This project was financially supported by the National Natural Science Foundation of China (32122075, 32072523).

1. Preparation for aniline blue staining
<ol>
	<li>Prepare the following reagents and tools for the experiment: a pollinator brush, tweezers, a pencil, sulfate paper, a pollination bag, zip lock bags, paper clips, formaldehyde, glacial acetic acid, absolute ethanol, centrifuge tubes, forceps, glue droppers, glass slides, coverslips, scalpels, and polyethylene glycol.</li>
	<li>Prepare in vitro germination medium containing 0.02% MgSO4, 0.01% KNO3, 0.03% Ca(NO3)2, 0.01% H3BO3, 20% PEG-4000, and 15% sucrose. Adjust the pH to 6.0-6.2 with KOH. Use a magnetic stirrer as PEG-4,000 is difficult to dissolve.</li>
	<li>Prepare Carnoy&#39;s fixative solution, which is absolute ethanol and glacial acetic acid mixed at a ratio of 3:1. Prepare FAA fixation solution, which is 40% formaldehyde, 80% ethanol, and glacial acetic acid (1:8:1). Prepare 4 M sodium hydroxide (NaOH) and aniline blue solution, which is 0.1% aniline blue in 0.1 M K3PO4. Use an amber bottle to store the aniline blue solution because it is light-sensitive.</li>
</ol>
2. Pollen collection
<ol>
	<li>Know the flowering period of the experimental trees (Ma jia pummelo in this study) in advance. Collect mature unopened flowers from the beginning of the flowering period to the peak flowering stage, and put them in a zip lock bag. The flowers can be stored at 4 &#176;C in the refrigerator for 24 h.</li>
	<li>Take the flowers to the lab. Use tweezers to collect the anthers, and place them in a Petri dish containing filter paper. Collect anthers from 20 to 30 flowers.</li>
	<li>Place the Petri dish containing the anthers in a 28 &#176;C oven for 24 h until the pollen grains dry. For the detailed flower organization, see Hu et al.9.</li>
	<li>Put the dried pollen into a 1.5 mL centrifuge tube. Save the pollen in an air-tight zip lock bag containing color-changing silica gel (desiccant). Close the bag, and label the bag with the name of the pollen variety and the date of storage. The dried pollen can be stored in a &#8722;20 &#176;C refrigerator for 96 weeks16.</li>
</ol>
3. In vitro pollen germination test
<ol>
	<li>Pour 300 &#181;L of liquid medium into a cell culture dish or the cap of a 2 mL centrifuge tube, and sprinkle the pollen evenly with the help of a pollination brush. Incubate the pollen at 28 &#176;C in a humidified dark environment for 12 h.</li>
	<li>Remove the top 1 mm of the tip of a 1,000 &#181;L pipette tip. Use the pipette to absorb the pollen with a small amount of culture solution and move it to the center of the microscope slide. Cover it with a coverslip. Observe the specimen with an inverted microscope using a 10x objective.</li>
	<li>Perform three independent replicates using approximately the same pollen density for each replicate17. Visually manage the amount of pollen, making sure that whole of the Petri dish is covered with pollen and that each Petri dish has an almost equal amount of pollen.</li>
	<li>The germinated pollen produces a pollen tube with a length approximately twice its diameter. Calculate the germination rate from 20 visual fields, which gives the percentage of germinated pollen in all pollen fields.</li>
</ol>
4. Pollination
<ol>
	<li>Choose a sunny day without wind for pollination. Select 10 fully developed buds about to open, peel back the petals carefully, and take care not to bruise them.</li>
	<li>Use a pollination brush to spread a sufficient amount of viable pollen onto the surface of the stigma, and take care not to damage the pistil. For self-pollination, use the pollen from the same plant/cultivar. For cross-pollination, use the pollen from a plant with a different genotype.</li>
	<li>Cover the pollinated flowers with a sulfate paper bag. Use a paper clip to seal the bag and to prevent pollination by genotypically distinct pollens.</li>
	<li>Write the name of the species and the number and time of pollination on the label. Hang the label on the branches near the pollinated flowers.</li>
</ol>
5. Sampling, fixation, and preservation
<ol>
	<li>Remove the pollination bags approximately 3-4 days after pollination, and collect the pollinated flowers in zip lock bags.</li>
