Localizing organomercury uptake and accumulation in zebrafish larvae at the tissue and cellular level

Edited by James Penner-Hahn, University of Michigan, Ann Arbor, MI, and accepted by the Editorial Board July 13, 2008
August 26, 2008
105 (34) 12108-12112


Using synchrotron x-ray fluorescence mapping, we have examined the uptake and localization of organic mercury in zebrafish larvae. Strikingly, the greatest accumulation of methyl and ethyl mercury compounds was highly localized in the rapidly dividing lens epithelium, with lower levels going to brain, optic nerve, and various other organs. The data suggest that the reported impairment of visual processes by mercury may arise not only from previously reported neurological effects, but also from direct effects on the ocular tissue. This novel approach is a powerful tool for directly investigating the molecular toxicology of heavy metals, and should be equally applicable to the study of a wide range of elements in developing embryos.
Toxic organic mercury compounds worry many communities worldwide, yet the detailed mechanisms underlying their transport and toxicity remain uncertain (1). These neurotoxic compounds are particularly insidious due to the latency in onset of toxic symptoms and have caused several devastating mass-poisonings of humans. Adults are affected, but exposure in utero has resulted in severe consequences such as microcephaly, cerebropalsy, seizures, and mental retardation. Methylmercury (MeHg) compounds are actively transported across cell membranes (2) although not, as originally thought, by molecular mimicry of methionine (3). Beyond this, however, our knowledge of the mechanisms underlying organic mercury (Hg) toxicity is fragmentary. MeHg-induced changes in cellular Ca2+ have been shown to be important (47), as has oxidative stress (4), and glutamate metabolism (8). In both yeast and human cells, overexpression of the ubiquitin-targeting enzyme Cdc34 confers protection against the cytotoxic effects of MeHg, which leads to the suggestion that an unknown protein containing a signal for ubiquitination by Cdc34 is involved in the development of the cytotoxic effects of MeHg (911).
Zebrafish (Danio rerio) are common model organisms for the study of embryonic development, and have found increasing use in vertebrate toxicology (12). While embryonic and larval zebrafish previously have been used to study the toxic effects of MeHg exposure at the whole body (13, 14) and at the molecular level (15), there are no available data on MeHg uptake and accumulation with respect to different tissues and organs. Such information is critical to properly integrate information on tissue and cell specific impacts of Hg with uptake and accumulation of the metal at the whole animal level. We present herein a novel approach to the investigation of heavy-metal toxicity using synchrotron x-ray fluorescence imaging (16) to directly image metal localization within zebrafish larvae and to observe remarkable differential accumulation of organic Hg using this technique. Strikingly, we find that both methyl and ethyl Hg derivatives are concentrated to the greatest degree in lens epithelium with lower levels present in the brain, optic nerve, and other organs. Our work suggests that direct effects on ocular tissue play a role in the disruption of visual processes by Hg (17). Furthermore, synchrotron x-ray fluorescence imaging is applicable to any heavy metal and is expected to have broad applications in vertebrate molecular toxicology including testing of new chelation therapy drugs.

