JBIC Journal of Biological Inorganic Chemistry

, Volume 23, Issue 7, pp 1119–1128 | Cite as

Emission of volatile halogenated compounds, speciation and localization of bromine and iodine in the brown algal genome model Ectocarpus siliculosus

  • Frithjof C. KüpperEmail author
  • Eric P. Miller
  • Stephen J. Andrews
  • Claire Hughes
  • Lucy J. Carpenter
  • Wolfram Meyer-Klaucke
  • Chiaki Toyama
  • Yasuyuki Muramatsu
  • Martin C. Feiters
  • Carl J. Carrano
Open Access
Original Paper
Part of the following topical collections:
  1. Alison Butler: Papers in Celebration of Her 2018 ACS Alfred Bader Award in Bioorganic or Bioinorganic Chemistry


This study explores key features of bromine and iodine metabolism in the filamentous brown alga and genomics model Ectocarpus siliculosus. Both elements are accumulated in Ectocarpus, albeit at much lower concentration factors (2-3 orders of magnitude for iodine, and < 1 order of magnitude for bromine) than e.g. in the kelp Laminaria digitata. Iodide competitively reduces the accumulation of bromide. Both iodide and bromide are accumulated in the cell wall (apoplast) of Ectocarpus, with minor amounts of bromine also detectable in the cytosol. Ectocarpus emits a range of volatile halogenated compounds, the most prominent of which by far is methyl iodide. Interestingly, biosynthesis of this compound cannot be accounted for by vanadium haloperoxidase since the latter have not been found to catalyze direct halogenation of an unactivated methyl group or hydrocarbon so a methyl halide transferase-type production mechanism is proposed.


Energy-dispersive X-ray analysis Halocarbons Methyl iodide Phaeophyta X-ray absorption spectroscopy 



Culture Collection of Algae and Protozoa


Dissolved organic matter


Energy-dispersive X-ray analysis


Electron paramagnetic resonance


Extended X-ray absorption fine structure




Inductively coupled plasma-mass spectrometry


National Institute of Standards and Technology


Phorbol myristate acetate


Vanadium bromoperoxidase


Vanadium iodoperoxidase


Vanadium haloperoxidase


X-ray absorption spectroscopy


Two centuries ago, the elements bromine and iodine were discovered in sea water and seaweed (Laminaria and Fucus) ashes, respectively [1, 2]. Due to their unique evolutionary history and phylogenetic distance from other important eukaryotic lineages [3, 4], brown algae present some remarkable chemical and physiological adaptations which are also reflected at the genome level [5], making them fascinating experimental models, not only for phycologists, but for a community of interdisciplinary researchers. This includes the function of iodide as extracellular antioxidant protecting the surface of Laminaria digitata (Hudson) Lamouroux against oxidative stress [6]. Indeed, L. digitata is the strongest iodine accumulator currently known among living organisms [7]. In fact, this constituted the first documented case of an inorganic antioxidant in a living system—and the chemically simplest antioxidant known. Key to the antioxidant function is the mobilization of iodide in the apoplast and its efflux into the surrounding seawater upon oxidative stress [6, 8]. In this context, communities of the giant kelp Macrocystis have been found to impact iodine speciation in coastal sea water [9]. The role of bromine is less well investigated, but a recent study [10] highlights that Laminaria accumulates bromide, which complements iodide as an antioxidant especially for the detoxification of superoxide, but with an overall more diverse role.

In contrast to kelps such as Laminaria and Macrocystis, which have a large size (up to around 3 m for Laminaria, and 70 m for Macrocystis, respectively), very complex morphology and which can attain a high standing stock in rocky coastal seabed ecosystems, Ectocarpus is a genus of filamentous brown algae with a worldwide distribution along temperate coastlines, and is a nuisance as a “fouling” organism on many man-made surfaces in the sea. It has some significant advantages as an experimental model compared to Laminaria or Macrocystis and constitutes one of the best-studied seaweeds [11, 12]: due to its small size of only a few mm or cm and its fast growth it can easily be cultivated in small volumes of seawater media both axenically and with associated bacteria [13]; it belongs to a sister group of the ecologically and economically very important kelps [14]; its entire, well-known life cycle can be completed within a few months in culture, many molecular tools are available, including mutant collections, microarrays and proteomics. It has also become the first seaweed of which the entire genome has been sequenced and thus offers unprecedented opportunities for study [5]. Features of its inorganic biochemistry include a remarkable, non-ferritin mode of iron storage [15]. Ectocarpus is also well studied with regards to its pathologies which include viruses [16, 17], fungi [18], oomycetes [18] and plasmodiophoraleans [19].

While brown algae of the genus Laminaria are the strongest accumulators of iodine in life on Earth [20, 21, 22] and a major contributor to the biogeochemical flux of iodine [23] and, to a lesser extent, of brominated and iodinated halocarbons to the atmosphere [24], hardly anything is known about Ectocarpus in this respect. This is remarkable considering the major role of brown algae in global halogen cycling, but also how well other areas of physiology and biochemistry have been studied in Ectocarpus. However, a single gene of a vanadium haloperoxidase (VHPO) and a number of enzymes related to halogen metabolism have been identified in the Ectocarpus genome. This includes at least three different families (21 loci) of haloacid dehalogenase (HAD) and two haloalkane dehalogenase families [5, 25]. It is tempting to hypothesize that the dehalogenases protect Ectocarpus against halocarbons which act as alkylating, toxic agents and are being produced by kelps and other potential substrate algae on which Ectocarpus occurs. The presence of a single VHPO gene, a bromoperoxidase, in the Ectocarpus genome contrasts with the presence of multigenic families of VHPOs, of both bromo- and iodoperoxidases in the kelps Laminaria [26, 27] and Macrocystis [28].

