This work was supported by the National Key Research and Development Program of China (No. 2019YFC1200503), by the National Science Foundation of China (No. 32072385), and by Youth Innovation Promotion Association CAS (2021084).

1. SBPH maintenance
<ol>
	<li>Rear the viruliferous and RSV-free SBPH individuals in a glass incubator (65 x 200 mm) with 5-6 rice (Oryza sativa cv. Nipponbare) seedlings per glass chamber in the laboratory. Grow the rice plants at 25 &#176;C under a 16 h light / 8 h dark photoperiod. 
	NOTE: The viruliferous and RSV-free SBPH individuals were initially caught in Jiangsu Province, China.</li>
	<li>Detect RSV in SBPH by dot-enzyme-linked immunosorbent assay (dot-ELISA) with a rabbit RSV-specific polyclonal antibody (see Table of Materials) raised against the RSV ribonucleoproteins (RNPs). 
	NOTE: For ensuring high offspring infection efficiency, viruliferous females were maintained separately, and 15% of their offspring were randomly tested for RSV infection. The details of dot-ELISA are described in steps 1.3-1.7.</li>
	<li>Homogenize single SBPH in 20 &#181;L of coating buffer (0.05 M Na2CO3-NaHCO3, pH 9.5). Spot 3 &#181;L of each on nylon membrane (see Table of Materials), and then dry the membrane at room temperature (RT).</li>
	<li>Incubate the membrane with 15 mL of blocking buffer (PBS + 3% skim milk) for 30 min at room temperature.</li>
	<li>Incubate the membrane with diluted primary rabbit-antibodies against RSV (1:10000) in 15 mL of PBS for 1 h at RT and wash the membrane three times with PBS for 5 min incubation each time.</li>
	<li>Incubate the membrane with 1.5 &#181;L of horseradish peroxidase-conjugated goat anti-rabbit antibodies (see Table of Materials) in 15 mL of PBS and wash three times with PBS for 5 min incubation each time.</li>
	<li>Develop the immunoblots with Enhanced HRP-DAB Chromogenic Kit (see Table of Materials) according to the protocols provided by the manufacturer.</li>
</ol>
2. Preparation of feeding chamber and artificial diet
<ol>
	<li>Weigh 2 g of sucrose powder and dissolve it in 40 mL of ddH2O to prepare 5% sucrose aqueous solution as the artificial diet.</li>
	<li>Filter the solution through a 0.22 &#181;m filter (see Table of Materials)&#160;to remove bacterial contamination and impurities.</li>
	<li>Starve 200 3rd-5th SBPH larvae for 3-5 h before introducing them into the chamber. 
	NOTE: 200 SBPH are prepared for one chamber; more SBPH should be prepared for several chambers.</li>
	<li>Prepare the glass cylinders as feeding chambers (Figure 1A). Cover one open end of the chamber with a paraffin membrane (see Table of Materials) before introducing the experimental insects. 
	NOTE: Each cylinder is 15.0 cm long and 2.5 cm in diameter. These glass cylinders are custom-made according to the experimental requirement.</li>
	<li>Transfer insects into a glass cylinder.</li>
	<li>Cover the other end of the chamber with stretched paraffin membrane (specifically, Parafilm M). Then, add 200 &#181;L of artificial diet to it. Finally, cover the liquid with another layer of stretched paraffin membrane. 
	NOTE: The paraffin membrane is stretched to about double its original area.</li>
	<li>Cover the chamber with aluminum foil, but leave the end with the artificial diet device exposed to the light.</li>
</ol>
3. Collection of SBPH saliva
<ol>
	<li>Collect artificial diet from RSV-free and viruliferous SBPH separately at the end of 24 h period.</li>
	<li>Cool the cylinder at 4 &#176;C to immobilize the insects.</li>
	<li>Uncover the outer film and collect the artificial diet liquid using a sterile pipette into 1.5 mL sterile tubes. Keep the collected saliva at -80 &#176;C until analysis. 
	NOTE: The collected saliva could be stored at -80 &#176;C for 1 year.</li>
	<li>Rinse the inner membrane with 50 &#181;L of fresh artificial diet three times by pipetting softly, and collect the artificial washing diet as described in step 3.3. Place the new artificial diet on the inner membrane and keep a freshly stretched Paraffin membrane on top.</li>
	<li>Repeat steps 3.2 and 3.3 for 5 days to 2 weeks. 
	NOTE: Count the survival rate of artificial feeding SBPH and ensure sufficient supplement of fresh SBPH according to the survival rate.</li>
	<li>Filter the collected samples through a 0.22 &#181;m filter&#160;unit to remove microbes and other contaminants.</li>
</ol>
4. Concentration of the collected saliva
<ol>
	<li>Transfer the collected saliva samples in a 0.5 mL 10 kD centrifugal filter (see&#160;Table of Materials)&#160;and spin at 5,500 x g at 4 &#176;C for 20 min. Collect the supernatant and make the final volume to 100 &#181;L.</li>
	<li>Measure the concentration of the collected saliva using an appropriate UV-Vis spectrophotometer following steps 4.3-4.6.</li>
	<li>Turn on the spectrophotometer and wash the pedestals three times with ddH2O.</li>
