the P and Palm domains [11,13,117,118]. A mutation in the cluster-binding site in Pol ε abolishes DNA binding and proofreading exonuclease activities [120].

The metallation of all class B DNA polymerases depends on the function of the CIA pathway. Mutations in the CIA machinery lead to the loss of clusters in DNA polymerases, resulting in reduced protein stability and activity that causes genome instability [11,74,77,78]. Although the precise roles of Fe-S clusters in these proteins remain enigmatic, increasing evidence suggests that they may function as redox switches controlling the speed of DNA synthesis. Faster synthesis is associated with the reduced form of the [4Fe-4S] cluster present in Pol δ. This may provide a rapid and reversible mechanism to respond to replication stress by slowing down DNA polymerase activity through cluster oxidation, allowing for the accumulation and activation of DNA repair factors at lesion sites. Once DNA is repaired, reductions in the polymerase cluster may enable replication to resume [121]. This model needs further validation.

The presence of the cluster in Pol α is somewhat controversial because structural data show Zn²⁺ bound at the position of the Fe-S cluster, which suggests that the presence of a cluster in Pol α might prevent its interaction with the corresponding B subunits [119,122,123,124,125,126]. Although no structures of Pol α showing a bound [4Fe-4S] cluster are currently available in the Protein Data Bank (PDB), it is not uncommon to find zinc replacing bona fide Fe-S clusters in protein structures due to the intrinsic sensitivity of these clusters to oxidative damage under aerobic conditions [39,117,127,128,129,130,131]. Resolving these discrepancies is essential to bridging a critical gap in our understanding of the role of Fe-S clusters in DNA polymerases.

3.1.2. DNA Primase

DNA replication begins with the synthesis of a short RNA primer of 8–12 nucleotides catalyzed by DNA primase. The primer is then extended by the catalytic subunit of Pol α [116] (Figure 2A). The primosome consists of four subunits: the catalytic subunit POLA1, the accessory subunit POLA2, and two primase subunits, the small catalytic subunit PRIM1 and the large regulatory subunit PRIM2. Each is located in the nucleoplasm [5], where DNA replication occurs. In addition to the cluster in POLA1, the primase complex contains an additional [4Fe-4S] cluster bound to the C-terminal domain of PRIM2, which is essential for primer synthesis [8,9,132]. None of the smaller non-catalytic subunits of Pol δ or Pol ε harbor Fe-S clusters [11]. PRIM2, which is present only in archaeal and eukaryotic primases [132,133,134], bridges the two catalytic domains of the primosome–POLA1 and PRIM1—thereby facilitating the transition from RNA primer synthesis to DNA elongation [132].

The metallation of PRIM2 relies on the CIA machinery [16,77,97,132]. It has been proposed that PRIM1 may act as an adaptor protein between PRIM2 and the CIA-targeting complex (Figure 1C), aiding in the client protein recognition necessary for cluster transfer [98,101]. The [4Fe-4S] cluster in PRIM2 is bound via a canonical four-Cys motif (C287, C367, C384, C424), located at the junction of the N-terminal and C-terminal domains, with each domain contributing two binding Cys residues [8,9,122,133,134,135] (Figure 1C). This arrangement suggests that the Fe-S cluster serves a structural rather than a redox-active role in the protein. However, one might question why such a complex cofactor would be used solely to preserve the structural integrity of a small domain. Recent evidence indicates that the [4Fe-4S] cluster in DNA primase functions as a redox switch, utilizing DNA charge transport to regulate the protein’s DNA binding activity [136,137]. Briefly, oxidized [4Fe-4S]³⁺ PRIM2 binds more tightly to DNA than its reduced counterpart [136,137]. Electrostatic analyses revealed that the oxidation/reduction of the cluster, along with the accumulation of negative charges on the newly synthesized primers, induces structural changes at the DNA–protein interface [138]. The redox switch model proposes that the redox state of the [4Fe-4S] cluster in DNA primase acts as a reversible switch for DNA binding. The oxidized cluster activates the primase, enhancing its binding to DNA to initiate primer synthesis. As the primase extends the RNA primer, Pol α interacts with the RNA/DNA primer template, and its [4Fe-4S] facilitates DNA-mediated redox signaling for primer truncation. This promotes PRIM2 dissociation by reducing its cluster, which allows for the handoff of the primed template to Pol α [136,137,139]. This is a model. It requires additional study and further research is needed to determine whether a DNA-mediated electron transfer relay coordinates the association, transfer, and dissociation steps of replication.

3.1.3. Fe-S Clusters and DNA Replication Origins

Nuclear receptor coactivator 4 (NCOA4) is the protein responsible for targeting ferritin for degradation during iron starvation, reintroducing iron into the labile iron pool. Under iron-repleted conditions, iron-bound NCOA4 is targeted for proteasomal degradation by the ubiquitin ligase HERC2, which maintains ferritinophagy at basal levels [140,141]. It was recently shown that NCOA4 binds a 3Fe-4S cluster [48]. The insertion of the Fe-S cluster in NCOA4 promotes its recognition by HERC2, triggering its degradation and inhibiting ferritinophagy [47]. How NCOA4 acquires the Fe-S cluster is unknown. In addition to its role in ferritinophagy, NCOA4 is a multifunctional protein also implicated in coordination between cellular iron availability and the DNA replication machinery [109]. Federico et al. demonstrated that under low-iron conditions, apo-NCOA4 binds to DNA replication origins, inhibiting their activation and preventing replication stress [109]. NCOA4 specifically interacts with MCM7, a subunit of the MCM2-7 helicase complex that unwinds DNA during replication. The binding of NCOA4 to the MCM complex hinders its helicase activity [46] (Figure 2B). This mechanism may allow mammalian cells to respond to low iron by delaying the initiation of DNA replication, an adaptive response when the pool of metallated Fe-S enzymes is limited. By finely tuning DNA replication to fluctuations in intracellular iron levels, this mechanism may help to safeguard genome stability during periods of diminished bioavailable iron and replicative stress [109].

3.1.4. Fe-S Clusters in Helicase Activity

Helicases are motor proteins that move along linear nucleic acid tracks using the energy from the hydrolysis of a nucleoside triphosphate. They function by destabilizing the hydrogen bonds between the complementary base pairs of unwinding DNA or RNA strands to generate the single-stranded intermediates required for processes like replication, repair, recombination, transcription, and splicing. Helicases are classified into six superfamilies, SF1-6 [142]. Of the about 24 DNA helicases present in human cells, only 5 are known to bind Fe-S clusters. Those belong to the two largest superfamilies of DNA helicases (SF1 and SF2) [143]. Yet not all members of those families bind a cluster. Four nuclear DNA helicases from the SF2 family (Rad3/XPD group) contain Fe–S clusters in mammalian cells: Fanconi Anemia (FA) protein J FANCJ, the Warsaw Breakage Syndrome protein ChlR1/DDX11, the telomere maintenance helicase RTEL1, and the component of the TFIIH complex involved in transcription and nucleotide excision repair XPD/ERCC2 [4,19,20,77,78,143]. XPD, the founding member of the Fe-S helicase family, is a central component of the general transcription factor TFIIH which plays major roles in transcription and nucleotide excision repair (NER), as discussed below. FANCJ participates in the homologous recombination pathway of DNA double-strand break repair, and it is part of the FA pathway proposed to function downstream of FANCD2. FANCJ also interacts with the breast cancer type 1 susceptibility protein BRCA1 and contributes to its DNA repair function. DDX11 has roles in the replication of DNA, establishment of sister chromatid cohesion, and maintenance of chromosome architecture. RTEL1 is essential for preserving telomere length, as discussed below. The SF1 helicase/nuclease DNA2 is the fifth helicase known to bind an Fe-S cluster in mammalian cells [23,143,144]. Although initially reported to be exclusively in the mitochondria [145], DNA2 also localizes to the nucleus [22]. DNA2 participates in the resolution of stalled replication forks during DNA synthesis [24] (Figure 2A). All of these Fe-S helicases have been linked to multiple cancers and hereditary disorders characterized by genome instability [4,21,146,147,148,149,150], and mutations resulting in loss of the cluster are associated with the onset of disease. For example, mutations adjacent to the Fe-S cluster-binding Cys residues (M299I and A349P) in FANCJ are linked to breast cancer and Fanconi anemia, respectively [15,151,152,153].

The four SF2 helicases bind a single cubane [4Fe-4S] cluster using a canonical 4Cys arrangement (Table 1) located within the helicase core at the N-terminus of the proteins [4,17]. The cluster binding to DNA2 is unusual, as one of the four Cys residues coordinating the cubane [4Fe-4S] is located a few hundred amino acids upstream of the other three (C136, C393, C396, C402) [24]. Structurally speaking, there are two different types of helicases: those that form ring-like hexameric structures and those that do not. None of the known Fe-S cluster-binding helicases form hexameric ring-like structures.

All known eukaryotic Fe-S cluster-containing helicases have a 5′-3′ polarity and unwind DNA substrates. Although the clusters are essential for the DNA unwinding activity of the SF2 helicases [17,146,153], they do not affect the DNA binding and ATP hydrolysis capacities [153]. In contrast, the loss of the cluster in DNA2 not only impairs DNA binding, but also impairs nuclease and ATPase activities [24]. The Fe-S helicases exhibit an apparent specialization for the resolution of non-canonical DNA structures, including G4-quadruplexes (G4s), D-loops, and R-loops [154]. However, not all G4-resolving helicases have been reported to bind an Fe-S cluster, such as the helicases BLM and WRN [146]. Non-canonical DNA structures play roles in genome organization, gene regulation, replication initiation and progression, recombination, and telomere metabolism [155,156]. However, unscheduled accumulation of these secondary DNA structures stalls replication forks, causing genome instability. FANCJ efficiently resolves G4 DNA structures to enable smooth DNA synthesis during replication [157,158,159] (Figure 2A). RTEL1 resolves R-loops and G4s [160,161,162] and binds to the rG4 formed by the Telomeric Repeat-Containing RNA (TERRA) for the maintenance of TERRA-containing R-loops, enhancing telomere stability [163]. DDX11 also resolves G4 DNA structures [164] to support sister chromatid cohesion [165]. And finally, G4 unwinding activity is also sustained by the archaeal XPD helicase [166]. Although it is clear, at least for FANCJ, that the presence of the Fe-S cluster is essential for this G4 unwinding activity [153], the mechanism by which the Fe-S cluster facilitates the unwinding of non-B DNA structures catalyzed by these helicases is still poorly understood.

The metallation of the helicases likely depends on the CIA machinery, as suggested by the interaction of MMS19 with XPD, FANCJ, DDX11, RTEL1, and DNA2 [16,21,77,78,100,111,167,168] and the decreased protein levels of these helicases when MMS19 is depleted [16,77,78]. The interaction of HSC20 with XPD and DDX11 has also been confirmed [84]. However, several questions remain regarding cluster insertion on those enzymes. For example, when the interaction of these helicases with other components of the CIA-targeting complex was examined, XPD, DNA2, and RTEL1 were found to interact with CIAO1 and CIAO2B [97]; however, FANCJ did not interact with CIAO1 [16]. CIAO1 is the CTC component that offers the docking site for CIA client proteins to interact with the CIA delivery system. In general, it is not known how these Fe-S proteins are recognized for cluster delivery as they all lack the C-terminal CTC recognition motif proposed to facilitate the recognition of CIA clients. Further investigation is required to identify the factors that mediate the delivery of clusters to all of the nuclear DNA helicases.

Surprisingly, Fe-S cluster binding in RNA helicases has not been described in mammalian cells. The only example of an RNA helicase binding an Fe-S cluster is the viral 5′-3′ SF1 helicase nsp13, the metallation of which depends on the host Fe-S cluster assembly machinery [128]. However, helicases that are currently annotated as Zn-binding proteins may in fact bind Fe-S clusters.

3.1.5. Fe-S Clusters and Telomere Maintenance

Telomeres, the protective caps at the ends of chromosomes that prevent the degradation and fusion of chromosomes, also rely on functional Fe-S cluster enzymes for their maintenance. The replication of telomeres is mechanistically challenging due to their repetitive nature and the presence of non-canonical DNA structures, such as G4s. Ultimately, this results in the shortening of telomeres during cell division. Key among the proteins involved in telomere maintenance is the ATP-dependent DNA helicase RTEL1, which unwinds G-quadruplex structures at telomeric DNA to facilitate telomere replication, preventing telomere loss [169] (Figure 2C). RTEL1 is located in the nucleoplasm and in nuclear speckles [5] and binds a redox-active cubane [4Fe-4S] cluster that is essential for its helicase activity. The cluster is bound at the N-terminus via a four-Cys arrangement (C145, C163, C172, C207) [20]. Although the role of the cluster in RTEL1 activity has not been determined, the physical interaction of RTEL1 with MMS19 [77,78] and the altered telomere length observed in MMS19 mutants [170] confirm that cluster insertion in RTEL1 depends on the CIA machinery.

3.2. Fe-S Clusters and DNA Repair

An estimate of 50,000 DNA-damaging events occur in our cells per day [171]. These lesions block replication fork progression, cause cell cycle arrest, and constitute a major source for genome instability. At least six major DNA repair systems are required to remove the distinct types of DNA damage: base excision repair (BER), nucleotide excision repair (NER), mismatch repair (MMR), homologous recombination (HR), non-homologous end joining (NHEJ), and translesion DNA synthesis (TLS) [172]. Fe-S clusters are integral components of the molecular machinery that repairs DNA to ensure that repairs DNA, thereby ensuring the precise (Figure 3). In addition to the Fe-S DNA polymerases and helicases discussed above, which are central to all DNA repair mechanisms [172,173], two Fe-S cluster DNA glycosylases, MUTYH and NTHL1, play central roles in the repair of oxidative DNA damage [174,175,176]. These enzymes cleave the glycosidic bond between bases and sugars to remove damaged or mispaired bases from the DNA duplex and act as initiators of BER [177]. MUTYH is specific for removing mismatched adenines and 8-dihydro-8-oxodeoxyguanine (8-oxoG) [178], and NTHL1 removes oxidatively damaged pyrimidines in DNA [179]. They bind a redox-active cubane [4Fe-4S] cluster in the catalytic domain using a four-cysteinyl motif located in a solvent-exposed loop, referred to as the Fe-S cluster loop (FCL) motif (Table 1) [28,180,181,182,183]. Although originally assumed to play purely structural roles, the Fe-S clusters in DNA glycosylases are required for DNA binding and affect the detection and removal of the damaged bases [183,184,185]. Electrochemical studies using purified proteins indicate that cluster oxidation in the bacterial DNA glycosylases EndoIII and MutY promotes an increase in binding affinity [186,187]. The hypothesis is that the cluster-binding motif is positioned for interaction with the DNA backbone [27,182,188] and the redox potential of the Fe-S clusters may aid glycosylases in scanning the DNA duplex for damaged bases [184]. Finally, although the metallation of these nuclear DNA glycosylases relies on the CIA machinery [111], they lack the C-terminal motif believed to guide the CIA-targeting complex to client apo-proteins.