The uptake of iodide from seawater in Laminaria involves VHPOs [21] and its strongest accumulation in this species seems to be linked to the presence of a particular VHPO subclass, the iodoperoxidases, specific for iodide oxidation [26, 27]. In Laminaria, most of the iodine is accumulated in the apoplast of the cortical cell layer [29].

The first line of defense against pathogens in Laminaria is an oxidative burst [30], which is considered a central element of eukaryotic defense in general [31] and which, in the case of kelp species, serves to control bacterial biofilms [32]. Among its triggers are oligoguluronates (GG; [30, 32]), bacterial lipopolysaccharides [33], prostaglandin A2 [34], methyl jasmonate, and polyunsaturated free fatty acids [35]. In L. digitata, early transcriptional defense responses are similar to those in land plants but also involve tightly regulated halogen metabolism which might play roles in more sophisticated chemical defense reactions including distance signaling [36, 37]. In brown algae, bromide and VHPOs have also been shown to catalyze oxidative cross-linking between cell wall polymers, suggesting a role in spore and gamete adhesion and cell-wall strengthening (for reviews see [38, 39]). Much less is known about defense mechanisms in Ectocarpus. Recent studies observed strong differences in susceptibility to infection by Eurychasma dicksonii among different Ectocarpus strains [40], and Eurychasma infection [41] as well as Cu stress [42] were found to result in overexpression of the single VHPO encoded in the Ectocarpus genome.

In our previous studies [6, 10, 43], X-ray absorption spectroscopy (XAS) has proven to be a suitable, non-invasive tool to probe the chemical state and solution environment of accumulated iodine and bromine in kelps. Likewise, we have used energy-dispersive X-ray analysis (EDX) for localization studies of Fe in Ectocarpus [44].

Following our recent study which included Ectocarpus among a number of taxa from various algal lineages [43], this study explores the speciation and localization of bromine and iodine metabolism in Ectocarpus. First iodine and bromine levels in Ectocarpus are determined using ICP-MS. Then the localization of bromine and iodine is investigated using energy-dispersive analysis of X-rays. K-edge XAS is applied to probe the stored chemical state of these elements under different physiological conditions in vivo (both in healthy cultures and in the presence of the oomycete Eurychasma dicksonii). Finally, the production of volatile halocarbons is explored.

Materials and methods

Biological material

A well-characterized strain of E. siliculosus which had served for sequencing the Ectocarpus genome [5] was obtained from the Culture Collection of Algae and Protozoa (CCAP 1310/4) and was used for all experiments described here. Ectocarpus was grown in half-strength Provasoli-enriched sea water as described previously [13, 45, 46], i.e. at 18.

Eurychasma-infected, unialgal host cultures were grown as described previously [13, 46], i.e. at 15 °C, illumination by 20 to 30 μE m−2 s−1 from daylight type fluorescent lamps for 16 h day−1. In a first experiment, about equal amounts (about 50 mg fresh weight each) of 6 Ectocarpus strains (Ec sil NZ Vic KU13/CCAP 1310/56, Ec fas Irl96 23-1n/CCAP 1310/342, Ec sil NZ4a3♀/CCAP 1310/47, Ec sil Vic88 12-1 lat. sp./CCAP 1310/185, Ec sil Vic Z2 lat. sp./CCAP 1310/191, Ec sil vic Z1 lat sp./CCAP 1310/192) infected by Eurychasma 96 [CCAP 4018/2, isolated from Shetland [18, 47] which had been cultured for about 6 months were pooled and used for XAS (below). In a second set of experiments, the following host–pathogen strain combinations were used: the host strains E. siliculosus (CCAP 1310/4, the genome model strain used by [5]), E. siliculosus (CCAP 1310/56, Ecsil NZ KU 1-3♂), E. fasciculatus (Ec fas Ros 007-04), Pylaiella littoralis (CCAP 1330/3, Pyl IR g), either without any Eurychasma infection, or infected with E. dicksonii Eury 96 (CCAP 4018/2) or E. dicksonii Eury 05 (CCAP 4018/1), respectively. Algal host strains (which included a close relative of Ectocarpus, Pylaiella littoralis) were initially cultured for 55 days on their own. Then, the above Eurychasma inocula were added and cultures were continued for another 30 days. Host–pathogen co-cultures were then harvested, placed in Kapton™ cells, and frozen in liquid nitrogen until use for XAS.

ICP-MS analysis of total iodine and bromine

Iodine and bromine contents in freeze-dried Ectocarpus filament samples were analyzed by ICP-MS after pyrohydrolysis separation as described previously [48, 49]. Concentrations obtained are given in ppm (i.e. μg halogen g−1 of freeze-dried material).

Energy-dispersive analysis of X-rays

Ectocarpus siliculosus was harvested from cultures prior to fixation, dehydration, and embedding. Filaments were fixed in a 0.1 M phosphate buffer solution containing 2% (w/v) paraformaldehyde, 1% (w/v) glutaraldehyde, and 1% (w/v) caffeine for 2 h. The fixed cells were then washed with 0.1 M phosphate buffer and dehydrated in successive ethanol baths of 30, 50, 75, 85, 95, and 100% (three times). The cells were then embedded in 1:1 (v/v) ethanol/LR White resin (LWR; EMS, Hatfield, PA, USA) for 3 h followed by 100% LWR overnight in gelatin capsules under vacuum. Sections of 3 μm were cut on a Leica EMUC6 microtome and deposited on glass slides. Slides were coated with platinum in a Quorum Technologies Q150T ES sputter coater. platinum-coated samples were analyzed under high vacuum in a Quanta 450 FEG environmental scanning electron microscope (ESEM) equipped with an Oxford Instruments INCA energy dispersive X-ray (EDX) microanalysis system.

