This work was funded by the National Natural Science Foundation of China (31971663) and the Young Elite Scientists Sponsorship Program by CAST (2020QNRC001).

1. Insect rearing
<ol>
	<li>Maintain&#160;the P. versicolora population in a growth chamber at the condition of 27 &#176;C and 70 &#177; 5% relative humidity with a photoperiod of 16 h light/8 h dark. Place them in perforated plastic boxes with tiled wet absorbing paper and feed them fresh poplar branches. Spray clean water on absorbing paper to maintain moisture and change the branches every two days.</li>
	<li>Isolate adults for oviposition after pupation. Feed them tender leaves to obtain more eggs.</li>
	<li>Collect newly laid eggs (within 24 h). Place the eggs on moist absorbing paper for 60 h to prepare axenic larvae. 
	&#8203;NOTE: Newly laid eggs will hatch after ~72 h. The best time to sterilize egg surfaces is the day before hatching; otherwise, the number of successfully hatched eggs will decrease.</li>
</ol>
2. Germ-free poplar culturing
<ol>
	<li>Prepare Murashige and Skoog (MS) medium and 1 mg/mL &#945;-naphthalene acetic acid (NAA) stock solutions (see the Table of Materials).</li>
	<li>In a biosafety hood, add 50 &#181;L of 1 mg/mL NAA stock solution to 500 mL of MS medium, shake to mix well, and pour ~50 mL per tissue culture container and wait for solidification.</li>
	<li>Prepare scalpels, alcohol lamp, and forceps; sterilize the scalpels and forceps in the alcohol lamp flame in the biosafety hood.</li>
	<li>Cut 3-4 cm stem segments with apical buds or lateral buds from poplar seedlings at ~1 month of growth (germ-free tissue-cultured seedlings), and insert them into the culture medium, one or two stem segments per container.</li>
	<li>Incubate these stem segments in a growth chamber at 25 &#176;C and 50 &#177; 10 cd light intensity with a photoperiod of 16 h light/8 h dark for approximately 30 days. Use the germ-free leaves to feed the axenic larvae.</li>
</ol>
3. Egg surface sterilization and rearing axenic larvae
<ol>
	<li>Autoclave Petri dishes, paintbrushes, filter paper, distilled water, and a Petri dish containing LB (Luria-Bertani) agar medium.</li>
	<li>Place leaves with adherent eggs in a Petri dish, carefully remove the eggs from the leaves using forceps, and transfer them to another Petri dish. 
	NOTE: This step should be performed very carefully because the eggs are adherent to the leaves.</li>
	<li>Wash these eggs with 75% ethanol for 8 min, and repeat the wash four times with sterile water.</li>
	<li>Transfer the eggs onto LB agar medium to preserve moisture for hatching. 
	NOTE: Using LB agar medium can help to verify whether the disinfected egg is germ-free.</li>
	<li>Place the Petri dish in a growth chamber and wait for the eggs to hatch within 24 h.</li>
	<li>In a biosafety hood, tile three pieces of wet filter paper in a Petri dish, place germ-free poplar leaves on the paper, collect the larvae and place them on the leaves, seal the Petri dish with parafilm, and incubate them in a growth chamber at 27 &#176;C and 70 &#177; 5% relative humidity with a photoperiod of 16 h light/8 h dark. 
	NOTE: The tissue-cultured leaves are delicate and can lose water quickly. Thus, using moist filter paper is necessary when transferring the leaves to the Petri dish.</li>
	<li>Change the leaves every two days.</li>
	<li>For the conventionally reared groups, transfer the eggs from the leaves to a Petri dish containing moist filter paper and feed these larvae with germ-free poplar leaves.</li>
</ol>
4. Verification of axenic larvae with culture-dependent assays
<ol>
	<li>Randomly select three 1st, 2nd, and 3rd instar larvae from the germ-free and conventionally reared groups.</li>
	<li>Dissect the 3rd instar larvae with sterile scissors and forceps under a stereomicroscope and collect their guts in microcentrifuge tubes. Collect intact 1st and 2nd instar larvae in tubes. 
	NOTE: Collect whole 1st and 2nd instar larvae as they are too small to dissect, and keep their guts intact.</li>
	<li>Add 100 &#181;L of phosphate-buffered saline and three steel balls to 1.5 mL tubes and grind the tissues into homogenates using a bead-beating homogenizer.</li>
	<li>Add 100 &#181;L of the homogenate to LB agar medium and plate using a sterile glass spreading rod.</li>
	<li>Place the plates at 37 &#176;C for 24 h and observe the bacterial colonies.</li>
</ol>
5. Verification of axenic individuals with culture-independent assays
<ol>
	<li>Extract the total DNA of tissues (obtained in section 4) using a DNA extraction kit.</li>
	<li>Measure DNA concentration using a spectrophotometer at 260 nm.</li>
	<li>Amplify the bacterial 16S rRNA gene using universal 16S rRNA primers with PCR. Set up the reaction system with 18 &#181;L of 1.1x PCR master mix (see the Table of Materials), 0.5 &#181;L of 27F primer (5&#39;-ACGGATACCTTGTTACGAC-3&#39;), 0.5 &#181;L of 1495R primer (5&#39;-ACGGATACCTTGTTACGAC-3&#39;) and 100 ng template DNA. Set the PCR conditions at 95 &#176;C for 3 min; 28 cycles of 95 &#176;C for 30 s, 55 &#176;C for 1 min, and 72 &#176;C for 1 min; followed by 72 &#176;C for 10 min. Store the PCR products at 4 &#176;C until further analysis.</li>
	<li>Mix the PCR products with nucleic acid dye and analyze them using electrophoresis on a 1% agarose gel in 1x TAE buffer. Use 10 &#181;L of a DNA marker as a reference.</li>
	<li>Observe the gel with a UV transilluminator and look for the target fragment around 1,500 bp.</li>
</ol>