Results and Discussion

The accumulation pattern of MeHg in live zebrafish was determined by exposing larvae at 3.5 days post fertilization (dpf) to waterborne MeHg-l-cysteine [CH3Hg-S-(l-Cys)] for 24 h before imaging. At the end of the exposure all larvae were alive although mostly moribund. During the imaging experiment the larvae were anesthetized with tricaine (ethyl 3-aminobenzoate methanesulfonate salt) and then immobilized in 1% low melting-point agarose gel. The elemental distribution maps were obtained by spatially rastering the live fish through a microfocused synchrotron x-ray beam at 10 μm and monitoring the Hg Lα, Ca and Zn Kα fluorescence intensities. Fig. 1 shows images of two intact MeHg-treated fish positioned with dorsal and lateral presentations to the x-ray beam. The images clearly show that Hg accumulation is neither homogenous, nor does it show the same distribution as either Zn or Ca. Regions of relatively high Hg include brain, gastrointestinal tract, and particularly the eye lens. Whereas the presence of Hg in the first two tissues is expected, the relatively high accumulation of MeHg in the eye lens is unprecedented. Both micro-Hg LIII near-edge spectra (18) of the lens region of treated fish and bulk spectra of large numbers of similarly treated fish were consistent with a local Hg coordination that was the same as the administered compounds (not illustrated), although we note that the high x-ray doses involved in recording near-edge spectra killed the fish. The highest Ca levels corresponded to the developing otoliths in the otic capsule. The Zn distribution reflects the general pigmentation pattern of the zebrafish, including high levels in the retina, with additional elevated Zn levels in the yolk sac.
Fig. 1.
X-ray fluorescence images of intact zebrafish. Dorsal (A, B, C) and lateral (D, E, F) views of 4.5-dpf living zebrafish larva after 24-h treatment with waterborne 100 μM MeHg-l-cysteine comparing optical images (A, D) with X-ray fluorescence images at 10 μm resolution of mercury alone (B, E), and colocalization of Hg (blue) with Zn (green) and Ca (red) (C, F).
More detailed quantitative information on Hg deposition was obtained by imaging transverse (≈6 μm) sections of Hg-exposed fish. We used newly hatched larvae (3.5 dpf) that were exposed to 0.2, 2, and 100 μM CH3Hg-S-(l-Cys) for 84, 36, and 24 h, respectively, or to 100 μM ethylmercury 2-thiosalicylate (thimerosal) for 20 h to additionally determine whether the accumulation pattern depends on the chemical form of the organic Hg. Following the exposures the larvae were fixed, embedded in methacrylate, and sectioned on a microtome. Of two adjacent sections, one was mounted on a glass slide and stained with methylene blue to obtain a histological image, and the other, intended for synchrotron x-ray fluorescence imaging, was placed on a plastic coverslip without any further processing [supporting information (SI) Fig. S1]. We collected 10-μm-resolution images of the spatial distribution of Hg, Zn, and Ca in zebrafish sections. Fig. 2 compares the elemental distributions in sections of exposed and control fish heads. The control fish showed no Hg signal, but displayed significantly higher Ca levels in the outer layers of the retina than the Hg exposed fish (Fig. 2), suggesting that MeHg might interfere with the natural Ca distribution as has been inferred by others (47). The images of Hg-exposed fish broadly resemble those of the whole live fish (Fig. 1), indicating that the procedures used to fix, embed, and section the specimen did not significantly modify the distributions of the elements investigated. Fig. 3 compares quantitative Hg distributions of representative sections of zebrafish larvae exposed to 100 μM CH3Hg-S-(l-Cys). We observe highly preferential accumulation of Hg in the outer layer of the eye lens, reaching ≈1 μg/cm2, a level more than four times higher than in other tissues examined. Another Hg-rich region in the eye, the outer layer of the retina, shows somewhat lower levels of Hg (≈0.12–0.21 μg/cm2), comparable with levels found in the various brain regions (BR ≈0.08–0.14 μg/cm2), and in the spinal cord (SC ≈0.12 μg/cm2). Liver, a known target organ for MeHg, shows Hg levels of ≈0.17 μg/cm2 (Fig. 3; LV, liver). Similar Hg levels are also observed in segmental muscles (MS) and in the yolk sac wall (Fig. 3; YL, yolk). Fig. 4 compares sections of zebrafish exposed to low levels of CH3Hg-S(l-Cys) (2 μM and 200 nM) indicating very similar spatial distributions and similar high accumulations of Hg. In fact, exposure to 200 nM MeHg-l-cysteine for 84 h gave higher accumulations in eye-lens epithelia, brain, liver, and muscle than did exposure to 100 μM for 24 h.
Fig. 2.
Elemental distributions in MeHg exposed and unexposed zebrafish. Comparison of elemental distributions for Hg, Ca, and Zn in sections of fish heads from 4.5-dpf larvae exposed for 24 h to 100 μM MeHg-l-cysteine (Upper) with those from control fish (Lower), measured using x-ray fluorescence imaging. Each section is paired with its respective histological image (Left). Quantities of the individual elements are plotted on the same (arbitrary) scale for Hg exposed and control sections (Hg scale from 0 to 0.96 μg/cm2).
Fig. 3.
Quantitative mercury distributions of treated zebrafish. Head and liver sections from 4.5-dpf zebrafish treated with waterborne 100 μM MeHg-l-cysteine for 24 h, comparing histological images with mercury distribution of the adjacent section; (BR) brain, (EL) eye lens, (LV) liver, (GT) gut, (KT) kidney tubule, (MS) skeletal muscle, (YL) yolk, (SC) spinal cord.
Fig. 4.
Quantitative mercury distributions of zebrafish treated with low levels of MeHg-l-cysteine. Head and liver sections from 5-dpf zebrafish larvae treated with waterborne 2 μM MeHg-l-cysteine for 36 h (A) and 7-dpf larvae treated with 200 nM MeHg-l-cysteine for 84 h (B) respectively. Histological images are compared with mercury distribution of the adjacent section; brain (BR), eye lens (EL), liver (LV), gut (GT), kidney tubule (KT), skeletal muscle (MS), spinal cord (SC).
