This work was supported by National Natural Science Foundation of China (32272530), the New Twenty Policies for University in Jinan Project (2021GXRC040), Major Scientific and Technological Innovation Projects in Shandong Province (2021TZXD002), and the Science and Education Integration Project of Qilu University of Technology (2022PYI009, 2022PY016, 2022PT105).

D. antiqua are obtained from the field of Fanzhen, Taian.
1. Preparation of sterile diets
<ol>
	<li>Peel off the outer layers of the scallions and discard the green leaves. Keep the white part of the scallions (Figure 1A) and wash them with sterile water, repeating the rinsing process three times. Cut the white part of the scallions into 1-2 cm diced pieces using scissors (see Table of Materials) sterilized with 75 % EtOH solution (mentioned in step 2.5) for later use.</li>
	<li>Weigh 50 g of the diced scallions and place them into a food processor (see Table of Materials). Add 50 mL of sterilized water and grind them in five times for 2 min each. All of the diced scallions should be grounded into a paste-like consistency (Figure 2A).</li>
	<li>Use tweezers to transfer 10 pieces 3rd instar larvae of D. antiqua into a 10 mL centrifuge tube containing 10 mL of 1x PBS, and grind them with a disposable grinding pestle (see Table of Materials) for about 5 min to obtain a larval grinding liquid.</li>
	<li>Pour the pasty scallion liquid and larval grinding liquid into a clean 500 mL conical flask (see Table of Materials). Wrap the conical flask mouth with two layers of transparent sealing film, and then place the flask in a shaking incubator (see Table of Materials) for fermentation at 25 &#176;C and 180 rpm for 12 h (Figure 2B).</li>
	<li>Cut out seven layers of square gauze with 10 cm sides (no need for sterility) with scissors and stack them together to form a simple filter device. Slowly pour the fermented scallion liquid onto the gauze and squeeze out the filtered liquid into a new 500 mL conical flask. Wrap the remaining scallion residue on the gauze in tin foil for later use.</li>
	<li>Filter the filtrate obtained in the previous step using a vacuum pump (see Table of Materials) and a Buchner funnel. Put the filter paper wetted by deionized water on the Buchner funnel, and then pour the filtrate and perform suction filtration for three times with different filter paper. Collect the final filtrate in the 500 mL conical flask.</li>
	<li>Perform another filtration using a 0.22 &#181;M filter bottle and a vacuum pump. Screw the cover of the suction filter flask in an ultra-clean bench (see Table of Materials) for further use. At this point, the fermented scallion residue and filtrate are obtained (Figure 2C). 
	NOTE: The filtered liquid obtained from this step will be sterile. (0.22 &#181;M filter bottle is a sterile vacuum filtration system for sterilizing the filtrate).</li>
	<li>Repeat the previous steps to obtain unfermented scallion residue and filtrate. Here the difference is that the fermentation is not needed.</li>
	<li>Weigh 0.24 g choline chloride and 0.56 g L-ascorbic acid (see Table of Materials) in a 10 mL centrifuge tube. Add 3 mL of deionized water and shake vigorously until they are completely dissolved. In the ultra-clean bench, use a new 5 mL syringe and attach three 0.22 &#181;M syringe filters (see Table of Materials) to sterilize the solution, and place it in a sterile 10 mL centrifuge tube for later use. 
	NOTE: Choline chloride is an essential neurotransmitter for the growth of insects, while L-ascorbic acid acts as a preservative.</li>
	<li>Weigh 2 g agar powder and 6 g TSB (see Table of Materials) and pour them into a 500 mL conical flask. Then add 100 mL of deionized water to prepare the culture medium. After rinsing the glass rod with deionized water, use it to stir the mixture until it is well mixed. Then, seal the conical flask using two layers of transparent sealing film.</li>
	<li>Wrap the fermented scallions and unfermented scallions (mentioned in step 1.5 and step 1.8) in aluminum foil and sterilize them, along with the culture medium, in an autoclave at 121 &#176;C for 20 min.</li>
	<li>In the ultra-clean bench, pour the sterilized choline chloride, L-ascorbic acid, and sterile filtrate into the culture medium while gently swirling to mix them thoroughly. Finally, add the fermented and unfermented scallions and shake vigorously by hand to obtain the sterile diets. Pour the diets into the 50 mL centrifuge tubes (sterile; vent cap with a 0.22 &#181;M PVDF hydrophobic membrane) (see Table of Materials) at an angle (Figure 2D). 
