I would like to thank Dr. Simone Gomes Ferreira (Microtomography Laboratory, University of Sao Paulo, Brazil) and Dr. Greg Lin (Center for Nanoscale Systems, Harvard University, USA) for their paramount help and indispensable user training for different microtomography systems and data analysis software. I also thank the staff at the EEB Greenhouse at the University of Connecticut (USA), especially Clinton Morse and Matthew Opel for providing the specimens of Viscum minimum. Dr. John Wenzel provided the opportunity and great help for the sampling of Pyrularia pubera. MSc. Carolina Bastos, MSc. Yasmin Hirao, and Talitha Motta greatly helped with the sampling of Scybalium fungiforme. MSc. Ariadne Furtado, and Drs. Fernanda Oliveira and Maria Aline Neves provided the reference for the use of phloxine B for the analysis of endophytic fungi. Video recording at the Vrije Universiteit Brussel was made possible through the help of Dr. Philippe Claeys, Dr. Christophe Snoeck, MSc. Jake Griffith, Dr. Barabara Veselka, and Dr. Harry Olde Venterink.&#160;Funding was provided by the Coordination for the Improvement of Higher Education Personnel (CAPES, Brazil) and the Harvard University Herbaria (USA).

1. Parasitic plant sample selection
<ol>
	<li>Collect the entire parasitic plant haustorium, including the attached host stem/root and segments of both proximal and distal ends of the parasitized host organ; the ideal length of each segment is equivalent to double the diameter of the haustorium. 
	NOTE: For lateral haustoria, include part of the parasite mother stem/root from which the haustorium was formed (Figure 1A,B,D). For endoparasites, collect a segment of the host stem/root in which signs of the parasite are visible (Figure 1B). In the case of terminal connections, the whole plant should be collected (Figure 1E).</li>
	<li>Submerge the whole sample in fixative solution (e.g., FAA) in a volumetric proportion of 1:10 (sample: fixative). Leave samples in fixative for at least 1 day, depending on the size of the sample27. 
	&#8203;NOTE: Samples can be stored in fixative before scanning or be transferred to a preserving solution (e.g., ethanol 70%). Fresh samples can also be used if the subsequent anatomical analysis is not warranted (see Discussion section). If working with fresh material, set up the apparatus for perfusion of the contrast solution, then collect the sample. The sample should not be allowed to dry out.</li>
</ol>
2. Application of contrasting solutions
<ol>
	<li>Chose the application method to be used. Use the vacuum method (step 2.3) for small (Figure 1A) or non-woody (Figure 1B) samples. Use the perfusion method for larger samples, provided it includes a segment of the host stem/root is (Figure 1A,C-E).</li>
	<li>Wear rubber gloves and other appropriate personal protection equipment (e.g., laboratory coat) when manipulating the sample regardless of the approach chosen in the following steps. 
	CAUTION: All contrasting solutions include heavy metal salts in their composition, and therefore should not be handled without adequate personal protection equipment and under a fume hood.</li>
	<li>For the vacuum method, follow the steps mentioned below.
	<ol>
		<li>Choose an appropriate vial and label it. The vial must be large enough to accommodate the sample and the contrasting solution, usually in a proportion of 1:10. Check instructions from the manufacturer to ensure that the vial can withstand low to moderate vacuum. 
		NOTE: Do not fill the vial to the brim, as the negative pressure (vacuum) can cause the liquid to spill.</li>
		<li>Place the sample in the vial with the contrasting solution (1% iodine or 3% phosphotungstate, see Table of Materials). Then place the vial in a vacuum chamber or desiccator connected to a vacuum pump. Remove the lid from the vial, then close the vacuum chamber or desiccator.</li>
		<li>Check that there are no cracks on the vacuum chamber/desiccator and that the vacuum pump has enough oil.</li>
		<li>Close the exhaustion valve of the pump to prevent air from scaping and open the exhaustion valve of the chamber/desiccator to force the air out.</li>
		<li>Turn on the pump and wait until the pressure reaches approximately 20&#34; Hg. 
		CAUTION: This process is usually fast, so do not leave the vacuum system unattended. 
		NOTE: While the metric system unit for pressure is Pascal (Pa), pressure gauges in most laboratory vacuum pumps display pressure in inches of mercury (&#34;Hg), pound per square inch (psi), or bars. 20&#34; Hg equals ca. 67.7 Pa, 10 psi, or 0.7 bar.</li>
		<li>Close the chamber/desiccator exhaust valve to prevent air from re-entering, then quickly turn off the pump.</li>
		<li>Leave the sample under vacuum for at least 2 h; larger samples require longer for the contrasting solution to penetrate it.</li>
		<li>After the desired period, remove the sample from the contrasting solution to prepare it for scanning.</li>
		<li>Slowly open the chamber/desiccator exhaustion valve to allow air to enter it.</li>
		<li>Wait for the pressure in the chamber/desiccator to be fully exhausted (i.e., pressure gauge reaches near 0), then carefully open it to retrieve the sample.</li>
		<li>Discard the contrasting solution appropriately and keep the sample in preparation for scanning.</li>
	</ol>
	</li>
	<li>For the perfusion method, follow the steps mentioned below.
	<ol>
		<li>Select a supply tank for the contrasting solution according to the size of the sample. Use a 50 mL syringe (with no needle or plunger) for small samples (Figure 2A) or a 1 L intravenous medical kit for large samples (Figure 2B).</li>
		<li>Connect one end of a transparent plastic tubing (see Table of Materials) to the supply tank, then connect the other end to a two-way or three-way valve. Connect a second tubing to a different outlet in the valve (Figure 2A,B).</li>