X-ray fluorescence spectroscopy

Ectocarpus filaments were washed briefly with 100 µM ethylenediaminetetracetic acid (EDTA) followed by distilled water to mitigate calcium ion interference in the fluorescence spectrum. Washed filaments were placed on Ultralene™ film (Spex Sample Prep, USA), allowed to dry, and then placed in the GEOCARS X-ray beam of the Advanced Photon Source at Argonne National Laboratory (Lemont, IL).

The selected region of interest was rastered at a resolution of 2 µm with an X-ray beam energy of 5.0 keV, flux of 1.206 × 1012 photons s−1, and dwell time of 20 ms pixel−1. The resulting iodine Lβ1 X-ray fluorescence signal was mapped using the Larch Mapviewer™ software version 0.9.32 [50]. By virtue of the intensity and broadness of the Ca Kβ1 signal, calcium ion interference precluded mapping of both the iodine Lα1 and Lβ1 fluorescent signals from untreated Ectocarpus filaments. Only after EDTA washing was the I Lβ1 signal resolved from Ca Kβ1, while the I Lα1 remained confounded by calcium due to proximity of their respective fluorescent energies.

X-ray absorption spectroscopy

Bromine and iodine K-edge XAS measurements and extended X-ray absorption fine structure (EXAFS) data reduction were carried out at the EMBL Outstation Hamburg at DESY, Germany as described previously [51].


Ectocarpus was sub-cultured into three 1 L flasks filled to 800 mL with sterilized coastal seawater enriched with Guillard’s F/2. Three identical flasks of the media (excluding the Ectocarpus culture) were incubated as controls. Incubation conditions were 20 °C at 100 μmol m−2 s−1 and a 12:12 light/dark cycle. Flasks were stoppered with sterilized cotton in hessian plugs and covered with foil.

For Ectocarpus emission experiments, the solid contents of a culture flask was divided into two 200 mL flasks, filled completely to eliminate headspace with purged, filtered seawater obtained from the Atlantic Ocean at 3000 m depth and sealed with glass stoppers. A third flask was prepared without Ectocarpus addition as a control. The flasks were stored at 15 °C in the dark, sampled and immediately analyzed every 2 h over a 12 h period. 10 mL sample aliquots were taken each time using autoclaved perfluoroalkoxy (PFA) tubing into borosilicate gas syringes, filtered through a standard glass-fiber filter (GFF) and 10 mL of blank seawater added to compensate and eliminate the headspace. The blank seawater used was continuously purged throughout the course of the experiment to minimize VOC content. At the end of the 12 h period the flasks were filtered and the solid mass of Ectocarpus recorded. This was then allowed to air dry and the dry weight was recorded. For investigating the effects of oxidative stress, the experiment was performed as described above except for the addition of 2 mM H2O2 to simulate oxidative stress.

10 mL sample aliquots were analysed using purge and trap coupled to thermal desorption–gas chromatography—time of flight mass spectrometry (P&T-TD-GC-TOFMS, Markes Unity2 CIA8 TDU, Agilent 6890, Markes BenchTof MS) with a custom built purge and trap unit [52]. The TOF–MS allowed high sensitivity monitoring of all ions from 35 to 500 amu simultaneously. Halocarbons were calibrated using NOAA standard SX-3570, accounting for the purge and trap efficiencies of each gas [52]. The control flask was analysed first at each time point to check for carry-over in the analysis and/or production in the blank seawater. No significant change in the analysed VOC content of the blank seawater was observed throughout the emission experiments.


Total halogen levels (ICP-MS)

Initially, halogen levels in cultured Ectocarpus were determined in order to establish whether there is any significant accumulation of iodine and bromine in this organism. Freeze-dried thalli of the brown alga Ectocarpus siliculosus cultivated in normal PES medium contained mean concentrations of 196 ± 11 ppm iodine and 738 ± 3 ppm bromine. In contrast, freeze-dried Ectocarpus siliculosus thalli cultivated in PES medium supplemented with 50 µM KI contained 804 ± 3 ppm iodine, but only 186 ± 3 ppm bromine.

Localization of halogens (EDX and XFS)

Next, the localization of halogens was explored in order to get first insight into potential function. Based on XFS and EDX, most iodine was concentrated in physode-like vesicles (Fig. 1), even though some was also detectable in the cytosol and/or apoplast (Fig. 2). The EDX signal from bromine (Fig. 2) was considerably weaker than that of iodine, showing some localization to the apoplast, cytosol, and possibly intracellular structures (Fig. 2).
Fig. 1

a Optical image of Ectocarpus filaments; b heat map of iodine Lβ1 fluorescence intensity with two regions of interest showing high iodine concentrations; c, d fluorescence spectra for regions of interest from b. Light and dashed vertical lines denote I Lα1 and Ca Kβ1 signals, respectively. Dark vertical lines represent the I Lβ1 signal. These are whole, intact filaments (not a section). The two circled regions of interest are probably physodes at the junction of the main filament and a branched filament. Scale bar = 10 μm

Fig. 2

SEM image of an E. siliculosus section (left), EDX map of iodine (middle), and EDX map of bromine (right). Scale bar = 5 µm. This is a cross-section of a filament that had been fixed, dehydrated, embedded in acrylic, sectioned with a microtome, and coated with platinum before performing EDX


XAS is the method of choice for determining the chemical environment and redox state of halogens in seaweeds [51]. The X-ray Absorption Near-Edge spectra (XANES) at the Br K-edge in Fig. 3 show that non-infected Ectocarpus (trace C) is quite similar to an aqueous solution of NaBr (trace D); it appears that Br is present in Ectocarpus as hydrated bromide [43]. The spectrum of Ectocarpus siliculosus infected with Eurychasma dicksonii (trace B) is subtly different and resembles more that of Br incorporated in an aromatic system, such as in the model compound 4-bromophenylalanine (trace A). The spectra show that in uninfected algae, bromine is present in its reduced anionic form, bromide, whereas upon infection prevailing over several months, incorporation of a major fraction into organic compounds occurs, concomitant with general senescence of the cultures. However, this was not detectable in experiments with a shorter time span of infective pressure.
Fig. 3