	&#8203;NOTE: After sterilization, the temperature of the culture medium should be maintained between 65-80 &#176;C to prevent rapid cooling and solidification when adding scallions or other ingredients. Besides, in each tube, pour approximately 10-15 mL of diets. Once the diets have solidified, store the tubes in a refrigerator at 4 &#176;C, if it is not be used immediately.</li>
</ol>
2. Acquisition of axenic eggs
<ol>
	<li>Laboratory rearing system for D. antiqua
	<ol>
		<li>Place a rearing cage in a 25 &#176;C internally LED illuminated incubator (24 h a cycle, the ratio of light to dark is 16:8) to rear male and female adults, allowing them to mate and lay eggs. Fill the water dispenser (Figure 1B) with water and seal it with moistened cotton wool to provide water for the adults.</li>
		<li>Place four 35 mm x 12 mm Petri dish bottoms in a 94 mm x 16 mm Petri dish lid. Wet cotton wool with ionized water and then place moist cotton wool at two of the Petri dish bottoms, and on top of the cotton wool, add one tablespoon of sucrose.</li>
		<li>Fill the other two Petri dish bottoms with two tablespoons of sucrose and yeast extract (see Table of Materials), which serve as food for the adult insects (Figure 1C). 
		NOTE: The sucrose and yeast extract mentioned in step 2.1.2 and step 2.1.3 do not need to be sterilized.</li>
		<li>Put a 94 mm x 16 mm Petri dish bottom containing moistened sand and a piece of unrooted, peeled garlic clove (Figure 1D) inside the fly cage to collect eggs. The female will be attracted by the smell of the garlic and lay eggs around it.</li>
	</ol>
	</li>
	<li>Collection of eggs
	<ol>
		<li>Take out the oviposition device, which is a 94 mm x 16 mm Petri dish containing sand and garlic mentioned above (step 2.14), from the cage. Pour the sand together with the garlic into a 500 mL beaker, and use tap water to flush the remaining eggs from the garlic into the beaker. At this point, the eggs are floating in the water.</li>
		<li>Take a 100-mesh sieve (see Table of Materials), and moisten it with tap water. Then pour the water containing the floating eggs from the beaker onto the sieve. The eggs will remain on the sieve.</li>
		<li>After the eggs naturally dry on the sieve, use a brush to gently sweep the eggs onto a Petri dish.</li>
	</ol>
	</li>
	<li>On the first day, flush the oviposition device to collect the eggs without disinfection or sterilization. On the second day at four o&#39;clock in the afternoon, flush the eggs again and collect them by sieving them with clean water. 
	NOTE: Collecting the eggs on the second day is to ensure consistent growth rate in the subsequent stages. Four o&#39;clock in the afternoon is the egg-laying peak for D. antiqua. Thus, it is ideal to collect the eggs at this time.</li>
	<li>Place the collected eggs onto a sterile cell sieve (see Table of Materials) in the ultra-clean bench (Figure 3A).</li>
	<li>Prepare the solution for egg sterilization
	<ol>
		<li>Take 5 mL of 5.2% NaClO (see Table of Materials) and transfer it into a sterilized 250 mL conical flask. Add 95 mL of sterilized water to make a total volume of 100 mL, resulting in a 0.26 % NaClO solution.</li>
		<li>To prepare a 75% EtOH solution, take 75 mL of 99.7 % EtOH and transfer it into a sterilized 250 mL conical flask. Add 25 mL of sterilized water to make a total volume of 100 mL, resulting in a 75% EtOH solution.</li>
	</ol>
	</li>
	<li>Pour 30 mL NaClO into a Petri dish and pour 30 mL EtOH into another Petri dish. Place the cell sieve containing eggs mentioned in step 2.4 into the dish containing NaClO. Soak it in the 0.26% NaClO solution for 0.5 min. Then transfer the sieve to the dish containing EtOH, allowing it to soak in the 75% EtOH solution for another 0.5 min.</li>
	<li>Repeat this process three times, alternating between 0.26% NaClO and 75% EtOH every 0.5 min. After that, axenic eggs will be obtained. At this point, the sieve containing the eggs should be submerged in EtOH. 
	NOTE: To ensure thorough sterilization, use a 1 mL pipette to aspirate NaClO and EtOH and disperse the eggs. Repeat the rinsing process multiple times. This step will further clean the surface of the eggs from residual bacteria and impurities, ensuring a clean and sterile surface.</li>
</ol>
3. Rearing of axenic larvae
<ol>
	<li>To transfer the sterilized eggs, use a pair of scissors that have been sterilized with an alcohol lamp to cut the end tip of a 1 mL pipette (Figure 3B). Ensure that the end opening of the cut pipette is larger than the diameter of the eggs (about 2 mm). Then, use the pipette to transfer the eggs from the EtOH solution (eggs together with the solution) (Figure 3C) to a centrifuge tube containing the sterile diets, and each tube should contain approximately 20-50 eggs (Figure 3D). Repeat this process until all the eggs have been transferred. 