Fig. 5 shows an overlay of a phase-contrast visible-light micrograph of a 100 μM section with its respective Hg distribution map, indicating that Hg also accumulates in the kidney tubules (KT <0.17 μg/cm2), dorsal aorta (DA ≈0.14–0.15 μg/cm2), the gut (GT ≈0.10–0.15 μg/cm2), and the pectoral fins (PF ≈0.08–0.13 μg/cm2). Slightly higher concentrations of Hg (up to 0.21 μg/cm2) were detected in the pericardial muscles, but no Hg was found in the heart (Fig. S2). Similar patterns of Hg accumulation, with the highest Hg at the periphery of the eye lens, were observed in tissues from larvae treated with another organomercury (C–Hg) thimerosal (Fig. S3). This suggests that the accumulation pattern does not vary strongly with the chemical form of organic Hg.
Fig. 5.
Hg distribution in the liver section from 4.5-dpf zebrafish treated with 100 μM MeHg-l-cysteine for 24 h. (A) The mercury map of the liver section and (B) the mercury map superimposed on the phase-contrast visible light micrograph of the same section to facilitate the localization of mercury within certain tissues; spinal cord (SC), skeletal muscle (MS), liver (LV), gut (GT), kidney tubule (KT), pectoral fin (PF), yolk (YL), dorsal aorta (DA). Ventral is facing the top of the page in these images.
Fig. 6 shows a 2.5-μm resolution image of the distribution of Hg in the eye section of MeHg-treated zebrafish compared with the histological image of the adjacent section. The distribution is fully consistent with all lower resolution images. Hg is preferentially accumulated in a very thin (<8 μm) layer at the periphery of the eye lens. Such a small dimension implicates a single cell layer in the accumulation of organic Hg in fish larvae. Some Hg is also clearly visible in the optic nerve, the brain, and the outer layer of the retina (Fig. 6B). In Fig. 6C the tricolor map overlaying Hg, Zn, and S distributions reveals that the Hg-rich single cell layer encloses the interior of the lens, which has a high S content, while Zn is found at elevated levels in the retina. Thus, Hg is concentrated at the periphery of the eye lens, which consists of the mitotically active lens epithelial cells. Throughout lens growth the epithelial cells continually divide and subsequently move inwards, differentiating into the concentric layers of fiber cells that become part of the dense inner lens core (19). During the differentiation process, the cell nucleus and other organelles degrade to provide the optical clarity that characterizes the mature lens tissue. High S content in the eye lens originates from the proteins (crystallins) rich in −SH groups (20) and glutathione molecules present mainly in the lens epithelium (21). The fiber cells in the zebrafish lens core are arranged in concentric shells, and thus the lens cross-section reveals the oldest tissue in the centre and the youngest at the outer rim. Due to the unique morphology and stability of the lens over the life of an organism, it has been suggested that the lens could potentially offer a historical record of Hg exposures affecting a fish through its lifetime (22). Our findings constitute a beginning step toward understanding the uptake and accumulation of organic Hg in the fish eye.
Fig. 6.
High resolution elemental distributions of zebrafish head. Head section from 4.5-dpf zebrafish larva treated with 100 μM MeHg-l-cysteine for 24 h. (A) histological image, (B) mercury distribution using x-ray fluorescence imaging at 2.5 μm resolution, and (C) Hg (green) superimposed on S (red) and Zn (blue). Quantities of the different elements are plotted on arbitrary scales.
The mechanism responsible for the preferential uptake of organic Hg in the eye lens epithelium is not known at present. However, since the overall development and morphology of the adult zebrafish lens is similar to that of other vertebrates, it may be hypothesized that a similar mechanism is also responsible for the accumulation of organic Hg in the mammalian eye. The accumulation of Hg in the eye region following MeHg exposure has been reported previously in several different species including rabbits (23) and rats (24), and there is increasing evidence for adverse effects of MeHg on both mammalian (17) and zebrafish (25) visual systems. Ocular manifestations of organic Hg exposure were also observed in victims of mass poisoning outbreaks in Iraq (26) and Japan (27), and a recent case of a patient with failing visual acuity was attributed to the patient's high intake of predatory fish (28). Visual defects, such as partial or complete loss of vision and constriction in peripheral visual fields, are common results of organic Hg poisoning and are known to reflect neuronal loss in certain parts of the brain (29). Our data using methyl- or ethylmercury-exposed zebrafish larvae reveal that a large accumulation of Hg can occur in the eye and especially in the lens epithelium. Thus, it is possible that Hg accumulation impairs the visual processes not only on a neurological level, but also by a more direct effect on the ocular tissue.
In summary, we have presented a novel approach to the investigation of heavy-metal molecular toxicology and localization in vertebrates using synchrotron x-ray fluorescence imaging, showing that organic Hg is preferentially taken up by the lens epithelial cells surrounding the lens core. The method has the advantage that it is nondestructive, unlike laser ablation mass-spectrometry imaging methods, which can in principal provide similar information (e.g., 30). Fluorescent probes of Hg2+ have also been described (31) and used for in vivo imaging of zebrafish (32, 33). Although powerful, these methods have the disadvantage that very low but significant fluorescence signals can be elicited in the presence of much more biologically abundant metal ions (such as Ca2+), and rely on secondary interaction of the metal with specific fluorescent probe molecules. In fact, our x-ray fluorescence imaging methods not only will detect the element of interest in any and every chemical form, but in certain cases variability in the near-edge spectrum should allow chemically-specific maps to be developed (16). We expect synchrotron x-ray fluorescence imaging of zebrafish to be a powerful tool for investigating molecular toxicology of heavy metals. The method is equally applicable to the study of other elements of concern, such as arsenic, selenium, thallium and lead. The technique also provides an ideal tool for investigating drugs such as chelation agents (34) and can be applied to the study of essential metals and other elements of interest during normal development.

