text:
- Research article
- Open Access
High-resolution quantitative imaging of mammalian and bacterial cells using stable isotope mass spectrometry
© Lechene et al.; licensee BioMed Central Ltd. 2006
- Received: 21 April 2006
- Accepted: 09 June 2006
- Published: 5 October 2006
Secondary-ion mass spectrometry (SIMS) is an important tool for investigating isotopic composition in the chemical and materials sciences, but its use in biology has been limited by technical considerations. Multi-isotope imaging mass spectrometry (MIMS), which combines a new generation of SIMS instrument with sophisticated ion optics, labeling with stable isotopes, and quantitative image-analysis software, was developed to study biological materials.
The new instrument allows the production of mass images of high lateral resolution (down to 33 nm), as well as the counting or imaging of several isotopes simultaneously. As MIMS can distinguish between ions of very similar mass, such as 12C15N- and 13C14N-, it enables the precise and reproducible measurement of isotope ratios, and thus of the levels of enrichment in specific isotopic labels, within volumes of less than a cubic micrometer. The sensitivity of MIMS is at least 1,000 times that of 14C autoradiography. The depth resolution can be smaller than 1 nm because only a few atomic layers are needed to create an atomic mass image. We illustrate the use of MIMS to image unlabeled mammalian cultured cells and tissue sections; to analyze fatty-acid transport in adipocyte lipid droplets using 13C-oleic acid; to examine nitrogen fixation in bacteria using 15N gaseous nitrogen; to measure levels of protein renewal in the cochlea and in post-ischemic kidney cells using 15N-leucine; to study DNA and RNA co-distribution and uridine incorporation in the nucleolus using 15N-uridine and 81Br of bromodeoxyuridine or 14C-thymidine; to reveal domains in cultured endothelial cells using the native isotopes 12C, 16O, 14N and 31P; and to track a few 15N-labeled donor spleen cells in the lymph nodes of the host mouse.
MIMS makes it possible for the first time to both image and quantify molecules labeled with stable or radioactive isotopes within subcellular compartments.
- Additional Data File
- Ionization Yield
- Ratio Image
- Mass Image
- Mass Line
Although there has been pioneering work using SIMS in biology [10–14], SIMS technology, until now, has presented irreconcilable tradeoffs [15] that have severely limited its use as a major discovery tool in biomedical research. To make secondary-ion methodology practicable for locating and measuring isotope tags in subcellular volumes, four major issues need to be addressed. First, to produce quantitative ultrastructural images, the technique must have sufficiently high spatial resolution, and quantitation and imaging must be associated. Second, because the quantitation of label involves measuring the excess of an isotope tag above its natural occurrence, and this excess is calculated by the ratio of two isotopes, the data from the two isotopes should be recorded simultaneously, in parallel, and from exactly the same region of the sample (that is, in register). This is to ensure that changes in instrument or sample conditions do not lead to errors in the calculated isotope ratios. Third, because nitrogen has an electron affinity of zero, N- ions do not form; nitrogen must therefore be detected as cyanide ions (CN-). Consequently, in order to use the stable isotope 15N as a label, the mass resolution of the instrument needs to be high enough to separate the ions 12C15N- and 13C14N-, which have the same mass number of 27, but do not have exactly the same atomic mass weight: differing by 0.00632 mass unit, less than 1 part in 4,000. Finally, the high mass resolution should not be at the expense of the secondary-ion current; the transmission of secondary ions from sample to detector needs to be high enough to allow data collection from sub-cubic-micrometer volumes and in a reasonable amount of time.
These four requirements necessitate a previously unattainable combination of instrumental capabilities, with the ability to collect large numbers of secondary ions at high mass resolution, parallel detection of several secondary ions, high lateral resolution, and high precision of measurements. A new generation of secondary-ion mass spectrometer has now been developed that can measure several ion masses in parallel, has a high mass resolution (mass/change in mass ratio of approximately 10,000) at high secondary-ion relative transmission (70–80%), and has a high lateral resolution, down to 33 nm [16, 17]. In this paper we present some biological applications of this new technology. These extraordinary capabilities allow us, for example, to image and measure in parallel the intracellular distribution of molecules labeled with the stable isotopes 15N or 13C because we can separate the isobaric species (that is, the species with the same atomic masses): 12C15N- from 13C14N- or 13C- from 12C1H-. Because the images are produced in parallel from the same sputtered volume they are in exact register with each other and these characteristics are necessary for obtaining quantitative atomic mass images. A quantitative mass image contains at each pixel a number of counts, which are a measure of the selected atomic mass and are directly proportional to the selected atomic mass abundance in the sample, at a location corresponding to the pixel address. Counts from several atomic masses, originating from the same location in the sample, can be recorded in parallel at the same pixel address, which allows us to derive meaningful isotope ratios. Isotope ratios are at the core of the methodology. When the sample has been labeled with a given isotope, a ratio higher than its natural abundance indicates the presence of the marker isotope at a particular location as well as measuring its relative excess. In addition, the high stabilities of the primary beam, the ion optics, the mass spectrometer and the detectors contribute to very precise measurements.
We have developed the use of this new generation of SIMS, together with tracer methods and quantitative image-analysis software, for locating and measuring molecules labeled with stable isotopes in subcellular compartments, a development that we call multi-isotope imaging mass spectrometry (MIMS). In this paper we present a range of examples showing how MIMS can be used to provide atomic mass images of biological specimens, and how in combination with stable isotope labeling it provides qualitative and quantitative information that is not possible to obtain with other methods.
Imaging of unlabeled cells and tissue sections
The high spatial resolution of MIMS is illustrated by its ability to image individual stereocilia, the mechanosensory organelles of the inner hair cells of the cochlea (Figure 2e; the field analyzed was 6 × 6 μm). The various intracellular structures are easy to identify by comparing the image with electron micrographs of the same structure. We estimate that a lateral resolution better than 33 nm can be achieved using a method derived from the 'knife-edge' technique (see [Additional data file 2]). Because the atomic mass image is formed using only a few atomic layers of the sample, the depth resolution (z resolution) can be smaller than 1 nm, much better than the resolution that would be provided by an exceptionally thin electron-microscopy section (10 nm). (In this paper, we define depth resolution as the minimum amount of material that needs to be sputtered to obtain an atomic mass image.) Studies of the same sections with both MIMS and electron microscopy – a technique developed for studying cosmic dust [19] – may help provide complementary structural observations.
We do not know from first principles the mechanism of contrast formation in atomic mass images from unstained samples. A striking observation of intense contrast observed with MIMS at mass 12C14N, shown in Figure 2g–i, may guide inquiries in this field. We were analyzing a mouse kidney when we observed a brightly contrasted structure, a circular 'snake', along the lining of the lumen of an artery. It is likely that this region is the 'elastica interna' as described in histology texts, a tissue layer that is usually visible only after the arterial tissue has been specially stained (Figure 2f). The MIMS images of the artery in Figure 2g–i were obtained without this special staining. The images indicate that this structure produces a high yield of 12C14N- ions in comparison with the other regions of the artery, and suggests a relationship between the image obtained and molecular composition and density.
MIMS can be used to visualize whole cells as well as sections. These images have a three-dimensional appearance, showing that MIMS can provide scanning atomic or molecular ion mass images of samples with relief, as does scanning electron microscopy. The lamellipodium of a well-spread endothelial cell imaged by MIMS at mass 12C14N is shown in Figure 2j; it appears as a light, sheet-like structure with darker lines radiating from the cytoplasm to the external border of the lamellipodium. In contrast, if actin polymerization is blocked by cytochalasin D treatment, the 12C14N mass image of an endothelial cell shows no lamellipodia but only thin, spike-like projections around the cell circumference, which are most probably retraction fibers (Figure 2k).
In conclusion, MIMS atomic mass images of CN- ions in biological samples are highly contrasted, even though they are obtained without any staining. The various structures are easy to identify down to a lateral resolution of approximately 33 nm, and the depth resolution can be as small as a few atomic layers. The largest single field that can be imaged is approximately 140 × 140 μm, but larger fields can be documented quickly by taking a series of images for 1–2 minutes each. There is no machine-specific requirement for the sample except that a vacuum must be sustained. Because MIMS is a surface-analysis method, one can use many kinds of samples: for example, tissue or cell sections embedded in a medium such as epon, or cells cultured directly on a support that can be brought into the analysis chamber and prepared with any usual histological technique. The thickness of the sample is not critical, provided that the electrical charges deposited can be dissipated. The surface analyzed does not have to be flat, and one can obtain SIMS images of three-dimensional samples.