	<li>Select the following options on the screen in proper order: Proteins | Protein A280 | Select Type | 1 Abs = 1 mg/mL. Then, check the checkbox Baseline Correction 340 nm.</li>
	<li>Load 2 &#181;L of 5% sucrose aqueous solution as blank, touch Blank at the bottom of the screen.</li>
	<li>After setting standards, load 2 &#181;L of the collected saliva for measurement. Read and record the protein concentration. 
	&#8203;NOTE: 1 mg of saliva proteins were finally collected in total at least.</li>
</ol>
5. Silver staining of saliva proteins
<ol>
	<li>Extract protein from the insect saliva samples using sample loading buffer (50 mM Tris-HCl pH 6.8, 10% glycerol, 2% SDS, 0.1% bromophenol blue, and 1% &#946;-mercaptoethanol). Then, fractionate it by 10% SDS-PAGE (see Table of Materials). Load the 5% sucroseaqueous solution treated in the same manner as a negative control.</li>
	<li>Load a 20 &#181;L aliquot of the sample onto an SDS-PAGE gel alongside a prestained marker. Run the gel for 15 min at 90 Volt, and then 50 min at 140 Volt.</li>
	<li>Fix the gel in 30% (vol/vol) ethanol, 10% (vol/vol) acetic acid for at least 30 min after electrophoresis.</li>
	<li>Rinse the gel twice with 20% (vol/vol) ethanol and water separately for 10 min each time.</li>
	<li>Sensitize the gel in 0.8 mM sodium thiosulfate for 1 min, and then rinse twice in water for 1 min each time.</li>
	<li>Immerse the gel in 12 mM silver nitrate for at least 1 h, and then dip it in deionized water for 10 s before transferring it to the developer solution.</li>
	<li>When the background of the gel is getting dark, immerse the gel in a stop solution (5% acetic acid) for at least 30 min to stop the reaction.</li>
	<li>Wash the gel twice with water for 30 min each time. Develop the image with the Detection System (see Table of Materials).</li>
</ol>
6. Protein detection by western blotting
<ol>
	<li>Detect the saliva mucin-like protein of SBPH (LssgMP) and RSV by western blots using specific antibodies, respectively.</li>
	<li>Treat insect saliva samples following step 5.1.</li>
	<li>Load a 20 &#181;L aliquot of the sample onto a 10% SDS-PAGE gel alongside a prestained marker and a 20 &#181;L RSV non-infected saliva sample as a negative control. Run the gel for 15 min at 90 Volt, and then for 50 min at 140 Volt.</li>
	<li>Mix 100 mL of 10x protein transfer buffer (wet) (see&#160;Table of Materials)&#160;with 900 mL of ddH2O to a work solution (1x), and then transfer proteins to a polyvinylidene difluoride membrane using protein transfer buffer (1x).</li>
	<li>Block the membrane in 5% skim milk with 0.01 M Tris-buffered saline with 0.05% Tween 20 (TBST) at room temperature (RT) for 1 h. 
	NOTE: In this protocol, mix 100 mL of 10x TBST (see&#160;Table of Materials)&#160;with 900 mL of ddH2O into the work solution.</li>
	<li>Incubate the membrane with primary rabbit antibodies against RSV or LssgMP (both 1:10000) diluted in TBST at RT for at least 2 h. 
	NOTE: The production of primary antibodies against RSV was mentioned above. A biotechnology company produced the rabbit anti-LssgMP polyclonal antibody against LssgMP peptide GIQFDSYSASDLTRC.</li>
	<li>Wash the membrane three times with TBST for 10 min of incubation each time.</li>
	<li>Incubate the membrane with horseradish peroxidase-conjugated goat anti-rabbit antibodies diluted in 1:10000 TBST.</li>
	<li>Develop the immunoblots with the enhanced chemiluminescence Western Blotting Detection System.</li>
</ol>
7. Detection of LssgMP expression pattern in SBPH
<ol>
	<li>Immobilize the insects at 4 &#176;C for 5 min.</li>
	<li>Wash the insects with 75% ethanol and ddH2O one by one, and then dissect the insects in pre-chilled TBS (0.01 M Tris-buffered saline).</li>
	<li>Dissect the insects from the abdomen while severing the forelegs of SBPH at the coxa-trochanter joint by forceps; wash the midgut and the salivary glands twice in TBS to remove any contamination from the hemolymph.</li>
	<li>Put five tissues into a 1.5 mL RNase-free tube to extract RNA. Consider each tube as one sample.</li>
	<li>Perform RNA extraction according to the manufacturer&#39;s protocols and Reverse-transcriptional PCR (RT-PCR) (see Table of Materials).</li>
	<li>Perform quantitative real-time PCR (qRT-PCR) to investigate the relative transcript expression levels of LssgMP in extracts of the whole body or various tissues of L. striatellus. 
	NOTE: The primer pairs used for gene amplification were LssgMP-q-F/LssgMP-q-R, SYBR Green-based qPCR was performed according to the manufacturer&#39;s protocol. The transcript level of L. striatellus translation elongation factor 2 (ef2) was quantified with primer pair ef2-q-F/ef2-q-R for the normalization of the cDNA templates. And the primer sequences are attached below:LssgMP-q-F: TCCGACCTCACCAGAGTTTACAG; LssgMP-q-R: GCTTCGTCCCAGGTACTGATTCC; ef2-q-F: GTCTCCACGGATGGGCTTT; ef2-q-R: ATCTTGAATTTCTCGGCATACATTT.</li>
</ol>

