Abstract
Cellular iron homeostasis is vital and maintained through tight regulation of iron import, efflux, storage and detoxification1,2,3. The most common modes of iron storage use proteinaceous compartments, such as ferritins and related proteins4,5. Although lipid-bounded iron compartments have also been described, the basis for their formation and function remains unknown6,7. Here we focus on one such compartment, herein named the ‘ferrosome’, that was previously observed in the anaerobic bacterium Desulfovibrio magneticus6. Using a proteomic approach, we identify three ferrosome-associated (Fez) proteins that are responsible for forming ferrosomes in D. magneticus. Fez proteins are encoded in a putative operon and include FezB, a P1B-6-ATPase found in phylogenetically and metabolically diverse species of bacteria and archaea. We show that two other bacterial species, Rhodopseudomonas palustris and Shewanella putrefaciens, make ferrosomes through the action of their six-gene fez operon. Additionally, we find that fez operons are sufficient for ferrosome formation in foreign hosts. Using S. putrefaciens as a model, we show that ferrosomes probably have a role in the anaerobic adaptation to iron starvation. Overall, this work establishes ferrosomes as a new class of iron storage organelles and sets the stage for studying their formation and structure in diverse microorganisms.
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Data availability
The mass spectrometry proteomics data have been deposited to the ProteomeXchange Consortium via the PRIDE67 partner repository with dataset identifier PXD017470. Ferrosome-associated proteins presented in Fig. 1a were identified from the data in Supplementary Table 11. The sequences, alignment and tree data used to generate Fig. 1g are provided as Supplementary Data 2. KEGG60 and IMG/M ER52 were used to collect data.
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Acknowledgements
We thank faculty at the EM-Lab at the University of California, Berkeley, for their assistance with TEM; A. Campos for assistance with the EDS measurements; M. Pang for assistance in culturing R. palustris for ferrosome production; K. LeGault and H. McCausland for their help with conjugations in R. palustris and M. magneticum; J. Gralnick (University of Minnesota) for providing S. putrefaciens strain CN-32; and the Coates laboratory and the Niyogi laboratory for sharing equipment. The QB3/Chemistry Mass Spectrometry Facility at the University of California, Berkeley, received support from the National Institutes of Health (shared instrumentation grant 1S10OD020062-01). Research reported in this publication was supported by funding from the National Institutes of Health (R01GM084122 and R35GM127114), the Office of Naval Research (N000141310421) and the Bakar Fellows Program. H.A.T. is supported by the National Science Foundation Graduate Research Fellowship Program under grant DGE 1752814. M.A. is supported by a grant through the Fondation pour la Recherche Médicale (ARF201909009123).
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C.R.G. and A.K. conceived and designed the study. C.R.G. performed all molecular cloning, genetic manipulation, TEM, cellular fractionations and sample preparations for LC–MS analyses. A.T.I. performed all LC–MS analyses. C.R.G. identified ferrosome-associated proteins with assistance from A.T.I. and H.A.T. C.R.G. carried out the bioinformatic analyses and tree construction. M.A. performed the EDS experiments and analysis. C.R.G. performed all growth assays with assistance from S.K. C.R.G. and A.K. prepared the manuscript with input from S.K., A.T.I., H.A.T. and M.A.
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Extended data figures and tables
Extended Data Fig. 1 Ferrosomes are visible by TEM in whole D. magneticus cells after transitioning from iron limited to iron replete conditions.
D. magneticus cells initially grown without iron (a) are shown 0.5 (b), 1.5 (c), and 6 (d) hours after addition of 100 μM ferric malate. (e) The maximum diameter of ferrosomes represented in b-d. Each data point represents one ferrosome and the bar indicates the mean maximum diameter in nm. Micrographs of D. magneticus one hour after adding low to high concentrations of ferric malate—1 μM (f), 10 μM (g), 100 μM (h), and 1 mM (i)—to iron-starved cells. (j) The maximum diameter of ferrosomes represented in f-i. Each data point represents one ferrosome and the bar indicates the mean maximum diameter in nm. Scale bars, 200 nm.
Extended Data Fig. 2 Isolation of ferrosomes and characteristics of associated proteins.
(a) Ferrosomes from ∆MAI D. magneticus (left) and magnetosomes from WT D. magneticus (right) form a pellet through 65% sucrose. Transmission electron micrographs of the ferrosome pellet (b) and the magnetosome pellet (c). Scale bars, 100 nm. (d-f) Membrane domain predictions of ferrosome-associated proteins in D. magneticus. DMR_28320 (a), DMR_28330 (b), and DMR_28340 (c) have 1, 5–6, and 0–2 putative transmembrane domains, respectively, as predicted by various methods analyzed through TOPCONS 1.065.
Extended Data Fig. 3 Multiple sequence alignment of FezB with characterized P1B-ATPases.
Conserved functional motifs in the actuator domain and the ATP-binding domain are indicated with blue and purple stars, respectively. The CxxC and histidine-rich metal binding sites in the cytoplasmic N-terminal domain of ZntA, CopA, and CopB are boxed. Transmembrane regions, predicted using TOPCONS 1.065, are underlined for each sequence. Putative metal-binding sites in the transmembrane domains are indicated with black stars.
Extended Data Fig. 4 WT and ∆fezBC D. magneticus strains make ferrosomes in iron replete medium when expressing fezABC in trans.
Transmission electron micrographs of WT (a) and ∆fezBC (b) strains with a control plasmid make magnetosomes (white carets) when grown in iron replete medium. When expressing fezABC in trans, both the WT (c) and ∆fezBC (d) strains make magnetosomes as well as ferrosomes when grown in iron replete medium. Areas of the cell containing one or more putative ferrosomes are indicated with yellow circles. Scale bars, 200 nm.
