We would like to thank Toshihiko Shiroishi for his assistance and for providing the research environment during this study. We are grateful to Kensuke Yanagi and Takato Izumi for advice on A. equina, and Masaatsu Tanaka for advice on the Harmothoe sp. specimen. We would like to thank the staff at Shimoda Marine Research Center, University of Tsukuba, and Misaki Marine Biological Station, The University of Tokyo for their help in sample collections. We would like to thank Editage (www.editage.jp) for English language editing. This work was supported by the JSPS Grant-in-Aid for Young Scientists (A) (JP26711022) to HN, and JAMBIO, Japanese Association for Marine Biology.

1. Fixation
<ol>
	<li>For Actinia equina, relax the animals in 10% MgCl2 seawater for about 15 min at room temperature. Transfer to 70% ethanol and store at room temperature.</li>
	<li>For Harmothoe sp., anesthetize the animals by placing them in ice-cold seawater for about 15 min. Transfer to 10% (v/v) formalin solution with seawater and store at room temperature.</li>
	<li>For Xenoturbella japonica, relax the animal using 7% MgCl2 in freshwater. Fix in 4% paraformaldehyde (PFA) in filtered seawater overnight. Place in 70% ethanol and store at 4 &#176;C. 
	CAUTION: PFA is hazardous and must be handled with care.</li>
</ol>
2. Staining
<ol>
	<li>Transfer the samples to 50% ethanol and store at room temperature for 15 h. Replace the 50% ethanol with 25% ethanol and store at room temperature for 2 h. 
	NOTE: This is not necessary for the Harmothoe sp. sample in 10% (v/v) formalin solution with seawater.</li>
	<li>Replace the solution with distilled water (DW) and store the samples in DW at room temperature for 2 h. Repeat this step three times.</li>
	<li>Prepare 25% Lugol solution by diluting the stock solution (below) to 25% with DW. Stock solution (100% Lugol solution) contains 10 g of KI and 5 g of I2, adjusted to 100 mL with DW. 
	NOTE: Lugol solution is light-sensitive, so store and handle the solution protected from light. Follow the regulations of each country and institution for iodine handling and waste disposal.</li>
	<li>Decant the DW from the samples and pour in the 25% Lugol solution. Stain for 24 h at room temperature.</li>
</ol>
3. Stage Mounting
<ol>
	<li>Prepare 0.5% agarose by dissolving 500 mg agarose in 100 mL of DW in a 250 mL conical flask in a microwave (800 W, about 1-3 min). Cool to about 30-40 &#176;C by keeping at room temperature. 
	CAUTION: Do not overheat or completely seal the flask when heating to prevent the agarose from boiling over.</li>
	<li>Mount large&#160;samples such as A. equina using a 50 mL tube. 
	NOTE: Use 50 mL tubes for mounting large samples that do not fit in a 1,000 &#956;L micropipette &#39;blue&#39; tip.
	<ol>
		<li>Place the stained sample in a 60-mm dish with DW to wash off excessive staining solution from the surface.</li>
		<li>Gently pour 5 mL of 0.5% agarose into a 50 mL tube and harden the agarose on ice. Be careful not to make bubbles in the agarose.</li>
		<li>Gently add 20 mL of 0.5% agarose to the 50 mL tube and place on ice until the agarose begins to harden. Place the specimen within the 0.5% agarose using forceps. Be careful not to make bubbles in the agarose.</li>
		<li>Adjust the position and orientation of the sample with forceps and harden the agarose on ice.</li>
		<li>Place clay on the microCT mounting stage and set the 50 mL tube on the clay (Figure 2A).</li>
	</ol>
	</li>
	<li>Mount small&#160;samples such as Harmothoe sp. and X. japonica using a 1,000 &#181;L micropipette &#39;blue&#39;&#160;tip. 
	NOTE: For smaller samples, 200 &#956;L micropipette &#8216;yellow&#8217; tip can be used. In this case, use 30 &#956;L of agarose for the plug and add 200 &#956;L of agarose or distilled water.
	<ol>
		<li>Draw up 100 &#181;L of 0.5% agarose into a 1,000 &#181;L micropipette &#39;blue&#39;&#160;tip and harden the agarose by holding the tip over&#160;ice, making a plug in the tip (Figure 2B-a).</li>
		<li>Decant the stained sample into a 60-mm dish without using forceps.</li>
		<li>Gently transfer the sample using ring tweezers into another 60-mm dish with DW to wash off excessive staining solution from the surface.</li>
		<li>Add 1,000 &#181;L of either&#160;0.5% agarose or&#160;DW into the plugged tip (step 3.3.1) using a micropipette. 
		NOTE: Use of 0.5% agarose is recommended, but use DW for fragile or precious samples in which agarose should be avoided.</li>
		<li>Gently transfer the sample from the 60-mm dish into the agarose or&#160;DW in the plugged tip using ring tweezers.</li>
		<li>Gently adjust the position and orientation of the sample with a petiolate needle or precision tweezers. Be careful not to make bubbles in the mounting medium. Make sure the sample is stable between the walls of the tip when DW is used as mounting medium (Figure 2D-b). Place the tip on ice to harden the agarose if it is used as mounting medium.</li>