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The authors declare that they have no conflicts of interest.

Successful rearing of insects on artificial diets was first reported in 1962 when Mittler and Dadd described the Paraffin membrane technique to hold an artificial diet29,30. And this method has been explored in many aspects of insect biology and behavior, for example, nutrient supplement, dsRNA feeding, and virus acquisition. Based on the requirements of saliva analysis, 5% sucrose is used as the general artificial diet to collect saliva of SBPH in this study. Fo...

Rice stripe virus (RSV), a negative-stranded RNA virus in the genus Tenuivirus, causes severe diseases in rice production in East Asia1,2,3. Transmission of RSV from infected rice plants to healthy ones depends on insect vectors, mainly Laodelphax striatellus, which transmits RSV in a persistent-propagative manner. SBPH acquires the virus after feeding on RSV-infected plants. Once inside the insect, RSV infects the midgut epithelial cell one day after feeding and then passes through the midgut barrier to penetrate the hemolymph. Subsequently, RSV spreads into different tissues via the hemolymph and then propagates. After a latent period of about 10-14 days post-acquisition, the virus inside the salivary gland can be transmitted to the healthy host plants via the secreted saliva while SBPH sucks sap from the phloem4,5,6,7,8,9,10. An efficient feeding process and various factors in the saliva are essential for the spread of RSV from the insect to the host plant.
Insect saliva secreted by salivary glands is believed to mediate insects, viruses, and host plants.Hemipteran insects usually produce two types of saliva: gelling saliva and watery saliva11,12,13. Gelling saliva is mainly secreted into the apoplasm to sustain the movement of the stylet among host cells and is also related to overcoming plant resistance and immune responses14,15,16,17. At the probing stage of feeding, insects intermittently secrete gelling saliva that immediately gets oxidized to form a surface flange. Then, single or branched sheaths encase the stylet to reserve a tubular channel18,19,20. The surface flange on the epidermis is presumed to facilitate penetration of the stylet by serving as an anchor point, while the sheaths around the stylet may provide mechanical stability and lubrication16,21,22,23. Nlshp was identified as an essential protein for salivary sheath formation and successful feeding of brown planthopper (Nilaparvata lugens, BPH). Inhibition of the expression of the structural sheath protein (SHP) secreted by the aphid Acyrthosiphon pisum reduced its reproduction by disrupting feeding from host sieve tubes24. Moreover, in some insect species, gel saliva factors are supposed to trigger plant immune responses by forming so-called herbivore-associated molecular patterns (HAMPs). In N. lugens, NlMLP, a mucin-like protein related to sheath formation, induces plant defenses against feeding, including cell death, the expression of defense-related genes, and callose deposition 25,26. Also, some gel saliva factors in aphids have been proved to trigger plant defense responses via gene-to-gene interactions similar to pathogen-associated molecular patterns12,15,27.