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The authors have no conflicts of interest to disclose.

Preparation of germ-free larvae and obtaining gnotobiotic larvae by reintroducing specific bacterial strains are powerful methods to elucidate the mechanisms underlying host-microbe interactions. Newly hatched larvae obtain gut microbiota in two main ways: vertical transmission from the mother to the offspring or horizontal acquisition from siblings and the environment34. The former can be fulfilled by parental transfer to the offspring through contamination of the egg surface35<...

Similar to mammals, the insect digestive tract is a cavity for food digestion and absorption. Most insects harbor diverse commensal bacteria that thrive in their guts and live on nutrition supplied by hosts1. The gut commensal community has a profound impact on multiple physiological processes in insects, including food digestion and detoxification2,3,4, nutrition and development5,6,7, defense against pathogens and parasites8,9,10,11, chemical communication12,13 and behaviors14,15. Intriguingly, some gut microbiota can be facultatively pathogenic or be manipulated by invading pathogens to aggravate infection, indicating that gut bacteria can be harmful in some cases16,17,18. Gut bacteria can also serve as a microbial resource for biotechnical applications and pest management. For example, lignocellulose-digesting bacteria from phytophagous and xylophagous insects were used to digest plant cells for developing biofuels19. The dispersal of engineered gut symbionts expressing bioactive molecules is a novel and promising tactic to manage agriculture and forestry pests and mosquitoes transmitting infectious diseases19,20,21, which can also be used to improve the fitness of beneficial insects22. Illustrating how a gut bacterium behaves in vivo is thus considered a priority to fully leverage its function and further exploit it for various applications.
Animals can harbor 1 to &#62;1000 symbiotic microbial species in the gut1. As a result, it is difficult to accurately verify how individual bacterial taxa or their assembly perform inside an animal, and whether the host or its microbial partners drive a specific function. Therefore, preparing axenic larvae to obtain gnotobiotic insects by mono- or multi-species colonization is necessary to investigate bacterial function and interaction with insects23. At present, administering antibiotic cocktails and sterilizing the surface of insect eggs are common methods to remove gut bacteria14,24,25,26. However, antibiotic diets cannot eliminate gut bacteria completely and have a negative effect on host insect physiology27,28. Consequently, the use of antibiotic-treated insects may obscure the true abilities of some gut bacteria. Fortunately, surface sterilization of eggs can negate this problem23,29, which has no or negligible effects on experimental insects. Furthermore, artificial diets cannot fully resemble natural insect food, and developing an artificial diet is a costly and labor-consuming process30,31.
The willow leaf beetle, Plagiodera versicolora&#160;(Laicharting) (Coleoptera: Chrysomelidae), is a widespread leaf-eating pest that mainly feeds on salicaceous trees, such as willows (Salix) and poplar (Populus L.)32,33. Here, the willow leaf beetle was used as a representative leaf-eating insect to develop a protocol to prepare and rear a germ-free insect. We exploited plant tissue culture to obtain germ-free poplar leaves to rear P. versicolora axenic larvae from sterilized eggs. The axenic status of P. versicolora larvae was verified via culture-dependent and culture-independent assays. This protocol can maintain axenic insects that better mimic the wild condition than insect rearing with an artificial diet. More importantly, this method is convenient at a very low cost, which increases the feasibility of obtaining axenic insects for future insect-gut microbiota interaction studies, especially for non-model insects without well-developed artificial diets.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.22 &#181;m syringe filters</td>