	<li>Immediately remove the petals, receptacles, and ovaries from the flowers, and immerse the stigmas fused to styles in a centrifuge tube containing a freshly prepared fixative solution. Incubate the stigmas and styles in the fixative solution overnight at 4 &#176;C.</li>
	<li>The next day, discard the fixative solution, and wash the stigmas and styles two to three times in 95% ethanol.</li>
	<li>Transfer the styles to a 70% ethanol solution. Ensure that the sample is completely immersed in the solution. The styles at this stage can be stored at 4 &#176;C for 1-2 months.</li>
</ol>
6. Aniline blue staining
<ol>
	<li>Wash the style samples stored in 70% ethanol with distilled water three to four times. Immerse in 4 M NaOH solution, seal, and incubate in a 65 &#176;C water bath for 60 min. During this step, the color of the style changes from yellow-white to orange-red.</li>
	<li>Soak the styles in distilled water for 30 min. Discard the distilled water, and wash the styles with distilled water three to four times or until the color of the style becomes yellow.</li>
	<li>Place the sample in a 10 mL tube, add the aniline blue until the sample has been dipped, and dye for 12 h in the dark.</li>
	<li>Observe the pollen tube growth with a fluorescence microscope.</li>
</ol>
7. Fluorescence microscopy
<ol>
	<li>Before observing the specimens, place the slide on a flat table, and add two to three drops of polyethylene glycol to the surface of the slide.</li>
	<li>Wash the style to be observed with distilled water. Use a scalpel to divide it into two halves along the longitudinal axis. Place one half of the style on the glass slide, and cover with a coverslip.</li>
	<li>Place the slide on the stage of the microscope above the aperture, and visualize using a 10x objective. Use the DAPI filter (excitation: 325-375 nm; emission: 435-485 nm). Observe five styles for each type of pollination. Observe the pollen tube growth.</li>
</ol>
8. PCR-based S genotype identification
<ol>
	<li>Extract the genomic DNA from the stigma sample using the CTAB method18.
	<ol>
		<li>Put the collected leaves into a 2 mL centrifuge tube, and snap-freeze in liquid nitrogen. Prepare HCl:EDTA:NaCl:H2O buffer at a ratio of 1:1:3:5 and a chloroform isoamyl alcohol mixture at a ratio of 24:1</li>
		<li>Add 10 mL of the prepared buffer, 0.2 g of CTAB, and 200 &#181;L of mercaptoethanol to a 50 mL centrifuge tube, and put them in a water bath at 65 &#176;C for 5 min until the solution becomes clear and transparent.</li>
		<li>Put the blades into a mortar, add the frozen samples, add liquid nitrogen, and grind. Put the ground sample into a 2 mL centrifuge tube, add 600 &#181;L of CTAB mixture, put it in a water bath at 65 &#176;C for 60 min, and mix it upside down every 30 min.</li>
		<li>Add 700 &#181;L of the chloroform isoamyl alcohol mixture (24:1), and mix upside down for 10 min. Centrifuge at 24 &#176;C at 12,000 x g for 10 min, pipette the supernatant, and transfer to a 1.5 mL centrifuge tube.</li>
		<li>Add 60 &#181;L of 5 M NaCl solution and 1 mL of absolute ethanol, and mix upside down. Freeze at &#8722;20 &#176;C for 30 min, and centrifuge at 24 &#176;C and 9,000 x g for 5 min.</li>
		<li>Discard the supernatant, add 1 mL of 70% ethanol solution, and leave at room temperature for 1-2 h. Centrifuge at 24 &#176;C, 9,000 x g for 5 min, discard the supernatant, aspirate the excess ethanol solution, and air-dry for 5 min.</li>
		<li>Add 100 &#181;L of sterile water to dissolve, measure the DNA concentration with a spectrophotometer, and freeze in a -4 &#176;C freezer for long-term storage.</li>
		<li>Configure the RT-PCR reaction system. Prepare the following reaction mix for 10 &#181;L containing 5 &#181;L of 2x PCRMix, 0.25 &#181;L each of forward and reverse primer, 1 &#181;L of DNA (100 ng/ &#181;L), and 3.5 &#181;L of H2O.</li>
	</ol>
	</li>
	<li>Set up the PCR program as per Table 1. The PCR program for all isoforms was 95 &#176;C for 5 min 32x (95 &#176;C for 30 s, 55 &#176;C for 30 s, and 72 &#176;C for 1 min) and 72 &#176;C for 5 min. Separate the products on 1.5% TAE-agarose gels and photograph9. Verify the specified S genotype using genomic DNA.</li>
</ol>