		<li>Secure the supply tank at an elevated position without disassembling the apparatus set up in the previous step. 
		NOTE: The vertical distance between the tank and the sample will dictate the solution perfusion pressure (Figure 2B). A distance of 20-50 cm is enough for small samples. For large samples, a distance of 1 m is more adequate.</li>
		<li>Close the three-way (Figure 2C) or two-way (Figure 2D) valve to prevent liquid from exiting the tubing system, then pour the contrasting solution into the supply tank. If using an intravenous medical kit, fill the bag with the solution and close the valve before securing the apparatus at an elevated position.</li>
		<li>Ensure that no large air bubbles are formed along the tubing system (Figure 2E). If necessary, allow the contrast solution to flow out of the tubing until the bubble is removed. Close the valve again and leave the apparatus in place.</li>
		<li>To prepare the sample for perfusion of the contrast solution, keep it submerged in liquid (water, ethanol, or fixative) and cut off the tip of the proximal end of the host stem/root in the sample (Figure 2F).</li>
		<li>Remove the sample from the liquid in which it was stored and wrap it in a paraffin film to avoid desiccation. Keep the sample nearby and ready to be connected to the apparatus.</li>
		<li>Carefully open the valve to allow the contrasting solution to flow slowly and fill the plastic tubing connected to the tank while holding the open end of the system at a slightly elevated position to prevent the contrasting solution from spilling. Again, ensure that no large air bubbles form along the tube.</li>
		<li>Connect the proximal end of the host stem/root in the sample to the open end of the tubing system (Figure 2B, magnified region). Avoid introducing air bubbles into the system during this step. If necessary, disconnect the sample from the apparatus and remove air bubbles from the system by allowing the solution to flow.</li>
		<li>While keeping the sample connected to the apparatus, place it inside a container to avoid leakage of the contrasting solution into the area where the experiment was set up. Use tubes of different diameters, plastic zip-ties, and valve adaptors (Figure 2G) to ensure all connections in the apparatus are well-fit, accommodate host branches of various sizes, and that the solution is not leaking out of the tubing system (Figure 2B, magnified region).</li>
		<li>Let the solution perfuse the sample for at least 2 h, or until the solution accumulates inside the container.</li>
		<li>Close the valve and carefully disconnect the sample from the apparatus. Drain the remainder of the solution into the container and dispose of it appropriately.</li>
		<li>Remove the paraffin film from the sample in preparation for scanning.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63423/63423fig02.jpg" /> 
Figure 2: Perfusion approach for contrast application. Small (A) and large (B) versions of the perfusion apparatus include a supply tank (st) and two plastic tubes (t1 and t2) connected by a valve (va). The proximal end of the host stem (H) bearing a parasite (P) attached to it via the haustorium (ha) is connected to the open end of the system (B, expanded). A three-way (C) or a two-way (D) valve is used to help prevent the formation of air bubbles inside the tubing system, which block the passage of contrasting solution (E). The tip of the proximal end of the host stem (H) is cut underwater to allow passage of the contrast solution (F). Zip-ties, valve adaptors, and tubing of different diameters help secure tighter connections and avoid leakage in the system (G). Figures 2B, D and F were created with BioRender. Scale bars = 2 cm. <a href="https://www.jove.com/files/ftp_upload/63423/63423fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Sample preparation and mounting
<ol>
	<li>Wash the sample by submerging it in water for 2 min. 
	CAUTION: Do not wash samples in the sink, as all contrasting solutions include heavy metal salts in their composition. Consider the water used for washing as a diluted contrasting solution and dispose of it appropriately.</li>
	<li>Place the sample in a paper towel at room temperature to allow excess water to evaporate for 2-5 min depending on the size of the sample. Alternatively, dry the sample slightly with the aid of a paper towel. Do not allow the sample to dry out completely.</li>
	<li>Wrap the sample in a paraffin film by stretching it to a thin layer. Avoid folding the paraffin film on top of the sample.</li>
	<li>Mount the wrapped sample onto a sample holder, keeping it stable and in position while it rotates&#160;during the scanning. Use adhesive tape, low-density foam, pipette tips, and/or clear plastic containers to secure the sample in place.</li>
	<li>Scan the sample and analyze the images following specific protocols and guidelines established for the micro-CT system available.</li>
</ol>

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R., 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=Automated+analysis+of+three-dimensional+xylem+networks+using+high-resolution+computed+tomography.">Automated analysis of three-dimensional xylem networks using high-resolution computed tomography.</a> New Phytologist. 191 (4), 1168-1179 (2011).</li><li>Lee, K., 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=Visualizing+plant+development+and+gene+expression+in+three+dimensions+using+optical+projection+tomography.">Visualizing plant development and gene expression in three dimensions using optical projection tomography.</a> Plant Cell. 18 (9), 2145-2156 (2006).</li></ol>

The author has nothing to disclose.