Materials and Methods

Animal Care and Embryo Collection.

Adult fish were kept at 28°C in carbon-filtered tap water, with a photoperiod of 14 h. Embryos were collected and staged following standard procedures (35). After collection, embryos and larvae were reared in 25-mL Petri dishes with system water changed daily.

Reagent and Treatment Solutions.

MeHg hydroxide (1M aqueous solution), l-cysteine, thimerosal, and tricaine were purchased from Strem Chemicals Inc., Alfa Aesar, and Sigma-Aldrich. The 1 M MeHg hydroxide solution was diluted in triple-distilled water to obtain a 40 mM stock solution. An equimolar stock solution of L-cysteine was prepared in 30 mM phosphate-buffered saline (PBS). A 10 mM stock solution of MeHg-l-cysteine complex was prepared by mixing suitable aliquots of 40 mM MeHg hydroxide and 40 mM l-cysteine solutions (molar ratio Hg:Cys of 1:1.2). The mixture was further diluted in 30 mM PBS as needed. Treatment solutions of MeHg-l-cysteine complex were made from dilutions of the stock in carbon-filtered tap water. A 10 mM stock solution of thimerosal was prepared in 30 mM PBS. The exposures were conducted in sterile 25-mL Petri dishes. All stock and treatment solutions were freshly made before each exposure.

MeHg and Thimerosal Treatment.

For the exposure, newly-hatched, 3.5-dpf larval zebrafish were placed in 25-mL Petri dishes containing 200 nM to 100 μM MeHg-l-cysteine or 100 μM thimerosal. Untreated control dishes were included in each experiment. Two replicate treatments (2 × 25 larvae) were carried out for each compound. Exposures were at 28°C for 24–84 and 20 h for MeHg-L-cysteine and thimerosal, respectively, and included the 10-h dark period. The water/treatment solutions were changed daily for exposures >24 h. After the exposure, the larvae were rinsed several times in fresh carbon-filtered water to remove any remaining Hg.