			<td>Millipore</td>
			<td>SLGP033RB</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mg/mL NAA stock solution</td>
			<td></td>
			<td></td>
			<td>a. Prepare 0.1 M NaOH solution (dissolve 0.8 g NaOH in 200 mL of distilled water). 
			b. Add 0.2 g NAA in a 250 mL beaker, add little 0.1 M NaOH solution until NAA dissolved, and adjust the final volume to 200 mL with distilled water. 
			c. Filter the solution to remove bacteria with a 0.22 &#181;m syringe filter and a 50 mL sterile syringe, subpackage the solution in 1.5 mL centrifuge tubes and restore at -20 &#176;C.</td>
		</tr>
		<tr>
			<td>1.5 mL microcentrifuge tubes</td>
			<td>Sangon Biotech</td>
			<td>F600620</td>
			<td></td>
		</tr>
		<tr>
			<td>10x PBS stock solution</td>
			<td>Biosharp Life Sciences</td>
			<td>BL302A</td>
			<td></td>
		</tr>
		<tr>
			<td>2 M KOH solution</td>
			<td></td>
			<td></td>
			<td>Dissolve 22.44 g KOH (molecular weight: 56.1) in 200 mL of distilled water and autoclave it for 20 min at 121 &#176;C.</td>
		</tr>
		<tr>
			<td>250 mL and 2,000 mL beakers</td>
			<td>Shubo</td>
			<td>sb16455</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL sterile syringes</td>
			<td>Jinta</td>
			<td>JT0125789</td>
			<td></td>
		</tr>
		<tr>
			<td>500 mL measuring cylinder</td>
			<td>Shubo</td>
			<td>sb1601</td>
			<td></td>
		</tr>
		<tr>
			<td>50x TAE stock solution</td>
			<td></td>
			<td></td>
			<td>a.&#160;Dissolve 242 g Tris and 18.612 g EDTA in 700 mL of distilled water. 
			b. Adjust pH to 7.8 with about 57.1 mL of acetic acid. 
			c. Adjust the final volume to 1,000 mL. 
			d. The stock solution was diluted to 1x TAE buffer when used.</td>
		</tr>
		<tr>
			<td>75% ethanol</td>
			<td>Xingheda trade</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#945;-naphthalene acetic acid (NAA)</td>
			<td>Solarbio Life Sciences</td>
			<td>86-87-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Absorbing paper</td>
			<td></td>
			<td></td>
			<td>22.3 cm x 15.3 cm x 9 cm</td>
		</tr>
		<tr>
			<td>Acetic acid</td>
			<td>Sinopharm Chemical Reagent Co. Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Agar</td>
			<td>Coolaber</td>
			<td>9002-18-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Biowest</td>
			<td>111860</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclave</td>
			<td>Panasonic</td>
			<td>MLS-3781L-PC</td>
			<td></td>
		</tr>
		<tr>
			<td>Bead-beating homogenizer</td>
			<td>Jing Xin</td>
			<td>XM-GTL64</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA extraction kit</td>
			<td>MP Biomedicals</td>
			<td>116560200</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Saiguo Biotech</td>
			<td>1340</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>Jiaojie</td>
			<td></td>
			<td>70 mm diameter</td>
		</tr>
		<tr>
			<td>Gel electrophoresis unit</td>
			<td>Bio-rad</td>
			<td>164-5052</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel Signal Green nucleic acid dye</td>
			<td>TsingKe</td>
			<td>TSJ003</td>
			<td></td>
		</tr>
		<tr>
			<td>Germ-free poplar seedlings</td>
			<td></td>
			<td></td>
			<td>Shan Xin poplar from Ludong University in Shandong Province</td>
		</tr>
		<tr>
			<td>Golden Star Super PCR Master Mix (1.1&#215;)</td>
			<td>TsingKe</td>
			<td>TSE101</td>
			<td></td>
		</tr>
		<tr>
			<td>Growth chamber</td>
			<td>Ruihua</td>
			<td>HP400GS-C</td>
			<td></td>
		</tr>
		<tr>
			<td>LB agar medium</td>
			<td></td>
			<td></td>
			<td>a. Dissolve 5 g tryptone, 5 g NaCl, 2.5 g yeast extract in 300 mL of distilled water. 
