We thank Drs. Alison Heffer and Yong Lu for setting up the microinjection apparatus and sharing their invaluable knowledge and experience with insect RNAi. This work was supported by the National Institutes of Health (R01GM113230 to L.P.).

1. Rearing of D. maculatus
NOTE: A breeding colony of D. maculatus was set up in the authors&#39; lab using adults and larvae purchased commercially. The species identity was verified using DNA barcoding53.
<ol>
	<li>To set up a new cage in the lab, spread a thin layer of wood shavings into a medium-size insect cage (30.5 x 19 x 20.3 cm3). Place a ~ 10 x 6 x 3 cm3 chunk of Styrofoam in the cage to let larvae hide for pupation. Add 20 - 50 beetles (either adults or late instar larvae). Beetles will hide in the wood shavings.</li>
	<li>Add wet cat food in a Petri dish or a weighing boat. Cover the cage with mesh cloth and place the cage in an incubator.</li>
	<li>For maintaining a breeding colony in the lab, set the temperature between 25 and 30 &#176;C. D. maculatus grows faster at higher temperatures, but this also promotes the growth of fungus and mites. To maintain a healthy colony, use 25 - 28 &#176;C for regular stock maintenance and 30 &#176;C for rapidly expanding the colony. The life cycle takes approximately three weeks to four months depending on environmental factors50 (Figure 2).</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54976/54976fig2.jpg" /> 
Figure 2: D. maculatus Lab Colony. Photograph of a typical D. maculatus insect cage is shown. Wood shavings are spread to let the beetles hide. Cat food is added in a small Petri dish or weighing boat. Styrofoam is placed into the cage as a refuge for final instar larvae to pupate. The cage shown here is 30.5 x 19 x 20.3 cm3 and houses a few hundred larvae, pupae, and adults. <a href="http://cloudfront.jove.com/files/ftp_upload/54976/54976fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="4">
	<li>Leave eggs in the cage to mature. To expand the colony, establish new cages with eggs (collected as described in Section 2), larvae, or adults, as above. D. maculatus lay eggs throughout the cage, especially near the food source.</li>
	<li>Replace wet cat food about twice a week. 
	NOTE: Due to the unpleasant odor of wet cat food, alternative food sources were also tested (see Discussion).</li>
</ol>
2. Embryo Collection
<ol>
	<li>To set up a collection, separate at least 50 males and 50 females from the colony. Young adults are best. Place adults in a mini-sized cage (17.8 x 10.2 x 12.7 cm3) without wood shavings. Add cat food in a weighing boat.</li>
	<li>Before starting collection, check the cage carefully for any embryos present and remove them. Put a stretched cotton ball into the cage, and leave the cage in a 30 &#176;C incubator. After the appropriate time window, the cotton ball will be ready for embryo collection.</li>
	<li>Fold a piece of black construction paper (A4 size) in half to create a crease, and then unfold.</li>
	<li>Remove the stretched cotton ball from the cage. Remove adults from the cotton ball and put them back into the cage.</li>
	<li>While pinching the stretched cotton ball very gently, tear it apart slowly into thin cotton filaments to let the eggs fall onto the black paper. 
	NOTE: D. maculatus embryos are fragile and hard to see in cotton. They can be easily crushed if holding the cotton ball too firmly.</li>
</ol>
3. dsRNA Preparation
<ol>
	<li>DNA Template Preparation for dsRNA Synthesis
	<ol>
		<li>Run 4 - 6 50 &#181;l PCR reactions using primers containing T7 promoter sequences at both 5&#39; ends to amplify DNA template according to manufacturer&#39;s protocol.</li>
		<li>Purify PCR product using a commercial PCR purification kit, following manufacturer&#39;s instructions. The ideal final concentration of DNA template is &#8805; 100 ng/&#181;l.</li>
	</ol>
	</li>
	<li>Injection Buffer Preparation
	<ol>
		<li>Prepare 100 ml of injection buffer (0.1 mM NaH2PO4, 5 mM KCl). Adjust pH to 6.8 with 5 M NaOH.</li>
		<li>Store aliquots at -20 &#176;C.</li>
	</ol>
	</li>
	<li>dsRNA Synthesis
	<ol>
		<li>Set up a reaction in a 0.2 ml PCR tube on ice and carry out an in vitro transcription reaction with T7 polymerase, following manufacturer&#39;s instructions, using ~ 500 ng template per reaction.</li>
		<li>Incubate the tube at 37 &#176;C overnight.</li>
		<li>Digest the DNA template with DNase, following manufacturer&#39;s instructions.</li>
		<li>Heat the tube to 94 &#176;C and hold at 94 &#176;C for 3 min.</li>
		<li>To anneal the dsRNA, slowly cool the tube by 1 &#176;C/min until it reaches 45 &#176;C after 1 hr. 