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The authors declare that they have nothing to disclose.

In fruit crops, both parthenocarpy and SI are important traits because they pave the way for seedless fruits - a trait that is highly appreciated by consumers. Self-incompatibility promotes the rejection of self-pollen and, thus, prevents inbreeding20. Among citrus, pummelo is a self-incompatible variety7. Almost 40% of all angiosperm species exhibit SI21. This trait prevents fruit setting, lowers the yield, and brings huge economic losses to growers...

Self-incompatibility (SI) is a genetically controlled mechanism present in approximately 40% of angiosperm species. In this process, the pistil rejects pollen from a plant with the same SI genotype and, thus, prevents self-fertilization1,2. Ma jia pummelo is a local variety in Jinagsu province, China, with the excellent qualities of large, pink fruit, a rich juice content, a sweet and sour taste, and a thick peel3. Although SI promotes outcrossing, it negatively impacts the yield and quality of fruits4 and necessitates suitable pollinizer trees with distinct SI genotypes for reliable fruit-setting rates and high yields. At present, there are two main types of SI, sporophytic self-incompatibility (SSI), represented by Brassicaceae, and gametophytic self-incompatibility (GSI), represented by Rosaceae, Papaveraceae, Rutaceae, and Solanaceae5,6,7,8.
Citrus is one of the most important fruit crops in the world. The S-RNase-based GSI system is found in many citrus accessions and negatively influences the fruit-setting rate9. In this system, SI is controlled by the S locus, a single polymorphic locus with two complex alleles that carry pistil S determinants and pollen S determinants7. The female determinant is the S ribonuclease (S-RNase), and the male determinant is the S locus F-box (SLF)7. The cells of the pistil secrete S-RNase proteins. The non-self S-RNases are recognized by the SLF proteins, which leads to the ubiquitination and degradation of the non-self S-RNases by the 26S proteasome pathway. In contrast, the self S-RNases are able to accumulate and inhibit pollen tube (PT) growth because they evade the SLF proteins and, therefore, are prevented from ubiquitinzation10,11,12,13.
Here, we report an in vivo technique that is useful for identifying S-genotypes and degrees of pollen compatibility and incompatibility. The protocol involves extracting total DNA from leaves and predicting the S genotype using S-specific primers. Moreover, aniline blue staining and fluorescence microscopy followed by hand pollination provide evidence for the degree of compatibility and incompatibility. The semi in vivo pollination procedure, which involves the manual pollination of flowers in the laboratory14,15,&#160;has also been adapted to assess the degrees of self-compatibility and incompatibility. However, we have also used field pollination followed by the bagging of flowers to avoid contamination from undesired pollen to allow the pollen tubes to develop in natural conditions. This protocol is simple and straightforward and provides the information necessary for the successful selection of suitable pollinizer trees.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>absolute ethanol</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10009218</td>
			<td></td>
		</tr>
		<tr>
			<td>Aniline blue</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Boric acid, H3BO3</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10004818</td>
			<td></td>
		</tr>
		<tr>
			<td>Brown bottle</td>
			<td>Labgic Technology Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium nitrate tetrahydrate, Ca(NO3 )2</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>80029062</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifugal tube</td>
			<td>Labgic Technology Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>centrifuge tubes</td>
			<td>Labgic Technology Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CTAB</td>
			<td>GEN-VIEW SCIENTIFIC INC</td>
			<td>57-09-0(CAS)</td>
			<td></td>
		</tr>
		<tr>
			<td>Dropping</td>
			<td>Jiangsu Songchang Medical Equipment Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid, EDTA</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td>10009617</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>LUXIANZI Biotechnology Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>formaldehyde</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10010018</td>
			<td></td>
		</tr>
		<tr>
			<td>Fully automatic sample fast grinder</td>
			<td>Shanghai Jingxin Industrial Development Co., Ltd</td>
			<td>Tissuelyser-96</td>
			<td></td>
		</tr>
		<tr>
			<td>glacial acetic acid</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10000218</td>