			b. Adjust the final volume to 500 mL, transfer the solution to a 1,000 mL conical flask, and add 7.5 g agar. 
			c. Autoclave the medium for 20 min at 121 &#176;C.</td>
		</tr>
		<tr>
			<td>Mini centrifuge</td>
			<td>DRAGONLAB</td>
			<td>D1008</td>
			<td></td>
		</tr>
		<tr>
			<td>MS basic medium</td>
			<td>Coolaber</td>
			<td>PM1121-50L</td>
			<td>M0245</td>
		</tr>
		<tr>
			<td>MS solid medium for germ-free poplar seedling culture</td>
			<td></td>
			<td></td>
			<td>a. Dissolve 4.43 g MS basic medium powder and 30 g sucrose in 800 mL of distilled water. 
			b. Adjust the pH to about 5.8 with 2 M KOH by a pH meter. 
			c. Adjust the final volume to 1,000 mL, separate into two parts, transfer into two 1,000 mL conical flasks, and add 2.6 g agar per 500 mL. 
			d. Autoclave for 20 min at 121 &#176;C.</td>
		</tr>
		<tr>
			<td>NanoDrop 1000 spectrophotometer</td>
			<td>Thermo Fisher Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Paintbrush</td>
			<td></td>
			<td></td>
			<td>1 cm width, used to collect the eggs</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>PM-996</td>
			<td></td>
		</tr>
		<tr>
			<td>PCR Thermal Cyclers</td>
			<td>Eppendorf</td>
			<td>6331000076</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Supin</td>
			<td></td>
			<td>90 mm diameter</td>
		</tr>
		<tr>
			<td>pH meter</td>
			<td>METTLER TOLEDO</td>
			<td>FE20</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes 0.2-2 &#181;L</td>
			<td>Gilson</td>
			<td>ECS000699</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes 100-1,000 &#181;L</td>
			<td>Eppendorf</td>
			<td>3120000267</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes 20-200 &#181;L</td>
			<td>Eppendorf</td>
			<td>3120000259</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes 2-20 &#181;L</td>
			<td>Eppendorf</td>
			<td>3120000232</td>
			<td></td>
		</tr>
		<tr>
			<td>Plant tissue culture container</td>
			<td>Chembase</td>
			<td>ZP21</td>
			<td>240 mL</td>
		</tr>
		<tr>
			<td>Plastic box</td>
			<td></td>
			<td></td>
			<td>2.35 L</td>
		</tr>
		<tr>
			<td>Potassium hydroxide (KOH)</td>
			<td>Sinopharm Chemical Reagent Co. Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Primers for amplifying the bacterial 16S rRNA gene</td>
			<td>Sangon Biotech</td>
			<td></td>
			<td>27-F: 5&#8217;-ACGGATACCTTGTTACGAC-3&#8217;, 1492R: 5&#8217;-ACGGATACCTTGTTACGAC-3&#8217;</td>
		</tr>
		<tr>
			<td>Sodium chloride (NaCl)</td>
			<td>Sinopharm Chemical Reagent Co. Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide (NaOH)</td>
			<td>Sinopharm Chemical Reagent Co. Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Steel balls</td>
			<td></td>
			<td>0.25 mm</td>
			<td>used to grind tissues</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>OLYMPUS</td>
			<td>SZ61</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sinopharm Chemical Reagent Co. Ltd</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trans2K plus II DNA marker</td>
			<td>Transgene Biotech</td>
			<td>BM121-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris base</td>
			<td>Biosharp Life Sciences</td>
			<td>1115</td>
			<td></td>
		</tr>
		<tr>
			<td>Tryptone</td>
			<td>Thermo Fisher Scientific</td>
			<td>&#160;LP0037</td>
			<td></td>
		</tr>
		<tr>
			<td>UV transilluminator</td>
			<td>Monad Biotech</td>
			<td>QuickGel 6100</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortexer</td>
			<td>Scilogex</td>
			<td>MX-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Willow branches</td>
			<td></td>
			<td></td>
			<td>Sha Lake Park, Wuhan, China</td>
		</tr>
		<tr>
			<td>Willow leaf beetle</td>
			<td></td>
			<td></td>
			<td>Huazhong Agricultural University, Wuhan, China</td>
		</tr>
		<tr>
			<td>Yeast extract</td>
			<td>Thermo Fisher Scientific</td>
			<td>LP0021</td>
			<td></td>
		</tr>
	</tbody>
</table>