	NOTE: The diameter of the pipette tip should be as large as possible to avoid breaking the eggs during transfer, which could lead to their death.</li>
	<li>Use a pipette to remove any excess ethanol from the tube. Place the centrifuge tube closed with lids inside a self-sealing bag, and place a piece of cotton at the opening of the bag to allow for air circulation. Finally, place the bag inside a 25 &#176;C incubator (relative humidity: 50%-70%; photoperiod: 16 h light: 8 h dark) for observation and cultivation.</li>
	<li>Approximately three days later, the axenic eggs in the diets start to hatch (Figure 4A). The hatched larvae will be active and the surface of the sterile diets will no longer be smooth. The larvae will start feeding on the diets.</li>
	<li>Continue to observe the larvae until they reach the stage of 2nd instar larvae. Keep monitoring their growth and development during this period.</li>
	<li>After 9-12 days, more than 80% of the eggs have developed into 3rd instar larvae (Figure 4B). This indicates successful growth and development of the larvae.</li>
</ol>
4. Validation of axenic larvae with culture-dependent assays
<ol>
	<li>Randomly select three axenic 3rd instar larvae from the centrifuge tube.</li>
	<li>Place the larvae in a 94 mm x 16 mm Petri dish containing 75% alcohol for cleaning. After that, transfer them to a container with sterile water for further rinsing.</li>
	<li>Use a disinfected dissecting forceps that has been sterilized with 75% EtOH solution to grasp the larvae (the head and tail are shown in Figure 5A). Use dissecting scissors to remove the head and tail ends (Figure 5B). Hold the tail of the larvae with the forceps and use another forceps to gently squeeze from the tail to the head and extract the internal organs (the result shown in Figure 5C). Place the internal organs in deionized water and gently pull off the intestine by pressing on the adipose tissue with forceps, and place the extracted internal (Figure 5D) in sterile water for further use.</li>
	<li>Place the intestine into a sterile 1.5 mL centrifuge tube, add 500 &#181;L of sterilized 1x PBS buffer, and use a disposable pestle to grind the intestinal tissue into a homogenate.</li>
	<li>Take 100 &#181;L of the homogenate and add it to a nutrient agar medium. Use an L-shaped spreader (see Table of Materials) for streaking the agar plate.</li>
	<li>Place the Petri dish in a biochemical incubator at 28 &#176;C and incubate for 72 h to observe the growth of colonies.</li>
</ol>
5. Validation of axenic larvae with culture-independent assays
<ol>
	<li>Extract total DNA from the gut tissue of axenic larvae using a DNA extraction kit (see Table of Materials) following manufacturer&#39;s instructions.</li>
	<li>Measure the DNA concentration using a UV spectrophotometer.</li>
	<li>Perform PCR reaction using universal bacterial 16S rRNA primers (1492R: 5&#39;- GGTTACCTTGTTACGACTT -3&#39;, 27F: 5&#39;- AGAGTTTGATCATGGCTCAG -3&#39;). The total reaction volume is 25 &#181;L, and it consists of 1 &#181;L DNA template, 0.5 &#181;L forward primer, 0.5 &#181;L reverse primer, 10.5 &#181;L ddH2O, and 12.5 &#181;L 2x Taq PCR Master Mix (see Table of Materials). Set the PCR reaction conditions as follows: initial denaturation at 94 &#176;C for 3 min; 35 cycles of denaturation at 94 &#176;C for 30 s, annealing at 55 &#176;C for 30 s, and extension at 72 &#176;C for 90 s; final extension at 72 &#176;C for 10 min. Store the PCR products at 4 &#176;C for further analysis.</li>
	<li>Perform PCR reaction using universal fungal ITS primers (ITS1: 5&#39;- TCCGTAGGTGAACCTGCGG -3&#39;, ITS4: 5&#39;- TCCTCCGCTTATTGATATGC -3&#39;). The total reaction volume is 25 &#181;L, and it consists of 1 &#181;L DNA template, 0.5 &#181;L forward primer, 0.5 &#181;L reverse primer, 10.5 &#181;L ddH2O, and 12.5 &#181;L 2x Taq PCR Master Mix. Set the PCR conditions at 95 &#176;C for 5 min; 35 cycles of 95 &#176;C for 30 s, 55 &#176;C for 1 min, and 72 &#176;C for 90 s; followed by 72 &#176;C for 10 min. Store the PCR products at 4 &#176;C for further analysis.</li>
	<li>Use Super Green nucleic acid dye (see Table of Materials) to mix with each of the two types of PCR products evenly, and analyze them by electrophoresis on a 1% agarose gel in 1x TAE buffer (diluted from 50x TAE, see&#160;Table of Materials). Use 3 &#181;L of DNA marker (see Table of Materials) as a reference.</li>
	<li>Use a UV transilluminator to observe the gel and locate the target fragment.</li>
</ol>