Br K-edge XANES. Bromine K-edge X-ray absorption edge spectra of Ectocarpus and relevant model compounds: a 4-bromophenylalanine (aq., from [51]); b Ectocarpus siliculosus infected with Eurychasma dicksonii; c non-infected Ectocarpus siliculosus, and d NaBr (10 mM, aq., from [51])

It was shown earlier [43, 51] that the Extended X-ray Absorption Fine Structure (EXAFS) of hydrated bromide could be simulated with a shell of oxygen atoms at approx. 3.3 Å; although the hydrogen atoms cannot be observed in EXAFS, it can be safely assumed that these oxygens are non-covalently bound to bromide by hydrogen bonds. For non-infected Ectocarpus this is the only contribution (Fig. 4a, b, bottom trace in both panels, attempts to simulate the weaker peaks in the Fourier transform did not lead to significant improvements). For infected Ectocarpus (Fig. 4a, b, top traces in both panels) the carbon peak in the Fourier transform is the strongest, and the pattern of peaks is reminiscent of that of aromatic amino acids such as 4-bromophenylalanine and 3,5-dibromotyrosine. The Br–C distance found in the simulation with both a phenyl group and a shell of H-bonded heteroatoms is 1.90 Ǻ, which is in between the ranges considered typical for bonds with sp2 (1.87–1.88 Ǻ) and sp3 (1.92–1.96 Ǻ) C; the good agreement between experiment and simulation for the other atoms of the phenyl ring strongly indicates sp2 C in this case (Table 1). For the incorporation of Br in organic compounds to be observed by XAS, it is necessary that the infection lasts for a prolonged duration of time (> 1 month in a batch culture); experiments in which the infection was left for only 3 weeks did not result in EXAFS other than that of hydrated bromide.
Table 1

EXCURVE results for bromine K-edge EXAFS (Fig. 4, Debye–Waller factors as 2σ22) in parentheses)

Ectocarpus siliculosus infected with Eurychasma dicksonii

Fit index 0.3985E−02

1.00 Ph (1 C at 1.89940 Å (0.00916); 2 C at 2.74504 Å (0.01912); 2 C at 4.28701 Å (0.02146); 1 C at 4.71337 Å (0.00297) + 1.43 O at 3.38615 Å (0.00302)

EF − 11.0613 eV, range 4.3623–350 eV

Br + sp2 C

Ectocarpus siliculosus

Fit index 0.3011E−02

7.53 O at 3.36691 Å (0.05396)

EF − 8.4995 eV, range 4.5913–350 eV


Fig. 4

Experimental (red) and simulated (black) k3-weighted Br K-edge EXAFS (a) and its Fourier transform (b) of (top traces) Ectocarpus siliculosus infected with Eurychasma dicksonii, and (bottom traces) non-infected Ectocarpus siliculosus. Simulation parameters are given in Table 1

The iodine K-edge spectra (Fig. 5) showed hardly any characteristic features. The EXAFS was so weak that it could not be reliably extracted. The XANES of non-infected Ectocarpus (Fig. 5, trace C) remotely resembles that of an aqueous solution of NaI (trace E) and probably represents iodide in the form of I. In its lack of features it resembles the XANES of lyophilized Laminaria [6] which contains iodide surrounded by a rather disordered shell of hydrogen-bonded heteroatoms, probably from biomolecules rather than water. Addition of 50 μM iodate (trace B) resulted in an increase of the edge step by a factor of approx. 2 prior to normalization (not shown). This addition did however not result in a significant change in the appearance of the edge, as might have been expected on the basis of the spectrum of pure iodate (trace A), suggesting that the added iodate is reduced to iodide upon contact with Ectocarpus. Addition of 50 μM iodide (trace D) resulted in a much larger edge step prior to normalization (not shown), viz. by a factor of 5. The resulting spectrum is featureless like that of the control, except for the more pronounced shoulder at 33,230 eV. In spite of the addition of a considerable amount of hydrated iodide, the features are not as sharp as that of the NaI solution (trace E). This implies that the iodide was taken up by Ectocarpus and that a relatively ordered hydration shell was replaced by biomolecules.
Fig. 5

I K-edge XANES. Iodine K-edge X-ray absorption edge spectra of Ectocarpus and relevant model compounds: a NaIO3 (aq., from [51]); b Ectocarpus siliculosus with 50 μM NaIO3 added; b, c Ectocarpus siliculosus control; d Ectocarpus siliculosus with 50 μM NaI added; and e NaI (20 mM, aq., from [51])

Emission of volatile compounds

In virtually all other seaweed species studied, halogen metabolism comes with the emission of volatile halogenated compounds [7, 24]. In the present study of Ectocarpus, the most predominant emission from the list of calibrated species (CH3I, CH2BrCl, CHCl3, CCl4, CH2Br2, CHBrCl2, CH2ICl, CHBr2Cl, CH2IBr, CHBr3) was methyl iodide (CH3I), which was markedly enhanced under oxidative stress conditions with a production rate of 1.16 × 10−5 pmol (g FW × min)−1 when exposed to 2 mM H2O2 compared to 5.98 × 10−6 pmol (g FW × min)−1 under control conditions during the first 2 h of the experiment (after which production leveled off, cf. Fig. 6). The % relative standard deviation for CH3I for this system is 9.4% [52]. No appreciable production of bromocarbons was observed. Many other non-halogenated species, mostly oxygenated, were detected (but calibrations were not available) based on similarities with NIST (National Institute of Standards and Technology) Mass Spectral Library spectra including acetone, isopropyl alcohol, 2-methyl propanol, butanal, 2-butanone, hexane, 3-methyl 2-butanone, 2-pentanone, pentanal, 3-methyl 2-pentanone, 3-methyl pentanal, 2-hexanone, hexanal, 4-methyl 2-hexanone, 3-heptanone, heptanal, 2-methyl 4-heptanone, 3-methyl 4-heptanone, 4-methyl 2-heptanone, 5-methyl 3-heptanone, 6-methyl 2-heptanone, 5-methyl 2-heptanone and 2-octanone.
Fig. 6