Quantitative labeling with stable isotopes
Having established that MIMS can be used to obtain atomic mass images of unstained biological objects, this led us to develop the unique feature of MIMS: the quantitative analysis of isotopes within subcellular compartments. We will now discuss how the technique can be used to measure the incorporation of isotopic tracers within compartments of sub-cubic-micrometer volume. To do this, the sample is labeled with stable isotopes such as 15N or 13C, which are present at much lower levels naturally than their counterpart 14N and 12C isotopes, and each isotope is then measured to determine whether it is present in an amount exceeding its natural abundance. Stable isotope labeling can be used, for example, to pursue the classic studies of Schoenheimer at the subcellular level [1].
The isotope abundance is measured by recording the secondary-ion currents (counts/time) obtained from a pair of isotopes, for example, 13C and 12C, calculating the ratio and then comparing it with their natural abundance ratio. In a control sample, which has not received an excess of the tracer isotope, the counts of each isotope are related to each other by their natural abundance. In other words, there will be a count of 13C or of 15N such that 13C/12C = 1.12%, or 15N/14N (measured as 12C15N/12C14N) = 0.367%, calculated from the values of their respective natural abundance. This means that when measured in parallel, all the analytical conditions being the same, the 15N (12C15N) count rate will be 272 times lower than the 14N (12C14N) count rate.
Quantitative labeling with 15N
The goal of our first experiments was to ensure that we could measure 15N/14N ratios equivalent to their natural abundance from tiny volumes of untreated control sample. Our first analyses were carried out using a stationary cesium (Cs+) primary ion beam. We counted in parallel the secondary ions 12C14N- and 12C15N- emitted from various areas smaller than 1 square micrometer in control samples of mouse tissues. We measured 12C15N/12C14N isotope ratios in control mouse tissue of 0.366% (standard error (SE) = 0.002, n = 12) in the cochlea, 0.368% (SE = 0.001, n = 14) in the kidney, and 0.368% (SE = 0.001, n = 6) in the intestine. These values are not statistically significantly different from the natural terrestrial value of the 15N/14N isotope ratio, 0.3673% [20]. This proved the feasibility of using this method on biological samples.
We then showed that we could measure the incorporation of a stable isotope label in an ultra-minute volume of biological material, as done for bulk tissue 60 years ago [1]. We fed mice a diet slightly enriched with 15N-L-leucine for a sufficient length of time (14 days) to result in total protein renewal in kidney and intestine. The 15N/14N isotope ratios determined using a stationary primary ion beam at various areas over the samples were equivalent to the 15N/14N ratio in the diet determined independently by combustion mass spectrometry analysis (intestine 4.45‰, SE = 0.05, n = 7; kidney 4.41‰, SE = 0.03, n = 12; diet 4.45‰, SE = 0.02, n = 7).
Using this method, only one location can be analyzed at a time and its precise position is difficult to ascertain in the absence of an image. With our instrument, we have developed a much more powerful but more complex method of isotope ratio imaging, where the isotope ratios are calculated from quantitative mass images obtained simultaneously from a set of isotopes. A quantitative mass image, as we call it, is the representation of an analyzed field in which each pixel is the address of a register at which the secondary-ion current of an isotope of interest has been recorded during analysis. Up to four secondary-ion currents, representative of four isotopes, can be recorded simultaneously at each pixel address with our instrument, for example 12C-, 13C-, 12C14N- and 12C15N-. A quantitative image of 256 × 256 pixels thus represents a set of (256 × 256 × 4) or 262,144 numbers. We call a group of pixel addresses a 'region of interest', and the first step in quantitation is to extract the values of counts/time/isotope from groups of pixels or from individual pixels. This allows us to measure many more regions from a single analytical field than we could do using a stationary beam, and also to associate quantitation and localization among cells and subcellular domains. All the mass imaging in the rest of this paper will refer to quantitative mass imaging.
The ratio images 12C15N-/12C14N- (Figure 3c) and 13C-/12C- (Figure 3f) result from the pixel-by-pixel division of the 12C15N- image by the 12C14N- image and of the 13C- image by the 12C- image, respectively. The contrast observed in the 12C15N-/12C14N- image is due to the excess of 15N in the area of the cochlea that has incorporated 15N derived from the 15 N-L-leucine. The internal control 13C-/12C- ratio image has no contrast because, in the absence of added 13C, the value of the ratio is equivalent to the natural ratio in any part of the analyzed field.
Using the quantitative images and the derived ratio images, and guided by the hue saturation intensity (HSI) transformation (see Materials and methods), we can now calculate a value for 15N incorporation into the main structures shown in Figure 3a–f. This can be expressed as percentage renewal by comparing the excess 15N in the tissue with the excess 15N in the diet (see Materials and methods). These values represent overall protein renewal among the different cochlear structures, as demonstrated for whole tissue in the classic work of Schoenheimer [1]. After 9 days of a 15N-L-leucine diet, the incorporation of 15N is markedly different among specific cell types. The outer hair cells have a 15N renewal of 52.5% ± 1.8 SD, not significantly different from that of the Deiter cells (47.2% ± 4.8 SD), or of the proximal part of the outer pillar cell above the basilar membrane (46.0% ± 5.7 SD), and of one population of tympanic border cells (48.3% ± 1.5 SD). The basilar membrane has a small 15N renewal (overall 21.1% ± 6.0 SD), not statistically different from part of the outer pillar cells at the level of the Deiter cells (18.4% ± 3.7 SD). Overall, the reticular lamina has a 15N renewal of 30.8% ± 8.9 SD, significantly higher than the basilar membrane and the distal outer pillar cells, and significantly lower than that of the outer hair cells. The lone outer hair-cell nucleus observed has a 15N renewal of 35.5%. Finally, we measured a second population of tympanic border cells with a 15N renewal significantly greater than in any other area (72.8% ± 2.5 SD). The internal control provided by the epon embedding medium had a 12C15N/12C14N isotope ratio of 0.365% ± 0.089 SD, equivalent to the natural abundance ratio and corresponding to a 15N renewal of 0%.
The unique power of MIMS is demonstrated by the quantitative imaging of subcellular structures at high resolution, revealing sub-cubic-micrometer-sized zones with high 15N renewal, and thus probably high protein renewal. In addition, the experiment showed that the same sample can be analyzed repetitively at a variety of spatial resolutions. We analyzed one of the bundles of stereocilia (Sb1, indicated by a white arrow in Figure 3a) at high resolution; we used a field of 3 × 3 μm, a beam size of about 35 nm, and 256 × 256 pixels (Figure 3g–l). Mass images of the 12C-, 13C-, 12 C14N- and 12C15N- ions were acquired in parallel. The 12C14N- image (Figure 3g) shows one bundle of stereocilia and a fraction of the cuticular plate of one hair cell, barely visible in the lower-resolution image in Figure 3a.
As in the cochlear analysis, but at a subcellular level, the 12C15N- image (Figure 3h) is similar in form to the 12C14N- image (Figure 3g) but has much lower counts; the total number of counts of 12C15N- and of 12C14N- are 8.36 × 104 and 1.61 × 107, respectively. The 12C- image (Figure 3j) has relatively poor contrast compared with the 12C14N- image (Figure 3g), as most of the 12C- ions arise from the embedding medium. The 13C- image (Figure 3k) is similar to the 12C- image, but with a much lower count; the total number of counts for 13C- and 12C- are 4.80 × 105 and 4.36 × 107, respectively. The pixel counts from the 13C- image include the fraction of 13C related to the 12C content by the natural ratio of 13C/12C. The ratio images 12C15N-/12C14N- (Figure 3i) and 13C-/12C- (Figure 3l) result from the pixel-by-pixel division of the 12C15N- image by the 12C14N- image and of the 13C- image by the 12C- image, respectively. The contrast observed in the 12C15N-/12C14N- image is due to the excess of 15N in the stereocilia, cuticular plate, and hair-cell areas that have incorporated 15N derived from the 15N-L-leucine. The internal control 13C-/12C- ratio image (Figure 3l) has no contrast, as in Figure 3f.
An HSI transformation of the 12C15N-/12C14N- ratio image of the stereocilia bundle in Figure 3i is shown in Figure 3m. The colors indicate the fractional excess 15N derived from the measured 12C15N-/12C14N- isotope ratios. The HSI image reveals small areas of high excess 15N located towards the tips of stereocilia or between stereocilia (magenta); within the stereocilia, close to these areas, there is minimal or no excess 15N, as indicated by the predominantly blue-green to blue color.
Calculated percent nitrogen renewal from stereocilia regions analyzed in Figure 3m
Tip or lateral links
Pixels (n)
% Renewal
√ Area (Å)
Pixels (n)
% Renewal
√ Area (Å)
We can thus measure with high precision in a single analyzed field a variety of values of 15N incorporation among different cell types, as calculated from the quantitative mass images in Figure 3a–c, or among subcellular structure over an area of 9 μm2 square, as calculated from the quantitative mass images in Figure 3g–i.
