Ecology

Quantification of changes in floristic similarityTo quantify changes in floristic similarity by naturalized flowering plant species, we extracted regional lists of alien species from the Global Naturalized Alien Flora (GloNAF) database45 and regional lists of native species from the Global Inventory of Floras and Traits (GIFT) database46. The GloNAF database contains lists of naturalized vascular plant taxa for 861 regions (countries or subnational administrative units), ranging in size from 0.03 to 6,864,961 km2 (median size is 15,152 km2) and covering >80% of the terrestrial ice-free surface globally47. GloNAF includes 13,803 plant taxa that, according to the original data sources, are alien plants and have established self-sustaining wild populations in the respective regions (i.e., are naturalized5). The GIFT database is a compilation of floras and checklists of predominantly native vascular plant species with an indication of their floristic status for more than 300,000 species across nearly 3000 regions with near global coverage46. We first selected regions that matched perfectly between GloNAF and GIFT. Additionally, we merged some GloNAF regions to match a larger GIFT region, and vice versa, by comparing the overlapping area of nested regions using the R package ‘sf’ (version 0.8-0)48.To ensure the highest data quality, and to be on the conservative side, we restricted our analysis to regions with complete or nearly complete checklists of both native and naturalized alien species. For GloNAF, we only included regions for which there was at least one species list judged to include more than 50% of the naturalized taxa for that region45. Although the judgment of species-list completeness is coarse and for most lists made by the GloNAF curators, it allows the exclusion of regions for which the data are obviously poor. For GIFT, we included a region only if at least one species list aimed to represent its entire native angiosperm flora. Our strict selection criteria resulted in a dataset including native and naturalized species for 658 non-overlapping regions, including 154 island regions, 503 mainland regions and one region including both islands and mainland areas (Chile). These regions covered all continents, except Antarctica, but there was low coverage for parts of Africa and Asia (Fig. 4).We restricted our analyses to flowering plants (angiosperms), which had the most complete species lists, and to species with accepted names in The Plant List24 (http://www.theplantlist.org/). We excluded species with an uncertain native/alien status or with a conflicting status, i.e., being native to a region according to GIFT but being alien to the same region according to GloNAF. Furthermore, since the native/alien status of many infraspecific taxa and hybrid taxa are less clear, we restricted our analyses to the species level (i.e., infraspecific taxa were assigned to the binomial species name), and we excluded hybrids. Our final dataset included 1,139,254 native species-by-region records for 189,110 species and 141,762 naturalized species-by-region records for 10,130 species.For all 216,153 possible pairwise combinations of the 658 regions, we quantified the taxonomic and phylogenetic similarities between their native floras (SimTaxnative, SimPhylnative), and between their floras including both native and naturalized alien species (SimTaxnative+naturalized, SimPhylnative+naturalized). As the regions vary largely in species richness (ranging from 11 to 13,720 species with a median of 1704), we used the Simpson similarity index for taxonomic similarity (Eq. 1)49, which is largely insensitive to species richness:50$${SimTax}=1-frac{{{min }}left(b,cright)}{a+{{min }}left(b,cright)}$$
(1)
Here a is the number of species common to both regions, b is the number of species that occur in the first region but not in the second and c is the number of species that occur in the second region but not in the first51. Likewise, we calculated the Simpson phylogenetic similarity index as phylogenetic similarity (Eq. 2) as implemented in the R package ‘betapart’ (version 1.5.1)52:$${SimPhyl}=1-frac{{{min }}left(B,Cright)}{A+{{min }}left(B,Cright)}$$