Iodomethane production by E. siliculosus without (triangles) and with (circles) 2 mM H2O2 treatment


Overall, the results of this study show that Ectocarpus has an active bromine and iodine metabolism. Both elements are actively accumulated from surrounding seawater, albeit (especially in the case of iodine) at much lower concentration factors than Laminaria [20, 21]: This study observed concentration factors of 714 for iodine and 2.6 for bromine (i.e. 2–3 and < 1 order of magnitude, respectively, based on fresh weight) compared to normal seawater. This compares to up to 5 orders of magnitude for iodine [21] and around 1 order of magnitude for bromine in Laminaria [10], and 3–4 orders of magnitude for iodine in Macrocystis [28]. Interestingly, artificial iodine supplementation (50 µM KI) in the culture medium resulted in a much lower bromine tissue concentration (186 ± 3 ppm) than in Ectocarpus cultures grown in medium without iodide supplementation (738 ± 3 ppm). This suggests that iodide and bromide compete for the same uptake and storage system, in which the sole V-bromoperoxidase detected in Ectocarpus [5, 25] is likely the key driver. Iodide is well established as being the preferred substrate of V haloperoxidases [53], viz. of both bromo- and iodoperoxidases [27]. Our recent work had already shown that Ectocarpus accumulates bromide [43], similar to morphologically more complex brown algae (Laminariales and Fucales). Here, XAS has shown that, like Laminaria [6], Ectocarpus also accumulates iodine in its reduced form, iodide, bound to H-bonding heteroatoms (O–H, N–H) in biomolecules, with the hydration shell removed, as in lyophilized Laminaria. Again, similar to Laminaria [29], the present study also shows that Ectocarpus accumulates bromide and iodide in the apoplast (iodine storage is exclusively apoplastic, while smaller amounts of bromine can also be detected in the cytosol—Fig. 1). Our observation that the addition of iodate to the Ectocarpus growth medium results in an increased (estimated from X-ray absorption intensity) uptake of the element as iodide implies a reduction, which is in line with recent observations concerning the iodine uptake by marine macroalgae [9]. Interestingly, the reductant could be iodide, to form elemental iodine, which may be considered besides HOI to be the neutral species that is formed as an intermediate in VHPO-mediated iodine uptake. According to Küpper & Kroneck [54] the equilibrium is towards disproportionation at high pH and temperature, so that one might argue that starting at lower T and pH with only IO3 and I, some I2 might be formed by comproportionation, and removed from the equilibrium by uptake in Ectocarpus and subsequent reduction, potentially explaining the enhanced iodine uptake upon addition of iodate.

The strong production of methyl iodide (CH3I), which was slightly increased under oxidative stress conditions, is remarkable and stands out among macroalgae. Production of this compound cannot be accounted for by the V bromoperoxidase present in Ectocarpus (which can, at least in theory, account for the production of CH2BrCl, CHCl3, CCl4, CH2Br2, CHBrCl2, CH2ICl, CHBr2Cl, CH2IBr, CHBr3), catalysis of which results in compounds with multiple halogenations on the same carbon atom. It is worth noting that some of the methyl ketones observed here (2-butanone, 3-methyl 2-butanone, 2-pentanone, 3-methyl 2-pentanone, 2-hexanone, 4-methyl 2-hexanone, 4-methyl 2-heptanone, 6-methyl 2-heptanone, 5-methyl 2-heptanone and 2-octanone) would be good starting materials for the haloform reaction [55, 56] that we proposed recently [43], which could account for the trihalomethanes listed above.

Instead, it is tempting to hypothesize that an S-adenosyl-methionine-dependent methyl transferase-type mechanism is operative here, similar to that described for methyl halide transferase, a class of enzymes which is responsible for chloromethane production in the higher plant Mesembryanthemum crystallinum [57] and other salt-tolerant higher plants [58]. Considering that halocarbon production in seaweeds has usually been associated with oxidative stress and, in particular, with hydrogen peroxide, it has to be highlighted that this reaction is independent of any reactive oxygen species:
$$ S{\text{-adenosyl-}} {\textsc{l}}{\text{-methionine}} + {\text{halide}} \to S{\text{-adenosyl-}} {\textsc{l}}{\text{-homocysteine}} + {\text{methyl halide}}. $$

It should be noted that (like for V haloperoxidases), iodide is the preferred substrate for methyl halide transferases, with usually much lower KM and higher Vmax values reported for iodide over bromide and chloride [58]. While a very moderate increase of methyl iodide production was observed in this study under oxidative stress conditions, this may arguably not be due to the direct participation of H2O2 in the biosynthetic reaction(s), but rather due to the mobilization of iodide from its apoplastic store due to H2O2, analogous to the situation in Laminaria [6], thus merely increasing substrate availability for a hypothetical methyl transferase-type enzyme. It should be noted that Ectocarpus genome annotation has at present not uncovered any S-adenosyl-methionine-dependent methyl transferase—but also that, at present, many open reading frames in the Ectocarpus genome encode for genes of still unknown function [25]. A BLAST search of the complete cDNA sequence of Beta maritima [AF084829; 58] against the Ectocarpus genome (11×) and latest (as of November 2017) cDNA database did not yield any significant matches (not shown). Other halogen-related enzymes that have been identified in the Ectocarpus genome include at least three different families (21 loci) of haloacid dehalogenase (HAD) and two haloalkane dehalogenases. The HADs belong to a large superfamily of hydrolases with diverse substrate specificity, including phosphatases and ATPases. The dehalogenase enzymes may contribute to protect Ectocarpus against halogenated compounds produced as defence metabolites by kelps [6] allowing it to successfully grow as an epiphyte or endophyte on kelp thalli [59, 60].