Live Sample Preparation.

Exposed and rinsed larvae were placed in a new Petri dish containing anesthetic solution (0.016% tricaine) (36). The anesthetized fish were transferred into a new dish containing 1% low melting point agarose gel (with 0.016% tricaine) for immobilization. Such mounted fish were alive for >5 h as evaluated by regular checks of heart beat using microscopy. Before imaging, a small block was cut out around the selected fish and mounted between two 6.3-μm films of polypropylene (SPEX CertiPrep.

Preparation of Sections.

For the tissue-section imaging, the larvae were fixed in 4% paraformaldehyde for 2 h at rt. The fixed larvae were dehydrated in a graded series (0%, 25%, 50%, 75%, and 100%) of ethanol in phosphate buffered saline + Tween 20 (PBST) buffer (30 mM PBS, 0.1% Tween 20) for 5 min each, and stored until needed in 100% ethanol at −20°C. For sectioning, the fixed and dehydrated larvae were rehydrated into PBST by 5 min washes in the reversed ethanol gradient. Selected larvae were properly oriented and embedded in 1% agarose gel. The blocks of gel containing the fish were cut out and dehydrated in 100% ethanol by gentle shaking for 5–8 h at 4°C. Next, the blocks were infiltrated overnight on a rotating stirrer at 4°C with JB-4 (Polysciences Inc.) catalyzed solution A (10 ml solution A: 0.125 g catalyst). The infiltration process with fresh infiltration solution continued on the following day for 5–6 h. The infiltrated samples were placed in embedding molds filled with a mixture of JB-4 solution B and fresh infiltration solution (1 ml solution B: 25 ml infiltration solution) and left overnight at 4°C to polymerize. Sections of 6-μm thickness were cut on a microtome using glass knives. Of two adjacent sections, one was mounted on a glass slide and stained with methylene blue, while the other, intended for synchrotron x-ray fluorescence imaging, was fixed on a Thermanox plastic coverslip (Gibco BRL) without any further processing. Before imaging, the plastic coverslip was mounted between the two 6.3-μm films of polypropylene.

X-ray Fluorescence Imaging.

X-ray fluorescence images were collected at the Stanford Synchrotron Radiation Laboratory (SSRL) using beamlines 9–3 and 2–3, and at the Canadian Light Source (CLS, Saskatoon, SK, Canada) using the Hard X-ray Microanalysis Beamline (HXMA). The incident x-ray energy was set to 12.7 keV and the Hg Lα1,2 and Ca, S, and Zn Kα1,2 fluorescence lines, as well as the intensity of the total scattered X-rays, were monitored using a silicon-drift Vortex detector (SII NanoTechnology USA Inc.).
Experiments on SSRL beamline 9–3 used a Si (220) double crystal monochromator and Rh-coated silicon mirrors for focusing and harmonic rejection, with the storage ring Stanford Positron Electron Accelerating Ring (SPEAR) containing 90–100 mA at 3.0 GeV. The microfocused beam of 10 μm was provided by a tapered glass capillary. The incident and transmitted x-ray intensities were measured with nitrogen-filled ion chambers. Samples were mounted at 90° to the incident x-ray beam and were spatially rastered in the microbeam using a Newport PM500 stage. Beam exposure was <1.5 sec per pixel for live zebrafish and ≈3.5 sec per pixel for sections.
Experiments on SSRL beamline 2–3 used a Si (220) double-crystal monochromator with harmonic rejection achieved by detuning one monochromator crystal to 50% peak intensity. The microfocused beam of 2.5 μm was provided by a Kirkpatrick-Baez mirror pair (Xradia Inc.) with the sample at 45° to the incident x-ray beam. Other experimental details were similar to those of SSRL beamline 9–3 (described above).
Experiments at the CLS HXMA beamline used a Si (220) double crystal monochromator and Rh-coated collimating and focusing beamline mirrors. Harmonic rejection was achieved by detuning one monochromator crystal to 50% peak intensity, with the storage ring operating at 200–250 mA at 2.9 GeV. The microfocused beam of 12 μm was provided by a Kirkpatrick-Baez mirror pair (fabricated at CLS), with the sample at 45° to the incident beam. Other experimental details were similar to those at SSRL (described above).