3.3. Fe-S Clusters, Transcription, and Nuclear RNA Transactions

Fe-S proteins are also involved in RNA processing pathways by modulating the activity of transcription factors, RNA polymerases, and RNA-modifying enzymes. These proteins can be found as subunits of the RNA polymerase complex III, the transcription factor IIH, the elongator complex, and the pre-mRNA processing complex in mammalian cells. Although Fe-S binding transcriptional regulators that sense intracellular iron levels have not been described in mammalian cells, examples from yeast will be reviewed below to illustrate the transfer of Fe-S clusters between regulators in the nucleus of eukaryotic cells.

3.3.1. Fe-S Cluster-Binding Fe-Dependent Transcriptional Regulators

Fe-dependent transcriptional regulators that bind Fe-S clusters and thereby sense intracellular iron levels have been described in yeast species but are not yet found in mammalian cells. In this discussion, we focus on fungal transcriptional regulators that sense intracellular iron levels, as their metallation appears to occur in the nucleus, suggesting the presence of dynamic Fe-S trafficking within the nuclear space. Although the iron regulatory mechanisms in the nonpathogenic fungi Saccharomyces cerevisiae and Schizosaccharomyces pombe are fundamentally distinct, both rely on Fe-S clusters as signals to regulate iron acquisition, storage, and utilization within the cell. In S. cerevisiae, the Fe-S cluster-binding transcriptional activators Aft1 and Aft2 regulate the expression of genes involved in iron uptake and use. Under iron-sufficient conditions, a heterodimeric [2Fe-2S] chaperone complex composed of a monothiol glutaredoxin (Grx3/4) and a BolA-like protein (Bol2) binds and delivers the cluster to the DNA-bound transcriptional activators, altering their oligomeric state and leading to their dissociation from DNA, thereby suppressing the transcription of iron uptake genes [189,190,191]. In S. pombe, the transcriptional repressors Fep1 and Php4 regulate the expression of iron uptake and iron utilization genes, respectively. Under iron-depleted conditions, Php4 accumulates in the nucleus and associates with the DNA-binding complex Php2/3/5 to suppress the expression of iron utilization genes. As iron levels increase, a [2Fe-2S] cluster-dependent interaction between Php4 and Grx4 promotes the dissociation of Php4 from the Php2/3/5 complex, leading to the de-repression of iron-usage gene expression [192,193,194]. In contrast, the transcriptional repressor Fep1 binds a [2Fe-2S] cluster under iron-sufficient conditions, which facilitates its binding to DNA and halts the expression of iron uptake genes [195,196]. Under iron-depleted conditions, the cluster is unidirectionally transferred from Fep1 to Grx4-Fra2, causing the apo form of Fep1 to detach from DNA, allowing for the expression of iron uptake genes [195]. The mechanism by which Fep1 acquires the cluster remains unclear. Notably, Fep1 localizes to the nucleus under both iron-repleted and iron-deficient conditions [197] and remains bound to DNA when iron is sufficient. This suggests that the transfer of the cluster between Fep1 and the Grx4-Fra2 chaperone system occurs within the nucleus.

The [2Fe-2S]-coordinating complex formed by the glutaredoxin protein Glrx3 and the BolA-like protein BolA2 is also present in the cytosol of mammalian cells, where it participates in the maturation of cytosolic Fe-S cluster proteins [61]. Although mammals, unlike fungi, do not have iron-responsive transcriptional regulators such as Aft1/2, Fep1, or Php4, it is tempting to infer from the fungal evidence that the Glrx3-[2Fe-2S]-BolA2 system may constitute a pool of bioavailable, exchangeable clusters that are transferred to client partners for signaling purposes. However, these proposed roles remain to be experimentally tested.

3.3.2. RNA Polymerase III

Pol III is a DNA-dependent RNA polymerase specialized for the transcription of tRNAs and other short, essential RNAs. It is responsible for transcribing approximately 15% of total cellular RNA, including tRNA, 5S ribosomal RNA, U6 spliceosomal RNA, and other small ubiquitous RNAs. The dysregulation of human Pol III is associated with tumor transformation, neurodegenerative and developmental disorders, and increased susceptibility to viral infections [198,199]. Pol III is a multi-subunit complex composed of seventeen subunits, with a ten-subunit catalytic core and a peripheral heterodimeric stalk [35,200]. RPC6 (POLR3F), the subunit that binds a cubane [4Fe-4S] cluster, is part of the peripheral heterotrimeric complex RPC3/RPC6/RPC7 that modulates Pol III dynamics during transcription initiation [36,37,201]. The Fe-S cluster binds to the C-terminus of RPC6 via the thiol groups of four Cys residues (C287, C290, C296, C307). The Fe-S cluster is sandwiched between RPC3, RPC7, and the RPC1 clamp, tethering the interactions between the catalytic core and the RPC3-RPC6-RPC7 heterotrimer, thereby stabilizing Pol III [37]. Interestingly, although the Fe-S binding residues of RPC6 are widely conserved in many organisms, they are missing in Saccharomyces cerevisiae [199]. Distinct from other RNA polymerases, Pol III exhibits functional activities in both the nucleus and the cytoplasm. The canonical function of Pol III involves transcribing tRNAs, 5S rRNAs, and other small RNAs within the nucleus. Interestingly, cytoplasmic Pol III plays a vital role as a viral DNA sensor, synthesizing RNA from AT-rich viral DNA templates and triggering innate immune responses [202,203]. The current model suggests that Pol III is assembled in the cytosol, with some of it translocated into the nucleus and some remaining in the cytosol to participate in viral DNA sensing and response [203]. However, the actual factors facilitating Fe-S cluster insertion in RPC6 remain to be elucidated.

3.3.3. Transcription Factor IIH

The general transcription factor TFIIH participates in transcription initiation and nucleotide excision repair [204]. It is required for promoter opening and the transition from the pre-initiation complex to active transcription. TFIIH is a protein complex composed of ten subunits divided into two subcomplexes: the core (XPB, p62, p52, p44, p34, p8) and the CDK-activating kinase (CDK7, cyclin H, MAT1) [205]. These subcomplexes are bridged by the helicase XPD, which connects p44 and MAT1. XPD (ERCC2) is a mostly nuclear helicase [5] that binds a [4Fe-4S] cluster via 4 Cys (C116, C134, C155, C190) in its N-terminal domain [4]. Mutations in XPD affect the interaction between XPD and p44, resulting in the dissociation of XPD from the TFIIH complex. This disruption results in defects in the phosphorylation of various transcriptional activators, causing aberrant transcriptional activation [206,207,208].

3.3.4. RNA-Modifying Proteins

In addition to capping and splicing, pre-mRNA undergoes 3’ end maturation before protein translation. The cleavage and polyadenylation specific factor complex (CPSF) plays a crucial role in this pre-mRNA processing step by adding a poly-A tail to the mRNA. CPSF consists of six different proteins that recognize the highly conserved polyadenylation signal in the pre-mRNA, cleave the 3’ end, and recruit additional polyadenylation factors. CPSF4, one of the subunits of the complex, binds one [2Fe-2S] cluster and four Zn²⁺ ions as cofactors. In vitro analyses have shown that both metal cofactors are necessary for recognizing the polyadenylation signal in pre-mRNA [39]. CPSF4 contains five CCCH domains which are known to bind Zn and Fe-S clusters. The binding site for the [2Fe-2S] cluster in CPSF4 exhibits flexibility due to redundant iron-binding sites, allowing the cluster to be bound to an alternative site when the preferred site is mutated [39,127]. It is currently unknown which CCCH domain of CPSF4 is the preferred binding site for the [2Fe-2S] cluster in vivo. It is of interest to determine whether such flexibility in cluster-binding sites has a meaningful functional effect on the biological role of the protein. Notably, CPSF4 is the only protein exclusively localized in the nucleus known to bind a [2Fe-2S] cluster. Although a physical interaction between CPSF4 and the Fe-S chaperone HSC20 has been described [84], the mechanism by which CPSF4 acquires its cluster is not well established.

The multifunctional elongator complex facilitates transcription by RNA Pol II [32], but is also responsible for the post-transcriptional modification of tRNAs at the wobble uridine (U₃₄), a crucial modification that enhances translation efficiency [34]. It is composed of two copies of each of its six individual protein subunits, ELP1-ELP6, that form two discrete subcomplexes known as ELP123 and ELP456 [209]. ELP3 is the catalytic subunit of the complex with tRNA acetyltransferase activity, which is necessary for multiple tRNA modifications, including the formation of 5-methoxycarbonylmethyl uridine (mcm5U) and its derivates [34]. ELP3 harbors a cubane [4Fe-4S] cluster bound to its N-terminus via three cysteines (C99, C109, C112) and an exchangeable S-adenosyl-L-methionine (SAM) (Figure 1D) [33,209,210,211]. Interestingly, ELP3 is the only elongator subunit that has orthologs present in eukaryotes, archaea, bacteria, and viruses [210]. ELP3 interacts with MMS19 [78,111], and a deficiency in ELP3, and other Fe-S proteins, was observed in cell lines derived from patients with biallelic loss of function in CIAO1 [212], confirming that ELP3 acquires its cluster from the CIA complex. As discussed above, ELP4 may serve as an adaptor protein between CTC and ELP3 (Figure 1D), exposing the C-terminal tryptophan motif recognized by the CIA-targeting complex for the delivery of [4Fe-4S] clusters to client proteins [98]. However, demonstration of the involvement of other CIA factors and ELP4 in facilitating the metallation of ELP3 is still lacking.

Humans have three other tRNA-modifying enzymes in the nucleus that bind Fe-S clusters. The SAM-dependent tRNA 4-demethylwyosine synthases, TYW1 and TYW1B, are involved in the wybutosine biosynthesis pathway and are responsible for forming imidazoline rings. This RNA modification is important for stabilizing the codon–anticodon interaction, maintaining the reading frame and preventing errors during translation [34,213]. Information reported in databases suggests that TYW1 localizes to the nucleus (reason for inclusion in Table 1). However, a recent publication shows that TYW1 mainly localizes in the cytosol in human cells [214], which is the compartment where the type of tRNA modification catalyzed by this enzyme is assumed to occur [215]. The third protein, CDK5RAP1, is a methylthiotransferase responsible for modifying four mitochondrial DNA-encoded tRNAs to optimize mitochondrial translation [216]. CDK5RAP1 is primarily located in the mitochondria but can also be found in the nucleoplasm and cytosol. Its RNA modification activity is not limited to mitochondrial tRNA, as there is evidence that it also introduces modification into nuclear RNA species [42]. These three proteins bind cubane [4Fe-4S] clusters via three cysteines and an exchangeable SAM (Table 1).

4. Fe-S Cluster Proteins in Mitosis

Once DNA is fully replicated, cells enter mitosis following an intricate and highly regulated process that ensures each new cell receives an exact copy of the genetic blueprint. It has long been established that MMS19 and CIA2B localize to the mitotic spindle during mitosis and that their knockdown disrupts the mitotic pathway, leading to poor alignment of chromosomes, improper segregation, the accumulation of nuclei with abnormal shapes, and the mislocalization of key mitotic factors [21]. However, it was only a few years ago that a mitotic factor was identified as binding an Fe-S cluster. The chromokinesin KIF4A, which is involved in precise chromosome segregation during mitosis, binds an Fe-S cluster [45] (Figure 2D). Molecular motor kinesins are the primary source of cellular forces and drive many types of intracellular movement along microtubule networks. KIF4A and KIF4B are the two variants of the kinesin family 4 present in humans. They share 98% homology in their DNA sequence with different chromosomal locations [217]. This ATP-dependent microtubule-based motor kinesin resides in the nucleoplasm during interphase and on the arms of condensed chromosomes during mitosis. KIF4A accumulates at the central spindle that connects the two daughter cells during the late stages of mitosis, from late anaphase through cytokinesis [44,218,219]. This stabilizes the bipolar mitotic spindle, which is essential for successful midzone formation and cytokinesis. In KIF4-deficient cells, the central spindle becomes disorganized, leading to cytokinesis failure [220,221], chromosome misalignment, spindle defects, and chromosome mis-segregation. This results in shorter and wider chromosomes, lagging chromosomes, aneuploidy, and ultimately tumor formation [219,222].

Unlike other motor kinesins, KIF4A resides in the nucleus during interphase where it performs additional non-mitotic functions. KIF4A has been proposed to play a role in the DNA damage response by modulating the BRCA2/Rad51 pathway. Its C-terminal cargo-binding domain interacts with the conserved C-terminal region of BRCA2, facilitating the translocation of BRCA2 to damaged DNA regions following injury [223]. Depletion of KIF4A results in significant slowing down of DNA replication and decreased homologous recombination [223]. KIF4 also acts as a modulator of large-scale chromatin architecture. It binds globally to chromatin and its depletion leads to chromatin decondensation and a loss of heterochromatin domains [222]. Overall, KIF4 deficiency has been associated with developmental delays and intellectual disabilities [224] and it is abnormally expressed in multiple cancers [217]. KIF4A has also been implicated in the infection process of hepatitis B and D viruses by regulating the transport of the NTCP receptor from the cytoplasm to the cell surface, where it functions as a receptor for viral entry [217,225].