		NOTE: Steps 3.3.2 through 3.3.5 can be performed in a thermocycler.</li>
		<li>To ethanol precipitate&#160;of dsRNA, add 280 &#181;l of RNase free water to 20 &#181;l of reaction to adjust to a final volume of 300 &#181;l. Add 30 &#181;l of 3 M NaOAc (pH 5.2) and 650 &#181;l of ethanol. Place tube in -20 &#176;C freezer overnight.</li>
		<li>After centrifuging at 15,682 x g for 30 min at 4 &#176;C, wash the dsRNA pellet with 70% ethanol. Dry pellet and dissolve in 20 &#181;l injection buffer.</li>
		<li>Measure the concentration of the dsRNA using a spectrophotometer at A260. The ideal concentration is 3 - 5 &#181;g/&#181;l. Check the quality by running 5 &#181;l of a 1:25 dilution of dsRNA on a 1% agarose gel at 90 V for 30 min. A single band of expected size will be readily visible.</li>
		<li>Store the dsRNA at -20 &#176;C or colder. 
		NOTE: For every RNAi knockdown experiment, include a control dsRNA, which would not be expected to impact the process being studied. gfp dsRNA is an excellent control for most experiments. Prepare and inject the control dsRNA side-by-side with the experimental dsRNA.</li>
	</ol>
	</li>
</ol>
4. Assembly of Microinjector and Micromanipulator
NOTE: See Figure 3.
<ol>
	<li>Connect nitrogen or other air supply to the pressure input of a pneumatic pump instrument. If a pneumatic pump is not available, apply pressure using a 50 ml syringe connected to fine tubing.</li>
	<li>Connect a foot switch to the remote connector of the pump to control the eject pressure flow.</li>
	<li>Attach a glass capillary holder to the eject pressure port of the pump with tube.</li>
	<li>Fix the capillary holder on a micromanipulator close to a dissecting microscope.</li>
</ol>
5. Embryonic RNAi
NOTE: Figure 4 shows a flowchart of D. maculatus embryonic RNAi.
<ol>
	<li>eRNAi Preparation
	<ol>
		<li>Prepare an embryo collecting Petri dish lid (90 mm diameter). Place a piece of black filter paper to cover the inside of the lid. Put a standard microscope slide on the black paper.</li>
		<li>Dilute dsRNA to appropriate concentration with injection buffer. Use 2 - 3 &#181;g/&#181;l for initial experiments. Always keep dsRNA on ice prior to injection.</li>
		<li>Add food coloring at 1:40 dilution to the dsRNA and pipette gently several times to mix well.</li>
	</ol>
	</li>
	<li>Collecting Embryos for Injection 
	NOTE: At 25 &#176;C, nuclei in embryos 6 - 8 hr After Egg Laying (AEL) migrate towards the egg periphery53. A cellular blastoderm is established within 8 - 10 hr AEL53. To ensure embryos prior to cellular blastoderm stage are used for injection, 0 - 3 hr AEL embryos are recommended for use53. Embryo collection, preparation, and injection usually take less than 2 hr if the glass capillaries are in good shape. Therefore, the whole process can be completed within 5 hr AEL.