Quantitative labeling with 13C
Despite the importance of free fatty acids (FFAs) for life, studies of their transport are difficult to extend to the cellular scale because no suitable methodology is available. Autoradiography cannot provide quantitative information on accumulation of FFAs in intracellular fat droplets, and fluorescently labeled FFAs may not accurately reflect the transport and metabolism of native FFAs [22, 23]. As a result, the mechanism that transports FFAs across a cell membrane remains uncertain (for recent reviews see [24–27]). Using quantitative mass imaging with MIMS we have directly studied the accumulation of 13C in cultured adipocytes incubated with 13C-labeled oleic acid (13C-OA; see [28] for further details). We measured a high level of 13C accumulation in intracellular lipid droplets. Quantitative MIMS images were obtained in parallel for 12C-, 13C-, 12C14N-, and the isobars 13C14N- and 12C15N-. The relative excess of 13C was measured at three different locations: outside the cell, inside the cell but outside the lipid droplets, and inside the lipid droplets.
The quantitative mass images of an adipocyte exposed to 13C-OA for 20 minutes are shown in Figure 4a–i. Images of the 12C-, 13C-, 12C14N- and 12C15N- ions or of the 12C-, 13C-, 12C14N- and 13C14N- ions were acquired in parallel. Images of the 12C- and 12C14N- ions (Figure 4a,d) show the cell histology. The mass image of the 13C- ion (Figure 4b) is similar to the 12C- ion image (Figure 4a) in form, but has a lower count rate. The pixel counts of the 13C- image include both the natural 13C and the supplementary 13C from the 13C-OA transported into the cell. This supplementary 13C is at a maximum at the intracellular lipid droplets, where FFAs accumulate. The mass image of the internal control 12C15N- ions (Figure 4e) is similar to the 12C14N- mass image, yet with a much lower count rate. Each pixel of the sample resulting in the 12C15N- image contains the fraction of 15N related to the 14N content by the natural ratio of 15N/14N. The enhanced contrast observed in the 13C-/12C- image (Figure 4c) is due to the excess 13C incorporated into the lipid droplets from the transported 13C-OA. The internal control 12C15N-/12C14N- ratio image (Figure 4f) has no contrast because in the absence of added 15N, the value of the ratio of 15N/14N is equivalent to the natural ratio across the analyzed field.
The HSI image of the 13C-/12C- ratio is shown in Figure 4g, and the same cell photographed by RDIC microscopy on the silicon chip before analysis with MIMS is shown in Figure 4h. The 13C-/12C- ratio, indirectly measured as the cyanide ion, 13C14N-/12C14N-, is shown in Figure 4i; this also shows accumulation of 13C- in the droplets. In contrast to the high 13C-/12C- ratios found in cells that were incubated with 13C-OA, cells washed with buffer solution after 13C-OA incubation had low 13C-/12C- ratios (images not shown). In cells not treated with 13C-OA, the value of the 13C-/12C- ratio measured under the same conditions was 1.15 ± 0.10%, not significantly different from the terrestrial 13C/12C value of 1.12%. An indication of the accuracy of these values was obtained from measurements of the 12C15N-/12C14N- ratio, whose value was 0.36 ± 0.01% in both washed and unwashed cells, in excellent agreement with the natural abundance of 0.37%. The cumulative values obtained from quantitative MIMS atomic mass images and extracted from the isotope ratio images are shown in Figure 4k.
We can remove material quickly from the sample surface in order to study a variety of depths within the cell. We refer to this as 'shaving' the sample. It is accomplished in conditions that give a high primary-ion beam current (such as by removing the objective diaphragm; see Figure 13 in the Discussion section). The results of such shaving are shown in Figure 4j. The adipocyte analyzed in Figure 4a–i was shaved using, for a few minutes, a primary beam current approximately 1,000-fold more intense than for the previous analysis. This uncovered a lipid droplet deeper in the cell with a very high 13C-/12C- ratio, as shown in the HSI image (Figure 4j). Finally, MIMS allows us to acquire hundreds of atomic mass image planes successively from the same cell, opening the door to full three-dimensional volume rendering. We have begun using this capability to study the distribution of 13C among the lipid droplets located within a single adipocyte after incubation with 13C-OA [29]. In conclusion, MIMS can be used to investigate lipid metabolism with high spatial and quantitative resolution [28]. Unlike other techniques, MIMS allows us to trace and to measure the movement of native FFAs at specific subcellular locations.
Quantitative labeling of prokaryotic cells with gaseous 15N
The ability to image and measure stable isotopes makes it easy and safe to apply MIMS to samples labeled with a gaseous precursor. Here we describe the application of MIMS to the study of nitrogen fixation in bacteria (Figure 5a–k). Teredinibacter turnerae is a diazotrophic (nitrogen-fixing) marine bacterium that can be isolated from the tissues of wood-boring marine bivalves (family Teredinidae) and grown in pure culture [30, 31]. Enterococcus faecalis is a bacterium that does not fix nitrogen. Both were cultured for 120 hours in a 15N atmosphere. Mass images of the 12C-, 13C-, 12C14N- and 12C15N- ions were acquired in parallel. T. turnerae is barely visible at mass 12C14N- (Figure 5a) but is seen as intensely labeled at mass 12C15N- (Figure 5b) because it has used gaseous 15N to build its molecular constituents. By contrast, E. faecalis is visible at mass 12C14N- (Figure 5a) but not at mass 12C15N- (Figure 5b) because it does not use gaseous nitrogen and therefore has not incorporated 15N above the natural ratio. The HSI image of the 12C15N-/12C14N- ratio (Figure 5c) shows E. faecalis with an isotope ratio equivalent to the natural isotope ratio (blue) and T. turnerae, which has incorporated an enormous quantity of 15N, with an isotope ratio at least 100 times higher (magenta).
MIMS can also be used to study the distribution of isotope tag incorporation within a bacterial population. The heterogeneity of nitrogen fixation among a population of T. turnerae is shown in Figure 5d, where the same field cultured for 32 hours in a 15N atmosphere is shown as a series of HSI 12C15N-/12C14N- ratio images. The different HSI panels reveal the level of 15N incorporation in bacteria using a compressed color scale as described in the legend to Figure 5. This analysis reveals the location and the distribution of ratio values, in other words of nitrogen fixation, among the bacteria within the analyzed field. Large differences in the amounts of 15N incorporation by T. turnerae cultured for 96 hours in a 15N atmosphere are demonstrated by the HSI images of the 12C15N-/12C14N- ratio (Figure 5e,f). Differences are visible among a few bacteria in contact with each other (Figure 5e) and even within a single bacterium (Figure 5f).
MIMS can detect and measure the function of intracellular bacteria within eukaryotic cells. This is shown by the quantitative imaging of the incorporation of 15N in T. turnerae living in the gill bacteriocytes of a shipworm (Lyrodus pedicellatus) raised under a 15N atmosphere, as shown in the HSI image of the 12C15N-/12C14N- ratio (Figure 5g). The bacteria in the bacteriocytes that have incorporated 15N are shown in colors between yellow and magenta (the shipworm tissue is blue). An internal control is the isotope ratio 13C-/12C- of the same field (Figure 5h), which shows a lack of contrast. The uniformity of the carbon ratio image eliminates the possibility of artifacts in the nitrogen ratio image as a result of morphologically or instrumentally induced isotope fractionation.
These quantitative images show that the MIMS method will be a powerful tool in the investigation of nitrogen fixation. It can also be used to study bacteria in natural environments and to explore the activity of diazotrophic symbionts in the tissues of plants and animals. It is worth noting that the size of an object can be estimated directly from the pixel signal intensity. This is illustrated with the flagellum visible on one T. turnerae cell (Figure 5i). For example, at mass 12C15N (Figure 5j), we have a mean of 1,473 counts per pixel on the bacterium (Figure 5k, inset, red box). A line profile of the counts per pixel across the flagellum of T. turnerae at mass 12C15N is shown in Figure 5k. The pixel crossed by the flagellum registered 66 counts. All conditions being approximately the same, the number of counts is proportional to the surface area of the material sampled in one pixel. In this particular image of 60 × 60 μm, 256 × 256 pixels, one pixel is equivalent to 234 nm covering an area of 54,756 nm2. If the length of the flagellum crosses a pixel, 66 counts would represent a width of (54,756 nm2/1,473 counts) × (66 counts/234 nm) = 10.5 nm; using the count values for the same pixels at mass 12C14N, the estimate is 10.6 nm, which is approximately the diameter of a T. turnerae flagellum.