Funding from the UK Natural Environment Research Council (NERC) through grants NE/D521522/1 (FCK), NE/F012705/1 (FCK), NE/K000454/1 (CH), and the Oceans 2025 (WP4.5) program FCK; the National Science Foundation (CHE-1664657) to CJC; and the MASTS pooling initiative (Marine Alliance for Science and Technology for Scotland, funded by the Scottish Funding Council and contributing institutions; grant reference HR09011) is gratefully acknowledged. We are also grateful to Claire M.M. Gachon (Scottish Association for Marine Science) for her assistance with the Ectocarpus/Eurychasma cultures. EPM and CJC are grateful to the Advanced Photon Source at Argonne National Laboratory for facilitating the GSECARS X-ray experiments and Tony Lanzirotti for his assistance on the GSECARS beamline and with subsequent data analysis. Furthermore, the authors are grateful for support from the European Community in the framework of the Access to Research Infrastructure Action of the Improving Human Potential Program to the EMBL Hamburg Outstation. Finally, we would like to thank Yuka Uchida (Department of Chemistry, Gakushuin University, Tokyo) for her help with the ICP-MS analyses mentioned here.


  1. 1.
    Balard AJ (1826) Ann Chim Phys (Paris) 32:337–384Google Scholar
  2. 2.
    Gay-Lussac L-J (1813) Ann Chim (Paris) 88:311–318Google Scholar
  3. 3.
    Baldauf SL (2003) Science 300:1703–1706CrossRefGoogle Scholar
  4. 4.
    Bhattacharya D, Stickel SK, Sogin ML (1991) J Mol Evol 33:525–536CrossRefGoogle Scholar
  5. 5.
    Cock JM, Sterck L, Rouzé P, Scornet D, Allen AE, Amoutzias G, Anthouard V, Artiguenave F, Aury J-M, Badger JH, Beszteri B, Billiau K, Bonnet E, Bothwell JHF, Bowler C, Boyen C, Brownlee C, Carrano CJ, Charrier B, Cho GY, Coelho SM, Collén J, Corre E, Da Silva C, Delage L, Delaroque N, Dittami SM, Doulbeau S, Elias M, Farnham G, Gachon CMM, Gschloessl B, Heesch S, Jabbari K, Jubin C, Kawai H, Kimura K, Kloareg B, Küpper FC, Lang D, Le Bail A, Leblanc C, Lerouge P, Lohr M, Lopez PJ, Martens C, Maumus F, Michel G, Miranda-Saavedra D, Morales J, Moreau H, Motomura T, Nagasato C, Napoli CA, Nelson DR, Nyvall-Collén P, Peters AF, Pommier C, Potin P, Poulain P, Quesneville H, Read B, Rensing SA, Ritter A, Rousvoal S, Samanta M, Samson G, Schroeder DC, Ségurens B, Strittmatter M, Tonon T, Tregear J, Valentin K, von Dassow P, Yamagishi T, Van de Peer Y, Wincker P (2010) Nature 465:617–621CrossRefGoogle Scholar
  6. 6.
    Küpper FC, Carpenter LJ, McFiggans GB, Palmer CJ, Waite TJ, Boneberg EM, Woitsch S, Weiller M, Abela R, Grolimund D, Potin P, Butler A, Luther GW III, Kroneck PMH, Meyer-Klaucke W, Feiters MC (2008) Proc Natl Acad Sci USA 105:6954–6958CrossRefGoogle Scholar
  7. 7.
    Küpper FC, Feiters MC, Olofsson B, Kaiho T, Yanagida S, Zimmermann MB, Carpenter LJ, Luther GW III, Lu Z, Jonsson M, Kloo L (2011) Angew Chem Int Ed 50:11598–11620CrossRefGoogle Scholar
  8. 8.
    Chance R, Baker AR, Küpper FC, Hughes C, Kloareg B, Malin G (2009) Estuar Coast Shelf Sci 82:406–414. CrossRefGoogle Scholar
  9. 9.
    Gonzales J, Tymon T, Küpper FC, Edwards MS, Carrano CJ (2017) PLoS ONE 12:e0180755. CrossRefPubMedPubMedCentralGoogle Scholar
  10. 10.
    Küpper FC, Carpenter LJ, Leblanc C, Toyama C, Uchida Y, Verhaeghe E, Maskrey B, Robinson J, Boneberg E-M, Malin G, Luther GW III, Kroneck PMH, Kloareg B, Meyer-Klaucke W, Muramatsu Y, Potin P, Megson IL, Feiters MC (2013) J Exp Bot 64:2653–2664CrossRefGoogle Scholar
  11. 11.
    Peters AF, Marie D, Scornet D, Kloareg B, Cock JM (2004) J Phycol 40:1079–1088CrossRefGoogle Scholar
  12. 12.
    Charrier B, Coelho SM, Le Bail A, Tonon T, Michel G, Potin P, Kloareg B, Boyen C, Peters AF, Cock JM (2008) New Phytol 177:319–332CrossRefGoogle Scholar
  13. 13.
    Müller DG, Gachon CMM, Küpper FC (2008) Cah Biol Mar 49:59–65Google Scholar
  14. 14.
    Silberfeld T, Leigh JW, Verbruggen H, Cruaud C, de Reviers B, Rousseau F (2010) Mol Phylogenet Evol 56:659–674. CrossRefPubMedGoogle Scholar
  15. 15.
    Böttger LH, Miller EP, Andresen C, Matzanke BF, Küpper FC, Carrano CJ (2012) J Exp Bot 63:5763–5772CrossRefGoogle Scholar
  16. 16.
    Müller DG, Kapp M, Knippers R (1998) Adv Virus Res 50:49–67CrossRefGoogle Scholar
  17. 17.
    Müller DG, Kawai H, Stache B, Lanka S (1990) Bot Acta 103:72–82CrossRefGoogle Scholar
  18. 18.
    Müller DG, Küpper FC, Küpper H (1999) Phycol Res 47:217–223CrossRefGoogle Scholar
  19. 19.
    Maier I, Parodi E, Westermeier R, Müller DG (2000) Protist 151:225–238CrossRefGoogle Scholar
  20. 20.
    Ar Gall E, Küpper FC, Kloareg B (2004) Bot Mar 47:30–37Google Scholar
  21. 21.
    Küpper FC, Schweigert N, Ar Gall E, Legendre JM, Vilter H, Kloareg B (1998) Planta 207:163–171CrossRefGoogle Scholar
  22. 22.
    Saenko GN, Kravtsova YY, Ivanenko VV, Sheludko SI (1978) Mar Biol 47:243–250CrossRefGoogle Scholar
  23. 23.
    McFiggans GB, Coe H, Burgess R, Allan J, Cubison M, Alfarra MR, Saunders R, Saiz-Lopez A, Plane JMC, Wevill DJ, Carpenter LJ, Rickard AR, Monks PS (2004) Atmos Chem Phys 4:701–713CrossRefGoogle Scholar
  24. 24.
    Carpenter LJ, Malin G, Küpper FC, Liss PS (2000) Glob Biogeochem Cycles 14:1191–1204CrossRefGoogle Scholar
  25. 25.