Data Analysis.

The data were processed using SMAK software (www-ssrl.slac.stanford.edu/∼swebb/smak.htm). Windowed Hg Lα1,2 and Zn Kα1,2 fluorescence data were corrected for scatter as previously described (37). The light element fluorescence signal in each image was corrected by calculating the average background intensity using the pixels outside the tissue and subtracting it from all of the pixels in the image. Quantities of Hg per pixel were calibrated using two certified highly uniform thin film standards on 6.3 μm-thick mylar substrates (Micromatter Co.), containing 16.3 and 17.1 μg/cm2 Au and TlCl, respectively. The scatter-corrected Au and Tl Lα1,2 fluorescence intensities were used to interpolate a Hg Lα1,2 fluorescence intensity and this was applied to the scatter-corrected Hg distribution maps to obtain the quantities of Hg per pixel in μg/cm2. Quantification was not possible for images of intact fish because the thickness is not known accurately. The attenuation of the incident and fluorescent x-rays is also non-trivial and means that a signal from the back of the sample is attenuated in a complex way by the sample. Sensitivity will depend both on the beamline used and on count-times per pixel, and was estimated using standard methods (38) to be 0.02 μg/cm2 for the quantified data collected on SSRL beamline 9–3. Similarly, signal-to-noise ratios were estimated, as peak signal intensity divided by the full-width half-maximum noise levels, to be ≈25 for typical data sets (SSRL, 9–3, 3.5 sec per pixel), although this improves with longer count times per pixel.


We thank Amy MacKay, Tomasz Korbas, Manuel Gnida, Roger Prince, Sam Webb, and Limei Zhang for their assistance, and William Talbot and Tuky Reyes for generously providing live zebrafish embryos for some of the experiments. We thank members of the George, Pickering, and Krone research groups, and staff at the Stanford Synchrotron Radiation Laboratory (SSRL) and at the Canadian Light Source (CLS). This work was supported by the CIHR Grant 34417, Canada Research Chairs (I.J.P. and G.N.G.), Jarislowsky Chair in Biotechnology (P.H.K.), and an NSERC Discovery Grant (P.H.K.). SSRL is supported by the DOE OBES. The SSRL Structural Molecular Biology Program is supported by the DOE OBER and by the NIH NCRR BTP. The CLS is supported by NSERC, NRC, CIHR, and the University of Saskatchewan.