Use of double labeling with bromodeoxyuridine and 15N-leucine to measure protein renewal
Because MIMS analysis sputters only a few atomic layers, a sample can be reanalyzed many times. This is illustrated by double-labeling studies of protein renewal and DNA replication in the mouse kidney after ischemia. We have previously shown [32] in the mouse that 30 minutes of bilateral renal ischemia, resulting in significant increases of blood urea nitrogen and creatinine, leads to protection of the mouse kidney against a subsequent ischemic insult 8 or 15 days later, even when the second ischemic period is extended to 35 minutes. Graded levels of time of initial ischemia resulted in graded levels of protection 8 days later. Bromodeoxyuridine (BrdU) and 15N-leucine were administered to mice subsequent to the first ischemia, in order to characterize the different response in cell proliferation in preconditioned and non-preconditioned kidneys exposed to ischemia on day 8 after the initial surgery. Quantitative MIMS images were recorded from thin sections of epon-embedded kidneys. The images were acquired in parallel at mass 12C14N to show a morphological overview, at mass 31P to view the cell nuclei and at mass 81Br to identify the nuclei undergoing DNA replication, as shown by BrdU incorporation. Protein renewal was calculated from parallel imaging at mass 12C14N and mass 12C15N.
Qualitative labeling with stable isotopes
An overlay of the 12C15N- signal in red with the 81Br- signal in green shows the co-localization of newly synthesized RNA and DNA in yellow (Figure 7c). This co-localization, visualized directly from the isotope images, avoids the complications potentially introduced by immunochemical methods [33]. Localization of newly synthesized RNA requires distinguishing a local excess of 15N over its natural occurrence. We carried out another MIMS analysis of the same cells to record the distributions of 12C14N- and 12C15N- in parallel (in our prototype instrument, we cannot simultaneously detect the isobars 12C14N- and 12C15N- together with 81Br- because of the steric hindrance of the electron multipliers). Overlaying the 12C14N- signal in red with the 12C15N- signal in green shows up the local excesses of 15N above its natural occurrence in yellow (Figure 7d). The yellow identifies the localization of 15N-uridine-labeled newly synthesized RNA within the nucleoli, along the nuclear envelope, and in the cytoplasm of the top cell.
Use of MIMS without isotope labeling to study gross differences in subcellular composition
Labeling with radioactive 14C
Use of MIMS to track individual cells in large cell populations
Following the transplantation of allogeneic tissues and organs, some donor-derived cells are known to leave the graft and present alloantigens to the host's immune system [35–37]. The exact nature and frequency of the donor cells infiltrating the host's lymphoid tissues are still unclear, however. The movement patterns of recipient immuno-competent cells that recognize the alloantigens are also unknown. These questions have remained unanswered because of the lack of sufficiently sensitive techniques for accurately tracking and visualizing small numbers of lymphocytes in tissues. Here, we use MIMS to track donor cells after transplantation.
We conclude that the MIMS technique allows the detection and frequency determination of rare cells infiltrating the recipient's lymphoid tissues after transplantation. The methodology can be extended to study solid organ transplants and the nesting of stem cells. It is important to note that the current techniques for tracking transplanted cells are fraught with difficulties [38]. Labeling DNA with stable isotopes should provide a method allowing long-term and physiological marking, the possibility of determining the generation of single cells from the amount of label within the cell, and the possibility of labeling both donor and recipient cells differently, enabling the unambiguous recognition of cell fusion or phagocytosis of donor cells by the recipient's cells [38].
We have shown that we can obtain quantitative atomic mass images using MIMS. Because these images are produced in parallel from the same sputtered volume they are in exact register with each other, which is necessary to derive meaningful isotope ratios. A lateral resolution that reaches 33 nm, in theory, and a high mass resolution (a mass/change in mass ratio of approximately 10,000) at high secondary-ion transmission (70–80%) allows us to image and quantitate molecules labeled with stable (or radioactive) isotopes within subcellular compartments. The high mass resolution allows us to distinguish the distribution of isobaric atomic species. For example, by separating 12C15N- from 13C14N- and 13C- from 12C1H-, we can image the intracellular distribution of molecules labeled with the stable isotopes 13C and 15N (such as 15N-thymidine and 13 C-uridine) in parallel and in the same preparation. Finally, the high stability of the primary beam, the mass spectrometer and the detectors contribute to precise quantification. These unique characteristics have allowed us to develop MIMS.
Instrumental advances
Instrument modifications
The objective column (OC) in the new SIMS instrument (see Figure 13), where the primary ions and secondary ions share the same path but in opposite directions, is a big step forward. It allows for a very short focal length for the ion-beam focusing lens (EOP) and for the secondary ion extracting lens (EOS) leading to a significant reduction in various aberrations. In addition, the incidence of the coaxial primary-ion and secondary-ion beams normal to each other allows us to reduce the geometrical dimensions of the instrument, leading to a significant reduction in various aberrations. The improvement rests on using the same lens for focusing the primary-ion beam, and for collecting the secondary ions that are formed. Secondary ions are emitted at a wide range of angles and energies. To collect them efficiently a strong electrostatic accelerating field is needed near the sample surface to gather their trajectories close to the surface normal. The conjugate action of the immersion lens and other lenses in the secondary optical column gives the secondary ion beam springing from the primary ion probe impact area the appropriate shape for matching the acceptance of the mass spectrometer tuned for high mass resolution. It should be emphasized that getting high mass resolution and high collection efficiency together is possible because the impact area of the primary-ion beam is very small (less than 0.1 μm2). This allows us to keep the beam small during its transport from the sample to the entrance of the spectrometer. From the general laws of optics, however, we can do this only for one position of the beam on the surface. As soon as the primary-ion beam moves, the secondary-ion beam emitted from the impact area is displaced as a whole in front of the spectrometer and a variable fraction of the beam (or no beam at all), depending upon the amplitude of the motion, enters the spectrometer.
To solve this problem, the machine uses an operation mode called 'dynamic transfer' to keep the secondary-ion beam stationary at the entrance of the spectrometer while the probe scans the surface. Dynamic transfer is achieved by the set of deviation plates, which move in synchrony with the scanning primary-ion beam. Because of the coaxial layout, those plates move both secondary and primary beams, but whereas they act on the secondary beam to align it along the optical axis, they act on the primary beam to make it rotate about the center of the aperture stop (see D1, Figure 13), which limits aperture aberrations on the probe. As a result, not only does the secondary beam stay stationary, which ensures a constant collection efficiency over the scanned area, but the intensity of the primary beam remains constant during the scanning; both of these avoid the 'vignetting', which is a gradual darkening of the corners of the ion images as the size of the field of view increases.
High mass resolution is obtained because the initial energy of ion spread has been cancelled at the level of the mass lines by coupling the electrostatic and magnetic prisms via a quadrupole lens. In addition, the elaborate ion optics allows the substantial reduction of second-order aperture aberrations, which improves the mass-resolving power (for example, at mass 27 the separation of 12C15N from 13C14N). This results from both the suppression of second-order aperture aberrations of the magnetic prism by tilting the entrance face of the magnetic prism, and the reduction of second-order aberrations of the electrostatic prism by placing a hexapole in front of the electrostatic prism (see Figure 13).
The mass spectrum of the secondary ions is displayed along the focal plane as a set of well-resolved mass lines. It is a discrete spectrum with wider spacing between ions separated by 1 mass unit and closer spacing when multiplets (isobaric species) occur. Because of the limited size of the magnet, however, the whole spectrum cannot be displayed at once. The ratio between the lower and the higher masses in the displayed range is about 20. By changing the strength of the magnetic field it is possible to choose the lower mass and, in consequence, the higher one. For instance, if 12 is the lower mass unit, the spectrum is displayed up to mass number 240.
A single ion impact on the first dynode of an electron multiplier ejects several secondary electrons, which are subsequently amplified by the other dynodes in a chain amplification process and generate a voltage pulse. Pulse heights depend upon the ion-electron conversion yield on the first dynode and upon the voltages applied on other dynodes. Pulse heights have a distribution that can be stretched by changing the high voltage. The distribution is usually depleted on the low-amplitude side, where one must account for the superposition with low-amplitude pulses from the electronic noise. A pulse-height discriminator eliminates low-amplitude contributions so as to select only signal pulses. Of course, a fraction of the signal is suppressed in that process, but this fraction is quite small, for example a few percent for CN-. The secondary-ion detection efficiency is set to be equivalent among the detectors in part by adjusting the rejection threshold and in part by adjusting the high voltage. Measurement of samples of known isotopic composition can be used to correct for any remaining differences in efficiency between detectors. Using the adjustments and tuning described above, we can measure the isotope ratios of secondary ions, which is at the core of MIMS methodology, in a precise and reproducible manner.