Structurally speaking, KIF4A features a highly conserved ATPase/motor domain at the N-terminus that binds to microtubules and provides the mechanochemical force for movement. This is followed by a central α-helical stalk domain and a C-terminal cargo-docking domain responsible for capturing cargoes [226,227]. The [4Fe-4S] cluster in KIF4A is bound to a conserved cysteine-rich domain located at the C-terminus [45]. However, it remains unclear which of the nine conserved Cys residues located in that region coordinate the Fe-S cluster. Whether the cluster is redox-active is also unknown.

In mitotic cells, KIF4A colocalizes with the CIA components CIAO2B and MMS19 at the spindle midzone and midbody between separating chromosomes [45] (Figure 2D). KIF4A also co-immunoprecipitates with CIAO2B and MMS19, confirming their physical interaction [45]. This interaction is enhanced after iron chelation, suggesting a more stable association between the CIA complex and the apo-client protein [45]. Similar enhancements in interactions between CIA and client proteins under iron-deprived conditions have been observed for other Fe-S proteins, including XPD, FANCJ, and Pol δ [78]. The mislocalization of the KIF4A variant lacking the conserved C-terminal cysteine domain that binds the cluster, along with the reduced association of endogenous KIF4A with the midzone/midbody following CIAO3 depletion, suggests that the Fe-S cluster in KIF4A modulates the proper subcellular localization of the protein at the mitotic spindle. This likely influences the protein–protein interactions essential for the mitotic process [45].

Although the cluster in KIF4A is quite labile in Fe radiolabeling experiments [55], its absence does not appear to affect the stability of the protein [45], as seen for other Fe-S cluster-binding proteins. This dynamic nature may be biologically relevant and allow for the assembly and disassembly of the Fe-S cluster in KIF4A to regulate the progression of mitosis [45], which could explain why the late-acting CIA factors colocalize at the midbody with components of the mitotic machinery. This suggests that the cluster is transferred to a folded client protein at the site of function, rather than co-translationally transferred. Further mechanistic studies are needed to elucidate the role of the Fe-S cluster in modulating KIF4A activity and whether the metallation of KIF4A occurs only after the breakdown of the nuclear membrane when cells enter mitosis, or whether KIF4A residing in the nucleus during interphase also binds an Fe-S cluster.

KIF4A and KIF4B are currently the only known components of the mitotic machinery that bind an Fe-S cluster. However, the significant mitotic defects observed when silencing the CIA, along with the colocalization of late-acting CIA factors with components of the mitotic machinery, suggest the potential existence of other, yet-to-be-discovered Fe-S cluster proteins that are essential for successful mitosis.

5. Other Fe-S Cluster Proteins in the Nucleus

The inventory of nuclear Fe-S cluster proteins presented in Table 1 includes some unexpected proteins whose functions are typically associated with other cellular compartments. These proteins are involved in post-translational modifications (ATE1, DPH1, and DPH2), nucleotide metabolism (XDH), the maturation of cytosolic Fe-S proteins (CIAPIN1, GLRX3, and BOLA2), immune modulation and restriction of viral replication (RSAD2), and mitochondrial function (SDHB, GLRX2, NFS1, and NFU1). It remains unclear why these proteins are also localized in the nucleus and whether they retain the function already characterized in other subcellular localizations or have alternative roles while in the nucleus.

6. Conclusions and Open Questions

About half of all known Fe-S binding proteins in human cells localize to the nucleus, with many identified only recently. A few decades ago, the presence of iron cofactors in the nucleus was considered unlikely due to the threat that chemically reactive iron ions pose to the genetic material. Yet the connection between iron cofactors and genome integrity is now well established. A large and increasing number of nuclear proteins require Fe-S clusters for full activity, particularly those involved in DNA replication and repair. However, the mechanistic benefit of incorporating Fe-S clusters into nucleic acid-binding proteins remains unclear. Do solvent-accessible, redox-active clusters play a role in sensing structural changes in DNA, e.g., DNA damage? According to ref. [228], failure to incorporate or maintain Fe-S clusters in these nuclear enzymes often leads to genome instability, which is linked to various diseases, including breast and ovarian cancer, Fanconi anemia, Xeroderma pigmentosum, and Cockayne, Warsaw breakage, and Hoyeraal–Hreidarsson syndromes.

Most nuclear Fe-S cluster proteins bind cubane [4Fe-4S] clusters using a canonical four-cysteinyl ligand structure and depend on the CIA machinery for insertion of their [4Fe-4S] clusters (Table S1). A systematic survey of the specific CIA subcomplexes and adaptors required for the metallation of each nuclear Fe-S protein is needed to fully understand how, when, and where Fe-S proteins acquire their clusters, as well as whether this process is regulated. Many nuclear Fe-S proteins appear to bind their clusters at solvent-exposed loops, and new evidence suggests that cluster transfer to fully folded proteins in complexes with adaptor proteins can occur. If co-translational insertion of clusters is not required for these proteins, is cluster insertion a process restricted to the cytoplasm, or does it also occur in the nucleus? CIA-targeting complexes may deliver clusters in situ when and where needed, as observed for the mitotic factor KIF4A, which colocalizes with MMS19 at the mitotic spindle. Solvent-exposed clusters are likely to be vulnerable to damage or disassembly, especially from cellular reactive oxygen or nitrogen species. The possibility of replacement or repair of damaged clusters for reactivating apo or damaged proteins raises important questions about whether dynamic trafficking of bioavailable Fe-S clusters occurs and whether an equilibrium exists between the apo and holo forms of these proteins. Cluster repair or replacement systems are likely to exist in multiple forms, as a single replacement system would not accommodate the variety of architectures for cluster-binding sites and stoichiometries exhibited by Fe-S proteins. Alternative systems for cluster replacement or repair could individually be needed for [2Fe-2S] proteins, [3Fe-4S] proteins, [4Fe-4S] proteins, and radical SAM enzymes.

One can speculate that these solvent-accessible clusters may allow for sensory roles, enabling dynamic cluster disassembly and reassembly in response to intracellular cues like redox status or iron levels. Thus, cells could modulate Fe-S enzyme activity according to their needs, similar to the cytosolic regulatory system of IRP1/ACO1. Fe-S trafficking systems, such as the Glrx3-[2Fe-2S]-BolA2 system that exchanges Fe-S clusters with nuclear iron-responsive transcriptional regulators in fungi, may also traffic a pool of bioavailable Fe-S clusters in the nucleus of mammalian cells. All of these hypotheses remain to be experimentally tested.

The discovery of Fe-S clusters in viral proteins, including the SARS-CoV-2 RNA-dependent RNA polymerase [129] and the helicase nsp13 [128], as well as the hepatitis B virus protein HBx, which localizes to the nucleus of mammalian cells during infection [229,230], indicates a broad role for Fe-S clusters in viral infection. It seems likely that additional nuclear Fe-S binding proteins will be identified, particularly among those automatically annotated in the proteome as Zn-binding proteins. We hope this review inspires further research on the unresolved questions regarding Fe-S clusters in human cells, potentially leading to promising avenues for the development of novel therapeutic strategies.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/inorganics12120316/s1, Table S1: Extended version of the inventory of Fe-S cluster binding proteins that localize to the nucleus of mammalian cells. The numbering of the residues implicated in cluster binding corresponds to the human proteins. The localization of the proteins reported here is based on consolidating information from references (when available) and from three different databases: The Human Protein Atlas (https://www.proteinatlas.org), The UniProt Knowledgebase (https://www.uniprot.org), and COMPARTMENTS (https://compartments.jensenlab.org). The asterisks (*) denote that Fe-S cluster ligands are shared between oligomeric partners from the CIA or ISC pathways. Question marks indicate unknown or debatable cluster binding residues, cluster types, or stoichiometries.

Author Contributions

Conceptualization, L.N.-A., A.L.-T. and C.C.P.; original draft preparation, L.N.-A. and A.L.-T.; L.N.-A. created the figures in BioRender.com; review and editing, L.N.-A., A.L.-T. and C.C.P. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the Intramural Research Program of the National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) at the National Institutes of Health (NIH).

Data Availability Statement

No new data were created or analyzed in this study. Data sharing is not applicable to this article.

Acknowledgments

The figures were created in BioRender. Novoa-Aponte, L. (2024) https://BioRender.com/m09w088 (accessed on 18 October 2024).

Conflicts of Interest

The authors declare no conflicts of interest.