			<td></td>
		</tr>
		<tr>
			<td>Grinding Tube</td>
			<td>Shanghai Jingxin Industrial Development Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoamyl alcohol</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td>10003218</td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropyl alcohol</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td>80109218</td>
			<td></td>
		</tr>
		<tr>
			<td>label</td>
			<td>M&#38;G Chenguang Stationery Co., Ltd.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Leica DMi8</td>
			<td>Shanghai Leica Co.,Ltd</td>
			<td>21903797</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium sulfate heptahydrate, MgSO4</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10013018</td>
			<td></td>
		</tr>
		<tr>
			<td>MICROSCOPE Cover glass</td>
			<td>Zhejiang Shitai Industrial Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td>10019318</td>
			<td></td>
		</tr>
		<tr>
			<td>paper clips</td>
			<td>M&#38;G Chenguang Stationery Co., Ltd.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pencil</td>
			<td>M&#38;G Chenguang Stationery Co., Ltd.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>pollinator brush</td>
			<td>Shanghai Yimei Plastics Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene glycol, PEG 6000</td>
			<td>Beijing Dingguo Changsheng Biotechnology Co., Ltd</td>
			<td>DH229-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene glycol, PEG-4000</td>
			<td>Guangzhou saiguo biotech Co., Ltd</td>
			<td>1521GR500</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium hydroxide, KOH</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10017008</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium nitrate, KNO3</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10017218</td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>Jiangsu Songchang Medical Equipment Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Slide</td>
			<td>Zhejiang Shitai Industrial Co., Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide, NAOH</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td>10019718</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sinopharm Chemical ReagentCo., Ltd</td>
			<td>10021418</td>
			<td></td>
		</tr>
		<tr>
			<td>sulfate paper</td>
			<td>Taizhou Jinnong Mesh Factory</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Thermostat water bath</td>
			<td>Shanghai Jinghong Experimental Equipment Co., Ltd</td>
			<td>L-909193</td>
			<td></td>
		</tr>
		<tr>
			<td>Trichloromethane</td>
			<td>Sinopharm Chemical Reagent Co.,Ltd</td>
			<td>10006818</td>
			<td></td>
		</tr>
		<tr>
			<td>Tripotassium phosphate tribasic trihydrate, K3PO4</td>
			<td>Shanghai Lingfeng Chemical Reagent Co.,Ltd</td>
			<td>20032318</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-HCl</td>
			<td>GEN-VIEW SCIENTIFIC INC</td>
			<td>1185-53-1</td>
			<td></td>
		</tr>
		<tr>
			<td>zip lock bags</td>
			<td>M&#38;G Chenguang Stationery Co., Ltd.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#946;-Mercaptoethanol</td>
			<td>GEN-VIEW SCIENTIFIC INC</td>
			<td>60-24-2(CAS)</td>
			<td></td>
		</tr>
	</tbody>
</table>

determination of self-(in)compatibility and inter-(in)compatibility relationships in citrus using manual pollination, microscopy, and s-genotype analyses

Citrus uses S-RNase-based self-incompatibility to reject self-pollen and, therefore, requires nearby pollinizer trees for successful pollination and fertilization. However, identifying suitable varieties to serve as pollinizers is a time-consuming process. To solve this problem, we have developed a rapid method for identifying pollination-compatible citrus cultivars that utilizes agarose gel electrophoresis and aniline blue staining. Pollen compatibility is determined based on the identification of S genotypes by extracting total DNA and performing PCR-based genotyping assays with specific primers. Additionally, styles are collected 3-4 days after manual pollination, and aniline blue staining is performed. Finally, the growth status of the pollen tubes is observed with a fluorescence microscope. Pollen compatibility and incompatibility can be established by observing whether the pollen tube growth is normal or suppressed, respectively. Due to its simplicity and cost-effectiveness, this method is an effective tool for determining the pollen compatibility and incompatibility of different citrus varieties to establish incompatibility groups and incompatibility relationships between different cultivars. This method provides information essential for the successful selection of suitable pollinizer trees and, thus, facilitates the establishment of new orchards and the selection of appropriate parents for breeding programs.