Spatial resolution
The parallel detection of different ions, the small size of the primary-ion beam, and the high efficiency of secondary-ion collection all contribute to the highly effective spatial resolution of MIMS, reaching 33 nm, 30-fold better than in previous instruments. Smaller beams are possible and may be useful for locating and measuring highly concentrated tags in volumes of the order of (20 nm)3. Primary sources with higher brightness could give smaller probes with sufficient current for practical analysis: they could be provided by a brighter Cs+ primary-ion source or by other types of sources [4].
Volume sputtered and detectability limit
The absolute sensitivity of SIMS is among the highest of current chemical analysis methods and the total amount of material needed for MIMS analysis is extremely small. Using MIMS, one can detect parts per billion of an isotope in a sub-cubic-micrometer volume of sample. Let us assume an analyzed field of 6 × 6 μm, a primary Cs+ beam current intensity of about 0.4 pA, a dwell time per pixel of 20 msecs, 256 × 256 pixels, a sputtering efficiency of five target atoms per Cs+ ion, an atomic layer thickness of 2.55 × 10-4 μm, and an atomic density of 6 × 1010 atoms per μm3: the total sputtered volume is then 0.27 μm3, and the sputtering rate is 0.4 nm/minute or 1.5 atomic layers/minute. For a probability of detection of 95%, three ions have to be emitted; assuming a useful ionization yield of 1 × 10-2 ions/atom, then the minimum detectable amount is 1.85 × 10-8 or approximately 20 parts per billion (see Additional data file 1).
Contrast formation
There is no theory using first principles that allows us to predict what happens when the surface of the sample is hit by a primary ion. Which atoms are most likely to escape?
What is the sputtering rate (the rate of removal of target atoms)? When a given atomic species is sputtered, what is the ionization yield (the fraction of atoms that are ionized)? How will the composition of the matrix affect the number of atoms sputtered, the ionization yield, and the formation of polyatomic clusters such as CN or CNH? What is the breaking pattern of large and small molecules in atoms and atomic clusters? How interdependent are the parameters mentioned? All these questions are yet to be answered. In addition, the choice of primary ions, so important in practice, remains mostly empirical. Indeed, in order to achieve the high ion yields required for trace analysis down to the low parts per billion range, the sample needs to be bombarded with or exposed to chemically reactive species, and O- or Cs+ are most commonly used. There is, however, a lack of detailed knowledge about the instantaneous surface composition, and all attempts to correlate observed ion yields with available ionization models should be considered highly questionable [39].
The contrast of a mass image is due to regional differences in secondary-ion currents emitted over the analyzed field. The main determinants of secondary-ion formation are the sputtering rate (the rate of removal of the target atom by the primary ions), the ionization yield (the fraction of sputtered atoms that are ionized) and the local concentration of atoms. These parameters cannot be derived directly from first principles, but we may safely assert the following. The sputtering rate in steady state is directly proportional to the primary beam current density at the sample surface and is dependent on the local atomic composition.
The ionization yield, in the negative secondary-ion emission mode, depends strongly on both the electron affinity of the elements or clusters under consideration and the concentration of the Cs+ ions that are implanted by the beam into the superficial atomic layers of the sample. To a first approximation, implanted primary-ion concentrations vary as the inverse of sputtering rates; this makes the ionization yield somewhat a function of the sputtering rate. However, we do not know the form of the function that links implanted Cs+ concentration to ionization yield.
Taking all these factors in consideration, we can predict the following. A low sputtering rate is generally associated with a higher concentration of implanted Cs+; lower sputtering rates should therefore lead to higher ionization yields. If ionization yields are strongly dependent on Cs+ concentrations, low sputtering rates may paradoxically lead to higher secondary-ion currents. In a situation in which the sputtering rates of different areas over the field of analysis are not the same, a given area may appear brighter because the sputtering rate is lower (with a relatively higher concentration of implanted Cs+) and the ionization yield is still sensitive to the concentration of implanted Cs+. Moreover, the ionization yield is likely to depend on local sputtering rates because, once a steady state has been reached, these local rates control the local concentrations of the implanted primary ions. As a consequence, the brightness of an image area depends upon the local concentration of the element under consideration, the implanted Cs+ concentration and the sputtering rate.
Without any contrast, images would not exist. Parallel mass images of different elements with different electron affinities convey a lot of information. For instance, a section of embedded cells bombarded with Cs+ gives 12C- images with low contrasts and, in parallel, 12C14N- images showing strong contrasts related to known cell structures (for instance, Figure 14, 12C- and 12C14N-). Let us first consider 12C- images. The local 12C- intensity depends on both the sputtering rate and the ionization yield. When analyzed in parallel, one would expect (but it is not demonstrated) ionization yields of 12C14N- ions, with their high electron affinity (3.6 eV), to saturate (to reach their maximum electron capture probability) more rapidly during Cs+ implantation than 12C- ions, whose electron affinity is much lower (1.2 eV). It follows that, if the ionization yield is strongly dependent on the concentration of implanted Cs+, at low concentrations of implanted Cs+ the 12C- ion emission should be more sensitive to small changes in the implanted Cs+ and may be more sensitive than 12C14N- ions to local differences or fluctuations in sputtering rate.
The low contrast that we observe in 12C- images of epon-embedded samples probably indicates that sputtering rates are evenly distributed across the field analyzed and that implanted Cs+ concentrations are approximately equivalent throughout the sample. The remaining contrasts are probably due to a slightly lower concentration of carbon in the biological tissue than in the embedding epon and to the formation of 12C14N atomic clusters that reduce the total number of 12C atoms available to form 12C- ions. If high contrasts were observed it would alert us to differences in sputtering rates or unexpected differences in atomic concentrations of carbon.
Because 12C- images indicate a homogeneity in the sputtering rate and a steady state of the implanted Cs+ concentration, the contrast observed on the CN- images acquired in parallel must reflect differences in local nitrogen concentration over the analyzed field and cannot be due to emission artifacts. The fact that, in many instances, interpretation of 12C14N- images is straightforward and allows us to recognize easily details of features in tissue or cell sections is a boon.
Finally, we wish to emphasize the strength of the quantitative information derived from the ratio of mass images acquired in parallel. All parameters being the same, atoms or clusters such as 26CN and 27CN are affected identically at a level of precision well beyond what is needed for our experiments, and the quantitative ratio images contain the true isotope ratio values, cancelling out sputtering, ionization and other effects that affect the emission of a single ion.
The relationship between contrast formation, lateral resolution and counting statistics
The lateral resolution is essentially a function of the size of the primary ion beam. The smaller the beam diameter, the higher the resolution; to be visually appreciated, however, the image has to have enough contrast; that is, the image has to contain enough counts. Higher counting rates will be obtained with a brighter primary ion beam, all other conditions being the same. The count rate does not affect the intrinsic resolution, but it is a main determinant of the counting statistics. For example, in a parallel set of 12C15N- and 12C14N- images, although a 12C15N- region in general contains far fewer counts than its 12C14N- homolog, it has the same lateral resolution. The variance of the 12C15N-/12C14N- isotope ratio, however, is essentially determined by the number of counts in the 12 C15N- region.
If the sample surface being studied is not flat but shows relief, we observe a three-dimensional appearance in the sample image. Here, the contrast may be due either to a differential implantation or to a differential sputtering that occurs because the primary beam attacks the sample surface at a variety of angles. It should be emphasized, however, that these phenomenon affect in the same proportion – at least to a first approximation – isotopes separated by 1 atomic mass unit. Thus, the variations in count rates cancel out (note that all relief effects disappear in the ratio images; see for example the adipocyte in Figure 4d,f), and the value of the isotope ratio still represents the local abundance of the individual isotopes with a precision of a few percent, which is entirely satisfactory for biological purposes. SIMS had previously been used for quantitation in the absence of imaging, and for imaging in the absence of quantitation. The advantage of MIMS is to combine high spatial resolution (that is, analyzing a much smaller amount of material than previously), with simultaneous (parallel) acquisition of information from several atomic masses, and with isobar separation at mass 27.
The ability to provide quantitative information is a unique attribute of MIMS. Imaging is a guide for exploiting the massive amount of unique quantitative information, which is based on the mining of the counts of secondary ions per pixel.