Extended Data Fig. 5 Sequence similarity network of proteins encoded by fez gene clusters and genes frequently found near fez gene clusters.
(a) Conserved fez gene clusters that encode FezB homologs. Conserved genes within the clusters are colored black. Gene clusters were identified using the “Gene cluster” tool in KEGG for each FezB homolog, in bold: Dde_0495, Dde_0498, Thimo_2900, vfu_A02104, SMUL_2748, RPA2333, KN400_3199, DMR_28330, and EUBELI_00578. The second copy of FezB in D. alaskensis, Dde_0498, is not shown because it is not part of a predicted conserved gene cluster. (b, c) Sequence similarity network highlighting the proteins encoded by ten genes upstream and downstream of 304 FezB homologs. Each node represents a protein and edges represent protein similarities that meet the specified e-value cutoff. (b) Network containing fez gene cluster-encoded proteins. Each group (labeled 1–8) contains one or more proteins encoded by conserved genes identified in (a) which are represented by black nodes and are labeled. Proteins or domains with an annotated function are labeled. Groups of proteins were further divided into subgroups which were used to identify proteins with GxxxG motifs in groups 2 and 5 and proteins with R-rich motifs in groups 1 and 3 (see Methods). The proteins represented in this network and their group/subgroup are listed in Supplementary Tables 3–6. (c) Network of proteins encoded by genes that are frequently found upstream and downstream of fez gene clusters. Only groups of more than 30 proteins are shown and the protein or domain annotation is labeled. Proteins with a known role in iron homeostasis are common and include iron transporters (FeoA, FeoB, outermembrane siderophore receptors, and some ABC transporters) and regulators (Fur and DtxR). The proteins represented in this network are listed in Supplementary Table 7.
Extended Data Fig. 6 Consensus motifs and characteristics of proteins with R-rich and GxxxG motifs.
Representative proteins encoded by fez gene clusters with (a) an R-rich motif or (b) a GxxxG motif. Logo shows the consensus motif for the subgroup or group of proteins to which the representative protein belongs. Predicted protein structure schematics show approximate location of the R-rich motif, putative transmembrane helices, and GxxxG motif for each protein (not to scale).
Extended Data Fig. 7 Transmission electron micrographs of S. putrefaciens and R. palustris.
WT S. putrefaciens (a, b) and R. palustris (c, d), ∆fezSp (e, f), ∆fezRp (g), ∆fezSp::fezSp (h, i), and ∆fezRp::fezRp (j). S. putrefaciens strains respiring fumarate in medium supplemented with 100 μM ferric malate (a, e, i) or 1 mM ferrous iron (b, f, j). R. palustris strains grown anaerobically (d, g, h) or aerobically (c). White arrows denote ferrosomes. Polyphosphate granules are indicated with white asterisks. Scale bars, 200 nm.
Extended Data Fig. 8 EDS spectra of S. putrefaciens and E. coli.
EDS spectrum of an S. putrefaciens ∆fezSp cell, which does not form ferrosomes. (b, c) EDS spectra of S. putrefaciens WT obtained from an area in the cell that contained ferrosomes (b) and an area that had no visible ferrosomes (c). The red asterisk indicates the iron peak associated with ferrosomes in WT S. putrefaciens. (d, e) Spectra of the background taken from areas of the S. putrefaciens WT (d) and ∆fezSp (e) grids that contained no cells. (f) An EDS spectrum of E. coli fezSp+ obtained from an area in the cell that had no visible ferrosomes. (g, h) Spectra of the background taken from areas of the E. coli cells with a control plasmid (g) or E. coli fezSp+ (h).
Extended Data Fig. 9 Effect of EDTA on the growth of S. putrefaciens.
(a) OD595 measurements over time of S. putrefaciens WT (navy) and ∆fezSp (yellow) grown aerobically with the indicated concentrations of EDTA. Each line is the mean of 3 individual cultures (technical replicates); error bars indicate s.d. (b) OD595 measurements over time of S. putrefaciens WT (navy) and ∆fezSp (yellow) grown anaerobically with the indicated concentrations of EDTA. Each line is the mean of 6 individual cultures (2 biological replicates with 3 technical replicates, with the exception of 150 μM EDTA which had 2 technical replicates); error bars indicate s.d. (c) Growth rate versus OD595 of the individual cultures shown in (b). Each circle represents the growth rate for an individual culture.
Supplementary information
Supplementary Table 1
P-type ATPases and their characteristics used to generate Fig. 2a.
Supplementary Table 2
Characteristics of bacteria and archaea with a FezB homologue.
Supplementary Table 3
Proteins belonging to group 1 (P-type ATPases).
Supplementary Table 4
Proteins belonging to group 2 (GXXXG motif).
Supplementary Table 5
Proteins belonging to group 3 (R-rich motif/DUF4405).
Supplementary Table 6
Other conserved proteins belonging to groups 4–8.
Supplementary Table 7
Proteins associated with fez gene clusters.
Supplementary Table 8
Bacterial strains used in this study.
Supplementary Table 9
Plasmids used in this study.
Supplementary Table 10
List of primers used in this study.
Supplementary Table 11
Relative protein quantification of proteins enriched with ferrosomes.
Supplementary Data 1
Raw data for S. putrefaciens growth assays used for Fig. 2e–g and Extended Data Fig. 9.
Supplementary Data 2
Sequences, alignment and tree data used to generate Fig. 1g.
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Grant, C.R., Amor, M., Trujillo, H.A. et al. Distinct gene clusters drive formation of ferrosome organelles in bacteria. Nature 606, 160–164 (2022). https://doi.org/10.1038/s41586-022-04741-x
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DOI: https://doi.org/10.1038/s41586-022-04741-x
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