Supporting Information

Supporting Information (PDF)
Supporting Information


TW Clarkson, L Magos, The toxicology of mercury and its chemical compounds. Crit Rev Toxicol 36, 609–662 (2006).
TA Simmons-Willis, AS Koh, TW Clarkson, N Ballatori, Transport of a neurotoxicant by molecular mimicry: the methylmercury-L-cysteine complex is a substrate for human L-type large neutral amino acid transporter (LAT) 1 and LAT2. Biochem J 367, 239–246 (2002).
R Hoffmeyer, et al., Molecular mimicry in mercury toxicology. Chem Res Toxicol 19, 753–759 (2006).
S Gasso, et al., Antioxidant compounds and Ca2+ pathway blockers differentially protect against methylmercury and mercuric chloride neurotoxicity. J Neurosci Res 66, 135–145 (2001).
E Vicente, et al., Cerebrospinal fluid S100B increases reversibly in neonates of methyl mercury-intoxicated pregnant rats. Neurotoxicology 25, 771–777 (2004).
TL Limke, SR Heidemann, WD Atchison, Disruption of intraneuronal divalent cation regulation by methylmercury: are specific targets involved in altered neuronal development and cytotoxicity in methylmercury poisoning? Neurotoxicology 25, 741–760 (2004).
C Johansson, et al., Neurobehavioural and molecular changes induced by methylmercury exposure during development. Neurotox Res 11, 241–260 (2007).
H Qu, T Syversen, M Aschnew, U Sonnewald, Effect of methylmercury on glutamate metabolism in cerebellar astrocytes in culture. Neurochem Intl 43, 411–416 (2003).
G-W Hwang, T Furuchi, A Naganuma, A ubiquitin-proteasome system is responsible for the protection of yeast and human cells against methylmercury. FASEB J 16, 709–711 (2002).
G-W Hwang, Y Ishida, A Naganuma, Identification of F-box proteins that are involved in resistance to methylmercury in Saccharomyces cerevisiae. FEBS Lett 580, 6813–6818 (2006).
T Furuchi, G-W Hwang, A Naganuma, Overexpression of the ubiquitin-conjugating enzyme Cdc34 confers resistance to methylmercury in Saccharomyces cerevisiae. Mol Pharmacol 61, 738–741 (2002).
AJ Hill, H Teraoka, W Heideman, RE Peterson, Zebrafish as a model vertebrate for investigating chemical toxicity. Toxicol Sci 86, 6–19 (2005).
JC Samson, J Shenker, The teratogenic effects of methylmercury on early development of the zebrafish, Danio rerio. Aquat Toxicol 48, 343–354 (2000).
JC Samson, R Goodridge, F Olobatuyi, JS Weis, Delayed effects of embryonic exposure of zebrafish (Danio rerio) to methylmercury (MeHg). Aquat Toxicol 51, 369–376 (2001).
P Gonzalez, Y Dominique, JC Massabuau, A Boudou, JP Bourdineaud, Comparative effects of dietary methylmercury on gene expression in liver, skeletal muscle, and brain of the zebrafish (Danio rerio). Environ Sci Technol 39, 3972–3980 (2005).
IJ Pickering, GN George, X-ray absorption spectroscopy imaging of biological tissues. AIP Conference Proceedings 882, 311–315 (2007).
AM El-Sherbeeny, JV Odom, JE Smith, Visual system manifestations due to systemic exposure to mercury. Cutan Ocul Toxicol 25, 173–183 (2006).
HH Harris, IJ Pickering, GN George, The chemical form of mercury in fish. Science 301, 1203 (2003).
R Dahm, HB Schonthaler, AS Soehn, J van Marle, GFJM Vrensen, Development and adult morphology of the eye lens in the zebrafish. Exp Eye Res 85, 74–89 (2007).
H Bloemendal, et al., Ageing and vision: structure, stability and function of lens crystallins. Prog Biophys Mol Biol 86, 407–485 (2004).
VN Reddy, Glutathione and its function in the lens – an overview. Exp Eye Res 50, 771–778 (1990).
SG Dove, Ontological changes in the crystallin composition of the eye lenses of the territorial damselfish Parma microlepis and their possible effects on trace-metal accumulation. Marine Biol 134, 653–663 (1999).
G DuVal, BR Grubb, PJ Bentley, Mercury accumulation in the eye following administration of methylmercury. Exp Eye Res 44, 161–164 (1987).
K Kairada, K Hirayama, A Yasutake, A Matsumura, R Okamura, Mercury accumulation in lens following single administration of methylmercury. Jpn J Ophthalmol 32, 275–280 (1988).
DN Weber, et al., Selenomethionine reduces visual deficits due to developmental methylmercury exposures. Physiol Behav 93, 250–260 (2008).
S Sabelaish, G Hilmi, Ocular manifestations of mercury poisoning. Bull World Health Organ 53, 83–86 (1976).
M Harada, Minamata disease: methylmercury poisoning in Japan caused by environmental pollution. Crit Rev Toxicol 25, 1–24 (1995).
M Saldana, CE Collins, R Gale, O Backhouse, Diet-related mercury poisoning resulting in visual loss. Brit J Ophthalmol 90, 1432–1434 (2006).
C Sanfeliu, J Sebastia, R Cristofol, E Rodriguez-Farre, Neurotoxicity of organomercurial compounds. Neurotox Res 5, 283–306 (2003).
BD Corbin, et al., Metal chelation and inhibition of bacterial growth in tissue abscesses. Science 319, 962–965 (2008).
S Yoon, EW Miller, Q He, PH Do, CJ Chang, A bright and specific fluorescent sensor for mercury in water, cells, and tissue. Angew Chem Int Ed 46, 6658–6661 (2007).
S-Y Ko, Y-K Yang, J Tae, I Shin, In vivo monitoring of mercury ions using a rhodamine-based molecular probe. J Am Chem Soc 128, 14150–14155 (2006).
B Tang, LJ Cui, KH Xu, LL Tong, GW Yang, LG An, A sensitive and selective near-infrared fluorescent probe for mercuric ions and its biological imaging applications. ChemBioChem 9, 1159–1164 (2008).
GN George, et al., Mercury binding to the chelation therapy agents DMSA and DMPS and the rational design of custom chelators for mercury. Chem Res Toxicol 17, 999–1006 (2004).
CB Kimmel, WW Ballard, SR Kimmel, B Ullmann, TF Schilling, Stages of embryonic development of the zebrafish. Dev Dyn 203, 253–310 (1995).
M Westerfield The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Danio rerio) (University of Oregon Press, Eugene, OR, 1995).
IJ Pickering, RC Prince, DE Salt, GN George, Quantitative, chemically-specific imaging of selenium transformation in plants. Proc Natl Acad Sci USA 97, 10717–10722 (2000).
M Mantler Handbook of Practical X-Ray Fluorescence Analysis, eds B Beckhoff, B Kanngieβer, N Langhoff, R Wedell (Springer, Berlin, 2006).