		<li>Cut the tip off a new 1,000 &#181;L micropipette &#39;blue&#39;&#160;tip (Figure 2B-b,c) and insert the tip of the plugged tip into the new tip.</li>
		<li>Place clay on the microCT mounting stage and set the tips with the sample inside on the clay (Figure 2C,D). 
		NOTE: The staining solution will start to wash off the sample once it is placed in DW, so proceed to the next scanning step promptly.</li>
	</ol>
	</li>
</ol>
4. MicroCT scanning
<ol>
	<li>Turn on the X-ray beam at 80 kV, 100 &#181;A.</li>
	<li>While observing the X-ray transmission image at the center of the screen (Figure 3A), move the stage so that the whole sample can be seen by clicking on the X and Z axis buttons (Figure 3A), and manually adjusting the Y axis knob on the mounting stage (Figure 3B). Set the contrast of the image so that the internal structures can be observed by adjusting the contrast conditions (Figure 3A: Image contrast).</li>
	<li>Adjust the orientation of the sample by changing the angle of the tube/tip in the clay (Figure 2A). Rotate the stage 90&#176; by setting the rotation axis (Figure 3A) to 90 and clicking on the relative movement button (Figure 3A). Perform the same maneuver four times to complete a full rotation. 
	NOTE: Manually turn off the X-ray beam each time the sample door is opened, unless the system turns it off automatically.</li>
	<li>Move the stage so that the sample is at the center of the view by clicking on the Z axis button (Figure 3A) and by manually adjusting the Y axis knob on the mounting stage (Figure 3B). Turn the stage by 90&#176; and do the same. Turn the stage 360&#176; and check that the sample is at the center of the view from all directions.</li>
	<li>Move the stage along the x-axis toward the X-ray beam source by clicking on the X axis button (Figure 3A) to enlarge the sample and adjust as needed so that it just fits in the view (Figure 3C).</li>
	<li>Turn the stage 360&#176; and adjust as needed so&#160;that the sample fits within view from all directions.</li>
	<li>Adjust the scanning conditions as shown in Table 1.</li>
	<li>Start scanning; it takes about 10 min.</li>
</ol>
5. Image reconstruction
<ol>
	<li>Start up the microCT system&#39;s accessory software (see the Table of Materials) and open the scanned data.</li>
	<li>Adjust differences in the rotation axis of the sample during scanning by clicking on the automatic shift value calculation button (Figure 4A: green box).</li>
	<li>Adjust the orientation of the image by rotating the orange arrows (Figure 4B). If the orientation was changed, repeat step 5.2.</li>
	<li>Click on the area tab (Figure 4C: magenta box) and trim areas where samples are not present (Figure 4C: yellow box).</li>
	<li>Click on the reconfiguration tab (Figure 4D: magenta box) and set the filters as follows to remove noise. Ring artifact reduce filter: Median filter -3; Noise elimination filter: Average filter-1.</li>
	<li>Perform reconfiguration by clicking on the reconfiguration button (Figure 4D: green box).</li>
	<li>Adjust image brightness and contrast by setting the black and white values as black value 0, white value 250 (Figure 4D: blue box).</li>
	<li>Save the reconstructed TIFF image dataset as an 8-bit TIFF by clicking on the save button. Rename TIFF files as following: Date_sample_resolution (&#181;m)_number.tiff. 
	NOTE: The original microCT datasets from this study are available in the Figshare repository, doi:10.6084/m9.figshare.767083736.</li>
</ol>
6. Data analyses
<ol>
	<li>Start up the data analysis software (see the Table of Materials) and enable importing of TIFF files by clicking on the Database icon (Figure 5A: magenta box) and turning off the box shown in Figure 5B.</li>
	<li>Click on the import icon (Figure 5C: magenta box), select the dataset saved in section 5, and click open.</li>
	<li>Click on the copy links button (Figure 5D) to import the data.</li>
	<li>Display the 2D cross-section by clicking on the 2D viewer icon (Figure 5C: blue box).</li>
	<li>Calibrate the dataset by clicking on the 3D viewer tab (Figure 5E: magenta box) and entering the resolution value at scanning (which was 0.018 in this study [Figure 5F]).</li>
	<li>Click on the brightness/contrast icon (Figure 5E green box). Adjust the brightness and contrast by moving the cursor inside the displayed 2D image and changing the window level and window width (Figure 5G).</li>
	<li>Check other cross-section images by moving the scrollbar (Figure 5G: box).</li>
	<li>Change the orientation of the cross-section by clicking on the orientation icon (Figure 5E: blue box) and check images at all orientations (Figure 5H).</li>
	<li>Click on the displayed image, and choose the export tab to store cross-section images.</li>
</ol>