References

Lill, R.; Broderick, J.B.; Dean, D.R. Special issue on iron-sulfur proteins: Structure, function, biogenesis and diseases. Biochim. Biophys. Acta 2015, 1853, 1251–1252. [Google Scholar] [CrossRef] [PubMed]
Braymer, J.J.; Freibert, S.A.; Rakwalska-Bange, M.; Lill, R. Mechanistic concepts of iron-sulfur protein biogenesis in Biology. Biochim. Et. Biophys. Acta (BBA)-Mol. Cell Res. 2021, 1868, 118863. [Google Scholar] [CrossRef] [PubMed]
Bak, D.W.; Weerapana, E. Proteomic strategies to interrogate the Fe-S proteome. Biochim. Biophys. Acta Mol. Cell Res. 2024, 1871, 119791. [Google Scholar] [CrossRef] [PubMed]
Rudolf, J.; Makrantoni, V.; Ingledew, W.J.; Stark, M.J.; White, M.F. The DNA repair helicases XPD and FancJ have essential iron-sulfur domains. Mol. Cell 2006, 23, 801–808. [Google Scholar] [CrossRef]
Thul, P.J.; Åkesson, L.; Wiking, M.; Mahdessian, D.; Geladaki, A.; Ait Blal, H.; Alm, T.; Asplund, A.; Björk, L.; Breckels, L.M. A subcellular map of the human proteome. Science 2017, 356, eaal3321. [Google Scholar] [CrossRef]
Consortium, T.U. UniProt: The universal protein knowledgebase in 2023. Nucleic Acids Res. 2023, 51, D523–D531. [Google Scholar] [CrossRef]
Binder, J.X.; Pletscher-Frankild, S.; Tsafou, K.; Stolte, C.; O’Donoghue, S.I.; Schneider, R.; Jensen, L.J. COMPARTMENTS: Unification and visualization of protein subcellular localization evidence. Database 2014, 2014, bau012. [Google Scholar] [CrossRef]
Klinge, S.; Hirst, J.; Maman, J.D.; Krude, T.; Pellegrini, L. An iron-sulfur domain of the eukaryotic primase is essential for RNA primer synthesis. Nat. Struct. Mol. Biol. 2007, 14, 875–877. [Google Scholar] [CrossRef]
Weiner, B.E.; Huang, H.; Dattilo, B.M.; Nilges, M.J.; Fanning, E.; Chazin, W.J. An iron-sulfur cluster in the C-terminal domain of the p58 subunit of human DNA primase. J. Biol. Chem. 2007, 282, 33444–33451. [Google Scholar] [CrossRef]
Starokadomskyy, P.; Gemelli, T.; Rios, J.J.; Xing, C.; Wang, R.C.; Li, H.; Pokatayev, V.; Dozmorov, I.; Khan, S.; Miyata, N. DNA polymerase-α regulates the activation of type I interferons through cytosolic RNA: DNA synthesis. Nat. Immunol. 2016, 17, 495–504. [Google Scholar] [CrossRef]
Netz, D.J.; Stith, C.M.; Stümpfig, M.; Köpf, G.; Vogel, D.; Genau, H.M.; Stodola, J.L.; Lill, R.; Burgers, P.M.; Pierik, A.J. Eukaryotic DNA polymerases require an iron-sulfur cluster for the formation of active complexes. Nat. Chem. Biol. 2012, 8, 125–132. [Google Scholar] [CrossRef] [PubMed]
Chea, J.; Zhang, S.; Zhao, H.; Zhang, Z.; Lee, E.Y.; Darzynkiewicz, Z.; Lee, M.Y. Spatiotemporal recruitment of human DNA polymerase delta to sites of UV damage. Cell Cycle 2012, 11, 2885–2895. [Google Scholar] [CrossRef] [PubMed]
Roske, J.J.; Yeeles, J.T. Structural basis for processive daughter-strand synthesis and proofreading by the human leading-strand DNA polymerase Pol ε. Nat. Struct. Mol. Biol. 2024, 1–11. [Google Scholar] [CrossRef]
Ræder, S.B.; Nepal, A.; Bjørås, K.Ø.; Seelinger, M.; Kolve, R.S.; Nedal, A.; Müller, R.; Otterlei, M. APIM-mediated REV3L–PCNA interaction important for error free TLS Over UV-induced DNA lesions in human cells. Int. J. Mol. Sci. 2018, 20, 100. [Google Scholar] [CrossRef]
Cantor, S.B.; Bell, D.W.; Ganesan, S.; Kass, E.M.; Drapkin, R.; Grossman, S.; Wahrer, D.C.; Sgroi, D.C.; Lane, W.S.; Haber, D.A. BACH1, a novel helicase-like protein, interacts directly with BRCA1 and contributes to its DNA repair function. Cell 2001, 105, 149–160. [Google Scholar] [CrossRef]
Seki, M.; Takeda, Y.; Iwai, K.; Tanaka, K. IOP1 protein is an external component of the human cytosolic iron-sulfur cluster assembly (CIA) machinery and functions in the MMS19 protein-dependent CIA pathway. J. Biol. Chem. 2013, 288, 16680–16689. [Google Scholar] [CrossRef]
Wu, Y.; Sommers, J.A.; Suhasini, A.N.; Leonard, T.; Deakyne, J.S.; Mazin, A.V.; Shin-Ya, K.; Kitao, H.; Brosz, R.M., Jr. Fanconi anemia group J mutation abolishes its DNA repair function by uncoupling DNA translocation from helicase activity or disruption of protein-DNA complexes. Blood J. Am. Soc. Hematol. 2010, 116, 3780–3791. [Google Scholar] [CrossRef]
Parish, J.L.; Rosa, J.; Wang, X.; Lahti, J.M.; Doxsey, S.J.; Androphy, E.J. The DNA helicase ChlR1 is required for sister chromatid cohesion in mammalian cells. J. Cell Sci. 2006, 119, 4857–4865. [Google Scholar] [CrossRef]
Simon, A.K.; Kummer, S.; Wild, S.; Lezaja, A.; Teloni, F.; Jozwiakowski, S.K.; Altmeyer, M.; Gari, K. The iron–sulfur helicase DDX11 promotes the generation of single-stranded DNA for CHK1 activation. Life Sci. Alliance 2020, 3. [Google Scholar] [CrossRef]
Landry, A.P.; Ding, H. The N-Terminal Domain of Human DNA Helicase Rtel1 Contains a Redox Active Iron-Sulfur Cluster. BioMed Res. Int. 2014, 2014, 285791. [Google Scholar] [CrossRef]
Ito, S.; Tan, L.J.; Andoh, D.; Narita, T.; Seki, M.; Hirano, Y.; Narita, K.; Kuraoka, I.; Hiraoka, Y.; Tanaka, K. MMXD, a TFIIH-independent XPD-MMS19 protein complex involved in chromosome segregation. Mol. Cell 2010, 39, 632–640. [Google Scholar] [CrossRef] [PubMed]
Duxin, J.P.; Dao, B.; Martinsson, P.; Rajala, N.; Guittat, L.; Campbell, J.L.; Spelbrink, J.N.; Stewart, S.A. Human Dna2 is a nuclear and mitochondrial DNA maintenance protein. Mol. Cell. Biol. 2009, 29, 4274–4282. [Google Scholar] [CrossRef] [PubMed]
Pokharel, S.; Campbell, J.L. Cross talk between the nuclease and helicase activities of Dna2: Role of an essential iron–sulfur cluster domain. Nucleic Acids Res. 2012, 40, 7821–7830. [Google Scholar] [CrossRef]
Mariotti, L.; Wild, S.; Brunoldi, G.; Piceni, A.; Ceppi, I.; Kummer, S.; Lutz, R.E.; Cejka, P.; Gari, K. The iron–sulphur cluster in human DNA2 is required for all biochemical activities of DNA2. Commun. Biol. 2020, 3, 322. [Google Scholar] [CrossRef]
Sparks, J.L.; Kumar, R.; Singh, M.; Wold, M.S.; Pandita, T.K.; Burgers, P.M. Human exonuclease 5 is a novel sliding exonuclease required for genome stability. J. Biol. Chem. 2012, 287, 42773–42783. [Google Scholar] [CrossRef]
Komine, K.; Shimodaira, H.; Takao, M.; Soeda, H.; Zhang, X.; Takahashi, M.; Ishioka, C. Functional complementation assay for 47 MUTYH variants in a MutY-disrupted Escherichia coli strain. Hum. Mutat. 2015, 36, 704–711. [Google Scholar] [CrossRef]
Chepanoske, C.L.; Golinelli, M.-P.; Williams, S.D.; David, S.S. Positively Charged Residues within the Iron–Sulfur Cluster Loop of E. coli MutY Participate in Damage Recognition and Removal. Arch. Biochem. Biophys. 2000, 380, 11–19. [Google Scholar] [CrossRef]
Nuñez, N.N.; Majumdar, C.; Lay, K.T.; David, S.S. Fe–S clusters and MutY base excision repair glycosylases: Purification, kinetics, and DNA affinity measurements. In Methods in Enzymology; Elsevier: Amsterdam, The Netherlands, 2018; Volume 599, pp. 21–68. [Google Scholar]
Ikeda, S.; Kohmoto, T.; Tabata, R.; Seki, Y. Differential intracellular localization of the human and mouse endonuclease III homologs and analysis of the sorting signals. DNA Repair. 2002, 1, 847–854. [Google Scholar] [CrossRef]
Kuo, C.-F.; McRee, D.E.; Fisher, C.L.; O’Handley, S.F.; Cunningham, R.P.; Tainer, J.A. Atomic structure of the DNA repair [4Fe-4S] enzyme endonuclease III. Science 1992, 258, 434–440. [Google Scholar] [CrossRef]
Carroll, B.L.; Zahn, K.E.; Hanley, J.P.; Wallace, S.S.; Dragon, J.A.; Doublié, S. Caught in motion: Human NTHL1 undergoes interdomain rearrangement necessary for catalysis. Nucleic Acids Res. 2021, 49, 13165–13178. [Google Scholar] [CrossRef]
Kim, J.-H.; Lane, W.S.; Reinberg, D. Human Elongator facilitates RNA polymerase II transcription through chromatin. Proc. Natl. Acad. Sci. USA 2002, 99, 1241–1246. [Google Scholar] [CrossRef] [PubMed]
Paraskevopoulou, C.; Fairhurst, S.A.; Lowe, D.J.; Brick, P.; Onesti, S. The elongator subunit Elp3 contains a Fe4S4 cluster and binds S-adenosylmethionine. Mol. Microbiol. 2006, 59, 795–806. [Google Scholar] [CrossRef]
Kimura, S.; Suzuki, T. Iron-sulfur proteins responsible for RNA modifications. Biochim. Biophys. Acta 2015, 1853, 1272–1283. [Google Scholar] [CrossRef] [PubMed]
Ramsay, E.P.; Abascal-Palacios, G.; Daiß, J.L.; King, H.; Gouge, J.; Pilsl, M.; Beuron, F.; Morris, E.; Gunkel, P.; Engel, C. Structure of human RNA polymerase III. Nat. Commun. 2020, 11, 6409. [Google Scholar] [CrossRef] [PubMed]
Blombach, F.; Salvadori, E.; Fouqueau, T.; Yan, J.; Reimann, J.; Sheppard, C.; Smollett, K.L.; Albers, S.V.; Kay, C.W.; Thalassinos, K. Archaeal TFEα/β is a hybrid of TFIIE and the RNA polymerase III subcomplex hRPC62/39. Elife 2015, 4, e08378. [Google Scholar] [CrossRef]
Li, L.; Yu, Z.; Zhao, D.; Ren, Y.; Hou, H.; Xu, Y. Structure of human RNA polymerase III elongation complex. Cell Res. 2021, 31, 791–800. [Google Scholar] [CrossRef]
Chen, W.; Qin, L.; Wang, S.; Li, M.; Shi, D.; Tian, Y.; Wang, J.; Fu, L.; Li, Z.; Guo, W. CPSF4 activates telomerase reverse transcriptase and predicts poor prognosis in human lung adenocarcinomas. Mol. Oncol. 2014, 8, 704–716. [Google Scholar] [CrossRef]
Shimberg, G.D.; Michalek, J.L.; Oluyadi, A.A.; Rodrigues, A.V.; Zucconi, B.E.; Neu, H.M.; Ghosh, S.; Sureschandra, K.; Wilson, G.M.; Stemmler, T.L.; et al. Cleavage and polyadenylation specificity factor 30: An RNA-binding zinc-finger protein with an unexpected 2Fe-2S cluster. Proc. Natl. Acad. Sci. USA 2016, 113, 4700–4705. [Google Scholar] [CrossRef]
Pritts, J.D.; Hursey, M.S.; Michalek, J.L.; Batelu, S.; Stemmler, T.L.; Michel, S.L. Unraveling the RNA Binding Properties of the Iron–Sulfur Zinc Finger Protein CPSF30. Biochemistry 2020, 59, 970–982. [Google Scholar] [CrossRef]
Young, A.P.; Bandarian, V. TYW1: A radical SAM enzyme involved in the biosynthesis of wybutosine bases. In Methods in Enzymology; Elsevier: Amsterdam, The Netherlands, 2018; Volume 606, pp. 119–153. [Google Scholar]
Reiter, V.; Matschkal, D.M.; Wagner, M.; Globisch, D.; Kneuttinger, A.C.; Muller, M.; Carell, T. The CDK5 repressor CDK5RAP1 is a methylthiotransferase acting on nuclear and mitochondrial RNA. Nucleic Acids Res. 2012, 40, 6235–6240. [Google Scholar] [CrossRef]
Pierrel, F.; Douki, T.; Fontecave, M.; Atta, M. MiaB protein is a bifunctional radical-S-adenosylmethionine enzyme involved in thiolation and methylation of tRNA. J. Biol. Chem. 2004, 279, 47555–47563. [Google Scholar] [CrossRef] [PubMed]
Lee, Y.M.; Lee, S.; Lee, E.; Shin, H.; Hahn, H.; Choi, W.; Kim, W. Human kinesin superfamily member 4 is dominantly localized in the nuclear matrix and is associated with chromosomes during mitosis. Biochem. J. 2001, 360, 549–556. [Google Scholar] [CrossRef] [PubMed]
Ben-Shimon, L.; Paul, V.D.; David-Kadoch, G.; Volpe, M.; Stümpfig, M.; Bill, E.; Mühlenhoff, U.; Lill, R.; Ben-Aroya, S. Fe-S cluster coordination of the chromokinesin KIF4A alters its subcellular localization during mitosis. J. Cell Sci. 2018, 131, jcs211433. [Google Scholar] [CrossRef]
Bellelli, R.; Castellone, M.D.; Guida, T.; Limongello, R.; Dathan, N.A.; Merolla, F.; Cirafici, A.M.; Affuso, A.; Masai, H.; Costanzo, V. NCOA4 transcriptional coactivator inhibits activation of DNA replication origins. Mol. Cell 2014, 55, 123–137. [Google Scholar] [CrossRef]
Kuno, S.; Iwai, K. Oxygen modulates iron homeostasis by switching iron sensing of NCOA4. J. Biol. Chem. 2023, 299. [Google Scholar] [CrossRef]
Zhao, H.; Lu, Y.; Zhang, J.; Sun, Z.; Cheng, C.; Liu, Y.; Wu, L.; Zhang, M.; He, W.; Hao, S. NCOA4 requires a [3Fe-4S] to sense and maintain the iron homeostasis. J. Biol. Chem. 2024, 300, 105612. [Google Scholar] [CrossRef]
Vinas-Castells, R.; Frias, A.; Robles-Lanuza, E.; Zhang, K.; Longmore, G.D.; Garcia de Herreros, A.; Diaz, V.M. Nuclear ubiquitination by FBXL5 modulates Snail1 DNA binding and stability. Nucleic Acids Res. 2013, 42, 1079–1094. [Google Scholar] [CrossRef]
Wang, H.; Shi, H.; Rajan, M.; Canarie, E.R.; Hong, S.; Simoneschi, D.; Pagano, M.; Bush, M.F.; Stoll, S.; Leibold, E.A. FBXL5 regulates IRP2 stability in iron homeostasis via an oxygen-responsive [2Fe2S] cluster. Mol. Cell 2020, 78, 31–41.e35. [Google Scholar] [CrossRef]
Bruening, W.; Prowse, A.H.; Schultz, D.C.; Holgado-Madruga, M.; Wong, A.; Godwin, A.K. Expression of OVCA1, a candidate tumor suppressor, is reduced in tumors and inhibits growth of ovarian cancer cells. Cancer Res. 1999, 59, 4973–4983. [Google Scholar]
Zhang, Y.; Zhu, X.; Torelli, A.T.; Lee, M.; Dzikovski, B.; Koralewski, R.M.; Wang, E.; Freed, J.; Krebs, C.; Ealick, S.E. Diphthamide biosynthesis requires an organic radical generated by an iron–sulphur enzyme. Nature 2010, 465, 891–896. [Google Scholar] [CrossRef]
Kwon, Y.T.; Kashina, A.S.; Varshavsky, A. Alternative splicing results in differential expression, activity, and localization of the two forms of arginyl-tRNA-protein transferase, a component of the N-end rule pathway. Mol. Cell. Biol. 1999, 19, 182–193. [Google Scholar] [CrossRef] [PubMed]
Van, V.; Brown, J.B.; O’Shea, C.R.; Rosenbach, H.; Mohamed, I.; Ejimogu, N.-E.; Bui, T.S.; Szalai, V.A.; Chacón, K.N.; Span, I. Iron-sulfur clusters are involved in post-translational arginylation. Nat. Commun. 2023, 14, 458. [Google Scholar] [CrossRef] [PubMed]
Du, Z.; Zhou, X.; Lai, Y.; Xu, J.; Zhang, Y.; Zhou, S.; Feng, Z.; Yu, L.; Tang, Y.; Wang, W. Structure of the human respiratory complex II. Proc. Natl. Acad. Sci. USA 2023, 120, e2216713120. [Google Scholar] [CrossRef]
Hao, Z.; Li, X.; Qiao, T.; Du, R.; Zhang, G.; Fan, D. Subcellular localization of CIAPIN1. J. Histochem. Cytochem. 2006, 54, 1437–1444. [Google Scholar] [CrossRef]
Banci, L.; Bertini, I.; Ciofi-Baffoni, S.; Boscaro, F.; Chatzi, A.; Mikolajczyk, M.; Tokatlidis, K.; Winkelmann, J. Anamorsin is a [2Fe-2S] cluster-containing substrate of the Mia40-dependent mitochondrial protein trapping machinery. Chem. Biol. 2011, 18, 794–804. [Google Scholar] [CrossRef]
Zhang, Y.; Yang, C.; Dancis, A.; Nakamaru-Ogiso, E. EPR studies of wild type and mutant Dre2 identify essential [2Fe–-2S] and [4Fe–-4S] clusters and their cysteine ligands. J. Biochem. 2016, 161, 67–78. [Google Scholar] [CrossRef]
Willems, P.; Wanschers, B.F.; Esseling, J.; Szklarczyk, R.; Kudla, U.; Duarte, I.; Forkink, M.; Nooteboom, M.; Swarts, H.; Gloerich, J. BOLA1 is an aerobic protein that prevents mitochondrial morphology changes induced by glutathione depletion. Antioxid. Redox Signal. 2013, 18, 129–138. [Google Scholar] [CrossRef]
Li, H.; Mapolelo, D.T.; Randeniya, S.; Johnson, M.K.; Outten, C.E. Human glutaredoxin 3 forms [2Fe-2S]-bridged complexes with human BolA2. Biochemistry 2012, 51, 1687–1696. [Google Scholar] [CrossRef]
Frey, A.G.; Palenchar, D.J.; Wildemann, J.D.; Philpott, C.C. A Glutaredoxin· BolA Complex Serves as an Iron-Sulfur Cluster Chaperone for the Cytosolic Cluster Assembly Machinery. J. Biol. Chem. 2016, 291, 22344–22356. [Google Scholar] [CrossRef]
Pandya, P.; Braiman, A.; Isakov, N. PICOT (GLRX3) is a positive regulator of stress-induced DNA-damage response. Cell. Signal. 2019, 62, 109340. [Google Scholar] [CrossRef]
Okuno, T.; Yamabayashi, H.; Kogure, K. Comparison of intracellular localization of Nubp1 and Nubp2 using GFP fusion proteins. Mol. Biol. Rep. 2010, 37, 1165–1168. [Google Scholar] [CrossRef] [PubMed]
Netz, D.J.; Pierik, A.J.; Stümpfig, M.; Mühlenhoff, U.; Lill, R. The Cfd1–Nbp35 complex acts as a scaffold for iron-sulfur protein assembly in the yeast cytosol. Nat. Chem. Biol. 2007, 3, 278–286. [Google Scholar] [CrossRef] [PubMed]
Stehling, O.; Netz, D.J.; Niggemeyer, B.; Rösser, R.; Eisenstein, R.S.; Puccio, H.; Pierik, A.J.; Lill, R. Human Nbp35 is essential for both cytosolic iron-sulfur protein assembly and iron homeostasis. Mol. Cell. Biol. 2008, 28, 5517–5528. [Google Scholar] [CrossRef] [PubMed]
Tong, W.-H.; Jameson, G.N.; Huynh, B.H.; Rouault, T.A. Subcellular compartmentalization of human Nfu, an iron–sulfur cluster scaffold protein, and its ability to assemble a [4Fe–4S] cluster. Proc. Natl. Acad. Sci. USA 2003, 100, 9762–9767. [Google Scholar] [CrossRef]
Cai, K.; Liu, G.; Frederick, R.O.; Xiao, R.; Montelione, G.T.; Markley, J.L. Structural/functional properties of human NFU1, an intermediate [4Fe-4S] carrier in human mitochondrial iron-sulfur cluster biogenesis. Structure 2016, 24, 2080–2091. [Google Scholar] [CrossRef]
Land, T.; Rouault, T.A. Targeting of a human iron–sulfur cluster assembly enzyme, nifs, to different subcellular compartments is regulated through alternative AUG utilization. Mol. Cell 1998, 2, 807–815. [Google Scholar] [CrossRef]
Fox, N.G.; Yu, X.; Feng, X.; Bailey, H.J.; Martelli, A.; Nabhan, J.F.; Strain-Damerell, C.; Bulawa, C.; Yue, W.W.; Han, S. Structure of the human frataxin-bound iron-sulfur cluster assembly complex provides insight into its activation mechanism. Nat. Commun. 2019, 10, 2210. [Google Scholar] [CrossRef]
Marelja, Z.; Stöcklein, W.; Nimtz, M.; Leimkühler, S. A novel role for human Nfs1 in the cytoplasm: Nfs1 acts as a sulfur donor for MOCS3, a protein involved in molybdenum cofactor biosynthesis. J. Biol. Chem. 2008, 283, 25178–25185. [Google Scholar] [CrossRef]
Lundberg, M.; Johansson, C.; Chandra, J.; Enoksson, M.; Jacobsson, G.; Ljung, J.; Johansson, M.; Holmgren, A. Cloning and expression of a novel human glutaredoxin (Grx2) with mitochondrial and nuclear isoforms. J. Biol. Chem. 2001, 276, 26269–26275. [Google Scholar] [CrossRef]
Lillig, C.H.; Berndt, C.; Vergnolle, O.; Lönn, M.E.; Hudemann, C.; Bill, E.; Holmgren, A. Characterization of human glutaredoxin 2 as iron–sulfur protein: A possible role as redox sensor. Proc. Natl. Acad. Sci. USA 2005, 102, 8168–8173. [Google Scholar] [CrossRef]
Kispal, G.; Csere, P.; Prohl, C.; Lill, R. The mitochondrial proteins Atm1p and Nfs1p are essential for biogenesis of cytosolic Fe/S proteins. EMBO J. 1999, 18, 3981–3989. [Google Scholar] [CrossRef] [PubMed]
Netz, D.J.; Mascarenhas, J.; Stehling, O.; Pierik, A.J.; Lill, R. Maturation of cytosolic and nuclear iron–sulfur proteins. Trends Cell Biol. 2014, 24, 303–312. [Google Scholar] [CrossRef] [PubMed]
Song, D.; Lee, F.S. A role for IOP1 in mammalian cytosolic iron-sulfur protein biogenesis. J. Biol. Chem. 2008, 283, 9231–9238. [Google Scholar] [CrossRef] [PubMed]