preparing and rearing axenic insects with tissue cultured seedlings for host-gut microbiota interaction studies of the leaf beetle

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain into an axenic insect is a powerful method to verify gut microbial function and elucidate the mechanisms underlying gut microbe-host interactions. Administering antibiotics or sterilizing egg surfaces are two commonly used methods to remove gut bacteria from insects. However, in addition to the potential adverse effects of antibiotics on insects, previous studies indicated that feeding antibiotics could not eliminate gut bacteria. Thus, germ-free artificial diets are generally employed to maintain axenic insects, which is a tedious and labor-intensive process that cannot fully resemble nutritional components in natural food. Described here is an efficient and simple protocol to prepare and maintain axenic larvae of a leaf beetle&#160;(Plagiodera versicolora). Specifically, surfaces of the beetle eggs were sterilized, following which germ-free poplar leaves were used to rear axenic larvae. The axenic status of the insects was further confirmed via culture-dependent and culture-independent assays. Collectively, by combining egg disinfection and germ-free cultivation, an efficient and convenient method was developed to obtain axenic P. versicolora, providing a readily transferable tool for other leaf-eating insects.

The life stages of P. versicolora are shown in Figure 1. The adult male is smaller than the adult female (Figure 1A). In the field, the beetle clusters its eggs on a leaf; here, four eggs were detached from a leaf (Figure 1B). The poplar stem segments and seedlings used for axenic insect rearing are shown in Figure 2. The gut of a 3rd instar larva is shown in Fig...

Watch this Scientific Journal Video about Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle at JoVE.com

Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle

To obtain an axenic insect, its egg surface is sterilized, and the hatched larva is subsequently reared using axenic leaves. This method provides an efficient way for axenic insect preparation without administering antibiotics or developing an artificial diet, which can also be applied to other leaf-eating insects.

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain ...

preparing-rearing-axenic-insects-with-tissue-cultured-seedlings-for

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Biology

2.4K Views.  Hubei University. To obtain an axenic insect, its egg surface is sterilized, and the hatched larva is subsequently reared using axenic leaves. This method provides an efficient way for axenic insect preparation without administering antibiotics or developing an artificial diet, which can also be applied to other leaf-eating insects.

Video: Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle

A Guide to Modern Quantitative Fluorescent Western Blotting with Troubleshooting Strategies

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins within complex biological samples. Since then, western blotting methodology has become a common component of the molecular biologists experimental repertoire. A cursory search of PubMed using the term &#8220;western blot&#8221; suggests that in excess of two hundred and twenty thousand published manuscripts have made use of this technique by the year 2014. Importantly, the last ten years have seen technical imaging advances coupled with the development of sensitive fluorescent labels which have improved sensitivity and yielded even greater ranges of linear detection. The result is a now truly Quantifiable Fluorescence based Western Blot (QFWB) that allows biologists to carry out comparative expression analysis with greater sensitivity and accuracy than ever before. Many &#8220;optimized&#8221; western blotting methodologies exist and are utilized in different laboratories. These often prove difficult to implement due to the requirement of subtle but undocumented procedural amendments. This protocol provides a comprehensive description of an established and robust QFWB method, complete with troubleshooting strategies.

The advancement of western blotting using fluorescence has allowed detection of subtle changes in protein expression enabling quantitative analyses. Here we describe a robust methodology for detection of a range of proteins across a variety of species and tissue types. A strategy to overcome common technical problems is also provided.

The late 1970s saw the first publicly reported use of the western blot, a technique for assessing the presence and relative abundance of specific proteins ...