The use of heavy metal solutions to improve plant tissue contrast has become a crucial step in sample preparation for micro-CT analysis. Several compounds commonly available in plant micro-morphology laboratories have been tested by Staedler et al., who recommend using phosphotungstate as the most effective agent in penetrating samples and increasing contrast index8. Results obtained here in the analysis of the haustorium of P. pubera corroborate this recommendation. In terms of contrast ...

High-resolution x-ray microcomputed tomography (micro-CT) is an imaging method in which multiple radiographs (projections) of a sample are recorded from different viewing angles and later used to provide a virtual reconstruction of the sample1. This virtual object can then be analyzed, manipulated, and segmented, allowing non-destructive exploration in three dimensions2. Initially designed for medical analyses and later for industrial applications, micro-CT also offers the advantage of visualizing inner organs and tissues without the need for invasive procedures3. Like other forms of imaging, micro-CT works with a trade-off between the field of view and pixel size, which means that high-resolution imaging of large samples is nearly unattainable4. Advances in using high-energy X-ray sources (i.e., synchrotron) and secondary optical magnification are constantly being made, allowing the smallest resolution to reach under 100 nm5,6. Nevertheless, longer scanning times are necessary for large samples, increasing the chance of artifacts due to sample movement or deformation inside the scanner. Furthermore, micro-CT is generally limited by natural density variations within the sample and how the sample interacts with X-rays. While a higher X-ray dose is best for penetrating denser samples, it is less efficient in capturing variations in density within and between the sample and its surrounding medium7. On the other hand, a lower X-ray dose offers less penetration power and often requires longer scanning times but more sensitivity in density detection7.
These restrictions have long hampered the use of microtomography for plant sciences, given that most plant tissues are composed of light (non-dense) tissue with low X-ray absorption8. The first applications of micro-CT were focused on mapping root networks within the soil matrix9,10. Later, plant structures with more significant differences in tissue density, such as wood, began to be explored. This has allowed investigations of xylem functionality11,12, development of complex tissue organizations13,14, and interactions among plants15,16,17. The analysis of soft and homogeneous tissue is becoming widespread due to contrast agents, which are now standard procedure&#160;in preparations for micro-CT scanning of plant samples. However, protocols for contrast introduction can have different results depending on sample volume, structural properties, and the type of solution used8. Ideally, the contrast agent should enhance distinction among different tissues, enable tissue/organ functionality evaluation, and/or provide biochemical information about a tissue18. Therefore, adequate sample treatment, preparation, and mounting before scanning become crucial for any micro-CT analysis.
Micro-CT of the parasitic plant haustorium 
Parasitic flowering plants represent a distinct functional group of angiosperms characterized by an organ known as haustorium19. This multicellular organ, a developmental hybrid between a modified stem and a root, acts on the host&#39;s attachment, penetration, and contact by a parasite20. For this reason, the haustorium is considered to &#34;embody the very idea of parasitism among plants&#34;21. A detailed understanding of this organ&#39;s development, structure, and functioning is crucial for parasitic plant ecology, evolution, and management studies. Nevertheless, parasitic plants&#39; overall complexity and highly modified structure and haustoria often hinder detailed analysis and comparison. Haustorium connections are also usually extensive and not homogenous in tissue and cell distribution (Figure 1). In this context, while working with small tissue fragments allows easier manipulation and higher resolution, it can lead to erroneous conclusions about the three-dimensional architecture of complex structures, such as the parasitic plant haustorium.
Although there is a vast literature on haustorium anatomy and ultrastructure for most parasitic plant species, the three-dimensional organization and the spatial relationship between parasite and host tissues remains poorly explored17. In a recent work by Masumoto et al.22, over 300 serial semi-thin microtome sections were imaged and reconstructed into a three-dimensional virtual object representing the haustorium of two parasite species. This method&#39;s excellent level of detail provides unprecedented insights into the haustorium&#39;s cellular and anatomical 3-D structure. However, such a time-consuming technique would forbid a similar analysis in parasites with more extensive haustorium connections. The use of micro-CT emerges as an excellent tool for three-dimensional analysis of complex and often bulky haustoria of parasitic plants. Although not a substitute for detailed anatomical sectioning and other complementary forms of microscopy analyses17,23, results obtained via micro-CT scanning, especially for large samples, can also serve as a guide to direct the sub-sampling of smaller segments, which can then be analyzed using other tools, such as confocal and electron microscopy, or re-analyzed with high-resolution micro-CT systems.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63423/63423fig01.jpg" /> 
Figure 1: Parasitic plants of different functional groups used in this protocol. Euphytoid parasite Pyrularia pubera (A), endoparasite Viscum minimum (B) with green fruits (dashed black circle), parasitic vine Cuscuta americana (C), mistletoe Struthanthus martianus (D), obligate root parasite Scybalium fungiforme (E). Segments of the host root (Hr) or stem (Hs) facilitate the application of contrast into the parasite haustorium (P). The presence of parasite mother root/stem (arrows) in the sample allows analysis of haustorium vessel organization. Rectangles indicate segments of the sample used for analysis. Scale bars = 2 cm. <a href="https://www.jove.com/files/ftp_upload/63423/63423fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