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

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+red+palm+weevil+Rhynchophorus+Ferrugineus+(Olivier)+(Coleoptera:+Dryophthoridae)+germ-free+larvae+for+host-gut+microbes+interaction+studies.">Preparation of red palm weevil Rhynchophorus Ferrugineus (Olivier) (Coleoptera: Dryophthoridae) germ-free larvae for host-gut microbes interaction studies.</a> Bio-Protocol. 9 (24), e3456 (2019).</li><li>Bavani, M. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sterilization+of+Lucilia+sericata+(Diptera:+Calliphoridae)+Eggs+for+maggot+debridement+therapy.">Sterilization of Lucilia sericata (Diptera: Calliphoridae) Eggs for maggot debridement therapy.</a> Journal of Medical Entomology. 59 (3), 1076-1080 (2022).</li><li>Han, L. Z., Li, S. B., Liu, P. L., Peng, Y. F., Hou, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+artificial+diet+for+continuous+rearing+of+Chilo+suppressalis+(Lepidoptera:+Crambidae).">New artificial diet for continuous rearing of Chilo suppressalis (Lepidoptera: Crambidae).</a> Annals of the Entomological Society of America. 105 (2), 253-258 (2012).</li><li>Bezerra, C. E. S., Amaral, B. B., Souza, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearing+Chrysoperla+externa+larvae+on+artificial+diets.">Rearing Chrysoperla externa larvae on artificial diets.</a> Neotropical Entomology. 46 (1), 93-99 (2017).</li><li>Feng, H. Q., Jin, Y. L., Li, G. P., Feng, H. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+of+an+artificial+diet+for+successive+rearing+of+Apolygus+lucorum+(Hemiptera:+Miridae).">Establishment of an artificial diet for successive rearing of Apolygus lucorum (Hemiptera: Miridae).</a> Journal of Economic Entomology. 105 (6), 1921-1928 (2012).</li><li>Hassan, B., Siddiqui, J. A., Xu, Y. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vertically+transmitted+gut+bacteria+and+nutrition+influence+the+immunity+and+fitness+of+Bactrocera+dorsalis+larvae.">Vertically transmitted gut bacteria and nutrition influence the immunity and fitness of Bactrocera dorsalis larvae.</a> Frontiers in Microbiology. 11, 596352 (2020).</li><li>Li, X. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+the+intestinal+bacterial+community+in+black+soldier+fly+larval+guts+and+its+influence+on+insect+growth+and+development.">Dynamics of the intestinal bacterial community in black soldier fly larval guts and its influence on insect growth and development.</a> Insect Science. 30 (4), 947-963 (2023).</li><li>Moran, N. A., McCutcheon, J. P., Nakabachi, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genomics+and+Evolution+of+heritable+bacterial+symbionts.">Genomics and Evolution of heritable bacterial symbionts.</a> Annual Review of Genetics. 42, 165-190 (2008).</li><li>Weinert, L. A., Araujo-Jnr, E. V., Ahmed, M. Z., Welch, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+incidence+of+bacterial+endosymbionts+in+terrestrial+arthropods.">The incidence of bacterial endosymbionts in terrestrial arthropods.</a> Proceedings of the Royal Society B-Biological Sciences. 282 (1807), 20150249 (2015).</li><li>Weeks, A. R., Turelli, M., Harcombe, W. R., Reynolds, K. T., Hoffmann, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+parasite+to+mutualist:+Rapid+evolution+of+Wolbachia+in+natural+populations+of+Drosophila.">From parasite to mutualist: Rapid evolution of Wolbachia in natural populations of Drosophila.</a> PLOS Biology. 5 (5), 997-1005 (2007).</li></ol>