For studying the saliva factors essential for insect feeding and/or pathogen transmission, it is necessary to analyze secreted saliva. Here, artificial feeding and collection methods to obtain sufficient amounts of saliva are described for further analysis. Using a medium containing only a single nutritional element, many saliva proteins were collected and analyzed by silver staining and western blotting. This method will be helpful in further research on factors in saliva that are essential for RSV transmission by SBPH.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10-KD centrifugal filter</td>
			<td>Merck Millipore</td>
			<td>R5PA83496</td>
			<td>For concentration</td>
		</tr>
		<tr>
			<td>10x Protein Transfer Buffer(wet)</td>
			<td>macGENE</td>
			<td>MP008</td>
			<td>Transfer buffer for western blotting</td>
		</tr>
		<tr>
			<td>10x TBST buffer</td>
			<td>Coolaber</td>
			<td>SL1328-500mL&#215;10</td>
			<td>Wash buffer for western blotting</td>
		</tr>
		<tr>
			<td>Azure c600 biosystems</td>
			<td>Azure Biosystems</td>
			<td>Azure c600</td>
			<td>Imaging system for western blotting and silver staining</td>
		</tr>
		<tr>
			<td>Color Prestained protein ladder</td>
			<td>GenStar</td>
			<td>M221-01</td>
			<td>Protein marker for western blotting</td>
		</tr>
		<tr>
			<td>ECL&#160;western blotting detection reagents</td>
			<td>GE Healthcare</td>
			<td>RPN2209</td>
			<td>Western blotting detection</td>
		</tr>
		<tr>
			<td>Enchanced HRP-DAB Chromogenic Kit</td>
			<td>TIANGEN</td>
			<td>#PA110</td>
			<td>Chromogenic reaction</td>
		</tr>
		<tr>
			<td>Horseradish peroxidase-conjugated goat anti-rabbit antibodies</td>
			<td>Sigma</td>
			<td>401393-2ML</td>
			<td>Polyclonal secondary antibody for western blotting</td>
		</tr>
		<tr>
			<td>Immobilon(R)-P Polyvinylidene difluoride membrane</td>
			<td>Merck Millipore</td>
			<td>IPVH00010</td>
			<td>Transfer membrane for western blotting</td>
		</tr>
		<tr>
			<td>iTaq Universal SYBR Green Supermix</td>
			<td>Bio-Rad</td>
			<td>1725125</td>
			<td>For quantitative real-time PCR (qRT-PCR)</td>
		</tr>
		<tr>
			<td>KIT,iSCRIPT cDNA SYNTHES</td>
			<td>Bio-Rad</td>
			<td>1708891</td>
			<td>For Reverse-transcriptional PCR (RT-PCR)</td>
		</tr>
		<tr>
			<td>Millex-GP Filter, 0.22 &#181;m</td>
			<td>Merck Millipore</td>
			<td>SLGP033RB</td>
			<td>For filtration</td>
		</tr>
		<tr>
			<td>Mini-PROTEAB TGX Gels</td>
			<td>Bio-Rad</td>
			<td>4561043</td>
			<td>For SDS-PAGE</td>
		</tr>
		<tr>
			<td>NanoDrop One</td>
			<td>Thermo Scientific</td>
			<td>ND-ONEC-W</td>
			<td>Detection of protein concentration</td>
		</tr>
		<tr>
			<td>Nylon membrane</td>
			<td>PALL</td>
			<td>T42754</td>
			<td>Membrane for dot-ELISA</td>
		</tr>
		<tr>
			<td>Parafilm M Membrane</td>
			<td>Sigma</td>
			<td>P7793-1EA</td>
			<td>Making artifical diet sandwichs</td>
		</tr>
		<tr>
			<td>Rabbit anti-LssgMP polyclonal antibody against LssgMP peptides</td>
			<td>Genstript</td>
			<td></td>
			<td>Rabbit primary anti-LssgMP polyclonal antibody for western blotting</td>
		</tr>
		<tr>
			<td>Rabbit anti-RSV polyclonal antibody</td>
			<td>Genstript</td>
			<td></td>
			<td>Rabbit primary anti-RSV polyclonal antibody for western blotting and dot-ELISA</td>
		</tr>
		<tr>
			<td>RNAprep pure Micro Kit</td>
			<td>TIANGEN</td>
			<td>DP420</td>
			<td>For RNA Extraction</td>
		</tr>
	</tbody>
</table>