Netz, D.J.; Pierik, A.J.; Stümpfig, M.; Bill, E.; Sharma, A.K.; Pallesen, L.J.; Walden, W.E.; Lill, R. A bridging [4Fe-4S] cluster and nucleotide binding are essential for function of the Cfd1-Nbp35 complex as a scaffold in iron-sulfur protein maturation. J. Biol. Chem. 2012, 287, 12365–12378. [Google Scholar] [CrossRef]
Stehling, O.; Vashisht, A.A.; Mascarenhas, J.; Jonsson, Z.O.; Sharma, T.; Netz, D.J.; Pierik, A.J.; Wohlschlegel, J.A.; Lill, R. MMS19 assembles iron-sulfur proteins required for DNA metabolism and genomic integrity. Science 2012, 337, 195–199. [Google Scholar] [CrossRef]
Gari, K.; León Ortiz, A.M.; Borel, V.; Flynn, H.; Skehel, J.M.; Boulton, S.J. MMS19 links cytoplasmic iron-sulfur cluster assembly to DNA metabolism. Science 2012, 337, 243–245. [Google Scholar] [CrossRef]
Balk, J.; Pierik, A.J.; Netz, D.J.A.; Mühlenhoff, U.; Lill, R. The hydrogenase-like Nar1p is essential for maturation of cytosolic and nuclear iron–sulphur proteins. EMBO J. 2004, 23, 2105–2115. [Google Scholar] [CrossRef]
SantaMaria, A.M.; Rouault, T.A. Regulatory and Sensing Iron–Sulfur Clusters: New Insights and Unanswered Questions. Inorganics 2024, 12, 101. [Google Scholar] [CrossRef]
Querci, L.; Piccioli, M.; Ciofi-Baffoni, S.; Banci, L. Structural aspects of iron-sulfur protein biogenesis: An NMR view. Biochim. Et. Biophys. Acta (BBA)-Mol. Cell Res. 2024, 1871, 119786. [Google Scholar] [CrossRef]
Maio, N.; Singh, A.; Uhrigshardt, H.; Saxena, N.; Tong, W.-H.; Rouault, T.A. Cochaperone binding to LYR motifs confers specificity of iron sulfur cluster delivery. Cell Metab. 2014, 19, 445–457. [Google Scholar] [CrossRef]
Li, K.; Tong, W.-H.; Hughes, R.M.; Rouault, T.A. Roles of the mammalian cytosolic cysteine desulfurase, ISCS, and scaffold protein, ISCU, in iron-sulfur cluster assembly. J. Biol. Chem. 2006, 281, 12344–12351. [Google Scholar] [CrossRef]
Kim, K.S.; Maio, N.; Singh, A.; Rouault, T.A. Cytosolic HSC20 integrates de novo iron–sulfur cluster biogenesis with the CIAO1-mediated transfer to recipients. Hum. Mol. Genet. 2018, 27, 837–852. [Google Scholar] [CrossRef] [PubMed]
Shi, H.; Bencze, K.Z.; Stemmler, T.L.; Philpott, C.C. A cytosolic iron chaperone that delivers iron to ferritin. Science 2008, 320, 1207–1210. [Google Scholar] [CrossRef] [PubMed]
Philpott, C.C. Coming into view: Eukaryotic iron chaperones and intracellular iron delivery. J. Biol. Chem. 2012, 287, 13518–13523. [Google Scholar] [CrossRef] [PubMed]
Philpott, C.C.; Ryu, M.-S.; Frey, A.; Patel, S. Cytosolic iron chaperones: Proteins delivering iron cofactors in the cytosol of mammalian cells. J. Biol. Chem. 2017, 292, 12764–12771. [Google Scholar] [CrossRef]
Philpott, C.C.; Patel, S.J.; Protchenko, O. Management versus miscues in the cytosolic labile iron pool: The varied functions of iron chaperones. Biochim. Et Biophys. Acta (BBA)-Mol. Cell Res. 2020, 1867, 118830. [Google Scholar] [CrossRef]
Patel, S.J.; Frey, A.G.; Palenchar, D.J.; Achar, S.; Bullough, K.Z.; Vashisht, A.; Wohlschlegel, J.A.; Philpott, C.C. A PCBP1–BolA2 chaperone complex delivers iron for cytosolic [2Fe–2S] cluster assembly. Nat. Chem. Biol. 2019, 15, 872–881. [Google Scholar] [CrossRef]
Adamec, J.; Rusnak, F.; Owen, W.G.; Naylor, S.; Benson, L.M.; Gacy, A.M.; Isaya, G. Iron-dependent self-assembly of recombinant yeast frataxin: Implications for Friedreich ataxia. Am. J. Hum. Genet. 2000, 67, 549–562. [Google Scholar] [CrossRef]
Bulteau, A.-L.; O’Neill, H.A.; Kennedy, M.C.; Ikeda-Saito, M.; Isaya, G.; Szweda, L.I. Frataxin acts as an iron chaperone protein to modulate mitochondrial aconitase activity. Science 2004, 305, 242–245. [Google Scholar] [CrossRef]
Cavadini, P.; O’Neill, H.A.; Benada, O.; Isaya, G. Assembly and iron-binding properties of human frataxin, the protein deficient in Friedreich ataxia. Hum. Mol. Genet. 2002, 11, 217–227. [Google Scholar] [CrossRef]
Bencze, K.Z.; Kondapalli, K.C.; Cook, J.D.; McMahon, S.; Millán-Pacheco, C.; Pastor, N.; Stemmler, T.L. The structure and function of frataxin. Crit. Rev. Biochem. Mol. Biol. 2006, 41, 269–291. [Google Scholar] [CrossRef] [PubMed]
Parent, A.; Elduque, X.; Cornu, D.; Belot, L.; Le Caer, J.-P.; Grandas, A.; Toledano, M.B.; D’autréaux, B. Mammalian frataxin directly enhances sulfur transfer of NFS1 persulfide to both ISCU and free thiols. Nat. Commun. 2015, 6, 5686. [Google Scholar] [CrossRef] [PubMed]
Bridwell-Rabb, J.; Fox, N.G.; Tsai, C.-L.; Winn, A.M.; Barondeau, D.P. Human frataxin activates Fe–S cluster biosynthesis by facilitating sulfur transfer chemistry. Biochemistry 2014, 53, 4904–4913. [Google Scholar] [CrossRef]
Balk, J.; Aguilar Netz, D.J.; Tepper, K.; Pierik, A.J.; Lill, R. The essential WD40 protein Cia1 is involved in a late step of cytosolic and nuclear iron-sulfur protein assembly. Mol. Cell. Biol. 2005, 25, 10833–10841. [Google Scholar] [CrossRef]
Stehling, O.; Mascarenhas, J.; Vashisht, A.A.; Sheftel, A.D.; Niggemeyer, B.; Rösser, R.; Pierik, A.J.; Wohlschlegel, J.A.; Lill, R. Human CIA2A-FAM96A and CIA2B-FAM96B integrate iron homeostasis and maturation of different subsets of cytosolic-nuclear iron-sulfur proteins. Cell Metab. 2013, 18, 187–198. [Google Scholar] [CrossRef]
Marquez, M.D.; Greth, C.; Buzuk, A.; Liu, Y.; Blinn, C.M.; Beller, S.; Leiskau, L.; Hushka, A.; Wu, K.; Nur, K. Cytosolic iron–sulfur protein assembly system identifies clients by a C-terminal tripeptide. Proc. Natl. Acad. Sci. USA 2023, 120, e2311057120. [Google Scholar] [CrossRef]
Paul, V.D.; Mühlenhoff, U.; Stümpfig, M.; Seebacher, J.; Kugler, K.G.; Renicke, C.; Taxis, C.; Gavin, A.-C.; Pierik, A.J.; Lill, R. The deca-GX3 proteins Yae1-Lto1 function as adaptors recruiting the ABC protein Rli1 for iron-sulfur cluster insertion. Elife 2015, 4, e08231. [Google Scholar] [CrossRef]
Odermatt, D.C.; Gari, K. The CIA Targeting Complex Is Highly Regulated and Provides Two Distinct Binding Sites for Client Iron-Sulfur Proteins. Cell Rep. 2017, 18, 1434–1443. [Google Scholar] [CrossRef]
Kassube, S.A.; Thomä, N.H. Structural insights into Fe–S protein biogenesis by the CIA targeting complex. Nat. Struct. Mol. Biol. 2020, 27, 735–742. [Google Scholar] [CrossRef]
Alhebshi, A.; Sideri, T.C.; Holland, S.L.; Avery, S.V. The essential iron-sulfur protein Rli1 is an important target accounting for inhibition of cell growth by reactive oxygen species. Mol. Biol. Cell 2012, 23, 3582–3590. [Google Scholar] [CrossRef]
Honarmand Ebrahimi, K.; Ciofi-Baffoni, S.; Hagedoorn, P.-L.; Nicolet, Y.; Le Brun, N.E.; Hagen, W.R.; Armstrong, F.A. Iron–sulfur clusters as inhibitors and catalysts of viral replication. Nat. Chem. 2022, 14, 253–266. [Google Scholar] [CrossRef]
Jang, S.; Imlay, J.A. Micromolar intracellular hydrogen peroxide disrupts metabolism by damaging iron-sulfur enzymes. J. Biol. Chem. 2007, 282, 929–937. [Google Scholar] [CrossRef]
Mettert, E.L.; Kiley, P.J. Fe–S proteins that regulate gene expression. Biochim. Et Biophys. Acta (BBA)-Mol. Cell Res. 2015, 1853, 1284–1293. [Google Scholar] [CrossRef] [PubMed]
Imlay, J.A. Iron-sulphur clusters and the problem with oxygen. Mol. Microbiol. 2006, 59, 1073–1082. [Google Scholar] [CrossRef]
Mettert, E.L.; Kiley, P.J. How is Fe-S cluster formation regulated? Annu. Rev. Microbiol. 2015, 69, 505–526. [Google Scholar] [CrossRef]
Lauble, H.; Kennedy, M.; Beinert, H.; Stout, C. Crystal structures of aconitase with isocitrate and nitroisocitrate bound. Biochemistry 1992, 31, 2735–2748. [Google Scholar] [CrossRef]
Federico, G.; Carrillo, F.; Dapporto, F.; Chiariello, M.; Santoro, M.; Bellelli, R.; Carlomagno, F. NCOA4 links iron bioavailability to DNA metabolism. Cell Rep. 2022, 40, 111207. [Google Scholar] [CrossRef]
Mayank, A.K.; Pandey, V.; Vashisht, A.A.; Barshop, W.D.; Rayatpisheh, S.; Sharma, T.; Haque, T.; Powers, D.N.; Wohlschlegel, J.A. An oxygen-dependent interaction between FBXL5 and the CIA-targeting complex regulates iron homeostasis. Mol. Cell 2019, 75, 382–393.e385. [Google Scholar] [CrossRef]
van Wietmarschen, N.; Moradian, A.; Morin, G.B.; Lansdorp, P.M.; Uringa, E.-J. The mammalian proteins MMS19, MIP18, and ANT2 are involved in cytoplasmic iron-sulfur cluster protein assembly. J. Biol. Chem. 2012, 287, 43351–43358. [Google Scholar] [CrossRef]
Puig, S.; Ramos-Alonso, L.; Romero, A.; Martínez-Pastor, M. The elemental role of iron in DNA synthesis and repair. Metallomics 2017, 9, 1483–1500. [Google Scholar] [CrossRef]
Troadec, M.-B.; Loréal, O.; Brissot, P. The interaction of iron and the genome: For better and for worse. Mutat. Res./Rev. Mutat. Res. 2017, 774, 25–32. [Google Scholar] [CrossRef] [PubMed]
Veatch, J.R.; McMurray, M.A.; Nelson, Z.W.; Gottschling, D.E. Mitochondrial dysfunction leads to nuclear genome instability via an iron-sulfur cluster defect. Cell 2009, 137, 1247–1258. [Google Scholar] [CrossRef] [PubMed]
Shi, R.; Hou, W.; Wang, Z.-Q.; Xu, X. Biogenesis of iron–sulfur clusters and their role in DNA metabolism. Front. Cell Dev. Biol. 2021, 9, 735678. [Google Scholar] [CrossRef]
Loeb, L.A.; Monnat, R.J., Jr. DNA polymerases and human disease. Nat. Rev. Genet. 2008, 9, 594–604. [Google Scholar] [CrossRef]
Ter Beek, J.; Parkash, V.; Bylund, G.O.; Osterman, P.; Sauer-Eriksson, A.E.; Johansson, E. Structural evidence for an essential Fe–S cluster in the catalytic core domain of DNA polymerase ϵ. Nucleic Acids Res. 2019, 47, 5712–5722. [Google Scholar] [CrossRef]
Jain, R.; Vanamee, E.S.; Dzikovski, B.G.; Buku, A.; Johnson, R.E.; Prakash, L.; Prakash, S.; Aggarwal, A.K. An iron–sulfur cluster in the polymerase domain of yeast DNA polymerase ε. J. Mol. Biol. 2014, 426, 301–308. [Google Scholar] [CrossRef]
Baranovskiy, A.G.; Lada, A.G.; Siebler, H.M.; Zhang, Y.; Pavlov, Y.I.; Tahirov, T.H. DNA polymerase δ and ζ switch by sharing accessory subunits of DNA polymerase δ. J. Biol. Chem. 2012, 287, 17281–17287. [Google Scholar] [CrossRef]
Lisova, A.E.; Baranovskiy, A.G.; Morstadt, L.M.; Babayeva, N.D.; Stepchenkova, E.I.; Tahirov, T.H. The iron-sulfur cluster is essential for DNA binding by human DNA polymerase ε. Sci. Rep. 2022, 12, 17436. [Google Scholar] [CrossRef]
Bartels, P.L.; Stodola, J.L.; Burgers, P.M.; Barton, J.K. A redox role for the [4Fe4S] cluster of yeast DNA polymerase δ. J. Am. Chem. Soc. 2017, 139, 18339–18348. [Google Scholar] [CrossRef]
Baranovskiy, A.G.; Babayeva, N.D.; Zhang, Y.; Gu, J.; Suwa, Y.; Pavlov, Y.I.; Tahirov, T.H. Mechanism of concerted RNA-DNA primer synthesis by the human primosome. J. Biol. Chem. 2016, 291, 10006–10020. [Google Scholar] [CrossRef]
Suwa, Y.; Gu, J.; Baranovskiy, A.G.; Babayeva, N.D.; Pavlov, Y.I.; Tahirov, T.H. Crystal structure of the human Pol α B subunit in complex with the C-terminal domain of the catalytic subunit. J. Biol. Chem. 2015, 290, 14328–14337. [Google Scholar] [CrossRef] [PubMed]
Klinge, S.; Núñez-Ramírez, R.; Llorca, O.; Pellegrini, L. 3D architecture of DNA Pol α reveals the functional core of multi-subunit replicative polymerases. EMBO J. 2009, 28, 1978–1987. [Google Scholar] [CrossRef] [PubMed]
Zhang, Y.; Baranovskiy, A.G.; Tahirov, T.H.; Pavlov, Y.I. The C-terminal domain of the DNA polymerase catalytic subunit regulates the primase and polymerase activities of the human DNA polymerase α-primase complex. J. Biol. Chem. 2014, 289, 22021–22034. [Google Scholar] [CrossRef]
Baranovskiy, A.G.; Tahirov, T.H. Elaborated action of the human primosome. Genes 2017, 8, 62. [Google Scholar] [CrossRef]
Pritts, J.D.; Michel, S.L. Fe-S clusters masquerading as zinc finger proteins. J. Inorg. Biochem. 2022, 230, 111756. [Google Scholar] [CrossRef]
Maio, N.; Raza, M.K.; Li, Y.; Zhang, D.-L.; Bollinger, J.M., Jr.; Krebs, C.; Rouault, T.A. An iron–sulfur cluster in the zinc-binding domain of the SARS-CoV-2 helicase modulates its RNA-binding and-unwinding activities. Proc. Natl. Acad. Sci. USA 2023, 120, e2303860120. [Google Scholar] [CrossRef]
Maio, N.; Lafont, B.A.; Sil, D.; Li, Y.; Bollinger, J.M., Jr.; Krebs, C.; Pierson, T.C.; Linehan, W.M.; Rouault, T.A. Fe-S cofactors in the SARS-CoV-2 RNA-dependent RNA polymerase are potential antiviral targets. Science 2021, 373, 236–241. [Google Scholar] [CrossRef]
Conlan, A.R.; Axelrod, H.L.; Cohen, A.E.; Abresch, E.C.; Zuris, J.; Yee, D.; Nechushtai, R.; Jennings, P.A.; Paddock, M.L. Crystal structure of Miner1: The redox-active 2Fe-2S protein causative in Wolfram Syndrome 2. J. Mol. Biol. 2009, 392, 143–153. [Google Scholar] [CrossRef]
Cutone, A.; Howes, B.D.; Miele, A.E.; Miele, R.; Giorgi, A.; Battistoni, A.; Smulevich, G.; Musci, G.; Di Patti, M.C.B. Pichia pastoris Fep1 is a [2Fe-2S] protein with a Zn finger that displays an unusual oxygen-dependent role in cluster binding. Sci. Rep. 2016, 6, 31872. [Google Scholar] [CrossRef]
Liu, L.; Huang, M. Essential role of the iron-sulfur cluster binding domain of the primase regulatory subunit Pri2 in DNA replication initiation. Protein Cell 2015, 6, 194–210. [Google Scholar] [CrossRef]
Baranovskiy, A.G.; Zhang, Y.; Suwa, Y.; Babayeva, N.D.; Gu, J.; Pavlov, Y.I.; Tahirov, T.H. Crystal structure of the human primase. J. Biol. Chem. 2015, 290, 5635–5646. [Google Scholar] [CrossRef] [PubMed]
Sauguet, L.; Klinge, S.; Perera, R.L.; Maman, J.D.; Pellegrini, L. Shared active site architecture between the large subunit of eukaryotic primase and DNA photolyase. PLoS ONE 2010, 5, e10083. [Google Scholar] [CrossRef] [PubMed]
Agarkar, V.B.; Babayeva, N.D.; Pavlov, Y.I.; Tahirov, T.H. Crystal structure of the C-terminal domain of human DNA primase large subunit: Implications for the mechanism of the primase-polymerase α switch. Cell Cycle 2011, 10, 926–931. [Google Scholar] [CrossRef] [PubMed]
O’Brien, E.; Holt, M.E.; Thompson, M.K.; Salay, L.E.; Ehlinger, A.C.; Chazin, W.J.; Barton, J.K. The [4Fe4S] cluster of human DNA primase functions as a redox switch using DNA charge transport. Science 2017, 355, eaag1789. [Google Scholar] [CrossRef]
O’Brien, E.; Salay, L.E.; Epum, E.A.; Friedman, K.L.; Chazin, W.J.; Barton, J.K. Yeast require redox switching in DNA primase. Proc. Natl. Acad. Sci. USA 2018, 115, 13186–13191. [Google Scholar] [CrossRef]
Amin, M.; Brooks, B.R. The oxidation of the [4Fe-4S] cluster of DNA primase alters the binding energies with DNA and RNA primers. Biophys. J. 2024, 123, 1648–1653. [Google Scholar] [CrossRef]
O’Brien, E.; Holt, M.E.; Salay, L.E.; Chazin, W.J.; Barton, J.K. Substrate binding regulates redox signaling in human DNA primase. J. Am. Chem. Soc. 2018, 140, 17153–17162. [Google Scholar] [CrossRef]
Mancias, J.D.; Wang, X.; Gygi, S.P.; Harper, J.W.; Kimmelman, A.C. Quantitative proteomics identifies NCOA4 as the cargo receptor mediating ferritinophagy. Nature 2014, 509, 105–109. [Google Scholar] [CrossRef]
Dowdle, W.E.; Nyfeler, B.; Nagel, J.; Elling, R.A.; Liu, S.; Triantafellow, E.; Menon, S.; Wang, Z.; Honda, A.; Pardee, G. Selective VPS34 inhibitor blocks autophagy and uncovers a role for NCOA4 in ferritin degradation and iron homeostasis in vivo. Nat. Cell Biol. 2014, 16, 1069–1079. [Google Scholar] [CrossRef]
Singleton, M.R.; Dillingham, M.S.; Wigley, D.B. Structure and mechanism of helicases and nucleic acid translocases. Annu. Rev. Biochem. 2007, 76, 23–50. [Google Scholar] [CrossRef]
Wu, Y.; Brosh, R.M., Jr. DNA helicase and helicase–nuclease enzymes with a conserved iron–sulfur cluster. Nucleic Acids Res. 2012, 40, 4247–4260. [Google Scholar] [CrossRef] [PubMed]
Yeeles, J.T.; Cammack, R.; Dillingham, M.S. An iron-sulfur cluster is essential for the binding of broken DNA by AddAB-type helicase-nucleases. J. Biol. Chem. 2009, 284, 7746–7755. [Google Scholar] [CrossRef] [PubMed]
Zheng, L.; Zhou, M.; Guo, Z.; Lu, H.; Qian, L.; Dai, H.; Qiu, J.; Yakubovskaya, E.; Bogenhagen, D.F.; Demple, B. Human DNA2 is a mitochondrial nuclease/helicase for efficient processing of DNA replication and repair intermediates. Mol. Cell 2008, 32, 325–336. [Google Scholar] [CrossRef] [PubMed]
Estep, K.N.; Brosh, R.M., Jr. RecQ and Fe–S helicases have unique roles in DNA metabolism dictated by their unwinding directionality, substrate specificity, and protein interactions. Biochem. Soc. Trans. 2018, 46, 77–95. [Google Scholar] [CrossRef]
Matsuzaki, K.; Borel, V.; Adelman, C.A.; Schindler, D.; Boulton, S.J. FANCJ suppresses microsatellite instability and lymphomagenesis independent of the Fanconi anemia pathway. Genes Dev. 2015, 29, 2532–2546. [Google Scholar] [CrossRef]
Capo-Chichi, J.M.; Bharti, S.K.; Sommers, J.A.; Yammine, T.; Chouery, E.; Patry, L.; Rouleau, G.A.; Samuels, M.E.; Hamdan, F.F.; Michaud, J.L. Identification and biochemical characterization of a novel mutation in DDX11 causing W arsaw breakage syndrome. Hum. Mutat. 2013, 34, 103–107. [Google Scholar] [CrossRef]
Deng, Z.; Glousker, G.; Molczan, A.; Fox, A.J.; Lamm, N.; Dheekollu, J.; Weizman, O.-E.; Schertzer, M.; Wang, Z.; Vladimirova, O. Inherited mutations in the helicase RTEL1 cause telomere dysfunction and Hoyeraal–Hreidarsson syndrome. Proc. Natl. Acad. Sci. USA 2013, 110, E3408–E3416. [Google Scholar] [CrossRef]
Falquet, B.; Ölmezer, G.; Enkner, F.; Klein, D.; Challa, K.; Appanah, R.; Gasser, S.M.; Rass, U. Disease-associated DNA2 nuclease–helicase protects cells from lethal chromosome under-replication. Nucleic Acids Res. 2020, 48, 7265–7278. [Google Scholar] [CrossRef]
Sommers, J.A.; Rawtani, N.; Gupta, R.; Bugreev, D.V.; Mazin, A.V.; Cantor, S.B.; Brosh, R.M. FANCJ uses its motor ATPase to destabilize protein-DNA complexes, unwind triplexes, and inhibit RAD51 strand exchange. J. Biol. Chem. 2009, 284, 7505–7517. [Google Scholar] [CrossRef]
Levran, O.; Attwooll, C.; Henry, R.T.; Milton, K.L.; Neveling, K.; Rio, P.; Batish, S.D.; Kalb, R.; Velleuer, E.; Barral, S. The BRCA1-interacting helicase BRIP1 is deficient in Fanconi anemia. Nat. Genet. 2005, 37, 931–933. [Google Scholar] [CrossRef]
Odermatt, D.C.; Lee, W.T.C.; Wild, S.; Jozwiakowski, S.K.; Rothenberg, E.; Gari, K. Cancer-associated mutations in the iron-sulfur domain of FANCJ affect G-quadruplex metabolism. PLoS Genet. 2020, 16, e1008740. [Google Scholar] [CrossRef] [PubMed]
