We have developed an X-ray absorption near edge structure spectroscopy method using fluorescence detection for visualizing in vivo coordination environments of metals in biological specimens. This approach, which we term fluorescence imaging XANES (φXANES), allows us to spatially depict metal-protein associations in a native, hydrated state whilst avoiding intrinsic chemical damage from radiation. This method was validated using iron-challenged Caenorhabditis elegans to observe marked alterations in redox environment.
Metal cofactors represent a fundamental component of biochemistry via their ability to facilitate electron transport and stabilize biomolecules. An important requirement for understanding the role of transition metals in normal biochemistry and disease processes is the determination of their coordination environment1,2. The synergy of synchrotron-based X-ray fluorescence microscopy (XFM) and X-ray absorption near edge structure (XANES) spectroscopy represents a powerful analytical approach for studying metal biochemistry at the micro-scale. This permits both quantitative mapping of metal distribution and profiling of the native coordination environment without the need for exogenous molecular probes3. We combined these two measurement strategies using the same synchrotron beamline to develop an imaging approach we have called fluorescence imaging (‘fi’, or φ for the Greek ‘phi’) φXANES.
XANES has traditionally been used in biology to profile coordination environments in fixed locations (‘point’ XANES)4, rather than functioning as a fine resolution imaging technique. In addition to lack of spatial information is the problem of extended exposure to the ionizing X-rays (>10 keV) that can damage the sample by disrupting chemical bonds. Photoreduction of redox metals can occur at doses around 107 Gy5,6. Exposure of XANES samples can be increased to 1010 Gy using cryogenic conditions (−100 °C)7, though samples are still susceptible to morphological damage8 and the requirement to maintain the specimen at low temperatures during preparation and measurement increases logistical complexity. Ideally, analysis of samples that remain hydrated and at physiological temperature is preferable. Here, we demonstrate the development of non-destructive φXANES imaging at standard laboratory conditions, validated in a Caenorhabditis elegans model of disrupted metal metabolism.
To determine the optimal conditions for φXANES (see Supplementary Note), we tested two experimental scenarios to establish the appropriate dose of radiation to which hydrated and anesthetized C. elegans could be exposed without inducing morphological changes and to avoid photoreduction of endogenous iron. These were: i) ‘high dose’ φXANES, where elemental maps of high statistical precision were obtained using long dwell and spatial oversampling in two directions with a symmetrical beam profile; and ii) ‘low dose’ φXANES, where a shorter dwell time was used along with a vertically-elongated beam (reducing the X-ray flux density), while undersampling in the vertical direction (for details of the focused beam see Supplementary Figure 1). Together, these measures reduced sampling time and localized beam exposure by a factor of 100. A representative whole-body XFM elemental map of calcium and iron distribution in a separate cryofixed and lyophilized specimen is presented for anatomical reference (Fig. 1a). For high dose φXANES, four hydrated adults were mapped using standard XFM parameters (Fig. 1b), with an anterior region containing iron-rich intestinal cells in one specimen selected for φXANES. Here, the region underwent XANES analysis spanning the iron K-edge incident energies (7100 to 7220 eV), exposing this region to an estimated 5 × 108 Gy, within the radiation dose range previously reported to stimulate photoreduction of iron6.
These four individuals were mapped again using standard XFM following φXANES, approximately 5 hours after the initial XFM scan. We performed intensity correlation analysis (ICA)9 on a region of interest representative of the pre and post-φXANES scanned area and found that distribution of pixel intensities significantly differed between the two maps (ICA quotient Q = 0.006; 0 = no correlation), demonstrating clear sample damage. In parallel, we selected an anatomically equivalent adjacent specimen that received ~106 Gy from the two XFM maps alone which, in contrast, maintained a consistent iron distribution (Q = 0.37; 0.5 = perfect correlation).
Low dose φXANES and XFM of a matching sample group were then examined (Fig. 1c), encompassing an additional four adults. These samples were exposed to 4 × 106 Gy, approximately 100-fold less than the high dose method. Iron spatial distribution pre- and post φXANES was maintained (Q = 0.41). Although low dose φXANES does sacrifice some spatial detail (5.6 μm2 versus 0.64 μm2 sampling area), the reduced radiation dose (<2 × 108 Gy) ensures photoreduction of iron6 and other metals10 is minimized while endogenous spatial distribution is maintained.
To demonstrate the potential of φXANES to profile bioinorganic chemistry in vivo, we examined a combined genetic and exogenously challenged model of severe iron dyshomeostasis. C. elegans lacking the iron-storage protein ferritin (both genes ftn-1 and ftn-2 are ablated via mutation; hereafter referred to as ferritin nulls) have increased oxidative load from elevated ferrous iron11, and have a shortened lifespan compared to wild type (Supplementary Figure 2). We designed two experimental paradigms, exposing both wild type and ferritin nulls to either basal iron levels via normal culturing conditions, or high iron through supplementation of their growth media. Adults were anesthetized and quantitatively mapped by XFM (Fig. 2a). Exposure to high iron increased levels in wild type animals compared to equivalent animals raised under basal conditions (one-way ANOVA with Tukey’s post hoc test; p < 0.001; Fig. 2b), whilst ferritin nulls raised on basal iron exhibited a decrease in total levels compared to wild type (p < 0.001). As ferritin is not involved in iron uptake, the reduced load is consistent with an inability to store iron12; ferritin nulls on high iron still displayed increased total body burden (p < 0.001).