(2)
Here A is the total length of the phylogenetic branches in the phylogenetic tree that are shared by the species of both regions, B is the total length of the phylogenetic branches that are shared only by the first region and C is the total length of the phylogenetic branches that are shared only by the second region51. To quantify changes in similarity due to naturalization of alien species, we calculated the degree of homogenization H (or differentiation, see below) for each pair of regions as$$H={ln}frac{{{Sim}}_{{native}+{naturalized}}+0.001}{{{Sim}}_{{native}}+0.001}$$
(3)
A small value of 0.001 was added to both similarities to avoid infinite values. A positive log-response ratio indicates homogenization (i.e., increased floristic similarity between two regions), and a negative one indicates differentiation (i.e., decreased floristic similarity). As an alternative to the Simpson similarity index, we also calculate the Sørensen similarity index, which additionally takes into consideration the nestedness of the floras in the paired regions51. As the results were not sensitive to the choice of similarity indices (Supplementary Fig. 14), we focused our analyses on the Simpson similarity index.To quantify phylogenetic similarity, we used a phylogenetic tree including all angiosperms with accepted names in The Plant List (Supplementary Fig. 2). The tree was developed based on the mega phylogeny of Smith and Brown53. We added missing species (n = 71,124, of which 733 are naturalized in other regions) with their accepted names in The Plant List to the root of their genus or families. For details on the development of the phylogenetic tree, see ref. 47.Quantification of geographic distances and climatic distancesWe calculated the pairwise geographic distance between regions as the distance between their geographic centroids using the R package ‘geosphere’ (version 1.5-10)54. We also calculated the nearest distance between the geographic borders of regions. However, since the distances between geographic centroids are highly correlated with distances between region borders (n = 216,153, r = 0.996, P  More

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The binding of [2Fe–2S] and [3Fe–4S] in clMagRThree conserved cysteines (C60, C124, and C126) of clMagR in a CXnCGC sequence motif (n is 63–65 in most cases) play critical roles in iron–sulfur cluster binding18 (Fig. 1a), which has been further validated by alanines substitution mutant clMagR3M (C60A, C124A, and C126A mutation of clMagRWT). Strep-tagged clMagRWT and clMagR3M were freshly prepared (labeled as “as-isolated”) and purified to homogeneity under aerobic conditions. The clMagRWT protein showed brown color and clMagR3M appeared colorless in the solution, indicating the presence or absence of iron–sulfur cluster, respectively. Consistently, the Ultraviolet–visible (UV–Vis) spectrum of as-isolated clMagRWT showed absorption from 300-to-600-nm region, and with an absorption peak at 325 and 415 nm, and a shoulder at 470 nm, whereas these absorption peaks were abolished in clMagR3M (Fig. 1b). Circular dichroism (CD) spectroscopy was applied to characterize the types of iron–sulfur cluster and their protein environments during cluster maturation42,43,44. As shown in Fig. 1c, clMagRWT shows distinct positive peaks at 371 nm and 426 nm and three negative peaks at 324 nm, 396 nm, and 463 nm, respectively, suggesting the presence of [2Fe–2S] cluster45. However, it is worth pointing out that [4Fe–4S] or [3Fe–4S] clusters usually exhibit negligible CD intensity compared to [2Fe–2S] as shown previously in NifIscA45,46, thus CD spectroscopy cannot exclude the existence of [4Fe–4S] or [3Fe–4S]. Electron paramagnetic resonance (EPR) spectroscopy was then used to analyze different states of as-isolated clMagRWT. The oxidized clMagRWT was S = 1/2 species, characterized by a rhombic EPR signal with g values at g1 = 2.016, g2 = 2.002, and g3 = 1.997 (Fig. 1d) which disappeared at 45 K, suggesting the presence of [3Fe–4S]1+ cluster47,48. After reduced with sodium dithionite (Fig. 1e), EPR signal from [2Fe–2S] cluster can be observed until the temperature increased to 60K49,50,51. Thus, two distinct iron–sulfur clusters were assigned by EPR spectroscopy of clMagRWT. Figure 1Characterization of iron–sulfur clusters in as-isolated clMagR. (a) Sequence alignment of MagR in eight representative species. Predicted secondary structures are shown in the upper lines, with two alpha-helices (orange cylinders) and seven beta-strands (green arrows). Conserved residues with iron–sulfur cluster binding properties are shown in the red background (100% conserved), indicated by stars. Other conserved residues are shown in the gray background and bold fonts. Species’ common name, Latin name and sequence ID in NCBI are listed as follows: Pigeon (Columba livia), XP_005508102.1*; Zebra finch(Taeniopygia guttata), XP_002194930.1*; Fly(Drosophila melanogaster), NP_573062.1*; Monarch butterfly(Danaus plexippus), AVZ24723.1*; Salmon(Salmo salar), XP_013999046.1*; Octopus(Octopus bimaculoides), XP_014786756.1*; Little brown bat(Myotis lucifugus), XP_006102189.1*; Human(Homo sapiens), NP_112202.2*. (b) UV–Vis absorption spectrum of as-isolated pigeon MagR (clMagRWT, black) and C60AC124AC126A substitution mutant (clMagR3M, red), indicating three cysteines contribute to the iron–sulfur cluster binding. SDS-PAGEs of protein preparation are shown as inserts, theoretical mass of the clMagR monomer and clMagR3M monomer were 16.41 kDa, 16.31 kDa, respectively. (c) CD spectrum of as-isolated clMagRWT(black) and clMagR3M(red). (d, e) X-band EPR spectrum of as-isolated clMagRWT at oxidized (d) and reduced status (e). The samples were frozen in TBS buffer and the spectrums were recorded at various temperatures (10 K, 25 K, 45 K, 60 K). (f) Low-temperature resonance Raman spectra of as-isolated clMagRWT. Spectra were recorded at 17 K using 488 nm laser excitation.Full size imageConsidering some iron–sulfur clusters in proteins are diamagnetic and therefore EPR silent, low-temperature Resonance Raman (RR) spectroscopy was then utilized as a probe to characterize those clusters52. With 488 nm excitation, the RR spectra of clMagRWT in the iron–sulfur stretching region (240–450 cm−1) show the presence of [3Fe–4S]1+ cluster (represented by two bridging modes at 286 and 347 cm−1, and one terminal modes at 364 cm−1) and [2Fe–2S]2+ cluster (represented by three iron–sulfur bridging mode at 293, 308 and 330 cm−1 and two terminal modes at 407 and 422 cm−1, as shown in Fig. 1f)52,53,54,55,56. Taking together, we conclude that as-isolated clMagRWT contains both cystine-ligated [2Fe–2S] cluster and [3Fe–4S] cluster.The assembly and conversion of [2Fe–2S] and [3Fe–4S] in clMagRIron–sulfur cluster assembly of IscA, an clMagR homology protein in bacteria, is mediated by cysteine desulfurase IscS2. To elucidate how iron–sulfur cluster assembles in clMagR, time-course experiment was performed, and UV–Vis absorption and CD spectrum were used to monitor the IscS-catalyzed iron–sulfur cluster assembly in clMagR (Fig. 2). No signal of the iron–sulfur cluster was recorded when the reaction begins (0 min), and then the characteristic visible absorption peak and CD spectrum of clMagRWT appeared after 5 min, indicating [2Fe–2S] cluster assembled. As the reaction proceeds, the UV–Vis absorption intensity increased and after 180 min the signal was dominated by a broad shoulder centered at 415 nm (Fig. 2a). Concomitantly, the CD spectrum of the [2Fe–2S] center decreased and then almost disappeared after 180 min, indicating that [2Fe–2S] had been converted to [3Fe–4S] clusters and the reconstitution finished (Fig. 2b).Figure 2Iron–sulfur cluster assembly on clMagR. (a, b) IscS-mediated iron–sulfur cluster assembly on clMagR monitored as a function of time by UV–Vis absorption (a) and CD spectroscopy (b). The spectra shown were taken with samples of pretreated clMagR to remove iron–sulfur clusters before reconstitution (apo-clMagR, 0 min, light green), incubated with IscS after 5 min (green), and after 180 min (dark green). (c, d) chemical reconstitution-mediated iron–sulfur cluster assembly on clMagR monitored as a function of time by UV–Vis absorption (c) and CD spectroscopies (d). The spectra shown were taken with samples of pretreated clMagR to remove iron–sulfur clusters before reconstitution (apo-clMagR, light green) and chemically reconstituted clMagR (chem re clMagR, purple). (e) X-band EPR spectrum of chemically reconstituted clMagRWT. The spectrum was recorded at 10 K. (f) Low-temperature resonance Raman spectra of chemically reconstituted clMagR. Protein and reagent concentrations are described in the Methods. Spectra were recorded at 17 K using 488 nm laser excitation.Full size imageIron–sulfur cluster assembly can be achieved by chemical reconstitution as well, since iron–sulfur apo-proteins are able to spontaneously form iron–sulfur clusters in vitro when supplied with iron and sulfide under reducing conditions1,43,57. With this approach, started with apo-clMagRWT, we successfully reconstituted [3Fe–4S] cluster in clMagR protein, confirmed by UV–Vis absorption and CD spectrum result (Fig. 2c,d). To further validate if [3Fe–4S] is the sole type of iron–sulfur cluster in clMagR after chemical reconstitution, EPR and low-temperature Resonance Raman spectroscopy were applied (Fig. 2e,f). The chemically reconstituted clMagRWT was S = 1/2 species, characterized by a rhombic EPR signal with g values at g1 = 2.017, g2 = 2.002, and g3 = 1.994 (Fig. 2e). The signal is assigned to a S = 1/2 [3Fe–4S]1+ cluster. The Low-temperature Resonance Raman spectrum showed an intense band at 346 cm−1 and additional bands at 406 and 420 cm−1, which demonstrated that chemically reconstituted clMagRWT only contains [3Fe–4S]1+ cluster (Fig. 2f).We further investigated if clMagR could serve as an iron–sulfur carrier protein to accept [2Fe–2S] cluster from scaffold protein such as IscU58. Briefly, 400 µM holo-IscU was mixed with 400 µM strep-tagged apo-clMagRWT and incubated for 180 min under reduced condition, then, after desalting and strep-tactin affinity column separation, UV–Vis absorption and CD spectroscopy were applied the iron–sulfur cluster transfer process (Fig. 3a). The intensity of UV–Vis spectrum decreased in IscU (Fig. 3b) but significantly increased in clMagR after reaction (Fig. 3d), indicating [2Fe–2S] cluster was transferred from IscU to clMagR59. Consistently, CD spectrum of IscU and clMagR also confirmed that [2Fe–2S] transfer occurred between IscU and clMagR (Fig. 3c,e). The resulting spectrum is very similar to that of the [2Fe–2S] intermediate assembled on IscS mediated reconstituted apo-clMagR (Fig. 2b).Figure 3clMagR serve as carrier protein to accept [2Fe–2S] cluster from IscU in vitro. (a) A cartoon schematically illustrates the experimental procedures of in vitro iron–sulfur cluster transfer from IscU to clMagR. (b, c) The UV–Vis absorption (b) and CD spectra (c) of IscU. IscU protein samples were taken before mixing with apo-clMagR (holo-IscU, black lines) and after incubated with apo-clMagR for 180 min (pink lines). (d, e) The UV–Vis absorption (d) and CD spectra (e) of clMagR. clMagR samples were taken before mixing with holo-IscU (apo-clMagR, light green lines) and after incubated with holo-IscU for 180 min (holo-clMagR, brown lines).Full size imageCys-60 is essential for clMagR to bind [3Fe–4S] cluster, not [2Fe–2S] clusterThree conserved cysteines (C60, C124, and C126) of clMagR play critical roles in iron–sulfur cluster binding, and the substitute mutation of these three residues abolished iron–sulfur binding (Fig. 1b,c)18. To elucidate if three cysteines bind [2Fe–2S] and [3Fe–4S] differently, single Cys-to-Ala substitutions (C60A, C124A, and C126A) were made and their iron–sulfur binding properties were characterized.Freshly purified as-isolated clMagRC60A showed light brown color, and [2Fe–2S] cluster binding was verified by UV–Vis absorption and CD spectrum (Fig. 4a,b). A typical protein-bound [2Fe–2S] cluster absorption peak at 325 nm and a shoulder at 415 nm are visible in UV–Vis absorption (Fig. 4a, light orange line). Consistently, the CD spectrum of as-isolated clMagRC60A mutant had a negative peak at 397 nm and a positive peak at 451 nm (Fig. 4b, light orange line), confirmed the [2Fe–2S] cluster binding, similar