The authors have no conflicts of interest to disclose.

Insects possess a highly complex gut microbiota20,21, necessitating the use of axenic insects inoculated with specific gut microbial strains for studying insect-microorganism interactions. The preparation of axenic insects is crucial for such research endeavors. Antibiotic treatment is a method used to eliminate gut microbiota. For example, Jung and Kim22 fed Spodoptera exigua with penicillin, while Raymond23</su...

Axenic animals, defined as animals in which no viable microorganisms or parasites can be detected, are valuable experimental models for studying host-microorganism interactions1,2. Insects, the largest group of invertebrates, can form symbiotic relationships with microorganisms3. Axenic insects can be used to study host-symbiont interactions in symbiotic systems4. For example, Nishide et al.5 established a practical sterile rearing procedure for the malodor worm Plautia stali, enabling reliable and rigorous analysis of host-symbiont interactions in model symbiotic systems. Axenic insects can be produced by sterilizing the egg stage and providing sterile food to the larvae and adults6,7. Axenic insects are of great significance and are widely used in biological research. For instance, a study conducted by Somerville et al.8 demonstrated that diamondback moths inoculated with Enterobacter cloacae improved the adaptability of transgenic males.
Delia antiqua Meigen is an economically important pest of onions and other Liliaceae crops worldwide, with its larvae damaging the bulbs of onions and other Liliaceae crops9. D. antiqua is mainly found in temperate climates and is widespread in onion-cultivation areas of the Americas, Europe, and Asia. If not properly controlled, it can cause crop losses in onions (Allium cepa L.), garlic (Allium sativum L.), shallots (Allium fistulosum L.), and leeks (Alliumchoenoprasum L.) ranging from 50% to 100%10,11. The larvae feed on the below-ground parts of plants, and this feeding causes the seedlings to wilt and eventually die. In addition, damaged plants can allow pathogens to enter, leading to bulb rotting12. Even if the plants are not completely consumed by the larvae, the damage they cause renders the onion plants unmarketable and results in economic losses.
Insects are closely associated with gut microbiota, and most insect guts contain a variety of symbiotic bacteria that thrive on the nutrients provided by the host13,14. Jing et al.15 showed that the primary function of the intestinal symbiotic community is to provide essential nutrients, followed by functions related to digestion and detoxification. In certain cases, gut bacteria can serve as a microbial resource for pest management purposes. Consequently, studying the individual gut bacteria&#39;s performances and specific functions within the body of D. antiqua is desirable. Therefore, preparing axenic larvae is particularly important for studying the interactions between specific bacterial strains and insects16. Currently, a commonly used method to eliminate insect gut bacteria is the use of an antibiotic combination to eradicate associated microbes17,18,19. Unlike using antibiotics alone, which can only reduce microbial numbers, axenic rearing of insects allows for control over the composition and quantity of microorganisms, enabling more accurate validation of gut microbiota functionality.
Thus, this article introduces a protocol for preparing and rearing axenic D. antiqua. Axenic larval food is obtained by utilizing high-temperature sterilization of natural diets combined with half-fermented foods. The eggs are sterilized following an experimental protocol to obtain axenic eggs, and finally, axenic larvae are cultured from the axenic eggs. The axenic rearing system was carried out for only one generation for the experiment. This will provide convenience for studying the interaction between insects and gut microbiota.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.22 &#956;M filter bottle</td>