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D., Giribet, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+micro-computed+tomography+as+a+minimally+invasive+tool+for+anatomical+study+of+bivalves+(Mollusca:+Bivalvia).">The use of micro-computed tomography as a minimally invasive tool for anatomical study of bivalves (Mollusca: Bivalvia).</a> Zoological Journal of the Linnean Society. , (2018).</li><li>Sasaki, T., Endo, K., Kogure, T., Nagasawa, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=3D+visualization+of+calcified+and+non-calcified+molluscan+tissues+using+computed+tomography.">3D visualization of calcified and non-calcified molluscan tissues using computed tomography.</a> Biomineralization. , 83-93 (2018).</li><li>Maeno, A., Tsuda, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Micro-computed+Tomography+to+Visualize+Vascular+Networks+in+Maize+Stems.">Micro-computed Tomography to Visualize Vascular Networks in Maize Stems.</a> Bio-protocol. 8 (1), 2682 (2018).</li><li>Nakano, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Correction+to:+A+new+species+of+Xenoturbella+from+the+western+Pacific+Ocean+and+the+evolution+of+Xenoturbella.">Correction to: A new species of Xenoturbella from the western Pacific Ocean and the evolution of Xenoturbella.</a> BMC Evolutionary Biology. 18, 83 (2018).</li><li>Maeno, A., Kohtsuka, H., Takatani, K., Nakano, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=MicroCT+files+from+'Microfocus+X-ray+computed+tomography+(microCT)+imaging+of+Actinia+equina+(Cnidaria),+Harmothoe+sp.+(Annelida),+and+Xenoturbella+japonica+(Xenacoelomorpha)'.">MicroCT files from 'Microfocus X-ray computed tomography (microCT) imaging of Actinia equina (Cnidaria), Harmothoe sp. (Annelida), and Xenoturbella japonica (Xenacoelomorpha)'.</a> figshare. , (2019).</li><li>Vickerton, P., Jarvis, J., Jeffery, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Concentration-dependent+specimen+shrinkage+in+iodine-enhanced+microCT.">Concentration-dependent specimen shrinkage in iodine-enhanced microCT.</a> Journal of Anatomy. 223 (2), 185-193 (2013).</li><li>Buytaert, J., Goyens, J., De Greef, D., Aerts, P., Dirckx, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Volume+shrinkage+of+bone,+brain+and+muscle+tissue+in+sample+preparation+for+micro-CT+and+light+sheet+fluorescence+microscopy+(LSFM).">Volume shrinkage of bone, brain and muscle tissue in sample preparation for micro-CT and light sheet fluorescence microscopy (LSFM).</a> Microscopy and Microanalysis. 20 (4), 1208-1217 (2014).</li><li>Sasov, A., Liu, X., Salmon, P. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Compensation+of+mechanical+inaccuracies+in+micro-CT+and+nano-CT.">Compensation of mechanical inaccuracies in micro-CT and nano-CT.</a> Proceedings of SPIE. 7078, 70781 (2008).</li></ol>

The authors have nothing to disclose.

Fixatives using formalin, such as the 10% (v/v) formalin solution in seawater used in this study, are known to preserve the morphology of diverse marine invertebrates and are often used for microCT imaging18,24,25,26,28,30,33. However, restrictions on the use of this chemical have become str...