two-layered membrane sandwich method for laodelphax striatellus saliva collection

Rice stripe virus (RSV), which causes significant economic loss of agriculture in East Asia, entirely depends on insect vectors for its effective transmission among host rice. Laodelphax striatellus (small brown planthopper, SBPH) is the primary insect vector that horizontally transmits RSV while sucking sap from the phloem. Saliva plays a significant role in insects' feeding behavior. A convenient method that will be useful for research on insects' saliva with piercing-sucking feeding behavior is described here. In this method, insects were allowed to feed on an artificial diet sandwiched between two stretched paraffin film layers. The diet containing the saliva was collected each day, filtered, and concentrated for further analysis. Finally, the quality of collected saliva was examined by protein staining and immunoblotting. This method was exemplified by detecting the presence of RSV and a mucin-like protein in the saliva of SBPH. These artificial feeding and saliva collection method will lay a foundation for further research on factors in insect saliva related to feeding behavior and virus transmission.

Schematics of artificial feeding installation and saliva collection 
Figure 1A depicts the glass cylinder (15 cm x 2.5 cm) used as a feeding chamber to collect the saliva. Firstly, the SBPH larvae were starved for several hours to improve the collection efficiency and then immobilized by chilling for 5 min. After the insects were transferred into the glass cylinder, both open ends of the chamber were covered with stretched Paraffin membrane. At one end, 200 &#181;L of 5...

Watch this Scientific Journal Video about Two-layered Membrane Sandwich Method for Laodelphax striatellus Saliva Collection at JoVE.com

Two-layered Membrane Sandwich Method for Laodelphax striatellus Saliva Collection

The present protocol describes a method to collect sufficient saliva from piercing-sucking insects using an artificial medium. This is a convenient method for collecting insect saliva and studying salivary function on insect feeding behavior and vector-borne virus transmission.

Two-layered Membrane Sandwich Method for Laodelphax striatellus Saliva Collection

Rice stripe virus (RSV), which causes significant economic loss of agriculture in East Asia, entirely depends on insect vectors for its effective ...

two-layered-membrane-sandwich-method-for-laodelphax-striatellus

Research

JoVE Journal

Biology

2.2K Views.  Chinese Academy of Sciences. The present protocol describes a method to collect sufficient saliva from piercing-sucking insects using an artificial medium. This is a convenient method for collecting insect saliva and studying salivary function on insect feeding behavior and vector-borne virus transmission.

Video: Two-layered Membrane Sandwich Method for Laodelphax striatellus Saliva Collection

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 ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

Use of a Robot for High-throughput Crystallization of Membrane Proteins in Lipidic Mesophases