			<td>Thermo Scientific</td>
			<td>450-0045</td>
			<td></td>
		</tr>
		<tr>
			<td>0.22 &#956;M Syringe Filter</td>
			<td>Biosharp</td>
			<td>BS-QT-011</td>
			<td></td>
		</tr>
		<tr>
			<td>100-mesh sieve</td>
			<td>Zhejiang Shangyu Jinding Standard Sieve Factory</td>
			<td>No Catalog numbers</td>
			<td></td>
		</tr>
		<tr>
			<td>1x PBS solution</td>
			<td>Solarbio</td>
			<td>P1020</td>
			<td></td>
		</tr>
		<tr>
			<td>2x Taq PCR Master Mix</td>
			<td>GENVIEW</td>
			<td>GR1113-1ML</td>
			<td></td>
		</tr>
		<tr>
			<td>5.2% NaClO solution</td>
			<td>Sinopharm Chemical Reagent Co., Ltd.</td>
			<td>80010428</td>
			<td></td>
		</tr>
		<tr>
			<td>500 mL Conical flask</td>
			<td>Thermo Scientific</td>
			<td>4103-0500</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL vented centrifuge tube</td>
			<td>JET BIOFIL</td>
			<td>BRT-011-050</td>
			<td></td>
		</tr>
		<tr>
			<td>50x TAE buffer</td>
			<td>GENVIEW</td>
			<td>GT1307</td>
			<td></td>
		</tr>
		<tr>
			<td>Agar powder</td>
			<td>Ding Guo</td>
			<td>DH010-1.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Biochemical incubator</td>
			<td>STIK</td>
			<td>21040121500010</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell sieve</td>
			<td>SAINING</td>
			<td>5022200</td>
			<td></td>
		</tr>
		<tr>
			<td>Choline chloride</td>
			<td>Sangon Biotech</td>
			<td>A600299-0100</td>
			<td></td>
		</tr>
		<tr>
			<td>ddH2O</td>
			<td>Ding Guo</td>
			<td>PER018-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable grinding pestle</td>
			<td>JET BIOFIL</td>
			<td>CSP-003-002</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA extraction kit</td>
			<td>Sangon Biotech</td>
			<td>B518221-0050</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA Marker</td>
			<td>Sangon Biotech</td>
			<td>B600335-0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol absolute</td>
			<td>Sinopharm Chemical Reagent Co., Ltd.</td>
			<td>10009218</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>NEWSTAR</td>
			<td>1087309025</td>
			<td></td>
		</tr>
		<tr>
			<td>Food processor</td>
			<td>Guangdong Midea Life Electric Appliance Manufacturing Co., Ltd.</td>
			<td>WBL25B26</td>
			<td></td>
		</tr>
		<tr>
			<td>Illuminated&#160; incubator</td>
			<td>Shanghai ESTABLISH Instrumentation Co., Ltd.</td>
			<td>A16110768</td>
			<td></td>
		</tr>
		<tr>
			<td>L-Ascorbic acid</td>
			<td>Sangon Biotech</td>
			<td>A610021-0100</td>
			<td></td>
		</tr>
		<tr>
			<td>L-shaped spreader</td>
			<td>SAINING</td>
			<td>6040000</td>
			<td></td>
		</tr>
		<tr>
			<td>Nutrient agar medium</td>
			<td>Hope Bio</td>
			<td>HB0109</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Bing Yu</td>
			<td>&#160;BY-103</td>
			<td>Purchase on Jingdong</td>
		</tr>
		<tr>
			<td>Shock incubator</td>
			<td>Shanghai Zhichu Instrument Co., Ltd.</td>
			<td>2020000014</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>GENVIEW</td>
			<td>CS326-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Super Green nucleic acid dye</td>
			<td>Biosharp</td>
			<td>BS355A</td>
			<td></td>
		</tr>
		<tr>
			<td>Super-clean table</td>
			<td>Heal Force</td>
			<td>AC130052</td>
			<td></td>
		</tr>
		<tr>
			<td>TSB</td>
			<td>Hope Bio</td>
			<td>HB4114</td>
			<td></td>
		</tr>
		<tr>
			<td>Vacuum pump</td>
			<td>Zhejiang Taizhou Seeking Precision Vacuum Pump Co., Ltd.</td>
			<td>22051031</td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast extract</td>
			<td>Thermo Scientific</td>
			<td>LP0021B</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The life stages of D. antiqua are depicted in Figure 4. The complete life cycle comprises eggs, larvae, pupa (Figure 4C), and adults (Figure 4D). They are cultivated in sterile centrifuge tubes, and their appearance and survival rate are indistinguishable from D. antiqua raised under non-axenic conditions. The growth and development times for each stage of D. antiqua can be found in Fi...

