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><tbody><tr><td>0.22 &amp;micro;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).&lt;br/&gt; 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.&lt;br/&gt; c. Filter the solution to remove bacteria with a 0.22 &amp;micro;m syringe filter and a 50 mL sterile syringe, subpackage the solution in 1.5 mL centrifuge tubes and restore at -20 &amp;deg;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 &amp;deg;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.&amp;nbsp;Dissolve 242 g Tris and 18.612 g EDTA in 700 mL of distilled water.&lt;br/&gt; b. Adjust pH to 7.8 with about 57.1 mL of acetic acid.&lt;br/&gt; c. Adjust the final volume to 1,000 mL.&lt;br/&gt; 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>&amp;alpha;-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&amp;times;)</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.&lt;br/&gt; b. Adjust the final volume to 500 mL, transfer the solution to a 1,000 mL conical flask, and add 7.5 g agar.&lt;br/&gt; c. Autoclave the medium for 20 min at 121 &amp;deg;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.&lt;br/&gt; b. Adjust the pH to about 5.8 with 2 M KOH by a pH meter.&lt;br/&gt; 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.&lt;br/&gt; d. Autoclave for 20 min at 121 &amp;deg;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 &amp;micro;L</td><td>Gilson</td><td>ECS000699</td><td></td></tr><tr><td>Pipettes 100-1,000 &amp;micro;L</td><td>Eppendorf</td><td>3120000267</td><td></td></tr><tr><td>Pipettes 20-200 &amp;micro;L</td><td>Eppendorf</td><td>3120000259</td><td></td></tr><tr><td>Pipettes 2-20 &amp;micro;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&amp;rsquo;-ACGGATACCTTGTTACGAC-3&amp;rsquo;, 1492R: 5&amp;rsquo;-ACGGATACCTTGTTACGAC-3&amp;rsquo;</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>&amp;nbsp;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

Research

JoVE Journal

Biology

2.4K Views.  Hubei University. De darmbacteriën kunnen meerdere fysiologische processen van insectengastheren beïnvloeden. Het voorbereiden en onderhouden van axeenlarven is een krachtig hulpmiddel om darmbacteriënfuncties te bestuderen. De belangrijkste voordelen van dit protocol zijn dat plantenweefselkweek wordt gebruikt om kiemvrije bladeren te verkrijgen om axeenbladetende insecten te achterhalen die zijn afgeleid van oppervlaktegesteriliseerde eieren, en dat deze methode niet wordt beperkt door kunstmatige diëten...

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

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. This is due to its reproductive behavior: females damage the external surface of fruits and vegetables when they lay eggs and the hatched larvae feed on their pulp. Wild C. capitata populations are traditionally controlled through insecticide spraying and/or eco-friendly approaches, the most successful being the Sterile Insect Technique (SIT). The SIT relies on mass-rearing, radiation-based sterilization and field release of males that retain their capacity to mate but are not able to generate fertile progeny. The advent and the subsequent rapid development of biotechnological tools, together with the availability of the medfly genome sequence, has greatly boosted our understanding of the biology of this species. This favored the proliferation of new strategies for genome manipulation, which can be applied to population control.
In this context, embryo microinjection plays a dual role in expanding the toolbox for medfly control. The ability to interfere with the function of genes that regulate key biological processes, indeed, expands our understanding of the molecular machinery underlying medfly invasiveness. Furthermore, the ability to achieve germ-line transformation facilitates the production of multiple transgenic strains that can be tested for future field applications in novel SIT settings. Indeed, genetic manipulation can be used to confer desirable traits that can, for example, be used to monitor sterile male performance in the field, or that can result in early life-stage lethality. Here we describe a method to microinject nucleic acids into medfly embryos to achieve these two main goals.

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. ...