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through macromolecular X-ray crystallography (MX). An essential ingredient of MX is a steady supply of ideally diffraction-quality crystals. The in meso or lipidic cubic phase (LCP) method for crystallizing membrane proteins is one of several methods available for crystallizing membrane proteins. It makes use of a bicontinuous mesophase in which to grow crystals. As a method, it has had some spectacular successes of late and has attracted much attention with many research groups now interested in using it. One of the challenges associated with the method is that the hosting mesophase is extremely viscous and sticky, reminiscent of a thick toothpaste. Thus, dispensing it manually in a reproducible manner in small volumes into crystallization wells requires skill, patience and a steady hand. A protocol for doing just that was developed in the Membrane Structural &amp; Functional Biology (MS&amp;FB) Group1-3. JoVE video articles describing the method are available1,4. 
The manual approach for setting up in meso trials has distinct advantages with specialty applications, such as crystal optimization and derivatization. It does however suffer from being a low throughput method. Here, we demonstrate a protocol for performing in meso crystallization trials robotically. A robot offers the advantages of speed, accuracy, precision, miniaturization and being able to work continuously for extended periods under what could be regarded as hostile conditions such as in the dark, in a reducing atmosphere or at low or high temperatures. An in meso robot, when used properly, can greatly improve the productivity of membrane protein structure and function research by facilitating crystallization which is one of the slow steps in the overall structure determination pipeline. 
In this video article, we demonstrate the use of three commercially available robots that can dispense the viscous and sticky mesophase integral to in meso crystallogenesis. The first robot was developed in the MS&amp;FB Group5,6. The other two have recently become available and are included here for completeness.	
An overview of the protocol covered in this article is presented in Figure 1. All manipulations were performed at room temperature (~20 &deg;C) under ambient conditions.

Herein is described a robotic approach to high-throughput crystallization of membrane proteins in lipidic mesophases for use in structure determination using macromolecular X-ray crystallography. Three robots capable of handling the viscous and sticky protein-laden mesophase integral to the method are introduced.

Structure-function studies of membrane proteins greatly benefit from having available high-resolution 3-D structures of the type provided through ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

A Practical and Novel Method to Extract Genomic DNA from Blood Collection Kits for Plasma Protein Preservation

Laboratory tests can be done on the cellular or fluid portions of the blood. The use of different blood collection tubes determines the portion of the blood that can be analyzed (whole blood, plasma or serum). Laboratories involved in studying the genetic basis of human disorders rely on anticoagulated whole blood collected in EDTA-containing vacutainer as the source of DNA for genetic / genomic analysis. Because most clinical laboratories perform biochemical, serologic and viral testing as a first step in phenotypic outcome investigation, anticoagulated blood is also collected in heparin-containing tube (plasma tube). Therefore when DNA and plasma are needed for simultaneous and parallel analyses of both genomic and proteomic data, it is customary to collect blood in both EDTA and heparin tubes. If blood could be collected in a single tube and serve as a source for both plasma and DNA, that method would be considered an advancement to existing methods. The use of the compacted blood after plasma extraction represents an alternative source for genomic DNA, thus minimizing the amount of blood samples processed and reducing the number of samples required from each patient. This would ultimately save time and resources.
The BD P100 blood collection system for plasma protein preservation were created as an improved method over previous plasma or serum collection tubes1, to stabilize the protein content of blood, enabling better protein biomarker discovery and proteomics experimentation from human blood. The BD P100 tubes contain 15.8 ml of spray-dried K2EDTA and a lyophilized proprietary broad spectrum cocktail of protease inhibitors to prevent coagulation and stabilize the plasma proteins. They also include a mechanical separator, which provides a physical barrier between plasma and cell pellets after centrifugation. Few methods have been devised to extract DNA from clotted blood samples collected in old plasma tubes2-4. Challenges from these methods were mainly associated with the type of separator inside the tubes (gel separator) and included difficulty in recovering the clotted blood, the inconvenience of fragmenting or dispersing the clot, and obstruction of the clot extraction by the separation gel.
We present the first method that extracts and purifies genomic DNA from blood drawn in the new BD P100 tubes. We compare the quality of the DNA sample from P100 tubes to that from EDTA tubes. Our approach is simple and efficient. It involves four major steps as follows: 1) the use of a plasma BD P100 (BD Diagnostics, Sparks, MD, USA) tube with mechanical separator for blood collection, 2) the removal of the mechanical separator using a combination of sucrose and a sterile paperclip metallic hook, 3) the separation of the buffy coat layer containing the white cells and 4) the isolation of the genomic DNA from the buffy coat using a regular commercial DNA extraction kit or a similar standard protocol.

We are describing a new method of isolating genomic DNA from whole blood collected for plasma/serology. After plasma collection, the compacted blood is usually discarded. Our novel method represents a significant improvement over existing methods and makes DNA and plasma available from a single collection, without requesting additional blood.

Laboratory tests can be done on the cellular or  fluid portions of the blood. The use of different blood collection tubes  determines the portion of the ...