Biological researchers generally have had to make thin sections and perform observations by light or electron microscopy in order to investigate the internal structures of opaque organisms. However, these methods are destructive and problematic when applied to rare or valuable specimens. Furthermore, several steps in the method, such as embedding and sectioning, are time consuming, and it can take several days to observe a sample, depending on the protocol. Moreover, when handling numerous sections, there is always a possibility of damaging or losing some sections. Tissue-clearing techniques are available for some specimens1,2,3,4,5 but are not yet applicable for many animal species.
To overcome these problems, some biologists have started using microfocus X-ray computed tomography (microCT) imaging6,7,8,9,10,11,12,13,14,15. In X-ray CT, the specimen is irradiated with X-rays from various angles that are generated from an X-ray source moving around the sample, and the transmitted X-rays are monitored by a detector that also moves around the sample. The X-ray transmission data obtained are analyzed to reconstruct cross-sectional images of the specimen. This method enables the observation of internal structures without destruction of the sample. Because of its safety and ease, it is commonly used in medical and dental applications, and CT systems can be found in hospitals and dental centers worldwide. Moreover, industrial X-ray CT is frequently used for observing non-medical samples for inspection and metrology in the industrial field. In contrast to medical CT, in which the X-ray source and the detectors are mobile, the two parts are fixed in industrial CT, with the sample rotating during scanning. Industrial CT generally produces higher resolution images than medical CT and is referred to as microCT (micrometer-level resolution) or nanoCT (nanometer-level resolution). Recently, research using microCT has rapidly increased in various fields of biology14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34.
Biological studies using CT initially targeted internal structures that mainly consist of hard tissue, such as bone. Advances in staining techniques using various chemical agents enabled the visualization of soft tissues in various organisms6,7,8,9,14,15,16,17,18,19,20,21,22,23,24,25,26,27,28,29,30,31,32,33,34. Of these reagents, iodine-based contrast agents are relatively safe, inexpensive, and can be used for the visualization of soft tissues in various organisms7,14. Concerning marine invertebrates, microCT has been widely used on such animals as molluscs6,25,32,33, annelids18,19,20,28, and arthoropods21,23,29,31. However, there have been few reports on other animal phyla, such as bryozoans6, xenacoelomorphs26, and cnidarians24,30. In general, there have been fewer studies using microCT on marine invertebrates than those on vertebrates. One major reason for this limited use on marine invertebrates is the vast diversity observed in these animals. Because of their diverse sizes, morphologies, and physiologies, each species reacts differently to different experimental procedures. Therefore, it is crucial during sample preparation to choose the most appropriate fixation and staining reagent, and to set conditions at each step, adjusted for each species. Similarly, it is also necessary to set the scanning configurations, such as the mounting method, voltage, current, mechanical magnifying rate, and the space resolution power, appropriately for each sample. To overcome this problem, a simple and comprehensible manual that covers all of the necessary steps, explains how each step can be adjusted depending on the specimen, and shows detailed examples from multiple samples is essential.
In the present study, we describe the microCT imaging protocol step-by-step, from sample fixation to data analysis, using three marine invertebrate species. Specimens of the sea anemone Actinia equina (Anthozoa, Cnidaria) were collected near the Misaki Marine Biological Station, University of Tokyo. They had a spherical, soft body that was about 2 cm in diameter (Figure 1A-C). Harmothoe sp. (Polychaeta, Annelida) samples were also collected near Misaki Marine Biological Station. They were slender worms that were about 1.5 cm in length, with tough chaetae present along the whole body (Figure&#160;1D). A Xenoturbella japonica35 (Xenoturbellida, Xenacoelomorpha) specimen was collected near Shimoda Marine Research Center, University of Tsukuba, during the 13th JAMBIO Coastal Organism Joint Survey. It was a soft-bodied worm that was about 0.8 cm in length (Figure 1E). Adjustments made for the conditions and configurations of each sample are explained in detail. Our study provides several suggestions on how to perform microCT imaging on marine invertebrates, and we hope that it will inspire biologists to utilize this protocol for their research.

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			<td>Wako Pure Chemical Industries</td>
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			<td>Comscantechno</td>
			<td>ScanXmate-E090S105</td>
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microfocus x-ray ct (microct) imaging of actinia equina (cnidaria), harmothoe sp. (annelida), and xenoturbella japonica (xenacoelomorpha)

Traditionally, biologists have had to rely on destructive methods such as sectioning in order to investigate the internal structures of opaque organisms. Non-destructive microfocus X-ray computed tomography (microCT) imaging has become a powerful and emerging protocol in biology, due to technological advancements in sample staining methods and innovations in microCT hardware, processing computers, and data analysis software. However, this protocol is not commonly used, as it is in the medical and industrial fields. One of the reasons for this limited use is the lack of a simple and comprehensible manual that covers all of the necessary steps: sample collection, fixation, staining, mounting, scanning, and data analyses. Another reason is the vast diversity of metazoans, particularly marine invertebrates. Because of marine invertebrates&#8217; diverse sizes, morphologies, and physiologies, it is crucial to adjust experimental conditions and hardware configurations at each step, depending on the sample. Here, microCT imaging methods are explained in detail using three phylogenetically diverse marine invertebrates: Actinia equina (Anthozoa, Cnidaria), Harmothoe sp. (Polychaeta, Annelida), and Xenoturbella japonica (Xenoturbellida, Xenacoelomorpha). Suggestions on performing microCT imaging on various animals are also provided.