Genetics

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

Developmental Biology

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and tomato. The susceptibility of M. sexta larvae to a variety of entomopathogenic bacterial species1-5, as well as the wealth of information available regarding the insect's immune system6-8, and the pending genome sequence9 make it a good model organism for use in studying host-microbe interactions during pathogenesis. In addition, M. sexta larvae are relatively large and easy to manipulate and maintain in the laboratory relative to other susceptible insect species. Their large size also facilitates efficient tissue/hemolymph extraction for analysis of the host response to infection.
The method presented here describes the direct injection of bacteria into the hemocoel (blood cavity) of M. sexta larvae. This approach can be used to analyze and compare the virulence characteristics of various bacterial species, strains, or mutants by simply monitoring the time to insect death after injection. This method was developed to study the pathogenicity of Xenorhabdus and Photorhabdus species, which typically associate with nematode vectors as a means to gain entry into the insect. Entomopathogenic nematodes typically infect larvae via natural digestive or respiratory openings, and release their symbiotic bacterial contents into the insect hemolymph (blood) shortly thereafter10. The injection method described here bypasses the need for a nematode vector, thus uncoupling the effects of bacteria and nematode on the insect. This method allows for accurate enumeration of infectious material (cells or protein) within the inoculum, which is not possible using other existing methods for analyzing entomopathogenesis, including nicking11 and oral toxicity assays12. Also, oral toxicity assays address the virulence of secreted toxins introduced into the digestive system of larvae, whereas the direct injection method addresses the virulence of whole-cell inocula.
The utility of the direct injection method as described here is to analyze bacterial pathogenesis by monitoring insect mortality. However, this method can easily be expanded for use in studying the effects of infection on the M. sexta immune system. The insect responds to infection via both humoral and cellular responses. The humoral response includes recognition of bacterial-associated patterns and subsequent production of various antimicrobial peptides7; the expression of genes encoding these peptides can be monitored subsequent to direct infection via RNA extraction and quantitative PCR13. The cellular response to infection involves nodulation, encapsulation, and phagocytosis of infectious agents by hemocytes6. To analyze these responses, injected insects can be dissected and visualized by microscopy13, 14.

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

The method described here utilizes direct injection of entomopathogenic bacteria into the hemocoel of Manduca sexta insect larvae. M. sexta is a commercially available and well-studied insect. Thus, this method represents a simple approach to analyzing host-bacterial interactions from the perspective of one or both partners.

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and ...

Immunology and Infection

The Insect Galleria mellonella as a Powerful Infection Model to Investigate Bacterial Pathogenesis

The study of bacterial virulence often requires a suitable animal model. Mammalian models of infection are costly and may raise ethical issues. The use of insects as infection models provides a valuable alternative. Compared to other non-vertebrate model hosts such as nematodes, insects have a relatively advanced system of antimicrobial defenses and are thus more likely to produce information relevant to the mammalian infection process. Like mammals, insects possess a complex innate immune system1. Cells in the hemolymph are capable of phagocytosing or encapsulating microbial invaders, and humoral responses include the inducible production of lysozyme and small antibacterial peptides2,3. In addition, analogies are found between the epithelial cells of insect larval midguts and intestinal cells of mammalian digestive systems. Finally, several basic components essential for the bacterial infection process such as cell adhesion, resistance to antimicrobial peptides, tissue degradation and adaptation to oxidative stress are likely to be important in both insects and mammals1. Thus, insects are polyvalent tools for the identification and characterization of microbial virulence factors involved in mammalian infections.
Larvae of the greater wax moth Galleria mellonella have been shown to provide a useful insight into the pathogenesis of a wide range of microbial infections including mammalian fungal (Fusarium oxysporum, Aspergillus fumigatus, Candida albicans) and bacterial pathogens, such as Staphylococcus aureus, Proteus vulgaris, Serratia marcescens Pseudomonas aeruginosa, Listeria monocytogenes or Enterococcus faecalis4-7. Regardless of the bacterial species, results obtained with Galleria larvae infected by direct injection through the cuticle consistently correlate with those of similar mammalian studies: bacterial strains that are attenuated in mammalian models demonstrate lower virulence in Galleria, and strains causing severe human infections are also highly virulent in the Galleria model8-11. Oral infection of Galleria is much less used and additional compounds, like specific toxins, are needed to reach mortality.
G. mellonella larvae present several technical advantages: they are relatively large (last instar larvae before pupation are about 2 cm long and weight 250 mg), thus enabling the injection of defined doses of bacteria; they can be reared at various temperatures (20 &deg;C to 30 &deg;C) and infection studies can be conducted between 15 &deg;C to above 37 &deg;C12,13, allowing experiments that mimic a mammalian environment. In addition, insect rearing is easy and relatively cheap. Infection of the larvae allows monitoring bacterial virulence by several means, including calculation of LD5014, measurement of bacterial survival15,16 and examination of the infection process17. Here, we describe the rearing of the insects, covering all life stages of G. mellonella. We provide a detailed protocol of infection by two routes of inoculation: oral and intra haemocoelic. The bacterial model used in this protocol is Bacillus cereus, a Gram positive pathogen implicated in gastrointestinal as well as in other severe local or systemic opportunistic infections18,19.