Isotope ratios
The isotope ratio values are the essential data needed to interpret the results, and the derivation of isotope ratios is an integral part of MIMS methodology. The absolute number of counts for a given atomic mass do not convey any information about the presence of an isotope excess. For example, the counts of 12C15N by themselves do not allow us to conclude anything about the presence of a 15N tag in the analyzed region. Only the ratio 15N/14N compared with the natural abundance of 15N is informative. The visual observation of contrast and/or brightness is no more informative, and may be misleading. In a given field, contrast is due to variations of the number of counts per pixel, which may have multiple undocumented causes. As discussed under 'Contrast formation' earlier, in addition to the local abundance of the isotope under consideration, the secondary-ion intensity depends on other parameters such as the number of atoms of Cs implanted and the quality of the matrix into which the isotope is introduced. Thus, the fact that a given area emits more than a neighboring one does not mean that it contains more of the isotope tag. High counts in 12C15N do not mean that there is an excess of 15N until they are normalized with respect of the counts of 14N by calculating the isotope ratio12C15N/12C14N. A region of high secondary-ion yield at mass 12C14N may have little incorporation of 15N (see, for example, the cochlear outer pillar cells (Figure 3a), which have a high secondary 12C14N yield and very low incorporation of 15N (Figure 3c and Additional data file 3). Conversely, a region with high 15N on the 12C15N mass image compared with the neighboring pixels may simply reflect a region with overall high yield of both 12C14N and 12C15N, even though their proportion is equivalent to the natural ratio. The isotope ratio values for regions of interest are the only information that may be used for evaluating where and how much a label such as 15N has been incorporated.
The power of the isotope ratio method is that all analytical parameters being the same (field analyzed, instrumental conditions, Cs+ implantation, matrix effect), one may assume that they affect two isotopes of the same atomic number – that is, isotopes that differ in general by 2 atomic mass units at most – in an identical fractional manner, so that any unknown effect will cancel out and that isotope ratio values will uniquely represent the relative abundance of one isotope with respect to the other.
Ensuring 'flatness' of the field, such that the isotope ratio values among regions of interest are equivalent regardless of their location in a control image field, is very important. It demonstrates that there is no differential fractionation among isobars: that is, that differences in isotope ratio among various areas are not the consequence of changes in transmission between a set of isotopes that are related to location within the field. This is why the 12C and 13C quantitative atomic mass images used to generate the 13C/12C ratio image (in Figure 3, for example) are an essential internal control. Because there is no 13C added, they control for the flatness of the field, which means: first, equivalence of the 13C/12C ratio at any location over the field analyzed; second, an absence of instrumental drift during analysis; third, the lack of some topographical effect; and fourth, the lack of a differential matrix effect. With all conditions being the same and because the secondary ions are detected in parallel from the same analyzed area, this ensures that measured high values of 15N/14N ratios do show an excess of 15N and cannot be an artifact.
We have developed several kinds of controls. Two essential controls in our quantitative imaging allow us to validate the analytical conditions. This validation is individual to the field being analyzed. The first control is to make sure that if no isotope has been added, the isotope ratio measured is equivalent to the natural abundance in a region of the field where no excess isotope is expected. When the sample is an epon-embedded section, this is measured on the epon, in a region of the analyzed field that does not contain tissue or cells.
The second essential control is to make sure that, for every field analyzed in quantitative imaging mode, a calculated excess of an isotope above its natural abundance is independent from its location in the analyzed field. This could happen if there was some instrumental fractionation of one isotope with respect to another, meaning that depending upon the location over the field, the transmission for one of the isotopes is somewhat different from another one. Control for the lack of fractionation is provided by the image derived from isotope ratios from quantitative mass images of a set of isotopes expected to be present at their natural abundance values and recorded over the same field, in the same instrumental condition and in parallel, simultaneously, with the isotope from which one expects an excess above its natural abundance at some location within the field. This is best performed by recording together in parallel the quantitative image of the experimental isotope and of an isotope that is expected to be present at the natural abundance level. The beauty of the 13C/12C ratio (as in the cochlear sample labeled with 15N, Figure 3) or of the 12C15N/12C14N compared with 13C14N/12C14N (as in an adipocyte labeled with 13C, Figure 4) is that they control for the very same field that is being analyzed, and in the same conditions. 'Normally' one is not fortunate enough to develop and use internal controls of such high power. These controls are ideal internal controls and are immensely more important than finding natural abundance value ratios in separate control samples where all conditions are different from the experimental measures.
We have developed MIMS, combining the use of a new SIMS instrument that simultaneously detects multiple isotopes with stable isotope labeling and quantitative image analysis. Using MIMS, it is possible to localize and measure precisely stable or radioactive isotopic tags in subcellular compartments in sub-cubic-micrometer volumes. Stable isotopes can be localized simultaneously at high resolution and measured with high precision, a unique combination. Studies with MIMS can use conventional tissue – and cell-preparation methods. The molecular tags can be any isotopes, stable or radioactive. One can take advantage of any molecule that has been labeled for mass spectrometry or nuclear magnetic resonance studies (2H, 13C, 15N, 17O, 18O, 33S) or for radioactive studies (3H and 14C). These features make MIMS methodology a powerful new way of viewing and measuring, in all fields of biomedical research, important parameters that were impossible or difficult to see or to measure with other methods.
Materials and methods
Data analysis was performed with an SGI O2 workstation, ISee Analytical Imaging Software (Inovision Corporation, Raleigh, NC), custom software within ISee Analytical Imaging Software, and Prophet statistical software (National Center for Research Resources, NIH).
Quantitative image data extraction
In the scanning mode, the number of counts for each secondary ion emitted from each pixel is stored in a computer file. Grayscale secondary-ion images for each of the selected secondary ions are obtained after analysis by reconstructing the data onto an array with a number of pixels equal to that of the data-acquisition array, with the pixel intensity in the image scaled according to the number of counts detected from that position for the ion. We developed a piece of software to extract 24-bit quantitative image files from hundreds of raw MIMS images, keeping the counts recorded in each pixel.
An essential feature of MIMS is the ability to locate and to quantitate regions of the sample that have accumulated an isotope, for example 15N, after the isotope has been provided externally. This is done by deriving ratio images. For example, the ratio image 12C15N-/12C14N- and the ratio image 13C-/12C- are derived from the pixel-by-pixel division of the respective parallel images. Quantitative images acquired with MIMS generate far more information than can be displayed using grayscale methods. To show high dynamic-range ratio images and to de-emphasize values resulting from data with few counts, we developed a method based on a hue saturation intensity transformation (HSI) [40, 41] of the ratio image; HSI is derived from a calcium ratio imaging procedure [42]. Think of the pixels in a MIMS image as equivalent to test tubes. A 256 × 256 pixel MIMS image would be an array of 65,536 test tubes from which we simultaneously count several isotopes (for example, the abundance of 12C, 13C, 14N and 15N). The HSI method allows us to identify among the 65,536 pixel test tubes (corresponding to areas of sub-micrometer size) those that contain significantly higher label counts. The main advantage of HSI is to color code the value of the ratios following a scale that can be extended or compressed, as well as taking full advantage of the fact that humans perceive many more differences in color than differences in shades of gray. For example, HSI enables us to identify regions of interest with significant excess of the 15N label by a means independent of visual recognition of expected histological structures. The use of HSI allows us to greatly simplify the first step of data analysis.
Another powerful piece of software automates the data analysis of sequences of raw images. After creating stacks of images from analyzed fields and drawing regions of interest, statistics for each of the regions in each of the images are automatically tabulated and exported to the Prophet software for statistical analysis.
Isotope ratios
Regions of interest were selected using HSI ratio images; secondary-ion counts for the same regions of interest were extracted from the corresponding unsmoothed images of 12C-, 13C-, 12C15N- and 12C14N- and the isotope ratio values from the derived images 13C-/12C- and 12C15N-/12C14N-. The 15N/14N ratios (or 13C-/12C-) for each region of interest were also determined by dividing the total counts of 12C15N- (or 13C-) over all pixels in that region by the total corresponding counts of 12C14N- (or 12C-). Importantly, none of the image manipulations that we perform alters the counts contained in the pixel or groups of pixels.
Nitrogen renewal
The percent nitrogen renewal was calculated for each region of interest as: 100 × (tissue 15N/14N ratio – control diet ratio)/(experimental diet ratio – control diet ratio), which gives the percentage of the labeled dietary nitrogen that has been incorporated into the sample. Zero percent represents a ratio equivalent to the terrestrial ratio; 100% represents a ratio identical to that in the experimental diet.
Labeling and tissue preparation
All experimental protocols used in this study were approved by the Harvard Medical Area Standing Committee on Animals, Protocol 02751. CBA/CaJ mice, 8 weeks old at the start of the experiments, were obtained from Jackson Laboratories (Bar Harbor, ME, USA).