We performed microCT imaging on A. equina (Anthozoa, Cnidaria), Harmothoe sp. (Polychaeta, Annelida), and X. japonica (Xenoturbellida, Xenacoelomorpha) after staining the samples with 25% Lugol solution. The staining successfully enhanced the contrast of the internal structures in all specimens, enabling observations of internal soft tissues (Figure 6). Together with past reports6,7,<sup class="x...

Watch this Scientific Journal Video about Microfocus X-ray CT microCT Imaging of Actinia equina Cnidaria, Harmothoe sp. Annelida, and Xenoturbella japonica Xenacoelomorpha at JoVE.com

Microfocus X-ray CT microCT Imaging of Actinia equina Cnidaria, Harmothoe sp. Annelida, and Xenoturbella japonica Xenacoelomorpha

Here, protocols for performing microfocus X-ray computed tomography (microCT) imaging of three marine invertebrate animals are explained in detail. This study describes steps such as sample fixation, staining, mounting, scanning, image reconstruction, and data analyses. Suggestions on how the protocol can be adjusted for different samples are also provided.

Microfocus X-ray CT (microCT) Imaging of Actinia equina (Cnidaria), Harmothoe sp. (Annelida), and Xenoturbella japonica (Xenacoelomorpha)

Traditionally, biologists have had to rely on destructive methods such as sectioning in order to investigate the internal structures of opaque organisms. ...

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8.9K Views.  National Institute of Genetics. Here, protocols for performing microfocus X-ray computed tomography (microCT) imaging of three marine invertebrate animals are explained in detail. This study describes steps such as sample fixation, staining, mounting, scanning, image reconstruction, and data analyses. Suggestions on how the protocol can be adjusted for different samples are also provided.

Video: Microfocus X-ray CT microCT Imaging of Actinia equina Cnidaria, Harmothoe sp. Annelida, and Xenoturbella japonica Xenacoelomorpha

Extracting Metrics for Three-dimensional Root Systems: Volume and Surface Analysis from In-soil X-ray Computed Tomography Data

Plant roots play a critical role in plant-soil-microbe interactions that occur in the rhizosphere, as well as processes with important implications to climate change and crop management. Quantitative size information on roots in their native environment is invaluable for studying root growth and environmental processes involving plants. X-ray computed tomography (XCT) has been demonstrated to be an effective tool for in situ root scanning and analysis. We aimed to develop a costless and efficient tool that approximates the surface and volume of the root regardless of its shape from three-dimensional (3D) tomography data. The root structure of a Prairie dropseed (Sporobolus heterolepis) specimen was imaged using XCT. The root was reconstructed, and the primary root structure was extracted from the data using a combination of licensed and open-source software. An isosurface polygonal mesh was then created for ease of analysis. We have developed the standalone application imeshJ, generated in MATLAB1, to calculate root volume and surface area from the mesh. The outputs of imeshJ are surface area (in mm2) and the volume (in mm3). The process, utilizing a unique combination of tools from imaging to quantitative root analysis, is described. A combination of XCT and open-source software proved to be a powerful combination to noninvasively image plant root samples, segment root data, and extract quantitative information from the 3D data. This methodology of processing 3D data should be applicable to other material/sample systems where there is connectivity between components of similar X-ray attenuation and difficulties arise with segmentation.

A methodology for obtaining visual and quantitative root structure information from X-ray computed tomography data acquired in-soil is presented.

Plant roots play a critical role in plant-soil-microbe interactions that occur in the rhizosphere, as well as processes with important implications to ...

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination control. Developments of some techniques aiming to reduce trace metal contamination in the last couple of decades have resulted in concentrations reported now being orders of magnitude lower than those in the past. These low concentrations often necessitate preconcentration of water samples prior to instrumental analysis of samples. Since contamination can appear in all phases of trace metal analyses, including sample collection (and during preparation of sampling containers), storage and handling, pretreatments, and instrumental analysis, specific care needs to be taken in order to reduce contamination levels at all steps. The effort to develop and utilize "clean techniques" in trace metal studies allows scientists to investigate trace metal distributions and chemical and biological behavior in greater details. This advancement also provides the required accuracy and precision of trace metal data allowing for environmental conditions to be related to trace metal concentrations in aquatic environments.
This protocol that is presented here details needed materials for sample preparation, sample collection, sample pretreatment including preconcentration, and instrumental analysis. By reducing contamination throughout all phases mentioned above for trace metal analysis, much lower detection limits and thus accuracy can be achieved. The effectiveness of "clean techniques" is further demonstrated using low field blanks and good recoveries for standard reference material. The data quality that can be obtained thus enables the assessment of trace metal distributions and their relationships to environmental parameters.

Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination ...