Methods for the Extraction of Endosymbionts from the Whitefly Bemisia tabaci

Bacterial symbionts form an intimate relationship with their hosts and confer advantages to the hosts in most cases. Genomic information is critical to study the functions and evolution of bacterial symbionts in their host. As most symbionts cannot be cultured in vitro, methods to isolate an adequate quantity of bacteria for genome sequencing are very important. In the whitefly Bemisia tabaci, a number of endosymbionts have been identified and are predicted to be of importance in the development and reproduction of the pests through multiple approaches. However, the mechanism underpinning the associations remains largely unknown. The obstacle partially comes from the fact that the endosymbionts in whitefly, mostly restrained in bacteriocytes, are hard to separate from the host cells. Here we report a step-by-step protocol for the identification, extraction and purification of endosymbionts from the whitefly B. tabaci mainly by dissection and filtration. Endosymbiont samples prepared by this method, although still a mixture of different endosymbiont species, are suitable for subsequent genome sequencing and analysis of the possible roles of endosymbionts in B. tabaci. This method may also be used to isolate endosymbionts from other insects.

Methods for the Extraction of Endosymbionts from the Whitefly Bemisia tabaci

Here, we present a protocol to isolate endosymbionts from the whitefly Bemisia tabaci through dissection and filtration. After amplification, the DNA samples are suitable for subsequent sequencing and study of the mutualism between endosymbionts and whitefly.

Bacterial symbionts form an intimate relationship with their hosts and confer advantages to the hosts in most cases. Genomic information is critical to ...

Application of Passive Head Motion to Generate Defined Accelerations at the Heads of Rodents

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular mechanisms behind the beneficial effects of exercise are poorly understood. Many physical workouts, particularly those classified as aerobic exercises such as jogging and walking, produce impulsive forces at the time of foot contact with the ground. Therefore, it was speculated that mechanical impact might be implicated in how exercise contributes to organismal homeostasis. For testing this hypothesis on the brain, a custom-designed ''passive head motion'' (hereafter referred to as PHM) system was developed that can generate vertical accelerations with controlled and defined magnitudes and modes and reproduce mechanical stimulation that might be applied to the heads of rodents during treadmill running at moderate velocities, a typical intervention to test the effects of exercise in animals. By using this system, it was demonstrated that PHM recapitulates the serotonin (5-hydroxytryptamine, hereafter referred to as 5-HT) receptor subtype 2A (5-HT2A) signaling in the prefrontal cortex (PFC) neurons of mice. This work provides detailed protocols for applying PHM and measuring its resultant mechanical accelerations at rodents' heads.

The present protocol describes a custom-designed ''passive head motion'' system, which reproduces mechanical accelerations at rodents' heads generated during their treadmill running at moderate velocities. It allows dissecting mechanical factors/elements from the beneficial effects of physical exercise.

Exercise is widely recognized as effective for various diseases and physical disorders, including those related to brain dysfunction. However, molecular ...

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, and parts of Africa over the last decade, becoming a devastating pest of various soft-skinned fruits in its invaded regions. Biological control, especially by means of self-perpetuating and specialized parasitoids, is expected to be a viable option for sustainable area-wide management of this highly mobile and polyphagous pest. Ganaspis brasiliensis Ihering (Hymenoptera: Figitidae) is a larval parasitoid that is widely distributed in East Asia, and has been found to be one of the most effective parasitoids of D. suzukii.
Following rigorous pre-introduction evaluations of its efficacy and potential non-target risks, one of the more host-specific genetic groups of this species (G1 G. brasiliensis) has been approved recently for introduction and field release in the United States and Italy. Another genetic group (G3 G. brasiliensis), which was also commonly found to attack D. suzukii in East Asia, may be considered for introduction in the near future. There is currently enormous interest in rearing G. brasiliensis for research or in mass-production for field release against D. suzukii. This protocol and associated video article describe effective rearing methods for this parasitoid, both on a small scale for research and a large scale for mass-production and field release. These methods may benefit further long-term research and use of this Asian-native parasitoid as a promising biological control agent for this global invasive pest.