    Cock JM, Sterck L, Ahmed S, Allen AE, Amoutzias G, Anthouard V, Artiguenave F, Arun A, Aury JM, Badger JH, Beszteri B, Billiau K, Bonnet E, Bothwell JH, Bowler C, Boyen C, Brownlee C, Carrano CJ, Charrier B, Cho GY, Coelho SM, Collen J, Le Corguille G, Corre E, Dartevelle L, Da Silva C, Delage L, Delaroque N, Dittami SM, Doulbeau S, Elias M, Farnham G, Gachon CMM, Godfroy O, Gschloessl B, Heesch S, Jabbari K, Jubin C, Kawai H, Kimura K, Kloareg B, Kuepper FC, Lang D, Le Bail A, Luthringer R, Leblanc C, Lerouge P, Lohr M, Lopez PJ, Macaisne N, Martens C, Maumus F, Michel G, Miranda-Saavedra D, Morales J, Moreau H, Motomura T, Nagasato C, Napoli CA, Nelson DR, Nyvall-Collen P, Peters AF, Pommier C, Potin P, Poulain J, Quesneville H, Read B, Rensing SA, Ritter A, Rousvoal S, Samanta M, Samson G, Schroeder DC, Scornet D, Segurens B, Strittmatter M, Tonon T, Tregear JW, Valentin K, Von Dassow P, Yamagishi T, Rouze P, Van de Peer Y, Wincker P, Ectocarpus Genome C (2012) In: Piganeau G (ed) Genomic insights into the biology of algae. Academic Press Ltd-Elsevier Science Ltd, London, pp 141–184CrossRefGoogle Scholar
  26. 26.
    Colin C, Leblanc C, Michel G, Wagner E, Leize-Wagner E, van Dorsselaer A, Potin P (2005) J Biol Inorg Chem 10:156–166CrossRefGoogle Scholar
  27. 27.
    Colin C, Leblanc C, Wagner E, Delage L, Leize-Wagner E, Van Dorsselaer A, Kloareg B, Potin P (2003) J Biol Chem 278:23545–23552CrossRefGoogle Scholar
  28. 28.
    Tymon TM, Miller EP, Gonzales JL, Raab A, Küpper FC, Carrano CJ (2017) J Inorg Biochem 177:82–88. CrossRefPubMedGoogle Scholar
  29. 29.
    Verhaeghe EF, Fraysse A, Guerquin-Kern J-L, Wu T-D, Devès G, Mioskowski C, Leblanc C, Ortega R, Ambroise Y, Potin P (2008) J Biol Inorg Chem 13:257–269CrossRefGoogle Scholar
  30. 30.
    Küpper FC, Kloareg B, Guern J, Potin P (2001) Plant Physiol 125:278–291CrossRefGoogle Scholar
  31. 31.
    Wojtaszek P (1997) Biochem J 322:681–692CrossRefGoogle Scholar
  32. 32.
    Küpper FC, Müller DG, Peters AF, Kloareg B, Potin P (2002) J Chem Ecol 28:2057–2081CrossRefGoogle Scholar
  33. 33.
    Küpper FC, Gaquerel E, Boneberg E-M, Morath S, Salaün J-P, Potin P (2006) J Exp Bot 57:1991–1999CrossRefGoogle Scholar
  34. 34.
    Zambounis A, Gaquerel E, Strittmatter M, Potin P, Küpper FC (2012) Algae 27:1–12CrossRefGoogle Scholar
  35. 35.
    Küpper FC, Gaquerel E, Cosse A, Adas F, Peters AF, Müller DG, Kloareg B, Salaün JP, Potin P (2009) Plant Cell Physiol 50:789–800. CrossRefPubMedGoogle Scholar
  36. 36.
    Cosse A, Potin P, Leblanc C (2009) New Phytol 182:239–250. CrossRefPubMedGoogle Scholar
  37. 37.
    Thomas F, Cosse A, Goulitquer S, Raimund S, Morin P, Valero M, Leblanc C, Potin P (2011) PLoS One 6(6):e21475. CrossRefPubMedPubMedCentralGoogle Scholar
  38. 38.
    La Barre S, Potin P, Leblanc C, Delage L (2010) Mar Drugs 8:988–1010. CrossRefPubMedPubMedCentralGoogle Scholar
  39. 39.
    Potin P, Leblanc C (2006) In: Smith AM, Callow JA (eds) Biological Adhesives. Springer Verlag, pp. 105-124Google Scholar
  40. 40.
    Gachon CMM, Strittmatter M, Müller DG, Kleinteich J, Küpper FC (2009) Appl Environ Microbiol 75:322–328CrossRefGoogle Scholar
  41. 41.
    Strittmatter M, Grenville-Briggs LJ, Breithut L, van West P, Gachon CMM, Kupper FC (2016) Plant Cell Environ 39:259–271. CrossRefPubMedGoogle Scholar
  42. 42.
    Ritter A, Umbertini M, Romac S, Gaillard F, Delage L, Mann A, Cock JM, Tonon T, Correa JA, Potin P (2010) Proteomics 10:2074–2088CrossRefGoogle Scholar
  43. 43.
    Küpper FC, Leblanc C, Meyer-Klaucke W, Potin P, Feiters MC (2014) J Phycol 50:652–664CrossRefGoogle Scholar
  44. 44.
    Miller EP, Böttger LH, Weerasinghe AJ, Crumbliss AL, Matzanke BF, Meyer-Klaucke W, Küpper FC, Carrano CJ (2014) J Exp Bot 65:585–594. CrossRefPubMedGoogle Scholar
  45. 45.
    Starr RC, Zeikus JA (1987) J Phycol 23((Suppl.)):1–47Google Scholar
  46. 46.
    Strittmatter M, Grenville-Briggs LJ, Breithut L, Van West P, Gachon CMM, Küpper FC (2016) Plant. Cell Environ 39:259–271. CrossRefGoogle Scholar
  47. 47.
    Küpper FC, Müller DG (1999) Nova Hedwig 69:381–389Google Scholar
  48. 48.
    Chai JY, Muramatsu Y (2007) Geostand Geoanal Res 31:143–150. CrossRefGoogle Scholar
  49. 49.
    Schnetger B, Muramatsu Y (1996) Analyst 121:1627–1631. CrossRefPubMedGoogle Scholar
  50. 50.
    Newville M (2013) J Phys Conf Ser 430:012007. CrossRefGoogle Scholar
  51. 51.
    Feiters MC, Küpper FC, Meyer-Klaucke W (2005) J Synchrotron Rad 12:85–93CrossRefGoogle Scholar
  52. 52.
    Andrews SJ, Hackenberg SC, Carpenter LJ (2015) Ocean Sci 11:313–321. CrossRefGoogle Scholar
  53. 53.
    Vilter H (1995) In: Sigel A, Sigel H (eds) Vanadium and its role in life. Marcel Dekker, New York, pp 325–362Google Scholar
  54. 54.
    Küpper FC, Kroneck PMH (2015) In: Kaiho T (ed) Iodine chemistry and applications. Wiley, Hoboken, pp 557–589Google Scholar
  55. 55.
    Theiler R, Cook JC, Hager LP, Studa JF (1978) Science 202:1094–1096CrossRefGoogle Scholar
  56. 56.
    Beissner RS, Guilford WJ, Coates RM, Hager LP (1981) Biochemistry 20:3724–3731. CrossRefPubMedGoogle Scholar
  57. 57.
    Wuosmaa AM, Hager LP (1990) Science 249:160–162. CrossRefPubMedGoogle Scholar
  58. 58.
    Ni X, Hager LP (1999) Proc Natl Acad Sci 96:3611–3615. CrossRefPubMedGoogle Scholar
  59. 59.
    Russell G (1983) Mar Ecol Prog Ser 13:303–304. CrossRefGoogle Scholar
  60. 60.
    Russell G (1983) Mar Ecol Prog Ser 11:181–187. CrossRefGoogle Scholar