The Insect Galleria mellonella as a Powerful Infection Model to Investigate Bacterial Pathogenesis

Oral and intra haemocolic infection of larvae of the greater wax moth Galleria mellonella is described. This insect can be used to study virulence factors of entomopathogenic as well as mammalian opportunistic bacteria. Rearing of the insects, methods of infection and examples of in vivo analysis are described.

The study of bacterial virulence often requires a suitable animal model. Mammalian models of infection are costly and may raise ethical issues. The use of ...

In vivo and In vitro Rearing of Entomopathogenic Nematodes (Steinernematidae and Heterorhabditidae)

Entomopathogenic nematodes (EPN) (Steinernematidae and Heterorhabditidae) have a mutualistic partnership with Gram-negative Gamma-Proteobacteria in the family Enterobacteriaceae. Xenorhabdus bacteria are associated with steinernematids nematodes while Photorhabdus are symbionts of heterorhabditids. Together nematodes and bacteria form a potent insecticidal complex that kills a wide range of insect species in an intimate and specific partnership. Herein, we demonstrate in vivo and in vitro techniques commonly used in the rearing of these nematodes under laboratory conditions. Furthermore, these techniques represent key steps for the successful establishment of EPN cultures and also form the basis for other bioassays that utilize these organisms for research. The production of aposymbiotic (symbiont&#8211;free) nematodes is often critical for an in-depth and multifaceted approach to the study of symbiosis. This protocol does not require the addition of antibiotics and can be accomplished in a short amount of time with standard laboratory equipment. Nematodes produced in this manner are relatively robust, although their survivorship in storage may vary depending on the species used. The techniques detailed in this presentation correspond to those described by various authors and refined by P. Stock&#8217;s Laboratory, University of Arizona (Tucson, AZ, USA). These techniques are distinct from the body of techniques that are used in the mass production of these organisms for pest management purposes.

In vivo and In vitro Rearing of Entomopathogenic Nematodes Steinernematidae and Heterorhabditidae

The goal of this presentation is to demonstrate in vivo and in vitro techniques for the rearing of entomopathogenic nematodes. In vivo methods consider the rearing of these nematodes with an insect host, whereas the in vitro methods utilize rich agar media.

Entomopathogenic nematodes (EPN) (Steinernematidae and Heterorhabditidae) have a mutualistic partnership with Gram-negative Gamma-Proteobacteria in the ...

Bioengineering

Culturing and Genetically Manipulating Entomopathogenic Nematodes

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic of their life cycle is the mutualistic association with the bacteria Photorhabdus and Xenorhabdus, respectively. The nematode parasites are able to locate and enter suitable insect hosts, subvert the insect immune response, and multiply efficiently to produce the next generation that will actively hunt new insect prey to infect. Due to the properties of their life cycle, entomopathogenic nematodes are popular biological control agents, which are used in combination with insecticides to control destructive agricultural insect pests. Simultaneously, these parasitic nematodes represent a research tool to analyze nematode pathogenicity and host anti-nematode responses. This research is aided by the recent development of genetic techniques and transcriptomic approaches for understanding the role of nematode secreted molecules during infection. Here, a detailed protocol on maintaining entomopathogenic nematodes and using a gene knockdown procedure is provided. These methodologies further promote the functional characterization of entomopathogenic nematode infection factors.

Entomopathogenic nematodes live in symbiosis with bacteria and together they successfully infect insects by undermining their innate immune system. To promote research on the genetic basis of nematode infection, methods for maintaining and genetically manipulating entomopathogenic nematodes are described.

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic ...