As micro-CT becomes an increasingly popular technique in plant sciences, there are guides, protocols, and literature dealing with sample scanning, three-dimensional reconstruction, segmentation, and analysis3,10,24. Thus, these steps will not be discussed here. As with any imagining technique, appropriate treatment and mounting of samples are a fundamental, albeit often being an overlooked procedure. For this reason, this protocol focuses on the preparation of haustorium samples for micro-CT scanning. More specifically, this protocol describes two approaches for introducing contrast agents into haustorium samples to improve visualization of different tissues and cell types in the haustorium, to facilitate the detection of parasitic tissue within the host root/stem, and&#160;to analyze parasite-host vascular connections in three dimensions. The preparations described here can also be adapted to the analysis of other plant structures.
Five species were used to better illustrate the convenience of the protocol described here. Each species represents one of the five functional groups of parasitic flowering plants, thus addressing specific points related to the functionality of each group. Pyrularia pubera (Santalaceae) was chosen to represent euphytoid parasites, which germinate in the ground and form multiple haustoria that connect the parasite to the roots of its hosts25. The haustoria created by these plants are often tenuous and easily torn apart from the host26 (Figure 1A), thus requiring a more delicate handling process. Endoparasites are represented here by Viscum minimum (Viscaceae). Species in this functional group are only visible outside the body of their hosts for short periods (Figure 1B) and live most of their life cycles as significantly reduced and mycelial-like strands of cells embedded within host tissues25. A third functional group comprises parasitic vines, which germinate on the ground but form only rudimentary roots, relying on multiple haustoria that attach to the stems of host plants25 (Figure 1C). Here, this functional group is represented by Cuscuta americana (Convolvulaceae). Contrary to parasitic vines, mistletoes germinate directly upon the branches of their host plants and develop either multiple or solitary haustoria25. The species chosen to illustrate this functional group is Struthanthus martianus (Loranthaceae), which forms various connections with the host branch (Figure 1D). Analysis of solitary mistletoe haustoria using a combination of micro-CT and light microscopy can be found in Teixeira-Costa &#38; Ceccantini17. Finally, obligate root parasites comprise species that germinate on the ground and penetrate the roots of host plants, upon which they are entirely dependent from the earliest growth stages25. These plants are represented here by Scybalium fungiforme (Balanophoraceae), which produce large tuber-like haustoria (Figure 1E).
All plant samples used in this protocol were fixed in a 70% formalin acetic acid alcohol (FAA 70). The fixation upon sampling is crucial for preserving plant tissues, especially if subsequent anatomical analyses are needed. In the case of parasitic plant haustorium, fixation is also essential, as this organ is often primarily composed of non-lignified parenchyma cells20. Detailed protocols for plant tissue fixation, including the preparation of fixative solutions, can be found elsewhere27. On the other hand, to a greater or lesser degree, fixatives can cause alterations of a sample&#39;s physical and chemical properties, rendering it unsuitable for specific biomechanical and histochemical analyses. Thus, fresh samples, i.e., non-fixated material collected immediately before preparation, can also be used with this protocol. Details on how to handle fresh samples and troubleshooting suggestions for fixated material are provided in the discussion section.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>3D X-ray microscope (XRM) system</td>
			<td></td>
			<td>Zeiss Versa 620</td>
			<td>used to scan Pyrularia pubera</td>
		</tr>
		<tr>
			<td>3D X-ray microscope + A2:D22</td>
			<td>Zeiss</td>
			<td>Versa 620</td>
			<td>Used for scanning the species P. pubera</td>
		</tr>
		<tr>
			<td>CT-Pro 3D software</td>
			<td>Nikon</td>
			<td>version XT 3.1.11</td>
			<td>Used for three-dimensional reconstruction of scans</td>
		</tr>
		<tr>
			<td>CT-Vox software</td>
			<td>Bruker</td>
			<td>version 3.3.1</td>
			<td>Used for analyses and acquisition of images and videos</td>
		</tr>
		<tr>
			<td>Dragonfly software</td>
			<td>Object Research Systems - ORS</td>
			<td>version</td>
			<td>Used for analyses and acquisition of images and videos</td>
		</tr>
		<tr>
			<td>Glass vials</td>
			<td>Glass Vials Inc. SE</td>
			<td>V2708C-FM-SP</td>
			<td>Sold by VWR - USA; make sure that vials are able to withstand vacuum at ca. 10 psi</td>
		</tr>
		<tr>
			<td>Inspect-X</td>
			<td>Zeiss</td>
			<td>version XT 3.1.11</td>
			<td>Used for controlling the Nikon X-Tek HMXST225 system</td>
		</tr>
		<tr>
			<td>Iodine solution 0.0282 N</td>
			<td>WR Chemicals BDH</td>
			<td>BDH7422-1</td>
			<td>Sold by VWR - USA</td>
		</tr>
		<tr>
			<td>Lead Nitrate II PA 500 g</td>
			<td>Vetec</td>
			<td>361.08</td>
			<td>Sold by SPLab</td>
		</tr>
		<tr>
			<td>Microtomography scanner</td>
			<td>Bruker</td>
			<td>Skyscan1176</td>
			<td>Used for scanning the species C. americana, S. martianus, and S. fungiforme</td>
		</tr>
		<tr>
			<td>Microtomography scanner</td>
			<td>Nikon</td>
			<td>X-Tek HMXST225</td>
			<td>Used for scanning the species V. minimum</td>
		</tr>
		<tr>
			<td>NRecon software</td>
			<td>Bruker</td>
			<td>version 1.0.0</td>
			<td>Used for three-dimensional reconstruction</td>
		</tr>
		<tr>
			<td>Phosphotungstic acid hydrate 3% in aqueous solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>101410-756</td>
			<td>Sold by VWR - USA</td>
		</tr>
		<tr>
			<td>Plastic film (Parafilm)</td>
			<td>Heathrow Scientific</td>
			<td>PM996</td>
			<td>Sold by VWR - USA</td>
		</tr>
		<tr>
			<td>Plastic IV bag 500 mL</td>
			<td>Taylor</td>
			<td>3478</td>
			<td>Sold by Fibra Cirurgica Produtos para Saude</td>
		</tr>
		<tr>
			<td>PVC tubing 3/4&#39;&#39;</td>
			<td>Nalge Nunc International</td>
			<td>SC63013-164</td>
			<td>Sold by VWR - USA</td>
		</tr>
		<tr>
			<td>Scanning system</td>
			<td></td>
			<td>Nikon X-Tek HMXST225</td>
			<td>used to scan Viscum minimum</td>
		</tr>
		<tr>
			<td>Scanning system</td>
			<td></td>
			<td>Bruker Skyscan 1176</td>
			<td>used to scan C. americana</td>
		</tr>
		<tr>
			<td>Scout-and-ScanTM software</td>
			<td>Zeiss</td>
			<td>version 16</td>
			<td>Used for controlling the Zeiss Versa 620 system and for three-dimensional reconstruction of scans</td>
		</tr>
		<tr>
			<td>Three-way valve</td>
			<td>ToToT</td>
			<td>DMTWVS-5</td>
			<td>Sold by Amazon USA</td>
		</tr>
		<tr>
			<td>Two-part syringe</td>
			<td>HSW Henke-Ject</td>
			<td>4850001000</td>
			<td>Used without the plunger</td>
		</tr>
		<tr>
			<td>Vacuum chamber</td>
			<td>Binder</td>
			<td>80080-434</td>
			<td>Sold by VWR - USA; includes pump and connecting tubes</td>
		</tr>
		<tr>
			<td>VG Studio Max software</td>
			<td>Volume Graphics</td>
			<td>version 3.0</td>
			<td>Used for analyses and acquisition of images and videos</td>
		</tr>
	</tbody>
</table>