Watch this Scientific Journal Video about Axenic Rearing of Delia antiqua to Detect the Impact of Intestinal Microbes on Its Growth and Development at JoVE.com

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.

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 ...

rearing-axenic-delia-antiqua-with-half-fermented-sterile-diets

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Biology

536 Views.  Qilu University of Technology (Shandong Academy of Sciences). 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.

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

Protocol for Mosquito Rearing (A. gambiae)

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night cycle. For this procedure, you'll need mosquito cages, 10% sterile sucrose solution, paper towels, beaker, whatman filter paper, glass feeders, human blood and serum, water bath, parafilm, distilled water, clean plastic trays, mosquito food (described below), mosquito net to cover the trays, vacuum, and a collection chamber to collect adults.

Protocol for Mosquito Rearing A. gambiae

This video illustrates the general techniques used to rear Anopheles gambiae in the laboratory. The methods for caring for laboratory mosquitoes are demonstrated through all stages of the organism's life cycle from larvae to pupae to blood-feeding adults.

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night ...

Culturing Caenorhabditis elegans in Axenic Liquid Media and Creation of Transgenic Worms by Microparticle Bombardment

In this protocol, we present the required materials, and the procedure for making modified&#160;C. elegans&#160;Habituation and Reproduction media (mCeHR). Additionally, the steps for exposing and acclimatizing C. elegans grown on E. coli to axenic liquid media are described. Finally, downstream experiments that utilize axenic C. elegans illustrate the benefits of this procedure. The ability to analyze and determine C. elegans&#160;nutrient requirement was illustrated by growing N2 wild type worms in axenic liquid media with varying heme concentrations. This procedure can be replicated with other nutrients to determine the optimal concentration for worm growth and development or, to determine the toxicological effects of drug treatments. The effects of varied heme concentrations on the growth of wild type worms were determined through qualitative microscopic observation and by quantitating the number of worms that grew in each heme concentration. In addition, the effect of varied nutrient concentrations can be assayed by utilizing worms that express fluorescent sensors that respond to changes in the nutrient of interest. Furthermore, a large number of worms were easily produced for the generation of transgenic C. elegans using microparticle bombardment.

Culturing Caenorhabditis elegans in Axenic Liquid Media and Creation of Transgenic Worms by Microparticle Bombardment

C. elegans is usually grown on solid agar plates or in liquid cultures seeded with E. coli. To prevent bacterial byproducts from confounding toxicological and nutritional studies, we utilized an axenic liquid medium, CeHR, to grow and synchronize a large number of worms for a range of downstream applications.