Data Processing Methods for 3D Seismic Imaging of Subsurface Volcanoes: Applications to the Tarim Flood Basalt

The morphology and structure of plumbing systems can provide key information on the eruption rate and style of basalt lava fields. The most powerful way to study subsurface geo-bodies is to use industrial 3D&#160;reflection seismological imaging. However, strategies to image subsurface volcanoes are very different from that of oil and gas reservoirs. In this study, we process seismic data cubes from the Northern Tarim Basin, China, to illustrate how to visualize sills through opacity rendering techniques and how to image the conduits by time-slicing. In the first case, we isolated probes by the seismic horizons marking the contacts between sills and encasing strata, applying opacity rendering techniques to extract sills from the seismic cube. The resulting detailed sill morphology shows that the flow direction is from the dome center to the rim. In the second seismic cube, we use time-slices to image the conduits, which corresponds to marked discontinuities within the encasing rocks. A set of time-slices obtained at different depths show that the Tarim flood basalts erupted from central volcanoes, fed by separate pipe-like conduits.

Three-dimensional (3D) reflection seismology is a powerful method for imaging subsurface volcanoes. By using industrial 3D seismological data from the Tarim Basin, we illustrate how to extract the sills and the conduits of subsurface volcanoes from seismic data cubes.

The morphology and structure of plumbing systems can provide key information on the eruption rate and style of basalt lava fields. The most powerful way ...

RGB and Spectral Root Imaging for Plant Phenotyping and Physiological Research: Experimental Setup and Imaging Protocols

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop cultivars against environmental stresses. An experimental protocol is presented for RGB and hyperspectral imaging of root systems. The approach uses rhizoboxes where plants grow in natural soil over a longer time to observe fully developed root systems. Experimental settings are exemplified for assessing rhizobox plants under water stress and studying the role of roots. An RGB imaging setup is described for cheap and quick quantification of root development over time. Hyperspectral imaging improves root segmentation from the soil background compared to RGB color based thresholding. The particular strength of hyperspectral imaging is the acquisition of chemometric information on the root-soil system for functional understanding. This is demonstrated with high resolution water content mapping. Spectral imaging however is more complex in image acquisition, processing and analysis compared to the RGB approach. A combination of both methods can optimize a comprehensive assessment of the root system. Application examples integrating root and aboveground traits are given for the context of plant phenotyping and plant physiological research. Further improvement of root imaging can be obtained by optimizing RGB image quality with better illumination using different light sources and by extension of image analysis methods to infer on root zone properties from spectral data.

An experimental protocol is presented for assessment of soil grown plant root systems with RGB and hyperspectral imaging. Combination of RGB image time series with chemometric information from hyperspectral scans optimizes insights into plant root dynamics.

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop ...

Stress Distribution During Cold Compression of Rocks and Mineral Aggregates Using Synchrotron-based X-Ray Diffraction

We report detailed procedures for performing compression experiments on rocks and mineral aggregates within a multi-anvil deformation apparatus (D-DIA) coupled with synchrotron X-radiation. A cube-shaped sample assembly is prepared and compressed, at room temperature, by a set of four X-ray transparent sintered diamond anvils and two tungsten carbide anvils, in the lateral and the vertical planes, respectively. All six anvils are housed within a 250-ton hydraulic press and driven inward simultaneously by two wedged guide blocks. A horizontal energy dispersive X-ray beam is projected through and diffracted by the sample assembly. The beam is commonly in the mode of either white or monochromatic X-ray. In the case of white X-ray, the diffracted X-rays are detected by a solid-state detector array that collects the resulting energy dispersive diffraction pattern. In the case of monochromatic X-ray, the diffracted pattern is recorded using a two-dimensional (2-D) detector, such as an imaging plate or a charge-coupled device (CCD) detector. The 2-D diffraction patterns are analyzed to derive lattice spacings. The elastic strains of the sample are derived from the atomic lattice spacing within grains. The stress is then calculated using the predetermined elastic modulus and the elastic strain. Furthermore, the stress distribution in two-dimensions allow for understanding how stress is distributed in different orientations. In addition, a scintillator in the X-ray path yields a visible light image of the sample environment, which allows for the precise measurement of sample length changes during the experiment, yielding a direct measurement of volume strain on the sample. This type of experiment can quantify the stress distribution within geomaterials, which can ultimately shed light on the mechanism responsible for compaction. Such knowledge has the potential to significantly improve our understanding of key processes in rock mechanics, geotechnical engineering, mineral physics, and material science applications where compactive processes are important.

We report detailed procedures for compression experiments on rocks and mineral aggregates within a multi-anvil deformation apparatus coupled with synchrotron X-radiation. Such experiments allow quantification of the stress distribution within samples, that ultimately sheds light on compaction processes in geomaterials.