Methods for Rearing the Parasitoid Ganaspis brasiliensis, a Promising Biological Control Agent for the Invasive Drosophila suzukii

Ganaspis brasiliensis-a larval parasitoid of Drosophila suzukii (a global invasive fruit crop pest)-has been approved or is considered for introduction into Europe and the United States for biological control of this pest. This article provides protocols for both small-scale and large-scale rearing of this parasitoid.

Native to East Asia, the spotted-wing drosophila, Drosophila suzukii (Matsumura) (Diptera: Drosophilidae), has established widely in the Americas, Europe, ...

Isolation, Characterization, and Therapeutic Application of Extracellular Vesicles from Cultured Human Mesenchymal Stem Cells

Extracellular vesicles (EVs) are heterogeneous membrane nanoparticles released by most cell types, and they are increasingly recognized as physiological regulators of organismal homeostasis and important indicators of pathologies; in the meantime, their immense potential to establish accessible and controllable disease therapeutics is emerging. Mesenchymal stem cells (MSCs) can release large amounts of EVs in culture, which have shown promise to jumpstart effective tissue regeneration and facilitate extensive therapeutic applications with good scalability and reproducibility. There is a growing demand for simple and effective protocols for collecting and applying MSC-EVs. Here, a detailed protocol is provided based on differential centrifugation to isolate and characterize representative EVs from cultured human MSCs, exosomes, and microvesicles for further applications. The adaptability of this method is shown for a series of downstream approaches, such as labeling, local transplantation, and systemic injection. The implementation of this procedure will address the need for simple and reliable MSC-EVs collection and application in translational research.

The present protocol describes the differential centrifugation for isolating and characterizing representative EVs (exosomes and microvesicles) from cultured human MSCs. Further applications of these EVs are also explained in this article.

Extracellular vesicles (EVs) are heterogeneous membrane nanoparticles released by most cell types, and they are increasingly recognized as physiological ...

Rearing Germ-Free Nasonia Wasps

Optimizing the Rearing Procedure of Germ-Free Wasps

Aseptic rearing technology is a method of culturing insects under sterile or almost sterile conditions, which can effectively eliminate the influence of external microorganisms on insect microbiota and thus promote the rapid development of insect microbiota research. Nasonia (wasp genus) is a parasitic wasp insect that has many advantages, such as a short lifespan, high genetic variation, easy operation, etc., and is widely used as an insect model system. Unlike antibiotic treatment, which can only reduce the number of microorganisms in animals, aseptic rearing techniques can control both the composition and quantity of microorganisms in animals, further facilitating the study of host-microbe interactions. However, previous versions of Nasonia rearing medium (NRM) have some defects and problems, such as a complex and time-consuming preparation process, easy contamination by bacteria or fungi, and short storage time. Therefore, this study solves these problems by optimizing the tools used in the NRM preparation process, storage conditions, and component ratios. The optimized medium could allow storage at -20 &#176;C for at least 3 months and eliminate the possibility of NRM contamination during feeding sterile wasps. This further improves the survival rate and health level of aseptic Nasonia, which is important for using Nasonia as a model for microbial research.

Author Spotlight: Optimizing the Rearing Procedure of Germ-Free Wasps  

Nasonia wasp embryos were dissected from Lucillia sericata pupae after parasitization for 12-24 h and washed with alcohol and 10% sodium hypochlorite solution to obtain germ-free embryos. After rearing the germ-free embryos&#160;and supplying them with Nasonia rearing medium to grow and develop in vitro, germ-free Nasonia adults were obtained.

Aseptic rearing technology is a method of culturing insects under sterile or almost sterile conditions, which can effectively eliminate the influence of ...

Germ-Free Wasp Embryo Collection and Rearing with Nasonia-Rearing Medium (NRM)

Germ-Free Wasp Embryo Collection and Rearing with Nasonia-Rearing Medium (NRM)  

Improved Methods for Imaging Maize Leaf Primordia

Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging

In maize (Zea mays) and other grasses (Poaceae), the leaf primordia are deeply ensheathed and rolled within the leaf whorl, making it difficult to study early leaf development. Here, we describe methods for preparing transverse sections and unrolled whole mounts of maize leaf primordia for fluorescence and confocal imaging. The first method uses a wire stripper to remove the upper portions of older leaves, exposing the tip of the leaf primordium and allowing its measurement for more accurate transverse section sampling. The second method uses clear, double-sided nano tape to unroll and mount whole-leaf primordia for imaging. We show the utility of the two methods in visualizing and analyzing fluorescent protein reporters in maize. These methods provide a solution to the challenges presented by the distinctive morphology of maize leaf primordia and will be useful for visualizing and quantifying leaf anatomical and developmental traits in maize and other grass species.