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.

The haustorium of parasitic plants is a complex organ comprising different tissues and cell types that intertwine and connect with the tissues of another plant, used as a host20. Micro-CT scanning can be leveraged to better understand this complex structure in a non-destructive and three-dimensional way when analyzing both small (Figure 1A-C) and large (Figure 1D,E) haustoria. To do ...

Watch this Scientific Journal Video about Leveraging Micro-CT Scanning to Analyze Parasitic Plant-Host Interactions at JoVE.com

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

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 ...

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1.8K Views.  Harvard University Herbaria. 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. 

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

A Technique to Screen American Beech for Resistance to the Beech Scale Insect (Cryptococcus fagisuga Lind.)

Beech bark disease (BBD) results in high levels of initial mortality, leaving behind survivor trees that are greatly weakened and deformed.&#160;The disease is initiated by feeding activities of the invasive beech scale insect, Cryptococcus fagisuga, which creates entry points for infection by one of the Neonectria species of fungus.&#160;Without scale infestation, there is little opportunity for fungal infection. Using scale eggs to artificially infest healthy trees in heavily BBD impacted stands demonstrated that these trees were resistant to the scale insect portion of the disease complex1.&#160;Here we present a protocol that we have developed, based on the artificial infestation technique by Houston2, which can be used to screen for scale-resistant trees in the field and in smaller potted seedlings and grafts.&#160;The identification of scale-resistant trees is an important component of management of BBD through tree improvement programs and silvicultural manipulation.

A Technique to Screen American Beech for Resistance to the Beech Scale Insect Cryptococcus fagisuga Lind.

Beech bark disease is initiated by feeding activities of the beech scale insect that create fungal entry points in the bark.&#160;Trees that are resistant to the scale insect are also disease resistant.&#160;Here we present the protocol we have developed to screen individual beech trees for beech scale resistance.

Beech bark disease (BBD) results in high levels of initial mortality, leaving behind survivor trees that are greatly weakened and deformed.&#160;The ...

RNAi-mediated Control of Aflatoxins in Peanut: Method to Analyze Mycotoxin Production and Transgene Expression in the Peanut/Aspergillus Pathosystem