Copyright information

© The Author(s) 2018

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

Authors and Affiliations

  • Frithjof C. Küpper
    • 1
    • 2
    Email author
  • Eric P. Miller
    • 3
  • Stephen J. Andrews
    • 4
  • Claire Hughes
    • 5
  • Lucy J. Carpenter
    • 4
  • Wolfram Meyer-Klaucke
    • 6
  • Chiaki Toyama
    • 7
  • Yasuyuki Muramatsu
    • 8
  • Martin C. Feiters
    • 9
  • Carl J. Carrano
    • 3
  1. 1.OceanlabUniversity of AberdeenNewburghScotland, UK
  2. 2.Dunstaffnage Marine LaboratoryScottish Association for Marine ScienceObanScotland, UK
  3. 3.Department of Chemistry and BiochemistrySan Diego State UniversitySan DiegoUSA
  4. 4.Wolfson Atmospheric Chemistry Laboratories, Department of ChemistryUniversity of YorkYorkUK
  5. 5.Environment DepartmentUniversity of YorkYorkUK
  6. 6.Department of Chemistry - Inorganic Chemistry, Faculty of ScienceUniversity of PaderbornPaderbornGermany
  7. 7.Geological Survey of JapanThe National Institute of Advanced Industrial Science and Technology (AIST)Tsukuba, IbarakiJapan
  8. 8.Department of Chemistry, Faculty of ScienceGakushuin UniversityTokyoJapan
  9. 9.Department of Organic Chemistry, Institute for Molecules and MaterialsRadboud University NijmegenNijmegenThe Netherlands

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