Collection and Analysis of Arabidopsis Phloem Exudates Using the EDTA-facilitated Method

The plant phloem is essential for the long-distance transport of (photo-) assimilates as well as of signals conveying biotic or abiotic stress. It contains sugars, amino acids, proteins, RNA, lipids and other metabolites. While there is a large interest in understanding the composition and function of the phloem, the role of many of these molecules and thus, their importance in plant development and stress response has yet to be determined. One barrier to phloem analysis lies in the fact that the phloem seals itself upon wounding. As a result, the number of plants from which phloem sap can be obtained is limited. One method that allows collection of phloem exudates from several plant species without added equipment is the EDTA-facilitated phloem exudate collection described here. While it is easy to use, it does lead to the wounding of cells and care has to be taken to remove contents of damaged cells. In addition, several controls to prove purity of the exudate are necessary. Because it is an exudation rather than a direct collection of the phloem sap (not possible in many species) only relative quantification of its contents can occur. The advantage of this method over others is that it can be used in many herbaceous or woody plant species (Perilla, Arabidopsis, poplar, etc.) and requires minimal equipment and training. It leads to reasonably large amounts of exudates that can be used for subsequent analysis of proteins, sugars, lipids, RNA, viruses and metabolites. It is simple enough that it can be used in both a research as well as in a teaching laboratory.

Collection and Analysis of Arabidopsis Phloem Exudates Using the EDTA-facilitated Method

Knowledge of the composition of the phloem sap as well as the mechanism of its loading and long-distance transport is essential for the understanding of long-distance signaling in plant development and stress/pathogen response and of assimilate transport. This manuscript describes the collection of phloem exudates using the EDTA-facilitated method.

The plant phloem is essential for the long-distance transport of (photo-) assimilates as well as of signals conveying biotic or abiotic stress. It ...

Microperfusion Technique to Investigate Regulation of Microvessel Permeability in Rat Mesentery

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular mechanisms regulating vascular permeability in cultured endothelial cell monolayers and the functional exchange properties of whole microvascular beds. A method to cannulate and perfuse venular microvessels of rat mesentery and measure the hydraulic conductivity of the microvessel wall is described. The main equipment needed includes an intravital microscope with a large modified stage that supports micromanipulators to position three different microtools: (1) a beveled glass micropipette to cannulate and perfuse the microvessel; (2) a glass micro-occluder to transiently block perfusion and enable measurement of transvascular water flow movement at a measured hydrostatic pressure, and (3) a blunt glass rod to stabilize the mesenteric tissue at the site of cannulation. The modified Landis micro-occlusion technique uses red cells suspended in the artificial perfusate as markers of transvascular fluid movement, and also enables repeated measurements of these flows as experimental conditions are changed and hydrostatic and colloid osmotic pressure difference across the microvessels are carefully controlled. Measurements of hydraulic conductivity first using a control perfusate, then after re-cannulation of the same microvessel with the test perfusates enable paired comparisons of the microvessel response under these well-controlled conditions. Attempts to extend the method to microvessels in the mesentery of mice with genetic modifications expected to modify vascular permeability were severely limited because of the absence of long straight and unbranched microvessels in the mouse mesentery, but the recent availability of the rats with similar genetic modifications using the CRISPR/Cas9 technology is expected to open new areas of investigation where the methods described herein can be applied.

The modified Landis technique enables paired measurement of the hydraulic conductivity of individual microvessels in the mesentery of normal and genetically modified rats under control and test conditions using microperfusion techniques. It provides a convenient method to evaluate mechanisms that regulate microvessel permeability and transvascular exchange under physiological conditions.

Experiments to measure the permeability properties of individually perfused microvessels provide a bridge between investigation of molecular and cellular ...

Ground State Depletion Super-resolution Imaging in Mammalian Cells

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to dramatic leaps in our understanding of how cells function. The development and availability of super-resolution microscopy has considerably extended the limits of optical resolution from ~250-20 nm. Biologists are no longer limited to describing molecular interactions in terms of colocalization within a diffraction limited area, rather it is now possible to visualize the dynamic interactions of individual molecules. Here, we outline a protocol for the visualization and quantification of cellular proteins by ground-state depletion microscopy&#160;for fixed cell imaging. We provide examples from two different membrane proteins, an element of the endoplasmic reticulum translocon, sec61&#946;, and a plasma membrane-localized voltage-gated L-type Ca2+ channel (CaV1.2). Discussed are the specific microscope parameters, fixation methods, photo-switching buffer formulation, and pitfalls and challenges of image processing.