Bharti, S.K.; Sommers, J.A.; George, F.; Kuper, J.; Hamon, F.; Shin-ya, K.; Teulade-Fichou, M.-P.; Kisker, C.; Brosh, R.M. Specialization among iron-sulfur cluster helicases to resolve G-quadruplex DNA structures that threaten genomic stability. J. Biol. Chem. 2013, 288, 28217–28229. [Google Scholar] [CrossRef] [PubMed]
Wulfridge, P.; Sarma, K. Intertwining roles of R-loops and G-quadruplexes in DNA repair, transcription and genome organization. Nat. Cell Biol. 2024, 26, 1025–1036. [Google Scholar] [CrossRef] [PubMed]
Hänsel-Hertsch, R.; Di Antonio, M.; Balasubramanian, S. DNA G-quadruplexes in the human genome: Detection, functions and therapeutic potential. Nat. Rev. Mol. Cell Biol. 2017, 18, 279–284. [Google Scholar] [CrossRef]
Wu, Y.; Shin-ya, K.; Brosh, R.M., Jr. FANCJ helicase defective in Fanconia anemia and breast cancer unwinds G-quadruplex DNA to defend genomic stability. Mol. Cell. Biol. 2008, 28, 4116–4128. [Google Scholar] [CrossRef]
London, T.B.; Barber, L.J.; Mosedale, G.; Kelly, G.P.; Balasubramanian, S.; Hickson, I.D.; Boulton, S.J.; Hiom, K. FANCJ is a structure-specific DNA helicase associated with the maintenance of genomic G/C tracts. J. Biol. Chem. 2008, 283, 36132–36139. [Google Scholar] [CrossRef]
Wu, C.G.; Spies, M. G-quadruplex recognition and remodeling by the FANCJ helicase. Nucleic Acids Res. 2016, 44, 8742–8753. [Google Scholar] [CrossRef]
Vannier, J.-B.; Sandhu, S.; Petalcorin, M.I.; Wu, X.; Nabi, Z.; Ding, H.; Boulton, S.J. RTEL1 is a replisome-associated helicase that promotes telomere and genome-wide replication. Science 2013, 342, 239–242. [Google Scholar] [CrossRef]
Uringa, E.-J.; Youds, J.L.; Lisaingo, K.; Lansdorp, P.M.; Boulton, S.J. RTEL1: An essential helicase for telomere maintenance and the regulation of homologous recombination. Nucleic Acids Res. 2011, 39, 1647–1655. [Google Scholar] [CrossRef]
Wu, W.; Bhowmick, R.; Vogel, I.; Özer, Ö.; Ghisays, F.; Thakur, R.S.; Sanchez de Leon, E.; Richter, P.H.; Ren, L.; Petrini, J.H. RTEL1 suppresses G-quadruplex-associated R-loops at difficult-to-replicate loci in the human genome. Nat. Struct. Mol. Biol. 2020, 27, 424–437. [Google Scholar] [CrossRef]
Ghisays, F.; Garzia, A.; Wang, H.; Canasto-Chibuque, C.; Hohl, M.; Savage, S.A.; Tuschl, T.; Petrini, J.H. RTEL1 influences the abundance and localization of TERRA RNA. Nat. Commun. 2021, 12, 3016. [Google Scholar] [CrossRef] [PubMed]
Wu, Y.; Sommers, J.A.; Khan, I.; de Winter, J.P.; Brosh, R.M. Biochemical characterization of Warsaw breakage syndrome helicase. J. Biol. Chem. 2012, 287, 1007–1021. [Google Scholar] [CrossRef] [PubMed]
van Schie, J.J.; Faramarz, A.; Balk, J.A.; Stewart, G.S.; Cantelli, E.; Oostra, A.B.; Rooimans, M.A.; Parish, J.L.; de Almeida Estéves, C.; Dumic, K. Warsaw Breakage Syndrome associated DDX11 helicase resolves G-quadruplex structures to support sister chromatid cohesion. Nat. Commun. 2020, 11, 4287. [Google Scholar] [CrossRef] [PubMed]
Gray, L.T.; Vallur, A.C.; Eddy, J.; Maizels, N. G quadruplexes are genomewide targets of transcriptional helicases XPB and XPD. Nat. Chem. Biol. 2014, 10, 313–318. [Google Scholar] [CrossRef] [PubMed]
Kou, H.; Zhou, Y.; Gorospe, R.C.; Wang, Z. Mms19 protein functions in nucleotide excision repair by sustaining an adequate cellular concentration of the TFIIH component Rad3. Proc. Natl. Acad. Sci. USA 2008, 105, 15714–15719. [Google Scholar] [CrossRef]
Seroz, T.; Winkler, G.S.; Auriol, J.; Verhage, R.A.; Vermeulen, W.; Smit, B.; Brouwer, J.; Eker, A.P.; Weeda, G.; Egly, J.-M. Cloning of a human homolog of the yeast nucleotide excision repair gene MMS19 and interaction with transcription repair factor TFIIH via the XPB and XPD helicases. Nucleic Acids Res. 2000, 28, 4506–4513. [Google Scholar] [CrossRef]
Ding, H.; Schertzer, M.; Wu, X.; Gertsenstein, M.; Selig, S.; Kammori, M.; Pourvali, R.; Poon, S.; Vulto, I.; Chavez, E. Regulation of murine telomere length by Rtel: An essential gene encoding a helicase-like protein. Cell 2004, 117, 873–886. [Google Scholar] [CrossRef]
Askree, S.H.; Yehuda, T.; Smolikov, S.; Gurevich, R.; Hawk, J.; Coker, C.; Krauskopf, A.; Kupiec, M.; McEachern, M.J. A genome-wide screen for Saccharomyces cerevisiae deletion mutants that affect telomere length. Proc. Natl. Acad. Sci. USA 2004, 101, 8658–8663. [Google Scholar] [CrossRef]
Friedberg, E.C.; Walker, G.C.; Siede, W.; Wood, R.D. DNA Repair and Mutagenesis; American Society for Microbiology Press: Washington, DC, USA, 2005. [Google Scholar]
Knijnenburg, T.A.; Wang, L.; Zimmermann, M.T.; Chambwe, N.; Gao, G.F.; Cherniack, A.D.; Fan, H.; Shen, H.; Way, G.P.; Greene, C.S. Genomic and molecular landscape of DNA damage repair deficiency across the cancer genome atlas. Cell Rep. 2018, 23, 239–254.e236. [Google Scholar] [CrossRef]
Fuss, J.O.; Tsai, C.-L.; Ishida, J.P.; Tainer, J.A. Emerging critical roles of Fe–S clusters in DNA replication and repair. Biochim. Et Biophys. Acta (BBA)-Mol. Cell Res. 2015, 1853, 1253–1271. [Google Scholar] [CrossRef]
Cunningham, R.P.; Asahara, H.; Bank, J.F.; Scholes, C.P.; Salerno, J.C.; Surerus, K.; Munck, E.; McCracken, J.; Peisach, J.; Emptage, M.H. Endonuclease III is an iron-sulfur protein. Biochemistry 1989, 28, 4450–4455. [Google Scholar] [CrossRef] [PubMed]
Hinks, J.A.; Evans, M.C.; De Miguel, Y.; Sartori, A.A.; Jiricny, J.; Pearl, L.H. An iron-sulfur cluster in the family 4 uracil-DNA glycosylases. J. Biol. Chem. 2002, 277, 16936–16940. [Google Scholar] [CrossRef] [PubMed]
Parker, A.; Gu, Y.; Lu, A.-L. Purification and characterization of a mammalian homolog of Escherichia coli MutY mismatch repair protein from calf liver mitochondria. Nucleic Acids Res. 2000, 28, 3206–3215. [Google Scholar] [CrossRef] [PubMed]
Jacobs, A.L.; Schär, P. DNA glycosylases: In DNA repair and beyond. Chromosoma 2012, 121, 1–20. [Google Scholar] [CrossRef]
McGoldrick, J.P.; Yeh, Y.-C.; Solomon, M.; Essigmann, J.M.; Lu, A.-L. Characterization of a mammalian homolog of the Escherichia coli MutY mismatch repair protein. Mol. Cell. Biol. 1995, 15, 989–996. [Google Scholar] [CrossRef]
Aspinwall, R.; Rothwell, D.G.; Roldan-Arjona, T.; Anselmino, C.; Ward, C.J.; Cheadle, J.P.; Sampson, J.R.; Lindahl, T.; Harris, P.C.; Hickson, I.D. Cloning and characterization of a functional human homolog of Escherichia coli endonuclease III. Proc. Natl. Acad. Sci. USA 1997, 94, 109–114. [Google Scholar] [CrossRef]
Lukianova, O.A.; David, S.S. A role for iron–sulfur clusters in DNA repair. Curr. Opin. Chem. Biol. 2005, 9, 145–151. [Google Scholar] [CrossRef]
Trasviña-Arenas, C.H.; Lopez-Castillo, L.M.; Sanchez-Sandoval, E.; Brieba, L.G. Dispensability of the [4Fe-4S] cluster in novel homologues of adenine glycosylase MutY. FEBS J. 2016, 283, 521–540. [Google Scholar] [CrossRef]
Guan, Y.; Manuel, R.C.; Arvai, A.S.; Parikh, S.S.; Mol, C.D.; Miller, J.H.; Lloyd, R.S.; Tainer, J.A. MutY catalytic core, mutant and bound adenine structures define specificity for DNA repair enzyme superfamily. Nat. Struct. Biol. 1998, 5, 1058–1064. [Google Scholar] [CrossRef]
Boal, A.K.; Yavin, E.; Lukianova, O.A.; O’Shea, V.L.; David, S.S.; Barton, J.K. DNA-bound redox activity of DNA repair glycosylases containing [4Fe-4S] clusters. Biochemistry 2005, 44, 8397–8407. [Google Scholar] [CrossRef]
Romano, C.A.; Sontz, P.A.; Barton, J.K. Mutants of the base excision repair glycosylase, endonuclease III: DNA charge transport as a first step in lesion detection. Biochemistry 2011, 50, 6133–6145. [Google Scholar] [CrossRef] [PubMed]
Porello, S.L.; Cannon, M.J.; David, S.S. A substrate recognition role for the [4Fe-4S] 2+ cluster of the DNA repair glycosylase MutY. Biochemistry 1998, 37, 6465–6475. [Google Scholar] [CrossRef] [PubMed]
Bartels, P.L.; Zhou, A.; Arnold, A.R.; Nuñez, N.N.; Crespilho, F.N.; David, S.S.; Barton, J.K. Electrochemistry of the [4Fe4S] cluster in base excision repair proteins: Tuning the redox potential with DNA. Langmuir 2017, 33, 2523–2530. [Google Scholar] [CrossRef] [PubMed]
Tse, E.C.; Zwang, T.J.; Barton, J.K. The oxidation state of [4Fe4S] clusters modulates the DNA-binding affinity of DNA repair proteins. J. Am. Chem. Soc. 2017, 139, 12784–12792. [Google Scholar] [CrossRef]
Bruner, S.D.; Norman, D.P.; Verdine, G.L. Structural basis for recognition and repair of the endogenous mutagen 8-oxoguanine in DNA. Nature 2000, 403, 859–866. [Google Scholar] [CrossRef]
Poor, C.B.; Wegner, S.V.; Li, H.; Dlouhy, A.C.; Schuermann, J.P.; Sanishvili, R.; Hinshaw, J.R.; Riggs-Gelasco, P.J.; Outten, C.E.; He, C. Molecular mechanism and structure of the Saccharomyces cerevisiae iron regulator Aft2. Proc. Natl. Acad. Sci. USA 2014, 111, 4043–4048. [Google Scholar] [CrossRef]
Ueta, R.; Fujiwara, N.; Iwai, K.; Yamaguchi-Iwai, Y. Iron-induced dissociation of the Aft1p transcriptional regulator from target gene promoters is an initial event in iron-dependent gene suppression. Mol. Cell Biol. 2012, 32, 4998–5008. [Google Scholar] [CrossRef]
Li, H.; Mapolelo, D.T.; Dingra, N.N.; Naik, S.G.; Lees, N.S.; Hoffman, B.M.; Riggs-Gelasco, P.J.; Huynh, B.H.; Johnson, M.K.; Outten, C.E. The yeast iron regulatory proteins Grx3/4 and Fra2 form heterodimeric complexes containing a [2Fe-2S] cluster with cysteinyl and histidyl ligation. Biochemistry 2009, 48, 9569–9581. [Google Scholar] [CrossRef]
Mercier, A.; Watt, S.; Bähler, J.; Labbé, S. Key function for the CCAAT-binding factor Php4 to regulate gene expression in response to iron deficiency in fission yeast. Eukaryot. Cell 2008, 7, 493–508. [Google Scholar] [CrossRef]
Vachon, P.; Mercier, A.; Jbel, M.; Labbé, S. The monothiol glutaredoxin Grx4 exerts an iron-dependent inhibitory effect on Php4 function. Eukaryot. Cell 2012, 11, 806–819. [Google Scholar] [CrossRef]
Dlouhy, A.C.; Beaudoin, J.; Labbé, S.; Outten, C.E. Schizosaccharomyces pombe Grx4 regulates the transcriptional repressor Php4 via [2Fe–2S] cluster binding. Metallomics 2017, 9, 1096–1105. [Google Scholar] [CrossRef] [PubMed]
Hati, D.; Brault, A.; Gupta, M.; Fletcher, K.; Jacques, J.F.; Labbe, S.; Outten, C.E. Iron homeostasis proteins Grx4 and Fra2 control activity of the Schizosaccharomyces pombe iron repressor Fep1 by facilitating [2Fe-2S] cluster removal. J. Biol. Chem. 2023, 299, 105419. [Google Scholar] [CrossRef] [PubMed]
Jacques, J.F.; Mercier, A.; Brault, A.; Mourer, T.; Labbe, S. Fra2 is a co-regulator of Fep1 inhibition in response to iron starvation. PLoS ONE 2014, 9, e98959. [Google Scholar] [CrossRef] [PubMed]
Pelletier, B.; Trott, A.; Morano, K.A.; Labbé, S. Functional characterization of the iron-regulatory transcription factor Fep1 from Schizosaccharomyces pombe. J. Biol. Chem. 2005, 280, 25146–25161. [Google Scholar] [CrossRef]
Ogunjimi, B.; Zhang, S.-Y.; Sørensen, K.B.; Skipper, K.A.; Carter-Timofte, M.; Kerner, G.; Luecke, S.; Prabakaran, T.; Cai, Y.; Meester, J. Inborn errors in RNA polymerase III underlie severe varicella zoster virus infections. J. Clin. Investig. 2017, 127, 3543–3556. [Google Scholar] [CrossRef]
Girbig, M.; Misiaszek, A.D.; Vorländer, M.K.; Lafita, A.; Grötsch, H.; Baudin, F.; Bateman, A.; Müller, C.W. Cryo-EM structures of human RNA polymerase III in its unbound and transcribing states. Nat. Struct. Mol. Biol. 2021, 28, 210–219. [Google Scholar] [CrossRef]
Wang, Q.; Li, S.; Wan, F.; Xu, Y.; Wu, Z.; Cao, M.; Lan, P.; Lei, M.; Wu, J. Structural insights into transcriptional regulation of human RNA polymerase III. Nat. Struct. Mol. Biol. 2021, 28, 220–227. [Google Scholar] [CrossRef]
Wang, Z.; Roeder, R.G. Three human RNA polymerase III-specific subunits form a subcomplex with a selective function in specific transcription initiation. Genes Dev. 1997, 11, 1315–1326. [Google Scholar] [CrossRef]
Chiu, Y.-H.; MacMillan, J.B.; Chen, Z.J. RNA polymerase III detects cytosolic DNA and induces type I interferons through the RIG-I pathway. Cell 2009, 138, 576–591. [Google Scholar] [CrossRef]
Tian, K.; Wang, R.; Huang, J.; Wang, H.; Ji, X. Subcellular localization shapes the fate of RNA polymerase III. Cell Rep. 2023, 42, 112941. [Google Scholar] [CrossRef]
Zurita, M.; Merino, C. The transcriptional complexity of the TFIIH complex. TRENDS Genet. 2003, 19, 578–584. [Google Scholar] [CrossRef] [PubMed]
Giglia-Mari, G.; Coin, F.; Ranish, J.A.; Hoogstraten, D.; Theil, A.; Wijgers, N.; Jaspers, N.G.; Raams, A.; Argentini, M.; Van Der Spek, P. A new, tenth subunit of TFIIH is responsible for the DNA repair syndrome trichothiodystrophy group A. Nat. Genet. 2004, 36, 714–719. [Google Scholar] [CrossRef] [PubMed]
Keriel, A.; Stary, A.; Sarasin, A.; Rochette-Egly, C.; Egly, J.-M. XPD mutations prevent TFIIH-dependent transactivation by nuclear receptors and phosphorylation of RARα. Cell 2002, 109, 125–135. [Google Scholar] [CrossRef] [PubMed]
Dubaele, S.; De Santis, L.P.; Bienstock, R.J.; Keriel, A.; Stefanini, M.; Van Houten, B.; Egly, J.-M. Basal transcription defect discriminates between xeroderma pigmentosum and trichothiodystrophy in XPD patients. Mol. Cell 2003, 11, 1635–1646. [Google Scholar] [CrossRef] [PubMed]
Compe, E.; Drané, P.; Laurent, C.; Diderich, K.; Braun, C.; Hoeijmakers, J.H.; Egly, J.-M. Dysregulation of the peroxisome proliferator-activated receptor target genes by XPD mutations. Mol. Cell. Biol. 2005, 25, 6065–6076. [Google Scholar] [CrossRef]
Abbassi, N.-e.-H.; Jaciuk, M.; Scherf, D.; Böhnert, P.; Rau, A.; Hammermeister, A.; Rawski, M.; Indyka, P.; Wazny, G.; Chramiec-Głąbik, A. Cryo-EM structures of the human Elongator complex at work. Nat. Commun. 2024, 15, 4094. [Google Scholar] [CrossRef]
Selvadurai, K.; Wang, P.; Seimetz, J.; Huang, R.H. Archaeal Elp3 catalyzes tRNA wobble uridine modification at C5 via a radical mechanism. Nat. Chem. Biol. 2014, 10, 810–812. [Google Scholar] [CrossRef]
Dalwadi, U.; Yip, C.K. Structural insights into the function of Elongator. Cell. Mol. Life Sci. 2018, 75, 1613–1622. [Google Scholar] [CrossRef]
Maio, N.; Orbach, R.; Zaharieva, I.T.; Töpf, A.; Donkervoort, S.; Munot, P.; Mueller, J.; Willis, T.; Verma, S.; Peric, S. CIAO1 loss of function causes a neuromuscular disorder with compromise of nucleocytoplasmic Fe-S enzymes. J. Clin. Investig. 2024, 134, e179559. [Google Scholar] [CrossRef]
Romero, A.M.; Martinez-Pastor, M.T.; Puig, S. Iron in Translation: From the Beginning to the End. Microorganisms 2021, 9, 1058. [Google Scholar] [CrossRef]
Sun, C.; Guo, R.; Ye, X.; Tang, S.; Chen, M.; Zhou, P.; Yang, M.; Liao, C.; Li, H.; Lin, B. Wybutosine hypomodification of tRNAphe activates HERVK and impairs neuronal differentiation. Iscience 2024, 27, 109748. [Google Scholar] [CrossRef] [PubMed]
Kessler, A.C.; Silveira d’Almeida, G.; Alfonzo, J.D. The role of intracellular compartmentalization on tRNA processing and modification. RNA Biol. 2018, 15, 554–566. [Google Scholar] [CrossRef] [PubMed]
Wei, F.Y.; Zhou, B.; Suzuki, T.; Miyata, K.; Ujihara, Y.; Horiguchi, H.; Takahashi, N.; Xie, P.; Michiue, H.; Fujimura, A.; et al. Cdk5rap1-mediated 2-methylthio modification of mitochondrial tRNAs governs protein translation and contributes to myopathy in mice and humans. Cell Metab. 2015, 21, 428–442. [Google Scholar] [CrossRef]
Sheng, L.; Hao, S.-L.; Yang, W.-X.; Sun, Y. The multiple functions of kinesin-4 family motor protein KIF4 and its clinical potential. Gene 2018, 678, 90–99. [Google Scholar] [CrossRef]
Lee, Y.M.; Kim, W. Kinesin superfamily protein member 4 (KIF4) is localized to midzone and midbody in dividing cells. Exp. Mol. Med. 2004, 36, 93–97. [Google Scholar] [CrossRef]
Mazumdar, M.; Sundareshan, S.; Misteli, T. Human chromokinesin KIF4A functions in chromosome condensation and segregation. J. Cell Biol. 2004, 166, 613–620. [Google Scholar] [CrossRef]
Zhu, C.; Jiang, W. Cell cycle-dependent translocation of PRC1 on the spindle by Kif4 is essential for midzone formation and cytokinesis. Proc. Natl. Acad. Sci. USA 2005, 102, 343–348. [Google Scholar] [CrossRef]
Kurasawa, Y.; Earnshaw, W.C.; Mochizuki, Y.; Dohmae, N.; Todokoro, K. Essential roles of KIF4 and its binding partner PRC1 in organized central spindle midzone formation. EMBO J. 2004, 23, 3237–3248. [Google Scholar] [CrossRef]
Mazumdar, M.; Sung, M.-H.; Misteli, T. Chromatin maintenance by a molecular motor protein. Nucleus 2011, 2, 591–600. [Google Scholar] [CrossRef]
Wu, G.; Zhou, L.; Khidr, L.; Guo, X.E.; Kim, W.; Lee, Y.M.; Krasieva, T.; Chen, P.-L. A novel role of the chromokinesin Kif4A in DNA damage response. Cell Cycle 2008, 7, 2013–2020. [Google Scholar] [CrossRef]
Kalantari, S.; Carlston, C.; Alsaleh, N.; Abdel-Salam, G.M.; Alkuraya, F.; Kato, M.; Matsumoto, N.; Miyatake, S.; Yamamoto, T.; Fares-Taie, L. Expanding the KIF4A-associated phenotype. Am. J. Med. Genet. Part A 2021, 185, 3728–3739. [Google Scholar] [CrossRef] [PubMed]
Gad, S.A.; Sugiyama, M.; Tsuge, M.; Wakae, K.; Fukano, K.; Oshima, M.; Sureau, C.; Watanabe, N.; Kato, T.; Murayama, A. The kinesin KIF4 mediates HBV/HDV entry through the regulation of surface NTCP localization and can be targeted by RXR agonists in vitro. PLoS Pathog. 2022, 18, e1009983. [Google Scholar] [CrossRef] [PubMed]
Mazumdar, M.; Misteli, T. Chromokinesins: Multitalented players in mitosis. Trends Cell Biol. 2005, 15, 349–355. [Google Scholar] [CrossRef]
Wu, G.; Chen, P.-L. Structural requirements of chromokinesin Kif4A for its proper function in mitosis. Biochem. Biophys. Res. Commun. 2008, 372, 454–458. [Google Scholar] [CrossRef]
Barton, J.K.; Silva, R.M.; O’Brien, E. Redox chemistry in the genome: Emergence of the [4Fe4S] cofactor in repair and replication. Annu. Rev. Biochem. 2019, 88, 163–190. [Google Scholar] [CrossRef]
Ueda, C.; Langton, M.; Chen, J.; Pandelia, M.-E. The HBx protein from hepatitis B virus coordinates a redox-active Fe-S cluster. J. Biol. Chem. 2022, 298, 101698. [Google Scholar] [CrossRef]
Henkler, F.; Hoare, J.; Waseem, N.; Goldin, R.D.; McGarvey, M.J.; Koshy, R.; King, I.A. Intracellular localization of the hepatitis B virus HBx protein. J. Gen. Virol. 2001, 82, 871–882. [Google Scholar] [CrossRef]