In this protocol, we present the required materials, and the procedure for making modified&#160;C. elegans&#160;Habituation and Reproduction media ...

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in agronomic practices resulted in serious outbreaks of stable flies in several countries. These outbreaks disrupted livestock production and human recreation resulting in public demands for increasing research and management efforts for this pest. A simple and inexpensive procedure for rearing stable flies for laboratory studies is presented. The procedure uses locally available diet components, equipment and supplies. The procedure can be adapted for rearing other muscoid flies including face fly (Musca autumnalis), horn fly (Haematobia irritans), and house fly (Musca domestica). The procedure produces stable fly puparia averaging 12.5 mg and ~35% egg to adult survival. Approximately 3000 flies are produced in each pan.

A procedure for rearing stable flies (Stomoxys calcitrans) is presented. The procedure uses locally available materials for diet components, equipment and supplies.

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in ...

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 ...

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.

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 ...

Enhancing Survival and Development of Marmosets Using a Hand-Rearing Approach

Hand-Rearing Method for Infant Marmosets

The common marmoset (Callithrix jacchus) is a small and highly social New World monkey with high reproduction rates, which has been proven to be a compelling non-human primate model for biomedical and neuroscience research. Some females give birth to triplets; however, the parents cannot raise all of them. To save these infants, we have developed a hand-rearing method for raising newborn marmosets. In this protocol, we describe the formula of the food, the time for feeding, the configuration of the temperature and humidity, as well as the adaptation of the hand-reared infants to the colony environment. This hand-rearing method significantly increases the survival rate of marmoset infants (without hand-rearing: 45%; with hand-rearing: 86%) and provides the opportunity to study the development of marmoset infants with similar genetic backgrounds raised in different postnatal environments. As the method is practical and easy to use, we anticipate that it could also be applied to other labs working with common marmosets.

Author Spotlight: Marmoset Research - Scope and Challenges 

Here, we describe a hand-rearing method for raising infant marmosets in an animal incubator. This method greatly increases the survival rate of marmoset infants, which provides the opportunity to study the development of marmoset infants with similar genetic backgrounds raised in different postnatal environments.

The common marmoset (Callithrix jacchus) is a small and highly social New World monkey with high reproduction rates, which has been proven to be a ...

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)  

Artificial Rearing and Heavy Metal Tolerance in Cabbage White Butterflies

Rearing the Cabbage White Butterfly (Pieris rapae) in Controlled Conditions: A Case Study with Heavy Metal Tolerance

The cabbage white butterfly (Pieris rapae) is an important system for applied pest control research and basic research in behavioral and nutritional ecology. Cabbage whites can be easily reared in controlled conditions on an artificial diet, making them a model organism of the butterfly world. In this paper, a manipulation of heavy metal exposure is used to illustrate basic methods for rearing this species. The general protocol illustrates how butterflies can be caught in the field, induced to lay eggs in greenhouse cages, and transferred as larvae to artificial diets. The methods show how butterflies can be marked, measured, and studied for a variety of research questions. The representative results give an idea of how artificial diets that vary in components can be used to assess butterfly performance relative to a control diet. More specifically, butterflies were most tolerant to nickel and least tolerant to copper, with a tolerance of zinc somewhere in the middle. Possible explanations for these results are discussed, including nickel hyper-accumulation in some mustard host plants and recent evidence in insects that copper may be more toxic than previously appreciated. Finally, the discussion first reviews variations to the protocol and directions for troubleshooting these methods, before considering how future research might further optimize the artificial diet used in this study. Overall, by providing a detailed video overview of the rearing and measurement of cabbage whites on artificial diets, this protocol provides a resource for using this system across a wide range of studies.

Author Spotlight: Investigating the Tolerance of Cabbage Butterflies to Urban Pollutants 

This paper presents a detailed protocol for rearing the cabbage white butterfly in controlled lab conditions with an artificial diet, which allows precise manipulations of early-life nutrition and toxin exposure. The representative results show how heavy metal toxicity can be assayed with this protocol.

The cabbage white butterfly (Pieris rapae) is an important system for applied pest control research and basic research in behavioral and nutritional ...