The Food and Agriculture Organization of the United Nations estimates that 25% of the food crops in the world are contaminated with aflatoxins. That represents 100 million tons of food being destroyed or diverted to non-human consumption each year. Aflatoxins are powerful carcinogens normally accumulated by the fungi Aspergillus flavus and A. parasiticus in cereals, nuts, root crops and other agricultural products. Silencing of five aflatoxin-synthesis genes by RNA interference (RNAi) in peanut plants was used to control aflatoxin accumulation following inoculation with A. flavus. Previously, no method existed to analyze the effectiveness of RNAi in individual peanut transgenic events, as these usually produce few seeds, and traditional methods of large field experiments under aflatoxin-conducive conditions were not an option. In the field, the probability of finding naturally contaminated seeds is often 1/100 to 1/1,000. In addition, aflatoxin contamination is not uniformly distributed. Our method uses few seeds per transgenic event, with small pieces processed for real-time PCR (RT-PCR) or small RNA sequencing, and for analysis of aflatoxin accumulation by ultra-performance liquid chromatography (UPLC). RNAi-expressing peanut lines 288-72 and 288-74, showed up to 100% reduction (p&#8804;0.01) in aflatoxin B1 and B2 compared to the control that accumulated up to 14,000 ng.g-1 of aflatoxin B1 when inoculated with aflatoxigenic A. flavus. As reference, the maximum total of aflatoxins allowable for human consumption in the United States is 20 ng.g-1. This protocol describes the application of RNAi-mediated control of aflatoxins in transgenic peanut seeds and methods for its evaluation. We believe that its application in breeding of peanut and other crops will bring rapid advancement in this important area of science, medicine and human nutrition, and will significantly contribute to the international effort to control aflatoxins, and potentially other mycotoxins in major food crops.

RNAi-mediated Control of Aflatoxins in Peanut: Method to Analyze Mycotoxin Production and Transgene Expression in the Peanut/Aspergillus Pathosystem

We demonstrate a method for the analysis of aflatoxins and transgene expression in peanut seeds that contain RNA-interference signals for silencing aflatoxin-synthesis genes in the fungus Aspergillus flavus. RNAi-mediated control of mycotoxins in plants has not been reported previously.

The Food and Agriculture Organization of the United Nations estimates that 25% of the food crops in the world are contaminated with aflatoxins. That ...

Experimental Column Setup for Studying Anaerobic Biogeochemical Interactions Between Iron (Oxy)Hydroxides, Trace Elements, and Bacteria

Fate and speciation of trace elements (TEs), such as arsenic (As) and mercury (Hg), in aquifers are closely related to physio-chemical conditions, such as redox potential (Eh) and pH, but also to microbial activities that can play a direct or indirect role on speciation and/or mobility. Indeed, some bacteria can directly oxidize As(III) to As(V) or reduce As(V) to As(III). Likewise, bacteria are strongly involved in Hg cycling, either through its methylation, forming the neurotoxin monomethyl mercury, or through its reduction to elemental Hg&#176;. The fates of both As and Hg are also strongly linked to soil or aquifer composition; indeed, As and Hg can bind to organic compounds or (oxy)hydroxides, which will influence their mobility. In turn, bacterial activities such as iron (oxy)hydroxide reduction or organic matter mineralization can indirectly influence As and Hg sequestration. The presence of sulfate/sulfide can also strongly impact these particular elements through the formation of complexes such as thio-arsenates with As or metacinnabar with Hg.
Consequently, many important questions have been raised on the fate and speciation of As and Hg in the environment and how to limit their toxicity. However, due to their reactivity towards aquifer components, it is difficult to clearly dissociate the biogeochemical processes that occur and their different impacts on the fate of these TE.
To do so, we developed an original, experimental, column setup that mimics an aquifer with As- or Hg-iron-oxide rich areas versus iron depleted areas, enabling a better understanding of TE biogeochemistry in anoxic conditions. The following protocol gives step by step instructions for the column set-up either for As or Hg, as well as an example with As under iron and sulfate reducing conditions.

Experimental Column Setup for Studying Anaerobic Biogeochemical Interactions Between Iron OxyHydroxides, Trace Elements, and Bacteria

Fate and speciation of arsenic and mercury in aquifers are closely related to physio-chemical conditions and microbial activity. Here, we present an original experimental column setup that mimics an aquifer and enables a better understanding of trace element biogeochemistry in anoxic conditions. Two examples are presented, combining geochemical and microbiological approaches.

Fate and speciation of trace elements (TEs), such as arsenic (As) and mercury (Hg), in aquifers are closely related to physio-chemical conditions, such as ...

Thermal Scanning Conductometry (TSC) as a General Method for Studying and Controlling the Phase Behavior of Conductive Physical Gels

The thermal scanning conductometry protocol is a new approach in studying ionic gels based on low molecular weight gelators. The method is designed to follow the dynamically changing state of the ionogels, and to deliver more information and details about the subtle change of conductive properties with an increase or decrease in the temperature. Moreover, the method allows the performance of long term (i.e. days, weeks) measurements at a constant temperature to investigate the stability and durability of the system and the aging effects. The main advantage of the TSC method over classical conductometry is the ability to perform measurements during the gelation process, which was impossible with the classical method due to temperature stabilization, which usually takes a long time before the individual measurement. It is a well-known fact that to obtain the physical gel phase, the cooling stage must be fast; moreover, depending on the cooling rate, different microstructures can be achieved. The TSC method can be performed with any cooling/heating rate that can be assured by the external temperature system. In our case, we can achieve linear temperature change rates between 0.1 and approximately 10 &#176;C/min. The thermal scanning conductometry is designed to work in cycles, continuously changing between heating and cooling stages. Such an approach allows study of the reproducibility of the thermally reversible gel-sol phase transition. Moreover, it allows the performance of different experimental protocols on the same sample, which can be refreshed to initial state (if necessary) without removal from the measuring cell. Therefore, the measurements can be performed faster, in a more efficient way, and with much higher reproducibility and accuracy. Additionally, the TSC method can be also used as a tool to manufacture the ionogels with targeted properties, like microstructure, with an instant characterization of conductive properties.