We report detailed procedures for performing compression experiments on rocks and mineral aggregates within a multi-anvil deformation apparatus (D-DIA) ...

Harvesting Venom Toxins from Assassin Bugs and Other Heteropteran Insects

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, and the majority of extant heteropterans retain this trophic strategy. Some heteropterans have transitioned to feeding on vertebrate blood (such as the kissing bugs, Triatominae; and bed bugs, Cimicidae) while others have reverted to feeding on plants (most Pentatomomorpha). However, with the exception of saliva used by kissing bugs to facilitate blood-feeding, little is known about heteropteran venoms compared to the venoms of spiders, scorpions and snakes.
One obstacle to the characterization of heteropteran venom toxins is the structure and function of the venom/labial glands, which are both morphologically complex and perform multiple biological roles (defense, prey capture, and extra-oral digestion). In this article, we describe three methods we have successfully used to collect heteropteran venoms. First, we present electrostimulation as a convenient way to collect venom that is often lethal when injected into prey animals, and which obviates contamination by glandular tissue. Second, we show that gentle harassment of animals is sufficient to produce venom extrusion from the proboscis and/or venom spitting in some groups of heteropterans. Third, we describe methods to harvest venom toxins by dissection of anaesthetized animals to obtain the venom glands. This method is complementary to other methods, as it may allow harvesting of toxins from taxa in which electrostimulation and harassment are ineffective. These protocols will enable researchers to harvest toxins from heteropteran insects for structure-function characterization and possible applications in medicine and agriculture.

Although many insects in the suborder Heteroptera (Insecta: Hemiptera) are venomous, their venom composition and the functions of their venom toxins are mostly unknown. This protocol describes methods to harvest heteropteran venoms for further characterization, using electrostimulation, harassment, and gland dissection.

Heteropteran insects such as assassin bugs (Reduviidae) and giant water bugs (Belostomatidae) descended from a common predaceous and venomous ancestor, ...

Investigation of Xenobiotics Metabolism In Salix alba Leaves via Mass Spectrometry Imaging

The method presented uses mass spectrometry imaging (MSI) to establish the metabolic profile of S. alba leaves when exposed to xenobiotics. Using a non-targeted approach, plant metabolites and xenobiotics of interest are identified and localized in plant tissues to uncover&#160;specific distribution patterns. Then, in silico prediction of potential metabolites (i.e., catabolites and conjugates) from the identified xenobiotics is performed. When a xenobiotic metabolite is located in the tissue, the type of enzyme involved in its alteration by the plant is recorded. These results were used to describe different types of biological reactions occurring in S. alba leaves in response to xenobiotic accumulation in the leaves. The metabolites were predicted in two generations, allowing the documentation of successive biological reactions to transform xenobiotics in the leaf tissues.

Investigation of Xenobiotics Metabolism In Salix alba Leaves via Mass Spectrometry Imaging

This method uses mass spectrometry imaging (MSI) to understand metabolic processes in S. alba leaves when exposed to xenobiotics. The method allows the spatial localization of compounds of interest and their predicted metabolites within specific, intact tissues.

The method presented uses mass spectrometry imaging (MSI) to establish the metabolic profile of S. alba leaves when exposed to xenobiotics. Using a ...

A Method to Preserve Wetland Roots and Rhizospheres for Elemental Imaging

Roots extensively interact with their soil environment but visualizing such interactions between roots and the surrounding rhizosphere is challenging. The rhizosphere chemistry of wetland plants is particularly challenging to capture because of steep oxygen gradients from the roots to the bulk soil. Here a protocol is described that effectively preserves root structure and rhizosphere chemistry of wetland plants through slam-freezing and freeze drying. Slam-freezing, where the sample is frozen between copper blocks pre-cooled with liquid nitrogen, minimizes root damage and sample distortion that can occur with flash-freezing while still minimizing chemical speciation changes. While sample distortion is still possible, the ability to obtain multiple samples quickly and with minimal cost increases the potential to obtain satisfactory samples and optimizes imaging time. The data show that this method is successful in preserving reduced arsenic species in rice roots and rhizospheres associated with iron plaques. This method can be adopted for studies of plant-soil relationships in a wide variety of wetland environments that span concentration ranges from trace-element cycling to phytoremediation applications.

We describe a protocol to sample, preserve, and section intact roots and the surrounding rhizosphere soil from wetland environments using rice (Oryza sativa L.) as a model species. Once preserved, the sample can be analyzed using elemental imaging techniques, such as synchrotron X-ray fluorescence (XRF) chemical speciation imaging.