Figure 2. The essential roles of nuclear Fe-S cluster-binding proteins in DNA replication, telomere maintenance, and mitosis. (A) A representation of a eukaryotic DNA replication fork. The mini-chromosome maintenance (MCM2-7) helicase complex unwinds DNA at replication origins, separating the double helix to generate the leading and lagging strands. All replicative DNA polymerases, Pol α, δ, and ε, bind a [4Fe-4S] cluster. DNA replication initiates with an RNA primer (orange) synthesized by DNA primase, with the regulatory subunit PRIM2 also binding a [4Fe-4S] cluster. Pol ε further extends the primer on the leading strand, while Pol δ does so on the lagging strand. The proliferating cell nuclear antigen sliding clamp PCNA enhances the processivity of DNA polymerases and facilitates DNA repair. The [4Fe-4S] cluster DNA helicases FANCJ and DDX11 unwind G4s to prevent stalling of the replisome. DNA lesions obstruct replication, triggering monoubiquitylation of PCNA, which recruits the DNA damage bypass DNA polymerase Pol ζ. The catalytic subunit of Pol ζ also binds a [4Fe-4S] cluster. Finally, the [4Fe-4S] cluster nuclease–helicase DNA2 cleaves exposed single-stranded DNA ends from Okazaki fragments and stalled replication forks. (B) Under Fe-sufficient conditions, the MCM helicase moves in opposite directions from the activated origin of replication. This results in the formation of two functional replication forks, enabling DNA synthesis. Under iron-depleted conditions, apo NCOA4 accumulates in the nucleus and binds to the helicase subunit MCM7, hindering MCM helicase activity and inactivating replication origins. This ensures that DNA synthesis only occurs when there is a sufficient pool of metallated Fe-S enzymes, thereby maintaining genomic integrity and preventing replication under suboptimal conditions. (C) The [4Fe-4S] cluster helicase RTEL1 unwinds G4s and R-loops at the telomeric repeats, facilitating telomere replication by telomerase and preventing telomere shortening. (D) During the late stages of mitosis (metaphase, anaphase, and telophase), the nuclear [4Fe-4S] cluster mitotic factor KIF4A binds the arms of condensed chromosomes. As a kinesin, KIF4A moves along microtubules to mobilize cargoes. KIF4A localizes at the central spindle, accumulating at the spindle midzone and midbody. The close-up image on the bottom illustrates the colocalization of the CIA targeting factors MMS19 and CIAO2B with KIF4A during mitosis, suggesting in situ metallation of KIF4A during function.