This article outlines a protocol for the detection of one or more plasma and/or intracellular membrane proteins using Ground State Depletion (GSD) super-resolution microscopy in mammalian cells. Here, we discuss the benefits and considerations of using such approaches for the visualization and quantification of cellular proteins.

Advances in fluorescent microscopy and cell biology are intimately correlated, with the enhanced ability to visualize cellular events often leading to ...

The CryoAPEX Method for Electron Microscopy Analysis of Membrane Protein Localization Within Ultrastructurally-Preserved Cells

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise subcellular localization of proteins is thus important for answering many biological questions. The quest for a robust label to identify protein localization combined with adequate cellular preservation and staining has been historically challenging. Recent advances in electron microscopy (EM) imaging have led to the development of many methods and strategies to increase cellular preservation and label target proteins. A relatively new peroxidase-based genetic tag, APEX2, is a promising leader in cloneable EM-active tags. Sample preparation for transmission electron microscopy (TEM) has also advanced in recent years with the advent of cryofixation by high pressure freezing (HPF) and low-temperature dehydration and staining via freeze substitution (FS). HPF and FS provide excellent preservation of cellular ultrastructure for TEM imaging, second only to direct cryo-imaging of vitreous samples. Here we present a protocol for the cryoAPEX method, which combines the use of the APEX2 tag with HPF and FS. In this protocol, a protein of interest is tagged with APEX2, followed by chemical fixation and the peroxidase reaction. In place of traditional staining and alcohol dehydration at room temperature, the sample is cryofixed and undergoes dehydration and staining at low temperature via FS. Using cryoAPEX, not only can a protein of interest be identified within subcellular compartments, but also additional information can be resolved with respect to its topology within a structurally preserved membrane. We show that this method can provide high enough resolution to decipher protein distribution patterns within an organelle lumen, and to distinguish the compartmentalization of a protein within one organelle in close proximity to other unlabeled organelles. Further, cryoAPEX is procedurally straightforward and amenable to cells grown in tissue culture. It is no more technically challenging than typical cryofixation and freeze substitution methods. CryoAPEX is widely applicable for TEM analysis of any membrane protein that can be genetically tagged.

This protocol describes the cryoAPEX method, in which an APEX2-tagged membrane protein can be localized by transmission electron microscopy within optimally-preserved cell ultrastructure.

Key cellular events like signal transduction and membrane trafficking rely on proper protein location within cellular compartments. Understanding precise ...

Neutron Spin Echo Spectroscopy as a Unique Probe for Lipid Membrane Dynamics and Membrane-Protein Interactions

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral budding, among other vital cellular processes. For efficient biological activity, cell membranes should be rigid enough to maintain the integrity of the cell and its compartments yet fluid enough to allow membrane components, such as proteins and functional domains, to diffuse and interact. This delicate balance of elastic and fluid membrane properties, and their impact on biological function, necessitate a better understanding of collective membrane dynamics over mesoscopic length and time scales of key biological processes, e.g., membrane deformations and protein binding events. Among the techniques that can effectively probe this dynamic range is neutron spin echo (NSE) spectroscopy. Combined with deuterium labeling, NSE can be used to directly access bending and thickness fluctuations as well as mesoscopic dynamics of select membrane features. This paper provides a brief description of the NSE technique and outlines the procedures for performing NSE experiments on liposomal membranes, including details of sample preparation and deuteration schemes, along with instructions for data collection and reduction. The paper also introduces data analysis methods used to extract key membrane parameters, such as the bending rigidity modulus, area compressibility modulus, and in-plane viscosity. To illustrate the biological importance of NSE studies, select examples of membrane phenomena probed by NSE are discussed, namely, the effect of additives on membrane bending rigidity, the impact of domain formation on membrane fluctuations, and the dynamic signature of membrane-protein interactions.

This paper describes the protocols for sample preparation, data reduction, and data analysis in neutron spin echo (NSE) studies of lipid membranes. Judicious deuterium labeling of lipids enables access to different membrane dynamics on mesoscopic length and time scales, over which vital biological processes occur.

Lipid bilayers form the main matrix of cell membranes and are the primary platform for nutrient exchange, protein-membrane&#160;interactions, and viral ...