For the experiments done here, mature flowers were selected, the anthers&#160;were collected, dried in an oven, and the pollen was germinated at 28&#176;C for 12 h. The pollen viability and germination rates were quantified as shown in Figure 1.
Citrus was manually pollinated, and the pollen compatibility and incompatibility were assessed using aniline blue staining and fluorescence microscopy. The compatible pollen could germinate on the surface of the stigma and...

Watch this Scientific Journal Video about Determination of Self-Incompatibility and Inter-Incompatibility Relationships in Citrus Using Manual Pollination, Microscopy, and S-Genotype Analyses at JoVE.

Determination of Self-Incompatibility and Inter-Incompatibility Relationships in Citrus Using Manual Pollination, Microscopy, and S-Genotype Analyses

This protocol provides a rapid method for determining pollen compatibility and incompatibility in citrus cultivars.

Determination of Self-(In)compatibility and Inter-(In)compatibility Relationships in Citrus Using Manual Pollination, Microscopy, and S-Genotype Analyses

Citrus uses S-RNase-based self-incompatibility to reject self-pollen and, therefore, requires nearby pollinizer trees for successful pollination and ...

determination-self-compatibility-inter-compatibility-relationships

Research

JoVE Journal

Biology

2.2K Views.  Huazhong Agricultural University. This protocol provides a rapid method for determining pollen compatibility and incompatibility in citrus cultivars.

Video: Determination of Self-Incompatibility and Inter-Incompatibility Relationships in Citrus Using Manual Pollination, Microscopy, and S-Genotype Analyses

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

Nano-fEM: Protein Localization Using Photo-activated Localization Microscopy and Electron Microscopy

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for protein localization, but subcellular context is often absent in fluorescence images. Immuno-electron microscopy, on the other hand, can localize proteins, but the technique is limited by a lack of compatible antibodies, poor preservation of morphology and because most antigens are not exposed to the specimen surface. Correlative approaches can acquire the fluorescence image from a whole cell first, either from immuno-fluorescence or genetically tagged proteins. The sample is then fixed and embedded for electron microscopy, and the images are correlated 1-3. However, the low-resolution fluorescence image and the lack of fiducial markers preclude the precise localization of proteins. 
Alternatively, fluorescence imaging can be done after preserving the specimen in plastic. In this approach, the block is sectioned, and fluorescence images and electron micrographs of the same section are correlated 4-7. However, the diffraction limit of light in the correlated image obscures the locations of individual molecules, and the fluorescence often extends beyond the boundary of the cell. 
Nano-resolution fluorescence electron microscopy (nano-fEM) is designed to localize proteins at nano-scale by imaging the same sections using photo-activated localization microscopy (PALM) and electron microscopy. PALM overcomes the diffraction limit by imaging individual fluorescent proteins and subsequently mapping the centroid of each fluorescent spot 8-10. 
We outline the nano-fEM technique in five steps. First, the sample is fixed and embedded using conditions that preserve the fluorescence of tagged proteins. Second, the resin blocks are sectioned into ultrathin segments (70-80 nm) that are mounted on a cover glass. Third, fluorescence is imaged in these sections using the Zeiss PALM microscope. Fourth, electron dense structures are imaged in these same sections using a scanning electron microscope. Fifth, the fluorescence and electron micrographs are aligned using gold particles as fiducial markers. In summary, the subcellular localization of fluorescently tagged proteins can be determined at nanometer resolution in approximately one week.

We describe a method to localize fluorescently tagged proteins in electron micrographs. Fluorescence is first localized using photo-activated localization microscopy on ultrathin sections. These images are then aligned to electron micrographs of the same section.