Each experimental group was mapped via φXANES using our optimized low dose parameters, scanning the iron K-edge (Fig. 2c). This range encompasses the characteristic pre-edge (~7115 eV), shoulder (~7124 eV), and crest (~7130 eV) features, arising from 1s → 3d, 1s → 4s and 1s → 4p electronic transitions, respectively. The precise energy of the pre-edge reflects the relative abundance of ferrous [Fe(II)] and ferric [Fe(III)] iron, and shifts to lower energies in the presence of increased Fe(II)13. When comparing the pooled XANES spectra (i.e. the mean for all pixels) for each measured individual, we observed that the centroid energy for the pre-edge transition in wild type (7114 eV; Fig. 2d) cultured on high iron was unchanged but the reduced intensity was indicative of an increase in the number of octahedral Fe(III) centres14, consistent with increased buffering of iron within ferritin, where it is arranged in such coordination geometry15. However, ferritin nulls, regardless of iron load, demonstrated a shift to lower centroid energies away from 7114 eV, indicating increased Fe(II). Comparing the relative intensity of the shoulder and crest features of the iron K-edge in each group further confirmed a disruption in the iron coordination environment. First-derivative iron XANES spectra exhibited a significant alteration in the cumulative Fe(III):total iron ratio between wild type and ferritin null groups (one-way ANOVA with Tukey’s post hoc test; p < 0.001; Supplementary Fig. 3a,b). This effect was independent of iron loading. In addition, we observed increased variability between ferritin nulls compared to wild type (Bartlett’s test for homogeneity of variances χ2 = 16.54; p < 0.001; Supplementary Fig. 3c), consistent with a homeostatic system in distress.
Potentially hundreds of individual iron-binding proteins contribute to the proteome (the ferroproteome), although even in microbes the precise number remains unclear16. These include proteins containing heme moieties, iron-sulfur clusters, ferrihydrite-like crystalline structures (as in ferritin), and multi-dentate ligands arising from specific amino acid conformations17. When examining the cumulative φXANES spectra, we are assessing the aggregate distribution of iron-protein coordination complexes in a whole organism. Spatial mapping by φXANES allows for individual tissue or cell types to be objectively assessed for changes to iron coordination in response to specific challenges at the μm scale. We applied principal component analysis (PCA) and k-means clustering (CA) as implemented in the Multivariate ANalysis Tool for Spectromicroscopy (MANTiS)18 package after tiling φXANES maps to directly compare spatial coordination states in wild type and ferritin nulls raised on high iron. Pixels with similar XANES spectra were assigned to six distinct regions of interest (ROIs), color coded in Fig. 2e as descending Fe(III):total iron. Of these six regions, ROIs 1 and 5 differed in proportions of total iron-containing pixels between genotypes (Supplementary Table 1). There was a systemic shift towards a lower Fe(III):total iron in ferritin nulls compared to wild type. The intestine consists of highly metabolically active cells in C. elegans, which in the ferritin null animals demonstrated the largest shift in iron redox balance, with ROI5 essentially replacing the spatial distribution of ROI1 (Fig. 2e). We confirmed that the absence of ferritin mapped to the changes in ROIs 1 and 5 by subtracting the XANES spectra of purified horse spleen ferritin (the ferrihydrite-like iron core of ferritin is a commonality in all species expressing ferritin homologues12) from each ROI (ΔXANES; Fig. 2f–h). While ROI1 (absent in ferritin nulls) shared stark similarities with the reference ferritin standard, ROI5 (markedly increased in ferritin nulls) showed XANES spectra that deviated significantly (Wilcoxon signed-rank test; W = 734; p < 0.05). Finally, the clear splitting in centroid energy of ferritin-absent ROI5 further supports the higher levels of Fe(II) in the ferritin null animals, consistent with the well characterized role of ferritin in buffering reactive ferrous iron as a redox-silenced mineralized Fe(III) species. For comparison, XANES spectra of additional iron-protein ligands (oxidized and reduced heme-containing cytochrome c) are shown in Supplementary Figure 4.
In summary, we have demonstrated that φXANES is a powerful method for mapping coordination environments in vivo, with no displacement of target elements and measurement dose well below previous studies of biological iron redox status inline with bulk XAS measurements, and without the need for cryogenic sample environment. φXANES in conjunction with PCA-CA is ideal for assessing changing coordination environments in tissue sections, small model organisms (including C. elegans and Drosophila melanogaster, which has previously been used for point XANES19) and cell culture. Although we validated this method using iron coordination, φXANES can be applied to any element to which XFM is sensitive, drugs that elicit a change in cellular redox environment, and longitudinal studies that require real-time assessment of changing coordination conditions in a biological system.
Methods and any associated references are available in the online version of this paper.
How to cite this article: James, S. A. et al. ϕXANES: In vivo imaging of metal-protein coordination environments. Sci. Rep. 6, 20350; doi: 10.1038/srep20350 (2016).
We thank Daryl L. Howard, David Paterson, and Peter Kappen (Australian Synchrotron) for experimental assistance and discussions. Parts of this research were undertaken on the XFM and XAS beamlines at the Australian Synchrotron. We acknowledge the Caenorhabditis Genetics Center of the US National Institutes of Health - Office of Research Infrastructure Programs (P40OD010440) for materials. The Australian Research Council Discovery Projects scheme (DP130100357), National Health and Medical Research Council and the Victorian Government’s Operational Infrastructure Support Program funded this work.