Thermal Scanning Conductometry TSC as a General Method for Studying and Controlling the Phase Behavior of Conductive Physical Gels

The kinetics of the cooling process defines the properties of ionic gels based on low molecular weight gelators. This manuscript describes the usage of thermal scanning conductometry (TSC), which obtains full control over the gelation process, along with in situ measurements of the samples' temperature and conductivity.

The thermal scanning conductometry protocol is a new approach in studying ionic gels based on low molecular weight gelators. The method is designed to ...

Ecosystem Fabrication (EcoFAB) Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A better mechanistic understanding of these complex plant-microbe interactions will be crucial to improving plant production as well as performing basic ecological studies investigating plant-soil-microbe interactions. Here, a detailed description for ecosystem fabrication is presented, using widely available 3D printing technologies, to create controlled laboratory habitats (EcoFABs) for mechanistic studies of plant-microbe interactions within specific environmental conditions. Two sizes of EcoFABs are described that are suited for the investigation of microbial interactions with various plant species, including Arabidopsis thaliana, Brachypodium distachyon, and Panicum virgatum. These flow-through devices allow for controlled manipulation and sampling of root microbiomes, root chemistry as well as imaging of root morphology and microbial localization. This protocol includes the details for maintaining sterile conditions inside EcoFABs and mounting independent LED light systems onto EcoFABs. Detailed methods for addition of different forms of media, including soils, sand, and liquid growth media coupled to the characterization of these systems using imaging and metabolomics are described. Together, these systems enable dynamic and detailed investigation of plant and plant-microbial consortia including the manipulation of microbiome composition (including mutants), the monitoring of plant growth, root morphology, exudate composition, and microbial localization under controlled environmental conditions. We anticipate that these detailed protocols will serve as an important starting point for other researchers, ideally helping create standardized experimental systems for investigating plant-microbe interactions.

Ecosystem Fabrication EcoFAB Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

This article describes detailed protocols for ecosystem fabrication of devices (EcoFABs) that enable the studies of plants and plant-microbe interactions in highly controlled laboratory conditions.

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A ...

An Optimized Rhizobox Protocol to Visualize Root Growth and Responsiveness to Localized Nutrients

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive harvest or expensive equipment. We present a customizable and affordable rhizobox method that allows the non-destructive visualization of root growth over time and is particularly well-suited to studying root plasticity in response to localized resource patches. The method was validated by assessing maize genotypic variation in plasticity responses to patches containing 15N-labeled legume residue. Methods are described to obtain representative developmental measurements over time, measure root length density in resource-containing and control patches, calculate root growth rates, and determine 15N recovery by plant roots and shoots. Advantages, caveats, and potential future applications of the method are also discussed. Although care must be taken to ensure that experimental conditions do not bias root growth data, the rhizobox protocol presented here yields reliable results if carried out with sufficient attention to detail.

Visualizing and measuring root growth in situ is extremely challenging. We present a customizable rhizobox method to track root development and proliferation over time in response to nutrient enrichment. This method is used to analyze maize genotypic differences in root plasticity in response to an organic nitrogen source.

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive ...

Composition and Distribution Analysis of Bioaerosols Under Different Environmental Conditions

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, we described a protocol for multiple analyses of the biological compositions in environmental PM. Five experiments are presented: (1) PM number monitoring by using a laser particle counter; (2) PM collection by using a cyclonic aerosol sampler; (3) PM collection by using a high-volume air sampler with filters; (4) culturable microbes collection by the Andersen six-stage sampler; and (5) detection of biological composition of environmental PM by bacterial 16SrDNA and fungal ITS region sequencing. We selected hazy days and a livestock farm as two typical examples of the application in this protocol. In this study, these two sampling methods, cyclonic aerosol sampler and filter sampler, showed different sampling efficiency. The cyclonic aerosol sampler performed much better in terms of collecting bacteria, while these two methods showed the same efficiency in collecting fungi. Filter samplers can work under low temperature conditions while cyclonic aerosol samplers have a sampling limitation for temperature. A solid impacting sampler, such as an Andersen six-stage sampler, can be used to sample bioaerosols directly into the culture medium, which increases the survival rate of culturable microorganisms. However, this method mainly relies on culture while more than 99% of microbes cannot be cultured. DNA extracted from the culturable bacteria collected by the Andersen six-stage sampler and samples collected by cyclonic aerosol sampler and filter sampler were detected by bacterial 16S rDNA and fungal ITS region sequencing.All the methods above may have wide application in many fields of study, such as environmental monitoring and airborne pathogen detection. From these results, we can conclude that these methods can be used under different conditions and may help other researchers further explore the health impacts of environmental bioaerosols.

Understanding the biological composition of environmental particulate matter is important for the study of its significant impacts on human health and disease spread. Here, we used three types of bioaerosol sampling methods and a biological analysis of airborne microbes to better explore airborne microbial communities under different environmental conditions.

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, ...