Mapping the distribution of proteins is essential for understanding the function of proteins in a cell. Fluorescence microscopy is extensively used for ...

Using High Resolution Computed Tomography to Visualize the Three Dimensional Structure and Function of Plant Vasculature

High resolution x-ray computed tomography (HRCT) is a non-destructive diagnostic imaging technique with sub-micron resolution capability that is now being used to evaluate the structure and function of plant xylem network in three dimensions (3D) (e.g. Brodersen et al. 2010; 2011; 2012a,b). HRCT imaging is based on the same principles as medical CT systems, but a high intensity synchrotron x-ray source results in higher spatial resolution and decreased image acquisition time. Here, we demonstrate in detail how synchrotron-based HRCT (performed at the Advanced Light Source-LBNL Berkeley, CA, USA) in combination with Avizo software (VSG Inc., Burlington, MA, USA) is being used to explore plant xylem in excised tissue and living plants. This new imaging tool allows users to move beyond traditional static, 2D light or electron micrographs and study samples using virtual serial sections in any plane. An infinite number of slices in any orientation can be made on the same sample, a feature that is physically impossible using traditional microscopy methods.
Results demonstrate that HRCT can be applied to both herbaceous and woody plant species, and a range of plant organs (i.e. leaves, petioles, stems, trunks, roots). Figures presented here help demonstrate both a range of representative plant vascular anatomy and the type of detail extracted from HRCT datasets, including scans for coast redwood (Sequoia sempervirens), walnut (Juglans spp.), oak (Quercus spp.), and maple (Acer spp.) tree saplings to sunflowers (Helianthus annuus), grapevines (Vitis spp.), and ferns (Pteridium aquilinum and Woodwardia fimbriata). Excised and dried samples from woody species are easiest to scan and typically yield the best images. However, recent improvements (i.e. more rapid scans and sample stabilization) have made it possible to use this visualization technique on green tissues (e.g. petioles) and in living plants. On occasion some shrinkage of hydrated green plant tissues will cause images to blur and methods to avoid these issues are described. These recent advances with HRCT provide promising new insights into plant vascular function.

High resolution x-ray computed tomography (HRCT) is a non-destructive diagnostic imaging technique that can be used to study the structure and function of plant vasculature in 3D. We demonstrate how HRCT facilitates exploration of xylem networks across a wide range of plant tissues and species.

High resolution x-ray computed tomography (HRCT) is a non-destructive diagnostic imaging technique with sub-micron resolution capability that is now being ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Determination of Fatty Acid Oxidation and Lipogenesis in Mouse Primary Hepatocytes

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid oxidation, or alternatively, synthesize new lipid via de novo lipogenesis. The net sum of these pathways is dictated by a number of factors, which in certain disease states leads to fatty liver disease. Excess hepatic lipid accumulation is associated with whole body insulin resistance and coronary heart disease. Tools to study lipid metabolism in hepatocytes are useful to understand the role of hepatic lipid metabolism in certain metabolic disorders.
In the liver, hepatocytes regulate the breakdown and synthesis of fatty acids via &#946;-fatty oxidation and de novo lipogenesis, respectively. Quantifying metabolism in these pathways provides insight into hepatic lipid handling. Unlike in vitro quantification, using primary hepatocytes, making measurements in vivo is technically challenging and resource intensive. Hence, quantifying &#946;-fatty acid oxidation and de novo lipogenesis in cultured mouse hepatocytes provides a straight forward method to assess hepatocyte lipid handling. 
Here we describe a method for the isolation of primary mouse hepatocytes, and we demonstrate quantification of &#946;-fatty acid oxidation and de novo lipogenesis, using radiolabeled substrates.

De novo lipogenesis and &#946;-fatty acid oxidation constitute key metabolic pathways in hepatocyte, pathways that are perturbed in several metabolic disorders, including fatty liver disease. Here we demonstrate isolation of mouse primary hepatocytes and describe quantification of &#946;-fatty acid oxidation and lipogenesis.

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid ...