Figure 3. The Fe-S cluster proteins within the six major DNA repair pathways: base excision repair (BER), nucleotide excision repair (NER), mismatch repair (MMR), homologous recombination (HR), non-homologous end joining (NHEJ), and translesion DNA synthesis (TLS). A schematic representation of DNA lesions and the corresponding repair pathways. The key proteins involved in each pathway are organized according Knijnenburg et al. [172]. DNA repair proteins that bind [4Fe-4S] clusters are shown in a blue square at the bottom. * While DDX11 and EXO5 are not formally categorized under HR, their functions align with this repair mechanism.

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Novoa-Aponte, L.; Leon-Torres, A.; Philpott, C.C. Guardians of the Genome: Iron–Sulfur Proteins in the Nucleus. Inorganics 2024, 12, 316. https://doi.org/10.3390/inorganics12120316

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Novoa-Aponte L, Leon-Torres A, Philpott CC. Guardians of the Genome: Iron–Sulfur Proteins in the Nucleus. Inorganics. 2024; 12(12):316. https://doi.org/10.3390/inorganics12120316

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Novoa-Aponte, Lorena, Andres Leon-Torres, and Caroline C. Philpott. 2024. "Guardians of the Genome: Iron–Sulfur Proteins in the Nucleus" Inorganics 12, no. 12: 316. https://doi.org/10.3390/inorganics12120316

APA Style

Novoa-Aponte, L., Leon-Torres, A., & Philpott, C. C. (2024). Guardians of the Genome: Iron–Sulfur Proteins in the Nucleus. Inorganics, 12(12), 316. https://doi.org/10.3390/inorganics12120316

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