Combined Size and Density Fractionation of Soils for Investigations of Organo-Mineral Interactions

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF relies on sequential density separation and sedimentation steps to isolate (1) the free light fraction (uncomplexed organic matter), (2) the occluded light fraction (uncomplexed organic matter trapped in soil aggregates) and (3) a variable number of heavy fractions (soil minerals and their associated organic matter) differing in composition. Provided that the parameters of the CSDF (dispersion energy, density cut-offs, sedimentation time) are properly selected, the method yields heavy fractions of relatively homogeneous mineral composition. Each of these fractions is expected to have a different complexing ability towards organic matter, rendering this a useful method to isolate and study the nature of organo-mineral interactions. Combining density and particle size separation brings an improved resolution compared to simple size or density fractionation methods, allowing the separation of heavy components according to both mineralogy and size (related to surface area) criteria. As is the case for all physical fractionation methods, it may be considered as less disruptive or aggressive than chemically-based extraction methods. However, CSDF is a time-consuming method and furthermore, the quantity of material obtained in some fractions can be limiting for subsequent analysis. Following CSDF, the fractions may be analyzed for mineralogical composition, soil organic carbon concentration and organic matter chemistry. The method provides quantitative information about organic carbon distribution within a soil sample and brings light to the sorptive capacity of the different, naturally-occurring mineral phases, thus providing mechanistic information about the preferential nature of organo-mineral interactions in soils (i.e., which minerals, what type of organic matter).

Combined size and density fractionation (CSDF) is a method to physically separate soil into fractions differing in texture (particle size) and mineralogy (density). The purpose is to isolate fractions with different reactivities towards soil organic matter (SOM), in order to better understand organo-mineral interactions and SOM dynamics.

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF ...

Xylem Water Distribution in Woody Plants Visualized with a Cryo-scanning Electron Microscope

A scanning electron microscope installed cryo-unit (cryo-SEM) allows specimen observation at subzero temperatures and has been used for exploring water distribution in plant tissues in combination with freeze fixation techniques using liquid nitrogen (LN2). For woody species, however, preparations for observing the xylem transverse-cut surface involve some difficulties due to the orientation of wood fibers. Additionally, higher tension in the water column in xylem conduits can occasionally cause artifactual changes in water distribution, especially during sample fixation and collection. In this study, we demonstrate an efficient procedure to observe the water distribution within the xylem of woody plants in situ&#160;by using a cryostat and cryo-SEM. At first, during sample collection, measuring the xylem water potential should determine whether high tension is present in the xylem conduits. When the xylem water potential is low (&#60; ca. &#8722;0.5 MPa), a tension relaxation procedure is needed to facilitate better preservation of the water status in xylem conduits during sample freeze fixation. Next, a watertight collar is attached around the tree stem and filled with LN2&#160;for freeze fixation of the water status of xylem. After harvesting, care should be taken to ensure that the sample is preserved frozen while completing the procedures of sample preparation for observation. A cryostat is employed to clearly expose the xylem transverse-cut surface. In cryo-SEM observations, time adjustment for freeze-etching is required to remove frost dust and accentuate the edge of the cell walls on the viewing surface. Our results demonstrate the applicability of cryo-SEM techniques for the observation of water distribution within xylem at cellular and subcellular levels. The combination of cryo-SEM with non-destructive in situ observation techniques will profoundly improve the exploration of woody plant water flow dynamics.

Observing the water distribution within the xylem provides significant information regarding water flow dynamics in woody plants. In this study, we demonstrate the practical approach to observe xylem water distribution in situ&#160;by using a cryostat and cryo-SEM, which eliminates artifactual changes in the water status during sample preparation.

A scanning electron microscope installed cryo-unit (cryo-SEM) allows specimen observation at subzero temperatures and has been used for exploring water ...

Sample Preparation Method of Scanning and Transmission Electron Microscope for the Appendages of Woodboring Beetle

This report described sample preparation methods that scanning and transmission electron microscope observations, demonstrated by preparing appendages of the woodboring beetle, Chlorophorus caragana Xie &#38; Wang (2012), for both types of electron microscopy. The scanning electron microscopy (SEM) sample preparation protocol was based on sample chemical fixation, dehydration in a series of ethanol baths, drying, and sputter-coating. By adding Tween 20 (Polyoxyethylene sorbitan laurate) to the fixative and the wash solution, the insect body surface of woodboring beetle was washed more cleanly in SEM. This study&#39;s transmission electron microscopy (TEM) sample preparation involved a series of steps including fixation, ethanol dehydration, embedding in resin, positioning using fluorescence microscopy, sectioning, and staining. Fixative with Tween 20 enabled penetrate the insect body wall of woodboring beetle more easily than it would had been without Tween 20, and subsequently better fixed tissues and organs in the body, thus yielded clear transmission electron microscope observations of insect sensilla ultrastructures. The next step of this preparation was determining the positions of insect sensilla in the sample embedded in the resin block by using fluorescence microscopy to increase the precision of target sensilla positioning. This improved slicing accuracy.

In order to observe ultrastructure of insect sensilla, scanning and transmission electron microscopy (SEM and TEM, respectively) sample preparation protocol were presented in the study. Tween 20 was added into the fixative to avoid sample deformation in SEM. Fluorescence microscopy was helpful for improving slicing accuracy in TEM.

This report described sample preparation methods that scanning and transmission electron microscope observations, demonstrated by preparing appendages of ...