to clMagRWT. However, in contrast to clMagRWT, chemical reconstitution failed to convert [2Fe–2S] cluster to [3Fe–4S] cluster in clMagRC60A. As shown in Fig. 4a,b (orange line), chemically reconstituted clMagRC60A showed similar and characteristic [2Fe–2S] UV–Vis absorption peaks and CD spectrum, but not [3Fe–4S] (Fig. 4a,b, orange lines), suggesting that C60A mutation abolished [3Fe–4S] cluster binding ability in clMagR.Figure 4Three conserved cysteines play different roles in iron–sulfur binding in clMagR. (a, b) Chemical reconstitution-mediated iron–sulfur cluster assembly on apo-clMagRC60A monitored by UV–Vis absorption (a) and CD spectroscopies (b). The samples of spectra shown are as-isolated clMagRC60A (light orange) and chemically reconstituted clMagRC60A (chem re clMagRC60A, orange). (c, d) chemical reconstitution-mediated iron–sulfur cluster assembly on clMagRC124A monitored by UV–Vis absorption (c) and CD spectroscopies (d). The samples of spectra shown are as-isolated clMagRC124A (light purple) and chemically reconstituted clMagRC124A (chem re clMagRC124A, purple). (e, f) chemical reconstitution-mediated iron–sulfur cluster assembly on pigeon clMagRC126A monitored by UV–Vis absorption (e) and CD spectroscopies (f). The samples of spectra shown are as-isolated clMagRC126A (light blue) and chemically reconstituted clMagRC126A (chem re clMagRC126A, blue). SDS-PAGE results were shown in the right of corresponding UV–Vis spectra as inserts (a, c, e). The theoretical mass of the clMagRC60A monomer, clMagRC124A monomer and clMagRC126A monomer were 16.38 kDa. (g, h) The UV–Vis absorption (c) and CD spectra (d) of clMagRC60A obtained by mixing apo-clMagRC60A and holo-IscU which was recorded before the addition of apo-clMagRC60A (dotted orange lines) and after incubation with apo-clMagRC60A for 180 min (orange lines). Protein and reagent concentrations are described in the Experimental procedures.Full size imageIn contrast, purified as-isolated clMagRC124A and clMagRC126A were colorless, and the binding of iron–sulfur clusters was barely detectable by UV–Vis and CD spectrum (Fig. 4c–f, light purple, and light blue lines, respectively). However, chemical reconstitution successfully reconstituted [3Fe–4S] cluster binding in both clMagRC124A and clMagRC126A (Fig. 4c–f, purple and blue lines, respectively). After chemical reconstitution, the UV–Vis absorption of both clMagRC124A and clMagRC126A mutants showed the signal of iron–sulfur cluster binding (Fig. 4c,e). Parallel CD spectrum studies confirmed both chemically reconstituted clMagRC124A and clMagRC126A have [3Fe–4S] cluster binding (Fig. 4d,f), similar to chemically reconstituted clMagRWT. The results demonstrated that Cys-124 and Cys-126 in clMagR play important roles in [2Fe–2S] cluster binding, thus, mutating these two residues lead to clMagR favors [3Fe–4S] binding.Considering clMagR can act as a carrier protein to accept iron–sulfur cluster from IscU (Fig. 3), it is worth testing if three cysteines play a different role in this process as well. Holo-IscU was mixed with apo-clMagR single cysteine mutants in a reduced state for 180 min. The apo status of all three mutants (labeled as apo-clMagRC60A, apo-clMagRC124A, and apo-clMagRC126A) had no iron–sulfur cluster binding before mixing with holo-IscU, as shown by negligible UV absorption and CD intensities (Fig. 4g,h and Supplementary Fig. 1a–d, dotted lines). After incubation with holo-IscU and separation of IscU and clMagR mutants, clMagRC60A showed distinct changes in UV–Vis absorption and CD spectrum (Fig. 4g,h). The UV–Vis absorption increased and showed better-resolved peaks at 322 nm, 410 nm, 504 nm (Fig. 4g, orange line), and parallel CD spectra had distinct positive peaks (319 nm, 355 nm, 445 nm, and 534 nm) and four negative peaks (333 nm, 392 nm, 477 nm, and 579 nm, Fig. 4h), indicating [2Fe–2S] cluster was transferred from IscU to clMagRC60A. Interestingly, clMagRC124A and clMagRC126A could also accept [2Fe–2S] cluster transferred from holo-IscU, though the binding efficiency is much lower than clMagRWT and clMagRC60A, as verified by UV–Vis and CD spectrum (Supplementary