Diets enriched in 15N-leucine
Standard L-leucine commercial protein-free amino acid diet (TD 86529, Harlan Teklad, Madison, WI, USA) was used as the control diet. The experimental diet was prepared by the manufacturer according to the same formula, but with 15N-L-leucine replacing 23% of the standard L-leucine (9 g 15N-L-leucine in the total of 38.8 g leucine, for a 3.5 kg batch of diet). To ensure a dietary steady state on these synthetic diets, mice were placed on the control amino-acid diet for 5 days before they were fed the experimental diet. Mice were sacrificed after 9 days or 22 days on the 15N-L-leucine diet. 15N-L-leucine (> 98% isotopically pure) was obtained from Cambridge Isotope Laboratories, Andover, MA, USA.
The 15N/14N nitrogen isotope ratios in the diets were measured by combustion mass spectrometry analysis (Metabolic Solutions, Nashua, NH, USA). The values were 3.657‰ (0.001 SE, n = 12) in the control diet (equivalent to the terrestrial value) and 13.972‰ (0.573 SE, n = 12) in the experimental diet.
Mouse cochlear tissue
After 9 or 22 days on the diet, deeply anesthetized mice underwent intracardiac perfusion with saline followed by fixative (2.5% glutaraldehyde and 1.5% paraformaldehyde in 0.065 M phosphate buffer). The temporal bones were removed, kept in fixative overnight at 4°C, decalcified (0.1 M EDTA with 0.4% glutaraldehyde) for 3 days, rinsed in deionized water, dehydrated in ethanol and propylene oxide, infused with propylene oxide/epon, and embedded in epon. Other tissues were treated similarly, but not decalcified [43].
Sections from the epon-embedded tissue, 0.5 mm in thickness, were cut with glass knives on a Sorvall ultramicrotome. The sections were deposited on (5 mm)2 silicon wafer chips, which provide a mechanically sturdy and adequately conductive support [44]. The silicon chips were then mounted in a custom-made sample holder for insertion into the MIMS instrument. The holder with the sections was placed in a vacuum oven for 3 days at 50°C before analysis and then introduced in the preanalysis chamber of the instrument in a 10-9 torr vacuum.
Teredinibacter turnerae
Cells of T. turnerae were grown in sealed 16 ml Hungate tubes containing 15.5 ml of simple salt medium (SBM amended with 0.5% sucrose without combined nitrogen [31]) allowing a headspace of 0.5 ml of atmospheric gas. After three days the headspace gas was removed by displacement with fresh medium and replaced with 0.5 ml of 15N gas (99%). Cells were then incubated at 22°C with gentle shaking for 0, 0.25, 0.5, 1, 2, 4, 8, 16, and 32 h. After incubation, cells were sedimented by centrifugation (12,000 g, 10 min, 4°C) and fixed by resuspension in 500 ml 3.7% formaldehyde in seawater for 30 min. Cells were then washed by sedimentation (14,000 g, 10 min, room temperature) and resuspended first in 50% ethanol and again in distilled water. Cells were finally resuspended in 10 μl of distilled water and 2 μl was applied to the silica wafer and air-dried at 37°C. Cells of Enterococcus faecalis were prepared identically except that they were grown in Tryptic Soy Broth medium (Bacto) at 37°C for 2.5 h (approximately eight doublings).
Shipworms, Lyrodus pedicellatus, were incubated in seawater in a 15N atmosphere, fixed in formaldehyde, embedded in epon, and sectioned for MIMS analysis.
Adipocytes were prepared and analyzed in MIMS using the same methods as Kleinfeld et al. [28].
Rat embryo fibroblasts
Rat embryo fibroblasts (REF52) were cultured in DME/F12 medium with 10% FCS. They were serum-deprived for 26 h after they reached 70–80% confluence; pulsed for 17 h with 10% FCS and 1 mM BrdU (1/10 of the concentration generally used in microfluorescence studies) and 1 mM 15N-uridine; fixed in their culture dish (using formaldehyde, glutaraldehyde, and picric acid); detached from plastic with propylene oxide; embedded in epon-Araldite; sectioned at 0.5 μm thickness; and mounted on polished Inox rod for MIMS analysis.
For 14C analysis, REF52 cells were cultured in DME/F12 media with 10% FCS. They were serum-deprived for 24 h after they reached 40% confluence; pulsed for 24 h with 10% FCS and 1 mCi 14C-thymidine at a concentration of 19.2 nanomol/ml; fixed in their culture dish using formaldehyde and glutaraldehyde; detached from plastic with propylene oxide; embedded in epon-Araldite; sectioned at 0.5 μm thickness; and mounted on silicon chips for analysis.
Tuning for quantitative mass image acquisition
Before a MIMS experiment, the sample is first viewed and photographed under a reflection differential interference contrast (RDIC) microscope with an automated stage, and the coordinates of multiple regions of interest are noted and used to find the same regions in the automated stage of the SIMS instrument. RDIC allows us to dispense with preparing and staining alternate sections for microscopy. The sample is placed in the analysis chamber under a vacuum of approximately 3 × 10-10 torr. A primary-ion beam of Cs+, after being shaped by the primary-ion column, is focused with the immersion lens and strikes the sample with an energy of 16 keV (see Figure 13). The primary-ion beam scans the region of interest on the stationary sample step-by-step in a raster pattern, generally of 256 × 256 pixels but the number of pixels can vary in practice from 32 × 32 to 1,024 × 1,024. The secondary ions emitted from the sample are focused with the immersion lens, shaped by the optics of the secondary-ion column, and sent through a hexapole in the double-sector mass spectrometer made up of an electrostatic prism followed by a magnetic prism. This permits first-order focusing of the secondary ions as narrow lines along the focal plane of the magnet. Four detector units are located at the focal plane and can be moved independently along it so as to select specific atomic masses. Four electron multipliers in the detector units detect the signals from the selected secondary ions simultaneously. The current and diameter of the primary-ion beam, the time the beam is directed at each pixel (the dwell time), and the size of the pixels can be adjusted according to the size of the area being analyzed.
For each analysis, we need to ensure first that we have implanted enough Cs atoms into the sample to maximize the secondary-ion counting statistics; second, we maximize the secondary-ion transmission; third, we have perfect mass spectra at all masses; and fourth, in control fields we obtain equivalent values of the isotope ratio over the entire image field. Tuning is first done in ion-counting mode, in which the secondary-ion count rate recorded by each detector is displayed numerically. No images are acquired. We ensure that the responses of the electron multiplier detectors (EM; see Figure 13) are equivalent. The primary Cs+ beam is defocused and kept stationary on a standard silicon sample. The secondary-ion current at mass 28Si is measured successively with each of the four EM detectors; the radius of the secondary-ion beam is set to enter the appropriate EM by changing the magnetic field strength of the magnetic prism. We adjust two parameters for each EM: the EM high voltage is set so that the pulse height distribution (PHD, the graphical representation of the distribution of the amplitude of the pulses generated by an EM at a particular voltage) of the EM output is similar for all four detectors; and the EM threshold voltage is set so that, in the absence of secondary-ion current, the noise of the counting system is smaller than 1 count per minute. When these conditions are fulfilled, the count rate for the same sample is the same among the four detectors, within ± 2.5%.
To begin tuning, we implant Cs atoms into the biological sample over a relatively large field (100–150 μm) at high primary-ion current (obtained by removing the diaphragm (D1, Figure 13) used to define the primary-beam aperture in the objective column (OC). After a period of several minutes, Cs atoms implanted into the sample enhance the secondary-ion yield so that the 12C14N- secondary-ion count rate reaches a steady state. Next, we ensure that the EM detectors are located in the correct position along the focal plane of the magnet (FP, Figure 13) for each mass. We maximize the count rate of each EM by moving the detector along the focal plane so as to maximize the secondary-ion count rate for the selected atomic mass.
We follow with the alignment of the secondary-ion beam so that the isobars 12C15N and 13C14N, and 13C and 12C1H, are well discriminated, indicating high mass resolution, while maintaining a high secondary-ion transmission (approximately 70–80%). An entrance slit (ES, Figure 13), whose width is a main determinant of the mass resolving power (MRP), is placed in the path of the secondary-ion beam. We maximize the count rate for a given mass by iteratively focusing the secondary-ion lens located in the objective column (E0S) and by adjusting the voltages of the deflection plates P2, P3, and CY within the secondary column. We then set an aperture slit (AS) and an energy slit (WS) to reduce angular and energy aberrations and thus to maximize the MRP. To evaluate the MRP, we scan the secondary-ion beam through the detectors selection slit (SS) using a set of deflection plates (DP) and we record a small portion of the mass spectrum by plotting the secondary-ion current versus the deflection voltage (Figure 15). The quadrupole and slit lens (LF4) are used to maximize the MRP. Then, for each detector, using the mass spectra plots, we set the voltage of the deflection plates (DP, Figure 13) so that the mass line of interest is centered in the selection slit (SS) of the EM (see Figure 13). This practically eliminates any spillover from neighboring isobars.