Information & Authors


Published in

Go to Proceedings of the National Academy of Sciences
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Proceedings of the National Academy of Sciences
Vol. 105 | No. 34
August 26, 2008
PubMed: 18719123


Submission history

Received: March 31, 2008
Published online: August 26, 2008
Published in issue: August 26, 2008


  1. methylmercury
  2. thimerosal
  3. x-ray fluorescence mapping
  4. eye lens


We thank Amy MacKay, Tomasz Korbas, Manuel Gnida, Roger Prince, Sam Webb, and Limei Zhang for their assistance, and William Talbot and Tuky Reyes for generously providing live zebrafish embryos for some of the experiments. We thank members of the George, Pickering, and Krone research groups, and staff at the Stanford Synchrotron Radiation Laboratory (SSRL) and at the Canadian Light Source (CLS). This work was supported by the CIHR Grant 34417, Canada Research Chairs (I.J.P. and G.N.G.), Jarislowsky Chair in Biotechnology (P.H.K.), and an NSERC Discovery Grant (P.H.K.). SSRL is supported by the DOE OBES. The SSRL Structural Molecular Biology Program is supported by the DOE OBER and by the NIH NCRR BTP. The CLS is supported by NSERC, NRC, CIHR, and the University of Saskatchewan.


This article is a PNAS Direct Submission. J.P.-H. is a guest editor invited by the Editorial Board.
This article contains supporting information online at www.pnas.org/cgi/content/full/0803147105/DCSupplemental.



Malgorzata Korbas
Department of Geological Sciences, University of Saskatchewan, 114 Science Place, Saskatoon, Saskatchewan S7N 5E2, Canada; and
Scott R. Blechinger
Department of Anatomy and Cell Biology, University of Saskatchewan, 107 Wiggins Road, Saskatoon, Saskatchewan S7N 5E5, Canada
Present address: Existing Substances Division, Health Canada, 4904D-269 Laurier Avenue, Ottawa, Ontario K1A 0K9, Canada.
Patrick H. Krone
Department of Anatomy and Cell Biology, University of Saskatchewan, 107 Wiggins Road, Saskatoon, Saskatchewan S7N 5E5, Canada
Ingrid J. Pickering
Department of Geological Sciences, University of Saskatchewan, 114 Science Place, Saskatoon, Saskatchewan S7N 5E2, Canada; and
Graham N. George§ [email protected]
Department of Geological Sciences, University of Saskatchewan, 114 Science Place, Saskatoon, Saskatchewan S7N 5E2, Canada; and


To whom correspondence should be addressed. E-mail: [email protected]
Author contributions: M.K., S.R.B., P.H.K., I.J.P., and G.N.G. designed research; M.K., S.R.B., I.J.P., and G.N.G. performed research; M.K. and G.N.G. analyzed data; M.K., P.H.K., I.J.P., and G.N.G. wrote the paper.

Competing Interests

The authors declare no conflict of interest.

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    Localizing organomercury uptake and accumulation in zebrafish larvae at the tissue and cellular level
    Proceedings of the National Academy of Sciences
    • Vol. 105
    • No. 34
    • pp. 12095-12636







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