Author Spotlight: Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging  

Maize leaf primordia are deeply ensheathed and rolled, making them difficult to study. Here, we present methods for preparing transverse sections and unrolled whole mounts of maize leaf primordia for fluorescence and confocal imaging.

In maize (Zea mays) and other grasses (Poaceae), the leaf primordia are deeply ensheathed and rolled within the leaf whorl, making it difficult to study ...

Studying Development of Axenic Delia antiqua and Ensuring Bacteria-Free Growth

Rearing Axenic Delia antiqua with Half-Fermented Sterile Diets

Axenic insects are obtained from sterile artificial rearing systems using sterile media. These insects, characterized by their small size, short growth cycle, and low feed requirements, are ideal for studying the relationship between microorganisms and hosts. The gut microbiota significantly influences the physiological characteristics of insect hosts, and introducing specific strains into axenic insects provides a method for verifying gut microbial functions. Delia antiqua, a threatening pest in the order Diptera, family Anthomyiidae, and genus Delia, primarily feeds on onions, garlic, leeks, and other vegetables of the family Liliaceae. Its larvae feed on the bulbs, causing rotting, wilting, and even death of entire plants. By rearing axenic larvae, follow-up studies can be conducted to observe the effects of intestinal microflora on the growth and development of D. antiqua. Unlike the method involving antibiotic elimination of associated microbes, this article presents a low-cost and high-efficiency approach to raising axenic D. antiqua. After surface sterilization of D. antiqua eggs, half-fermented sterile diets were used to raise larvae, and the axenic state of D. antiqua was verified through culture-dependent and culture-independent assays. In conclusion, the combination of insect egg sterilization and the preparation of sterile diets for larval culture has enabled the development of an efficient and simple method for obtaining axenic D. antiqua. This method provides a powerful approach to studying insect-microflora interactions.

Author Spotlight: Axenic Rearing of Delia antiqua to Detect the Impact of Intestinal Microbes on Its Growth and Development 

A simple procedure for rearing axenic Delia antiqua with half-fermented sterile diets is described. Only one Wolbachia strain was detected in each instar of axenic D. antiqua using PCR.

Axenic insects are obtained from sterile artificial rearing systems using sterile media. These insects, characterized by their small size, short growth ...

Optimizing Pinewood Nematode Infection in Pinus pinaster

Infection of In Vivo and In Vitro Pines with the Pinewood Nematode Bursaphelenchus xylophilus and Isolation of Induced Volatiles

The pinewood nematode (PWN) is a phytoparasite that causes pine wilt disease (PWD) in conifer species. This plant parasitic nematode has heavily contributed to pine deforestation in Asian countries, e.g., Japan, China, and Korea. Over the last two decades, in Europe, Portugal and Spain have been greatly affected. Research on the mechanisms of PWN infection and/or PWD progression in susceptible host species relies on the controlled infection of pine seedlings under greenhouse conditions. This technique is laborious and mobilizes substantial economic and human resources. Additionally, it can be prone to variability that results from the genetic diversity associated with some pine species but also from the interference of external factors. As an alternative, in vitro co-cultures of pine with PWNs offer a more advantageous system for studying biochemical changes since they a) allow controlling single environmental or nutritional variables, b) occupy less space, c) require less time to obtain, and d) are&#160;free from contamination or from host genetic variation. The following protocol details the standard in vivo PWN infection of Pinus pinaster, the maritime pine, and the establishment of the novel in vitro co-cultures of pine shoots with the PWN as an improved methodology to study this phytoparasite influence on pine volatiles. PWN-induced volatiles are extracted from in vivo and in vitro infected pines by hydrodistillation and distillation-extraction, and the emitted volatiles are captured by solid phase microextraction (SPME), using fiber or packed column techniques.

Author Spotlight: In Vitro Co-Culture System of Pine Shoots and Pinewood Nematode for Studying Host Volatile Response

The protocol describes the in vivo and in vitro pinewood nematode infection of Pinus pinaster and their volatilome analysis through Gas Chromatography (GC) and GC coupled to Mass Spectrometry (GC-MS).