We then switch to imaging mode, which displays 'live' scanning ion images from two selected masses. After adjusting the size of the analyzed field, the last steps aim to balance the tuning and to focus the primary-ion beam. We ensure that the analyzed field is balanced, that is, that the secondary-ion intensity emitted by a small fraction of the field is equivalent over the entire field, with iterative small voltage adjustments of the secondary-ion objective lens (EOS), and of a series of deflection plates (P2, P3 and CY) within the secondary column. We optimize the focus by adjusting the voltage of the primary-ion objective lens (EOP, Figure 13). To analyze at the highest possible spatial resolution (approximately 35 nm), we increase the demagnification of the primary-ion source by increasing the voltage of the lens L1 (see Figure 13) in the primary column. We may then have to slightly readjust the tuning of the secondary-ion beam. Finally, before starting analysis, we adjust the number of pixels in the image (256 × 256 up to 2,048 × 2,048) such that the beam diameter is approximately twice the pixel size, and we ensure that the acquisition time is long enough to obtain sufficient counting statistics for masses with low count rates (for example 13C and 12C15N).
After a sample is introduced into the MIMS analysis chamber, we can set the instrument to acquire qualitative MIMS images – in general at mass 12C14N, which gives a good anatomical view – within 30 minutes. Significantly more time is required, however, for the optimization of the many instrumental parameters (tuning) that is needed to record quantitative mass images suitable for extracting meaningful quantitative data. For new types of samples it may take hours to a day. Yet the physics is robust, and further tuning on an identical type of sample does not take more than 1 to 2 hours.
Additional data files
The following files are available: Additional data file 1, containing an explanation of how the volume sputtered, the useful yield, and the detectability limit are calculated; Additional data file 2, containing an explanation of how the lateral resolution is calculated; Additional file 3, describing data reduction from labeled and control mouse cochlear sections; Additional data file 4, describing the relationship between the brightness of an image and the number of counts; Additional data file 5, comparing MIMS with autoradiography; Additional data file 6, containing an explanation of the shape of mass line peaks in MIMS; and Additional data file 7, containing more details on Figure 2e.
Development of the new SIMS instrument was supported by ONERA, CNRS, Université Paris Sud and Cameca (France). We thank Jean-Daniel Sraer for his help in obtaining the prototype of the new SIMS machine and M.J. Karnovsky for his long-time support. We thank Hjalmar Brismar and Anne Mudge for overall comments, Edmund Mroz and Tae Gee Jung for providing mouse cochlear samples, Louise Trakimas for her most expert histological help. Molly Palmer for her patience, efficiency and dedication in working on the manuscript, and Anna Sharman for her comprehensive editorial attention. This work was supported in part by NIH grants P41RR14579, P41EB001974, DC00033, DC03463, RO1DC04179, P41RR14579, P41EB001974, R37DK39773, RO1EY12963, RO1GM47214, and RO1DK58762 and by NSF/IBN grant IBN-998298.
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Thursday, Sept. 17, 2009 01:22 pm
Springfield Celts looking for new young ruggers
Most people have no idea that the capital city has its very own rugby team.
"The Celts used to be bigger in Springfield, and then we had a waning period and the interest came down," says Brian Foreman, a member of the Springfield Celts since 1998. "People still say, 'Springfield has a rugby team?'
"Yeah, they've been around since 1975."
The Springfield Celts rugby club, a nonprofit organization with roughly 25 members, plays every spring and fall at Kennedy Park across from the Abraham Lincoln Capital Airport. Most players like Foreman, a 35-year-old chemist for the Illinois Department of Transportation, played high school and college football. So far this season they've competed in three matches — against the St. Louis Royals, Southern Illinois University and Illinois State University — but they're more focused lately on recruiting new, young talent to their sport.
Jim Carlberg, the Celts' head coach, has been with the team since 1984. Rugby, an 80-minute, full-contact sport that combines elements of football, basketball, soccer and chess, continues to rise in popularity across the United States. But for some reason, he says, it hasn't quite caught on in Springfield.
"The team has grown through good numbers and almost folded through the past 34 years," Carlberg says. "We've seen highs and lows."
The Celts have had trouble attracting players in their 20s — making it difficult to grow as an organization and to compete against younger and stronger teams. Carlberg seeks to remedy this problem by building a new youth rugby program in Springfield and the surrounding area.
He envisions 6-to-12-year-olds joining a rookie program, where they'd play non-contact rugby and focus on learning the sport's skills. High school athletes, who normally play soccer or football in the fall, would filter into a Jr. Celts rugby program in the spring to learn advanced ball handling and tackling. Not only would young athletes learn a new set of skills, Carlberg says, but they'd also stay in shape for their primary sports.
"I've talked to a lot of high school rugby coaches in the Illinois area, and their corresponding high school coaches love the rugby program," he explains. "Kids playing in the fall after spring rugby are double the capabilities."
The Illinois Youth Rugby Association has already shown interest in starting a youth rugby club in the Springfield area, Carlberg says, so now he needs local high schools to help draft players for next spring. The Celts' five-year goal would be to net these players after they move up to college rugby and then return to Springfield.
"The Celts would get a kid who has 10 to 15 years of rugby under his belt, but only be 22 years old," Carlberg says. "That's the ultimate player."
The Springfield Celts Rugby Club started outreach efforts in the beginning of August, when they teamed up with the Illinois Inter-Agency Athletic Association to sponsor a rugby skills camp for at-risk youth from six children's homes. Last week the team also passed out fliers and talked with students at Lincoln Land Community College.
For now, no experience is needed to play with the Celts — new recruits just need a good pair of spikes, a tough pair of shorts and a mouthpiece.
"The only thing required is the ability to come out and want to play," Carlberg says. "It doesn't matter if you're big, short, skinny, tall. We train from the ground-up."
For more information, go to www.celtsrugby.com.
Contact Amanda Robert at firstname.lastname@example.com.

subset: commoncrawl


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How You Can Overcome Panic Attacks
Overcoming Panic Attacks
It can be unsettling and overwhelming to cope with panic attacks. These are some things you can try to cut those fearing panic attacks down to size. They don't need to be a part of you.
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Remember To Breathe
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Learn relaxation techniques that you can use for the onset of a panic attack starts.
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Try to do this process ten times and you should start to feel better.
Use positive dialogue and reassuring images to talk yourself out of an oncoming panic attack. Know that you will not last forever. Tell yourself to stay in control.
One of the best ways to control a panic attack is by using breathing techniques.
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Cognitive Behavior Therapy And Other Courses Can Help
Don't become more anxious at the thought of a panic attack.
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These professional therapists have used various treatments and therapy to aid many people who suffer from panic attacks, and could help you too.
Do some online research to find practitioners who specialize in the treatment of anxiety and panic disorders, and make sure the accredited doctor you go to is experienced in treating panic disorders and anxiety.
This method will enable you be able to face your fears!
Allow yourself to succumb to the remedies that will help your anxiety and engage in healing to happen.
Understand what it is that is causing your panic attacks. Identify the problem and start to address them immediately.
Take Time Out To Enjoy Yourself
Begin to bring some more enjoyment into your life. Take yourself out to watch a movie.
Go for a walk in the park or meet up with a friend. Laugh as much as you can. There are many ways to bring on laughter, such as reading comic books or watching a funny movie.
By surrounding yourself with positive friends, you'll always have someone to depend on to cheer you up when you're feeling down, you will find that they help to lift your spirits when you are struggling with things.
Be sure to get together with your friends and loved ones often for extra support.
Try helpful breathing techniques or relaxing music instead, and attempt to calm your breathing.
Fighting off an attack may make it worse and make it last longer.
Rather than focusing on coping with each individual attack, focus on the behaviors that stop them from happening entirely. Consider doing meditation, doing meditation or going to a yoga class.
Do whatever works best for you.
Never try to self-medicate when you have panic attacks. Alcohol and drugs can help you relax, but can worsen your symptoms.
Consult with a professional about the best course of panic attacks.
You have to work until you're worn out and push yourself much harder than you exercise. If you become accustomed to a certain exercise to the point it is no longer a challenge that leads to exhaustion, then move on to something new that will.
Good exercise is not just good for the physical body, but for your spirit as well.
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There are resources out there for people with panic attacks. This non-profit organization has dedicated itself to treating, therapy and cure of anxiety and panic disorders.
You can find the help you have been needing! The "Panic Away" Course has helped tens of thousands with anxiety – and they gurantee it to work.
Hopefully these tips will empower you to fight back against panic. Keep in mind that your negative feelings are oftentimes connected to your panic attacks.
You know that you are able to do this.
Taking positive steps to overcome panic attacks and getting them under control is time well spent on getting your life back on an even keel.
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