Roots extensively interact with their soil environment but visualizing such interactions between roots and the surrounding rhizosphere is challenging. The ...

Leveraging Micro-CT Scanning to Analyze Parasitic Plant-Host Interactions

Micro-CT scanning has become an established tool in investigating plant structure and function. Its non-destructive nature, combined with the possibility of three-dimensional visualization and virtual sectioning, has allowed novel and increasingly detailed analysis of complex plant organs. Interactions among plants, including between parasitic plants and their hosts, can also be explored. However, sample preparation before scanning becomes crucial due to the interaction between these plants, which often differ in tissue organization and composition. Furthermore, the broad diversity of parasitic flowering plants, ranging from highly reduced vegetative bodies to trees, herbs, and shrubs, must be considered during the sampling, treatment, and preparation of parasite-host material. Here two different approaches are described for introducing contrast solutions into the parasite and/or host plants, focusing on analyzing the haustorium. This organ promotes connection and communication between the two plants. Following a simple approach, details of haustorium tissue organization can be explored three-dimensionally, as shown here for&#160;euphytoid, vine, and mistletoe parasitic species. Selecting specific contrasting agents and application approaches also allow detailed observation of endoparasite spread within the host body and detection of direct vessel-to-vessel connection between parasite and host, as shown here for an obligate root parasite. Thus, the protocol discussed here can be applied to the broad diversity of parasitic flowering plants to advance the understanding of their development, structure, and functioning.

Micro-CT is a non-destructive tool that can analyze plant structures in three dimensions. The present protocol describes the sample preparation to leverage micro-CT to analyze parasitic plant structure and function. Different species are used to highlight the advantages of this method when coupled with specific preparations.

Micro-CT scanning has become an established tool in investigating plant structure and function. Its non-destructive nature, combined with the possibility ...

Isolation and Identification of Waterborne Antibiotic-Resistant Bacteria and Molecular Characterization of their Antibiotic Resistance Genes

The development and spread of antibiotic resistance (AR) through microbiota associated with freshwater bodies is a major global health concern. In the present study, freshwater samples were collected and analyzed with respect to the total bacterial diversity and AR genes (ARGs) using both conventional culture-based techniques and a high-throughput culture-independent metagenomic approach. This paper presents a systematic protocol for the enumeration of the total and antibiotic-resistant culturable bacteria from freshwater samples and the determination of phenotypic and genotypic resistance in the culturable isolates. Further, we report the use of whole metagenomic analysis of the total metagenomic DNA extracted from the freshwater sample for the identification of the overall bacterial diversity, including non-culturable bacteria, and the identification of the total pool of different ARGs (resistome) in the water body. Following these detailed protocols, we observed a high antibiotic-resistant bacteria load in the range of 9.6 &#215; 105-1.2 &#215; 109 CFU/mL. Most isolates were resistant to the multiple tested antibiotics, including cefotaxime, ampicillin, levofloxacin, chloramphenicol, ceftriaxone, gentamicin, neomycin, trimethoprim, and ciprofloxacin, with multiple antibiotic resistance (MAR) indexes of &#8805;0.2, indicating high levels of resistance in the isolates. The 16S rRNA sequencing identified potential human pathogens, such as Klebsiella pneumoniae,&#160;and opportunistic bacteria, such as Comamonas spp., Micrococcus spp., Arthrobacter spp., and Aeromonas spp. The molecular characterization of the isolates showed the presence of various ARGs, such as blaTEM, blaCTX-M (&#946;-lactams), aadA, aac (6&#39;)-Ib (aminoglycosides), and dfr1 (trimethoprims), which was also confirmed by the whole metagenomic DNA analysis. A high prevalence of other ARGs encoding for antibiotic efflux pumps-mtrA, macB, mdtA, acrD, &#946;-lactamases-SMB-1, VIM-20, ccrA, ampC, blaZ, the&#160;chloramphenicol acetyltransferase gene catB10, and&#160;the rifampicin resistance gene rphB-was also detected in the metagenomic DNA. With the help of the protocols discussed in this study, we confirmed the presence of waterborne MAR bacteria with diverse AR phenotypic and genotypic traits. Thus, whole metagenomic DNA analysis can be used as a complementary technique to conventional culture-based techniques to determine the overall AR status of a water body.

Here, we present a detailed protocol for the isolation and identification of antibiotic-resistant bacteria from water and the molecular characterization of their antibiotic resistance genes (ARGs). The use of culture-based and non-culture-based (metagenomic analysis) techniques provides complete information about the total bacterial diversity and the total pool of different ARGs present in freshwaters from Mumbai, India.

The development and spread of antibiotic resistance (AR) through microbiota associated with freshwater bodies is a major global health concern. In the ...