Fig. 1a–d). It seems that clMagRC60A accept [2Fe–2S] cluster from scaffold protein IscU more effectively compared with clMagRC124A and clMagRC126A. And after incubation with clMagR mutants, UV–Vis absorption of IscU significantly decreased, confirmed that iron–sulfur cluster transfer occurred in between holo-IscU and three clMagR mutants (Supplementary Fig. 1e).Again, our data demonstrated that three conserved cystines of clMagR played different roles on the iron–sulfur cluster binding, and especially Cys-60 is essential for clMagR to bind [3Fe–4S] cluster, not [2Fe–2S] cluster. Therefore, it is possible to obtain a [2Fe–2S] cluster binding only clMagR by mutating Cys-60. Thus, we labeled clMagR protein samples based on their iron–sulfur cluster in later experiments. For example, we labeled the chemically reconstituted clMagRWT as [3Fe–4S]-clMagRWT, and clMagRC60A that accepted [2Fe–2S] cluster from holo-IscU as [2Fe–2S]-clMagRC60A, to investigate the magnetic property of clMagR when it binds different iron–sulfur clusters.[3Fe–4S]-clMagR shows different magnetic properties from [2Fe–2S]-clMagRMagR has been reported as a putative magnetoreceptor and exhibits intrinsic magnetic moment experimentally and theoretically when forms complex with cryptochrome (Cry)18,20,21. To elucidate if different iron–sulfur clusters binding in clMagR have different magnetic features and respond to external magnetic fields differently, we obtained [3Fe–4S] and [2Fe–2S] bound only clMagR protein by chemical reconstitution of clMagRWT (as [3Fe–4S]-clMagRWT) and holo-IscU incubated and re-purified clMagRC60A (as [2Fe–2S]-clMagRC60A), respectively, and measured the magnetic moment of these proteins with Superconducting Quantum Interference Device (SQUID) magnetometry. SQUID is a highly sensitive magnetometry to measure extremely subtle magnetic fields and to study the magnetic properties of a range of samples, including extremely low magnetic moment biological samples. Therefore, it has been regularly used as a first test to identify the specific kind of magnetism of a given specimen, such as ferromagnetic, antiferromagnetic, paramagnetic or diamagnetic, by measuring at different temperatures and external magnetic field strength. For example, B-DNA was identified as paramagnetic under low temperature by SQUID60.Purified clMagR3M was utilized as a control since it had no iron–sulfur cluster binding due to lack of cysteine residues (Fig. 1b,c). The magnetic measurement was done at different temperatures (5 K and 300 K) and MH curves (magnetization (M) curves measured versus applied fields (H)) were generated for three proteins to reflect the protein magnetic anisotropy. The MH curves of clMagR3M clearly exhibited diamagnetic property at both 5 K and 300 K, suggesting that magnetism of clMagR is dependent on the iron–sulfur cluster (Fig. 5a,b, red lines). In contrast, [3Fe–4S]-clMagRWT showed superparamagnetic behavior at 5 K which has saturation magnetization (MS) at 2 T about 0.22771 emu/g protein (Fig. 5a, purple line), [2Fe–2S]-clMagRC60A is paramagnetic at 5 K (Fig. 5a, orange line). Interestingly, at higher temperature such as 300 K, [2Fe–2S]-clMagRC60A is diamagnetic while [3Fe–4S]-clMagRWT is paramagnetic (Fig. 5b, orange line and purple line). The different magnetism, as well as the different saturation magnetization of clMagR with different iron–sulfur binding, are clearly important features of this putative magnetoreceptor, and worth further investigation and validation in vivo in the future.Figure 5[3Fe–4S]-clMagRWT shows different magnetic properties from [2Fe–2S]-clMagRC60A. (a) Field-dependent magnetization curves (MH) at 5 K for [2Fe–2S]-clMagRC60A (orange), [3Fe–4S]-clMagRWT (chem re clMagRWT, purple), and clMagR3M (red). The magnetic susceptibility of [2Fe–2S]-clMagRC60A is 2.27749E−6 and the magnetic susceptibility of clMagR3M is − 4.0438E−7. (b) Field-dependent magnetization curves (MH) at 300 K for [2Fe–2S]-clMagRC60A (orange), [3Fe–4S]-clMagRWT (chem re clMagRWT, purple), and clMagR3M (red). And the magnetic susceptibility is − 1.83638E−7, 5.93483E−8, − 3.26432E−7, respectively.Full size image More