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a versatile analytical tool for characterizing viscoelastic mechanical properties for a variety of materials ranging from hard (plastic, glass, metal, etc.) surfaces to cells on any substrate. It has been widely used to measure the stiffness of cells, but less frequently used to measure the stiffness of aortas. In this paper, we will describe the procedures for using AFM in contact mode to measure the ex vivo elastic modulus of unloaded mouse arteries. We describe our procedure for isolation of mouse aortas, and then provide detailed information for the AFM analysis. This includes step-by-step instructions for alignment of the laser beam, calibration of the spring constant and deflection sensitivity of the AFM probe, and acquisition of force curves. We also provide a detailed protocol for data analysis of the force curves.

Measuring the Stiffness of Ex Vivo Mouse Aortas Using Atomic Force Microscopy

We present detailed protocols for isolation of aortas from mouse and measurement of their elastic modulus using atomic force microscopy.

Arterial stiffening is a significant risk factor and biomarker for cardiovascular disease and a hallmark of aging. Atomic force microscopy (AFM) is a ...

Biological Sample Preparation by High-pressure Freezing, Microwave-assisted Contrast Enhancement, and Minimal Resin Embedding for Volume Imaging

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the imaging modality in a focused ion beam scanning electron microscope (FIB-SEM), which is used to obtain stacks of sequential images for 3D reconstruction and modelling. High-pressure freezing (HPF) allows close to native structural preservation, but the subsequent freeze substitution often does not provide sufficient contrast, especially for a bigger specimen, which is needed for high-quality imaging in the SEM required for 3D reconstruction. Therefore, in this protocol, after the freeze substitution, additional contrasting steps are carried out at room temperature. Although these steps are performed in a microwave, it is also possible to follow traditional bench processing, which requires longer incubation times.The subsequent embedding in minimal amounts of resin allows for faster and more precise targeting and preparation inside the FIB-SEM. This protocol is especially useful for samples that require preparation by high-pressure freezing for a reliable ultrastructural preservation but do not gain enough contrast during the freeze substitution for volume imaging using FIB-SEM. In combination with the minimal resin embedding, this protocol provides an efficient workflow for the acquisition of high-quality volume data.

Here we present a protocol for combining two sample processing techniques, high-pressure freezing and microwave-assisted sample processing, followed by minimal resin embedding for acquiring data with a focused ion beam scanning electron microscope (FIB-SEM). This is demonstrated using a mouse tibial nerve sample and Caenorhabditis elegans.

The described sample preparation technique is designed to combine the best quality of ultrastructural preservation with the most suitable contrast for the ...

Quantitative Analysis of Aspergillus nidulans Growth Rate using Live Microscopy and Open-Source Software

It is well established that colony growth of filamentous fungi, mostly dependent on changes in hyphae/mycelia apical growth rate, is macroscopically estimated on solidified media by comparing colony size. However, to quantitatively measure the growth rate of genetically different fungal strains or strains under different environmental/growth conditions (pH, temperature, carbon and nitrogen sources, antibiotics, etc.) is challenging. Thus, the pursuit of complementary approaches to quantify growth kinetics becomes mandatory in order to better understand fungal cell growth. Furthermore, it is well-known that filamentous fungi, including Aspergillus spp., have distinct modes of growth and differentiation under sub-aerial conditions on solid media or submerged cultures. Here, we detail a quantitative microscopic method for analyzing growth kinetics of&#160;the model fungus Aspergillus nidulans, using live imaging in both submerged cultures and solid media. We capture images, analyze, and quantify growth rates of different fungal strains in a reproducible and reliable manner using an open source, free software for bio-images (e.g., Fiji), in a way that does not require any prior image analysis expertise from the user.

Quantitative Analysis of Aspergillus nidulans Growth Rate using Live Microscopy and Open-Source Software

We present a label-free live imaging protocol using transmitted light microscopy techniques to capture images, analyze and quantify growth kinetics of the filamentous fungus A. nidulans in both submerged cultures and solid media. This protocol can be used in conjunction with fluorescence microscopy.

It is well established that colony growth of filamentous fungi, mostly dependent on changes in hyphae/mycelia apical growth rate, is macroscopically ...