Establishment and Maintenance of Gnotobiotic American Cockroaches (Periplaneta americana)

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment and maintenance of gnotobiotic American cockroaches (Periplaneta americana). This approach includes built-in sterility checks for ongoing quality control. Gnotobiotic insects are defined here as cockroaches that still contain their vertically transmitted endosymbiont (Blattabacterium) but lack other microbes that normally reside on their surface and in their digestive tract. For this protocol, egg cases (oothecae) are removed from a (nonsterile) stock colony and surface sterilized. Once collected and sterilized, the oothecae are incubated at 30 &#176;C for approximately 4&#8722;6 weeks on brain-heart infusion (BHI) agar until they hatch or are removed due to contamination. Hatched nymphs are transferred to an Erlenmeyer flask containing a BHI floor, sterile water, and sterile rat food. To ensure that the nymphs are not housing microbes that are unable to grow on BHI in the given conditions, an additional quality control measure uses restriction fragment-length polymorphism (RFLP) to test for nonendosymbiotic microbes. Gnotobiotic nymphs generated using this approach can be inoculated with simple or complex microbial communities and used as a tool in gut microbiome studies.

Establishment and Maintenance of Gnotobiotic American Cockroaches Periplaneta americana

This protocol is used in establishing and maintaining gnotobiotic American cockroaches (Periplaneta americana) by surface sterilizing the egg cases (oothecae) prior to hatching. These gnotobiotic insects contain their vertically transmitted Blattabacterium endosymbionts but have axenic guts.

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment ...

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human food. Outside its native range, culturing G. bimaculatus is feasible due to its dietary plasticity, rapid reproductive cycle, lack of diapause requirement, tolerance for high-density rearing, and robustness against pathogens. Thus, G. bimaculatus can be&#160;a versatile model for studies of insect physiology, behavior, embryology, or genetics.
Cultural parameters, such as stocking density, within-cage refugia, photoperiod, temperature, relative humidity, and diet, all impact cricket growth, behavior, and gene expression and should be standardized. In the burgeoning literature on farming insects for human consumption, these crickets are frequently employed to evaluate candidate feed admixtures derived from crop residues, food-processing byproducts, and other low-cost waste streams.
To support ongoing experiments evaluating G. bimaculatus growth performance and nutritional quality in response to variable feed substrates, a comprehensive set of standard protocols for breeding, upkeep, handling, measurement, and euthanasia in the laboratory was developed and is presented here. An industry-standard cricket feed has proven nutritionally adequate and functionally appropriate for the long-term maintenance of cricket breeding stocks, as well as for use as an experimental control feed. Rearing these crickets at a density of 0.005 crickets/cm3 in screen-topped 29.3 L polyethylene cages at an average temperature of 27 &#176;C on a 12 light (L)/12 dark (D) photoperiod, with moistened coconut coir serving both as hydration source and oviposition medium has successfully sustained healthy crickets over a 2-year span. Following these methods, crickets in a controlled experiment yielded an average mass of 0.724 g 0.190 g at harvest, with 89% survivorship and 68.2% sexual maturation between stocking (22 days) and harvest (65 days).

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

This paper outlines basic methods to standardize important factors such as density, feed availability, hydration source, and environmental controls for the long-term rearing of laboratory cultures of the edible cricket, Gryllus bimaculatus.

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human ...

An Experimental Study on Colorado Potato Beetle Hibernation Under Natural Conditions

One of the major pests of potato Solanum tuberosum L. in the temperate zone is the insect Colorado potato beetle (CPB). Most studies on the immunity and diseases of the CPB are conducted during active feeding stages. Nonetheless, there are fewer studies on resting stages, although these beetles spend most of their life cycle in a state of winter diapause (hibernation). In this work, a method for investigating CPB hibernation under natural conditions was developed and tested, offering an opportunity to collect a sufficient number of individuals in winter. In this article, CPB survival was assessed, and infectious agents at different stages of hibernation were identified. CPB mortality increased during the hibernation, reaching a maximum in April-May. Entomopathogenic fungi (Beauveria, Isaria, and Lecanicillium) and bacteria Bacillus, Sphingobacterium, Peribacillus, Pseudomonas, and Serratia were isolated&#160;from the dead insects. The survival rate of the beetles for the entire hibernation period was 61%. No frozen or desiccated beetles were found, indicating the success of the presented method.

Here we present a method for studying Colorado potato beetle hibernation under the natural conditions of the temperate zone as well as a technique for collecting beetles in winter. This method allows to obtain a desired number of overwintering individuals for various analyses at any stage of hibernation.

One of the major pests of potato Solanum tuberosum L. in the temperate zone is the insect Colorado potato beetle (CPB). Most studies on the immunity and ...

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 ...

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

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