	<ol>
		<li>Collect embryos as described in steps 2.2-2.5.</li>
		<li>Tap the black construction paper to transfer embryos into collecting Petri dish lid.</li>
		<li>Stick double-sided tape along the edge of the slide.</li>
		<li>Using a paint brush, align the embryos on the tape perpendicular to the slide edge with the anterior or posterior end at the edge. Note that the anterior and posterior ends of early embryos are not readily distinguishable, thus on average half will be injected at the posterior end which is ideal.</li>
	</ol>
	</li>
	<li>Loading the dsRNA into the Glass Capillary
	<ol>
		<li>Take up 2 - 4 &#181;l of dsRNA into a 20 &#181;l microloader pipette tip.</li>
		<li>Insert the tip into a pre-pulled glass capillary (referred to as needle) and gently pipette the dsRNA solution into the capillary. Prepare needles from commercial glass capillaries using a micropipette puller. An example of an ideal pulled needle is shown in Figure 5. The length and taper of needles may need to be optimized after injection trials.</li>
		<li>Fix the glass capillary into the glass capillary holder.</li>
		<li>Assemble the capillary holder into the micromanipulator.</li>
	</ol>
	</li>
	<li>Injecting dsRNA into Embryos
	<ol>
		<li>Carefully transfer the slide with embryos onto the stage of the dissecting microscope.</li>
		<li>Move the slide to bring one embryo to the center of the field and focus microscope.</li>
		<li>Position the tip of the capillary with the micromanipulator while looking into the dissecting microscope. Bring the tip close to the end of the first embryo in the row.</li>
		<li>Switch on the nitrogen or other air supply.</li>
		<li>Set the solenoid input selector switch to vacuum.</li>
		<li>Adjust the eject pressure regulator to 10 - 15 psi. Adjust the pressure depending on the apparatus.</li>
		<li>Open the capillary if necessary. Once the tip is open, the colored dsRNA solution will fill the tip.</li>
		<li>Move the capillary tip forward to puncture the embryo. If the pressure setting is ideal, no embryonic fluid will flow into the capillary.</li>
		<li>Step on the foot switch to eject dsRNA solution into the embryo until an appropriate amount of solution is ejected. Figure 5 shows examples of embryo morphology after appropriate and inappropriate amounts of solution have been injected.</li>
		<li>Keep the capillary tip inside the embryo for ~ 2 sec, and then remove it.</li>
		<li>Move the capillary to the next embryo and repeat steps 5.4.8 - 5.4.10 until all embryos are injected. Change glass capillary if it gets clogged or breaks.</li>
	</ol>
	</li>
	<li>Post-injection Recovery and Incubation
	<ol>
		<li>After injection, place the slide in a Petri dish.</li>
		<li>Add a wet cotton ball to the Petri dish. Do not let the cotton ball touch the slide.</li>
		<li>Cover the Petri dish and wrap it with sealing film. Label the dish with the name of the dsRNA, concentration, date, time, and number of injected embryos.</li>
		<li>Place the Petri dish in a 30 &#176;C incubator until the embryos hatch.</li>
	</ol>
	</li>
</ol>
6. Parental RNAi
<ol>
	<li>Isolating Pupae for pRNAi
	<ol>
		<li>Collect pupae from the colony.</li>
		<li>Sort male and female pupae under microscope (see Figure 6). Keep them in two separate Petri dishes in a 30 &#176;C incubator to ensure that no mating occurs prior to injection.</li>
		<li>Check every day for eclosed adults. Pupation usually takes 5 - 7 days at 30 &#176;C.</li>
		<li>Transfer eclosed males and females (see Figure 6) to two separate Petri dishes and feed them cat food every other day until they are ready for injection.</li>
		<li>Proceed to injection once there are enough females and males at appropriate age. Females 4 to 8 days post-eclosion are best. For a typical analysis, 8 - 12 females are used. An equal number of males are needed for mating54.</li>
	</ol>
	</li>
	<li>Injecting dsRNA into Female Abdomen
	<ol>
		<li>Prepare dsRNA on ice (5.1.2 and 5.1.3).</li>
		<li>Attach a 32-gauge needle to a 10 &#181;l syringe.</li>
		<li>Anaesthetize females on CO2 stage.</li>
		<li>Load dsRNA solution into the syringe without taking up any air.</li>
		<li>Hold a female ventral side up with one hand and hold the syringe with the other.</li>
		<li>Gently penetrate the segmacoria (membrane) between sternites 2 and 3 with the tip of the needle. Figure 7 shows a female during injection. If the needle does not penetrate the tissue easily, angle the needle upward and slowly press down. Insert approximately 2 mm of the needle into the body.</li>
		<li>Check the syringe scale while slowly pushing the plunger, injecting ~ 2 &#181;l per female.</li>
		<li>After injecting, hold the needle still for at least 5 sec before removing it from the female&#39;s abdomen.</li>
		<li>Transfer the injected females to a Petri dish (90 mm diameter) and feed with cat food.</li>
		<li>Label the Petri dish with the dsRNA name, amount injected, date, and number of females.</li>
	</ol>
	</li>
	<li>Post-injection Recovery and Mating
	<ol>
		<li>Incubate the Petri dish in a 30 &#176;C incubator.</li>
		<li>After 24 hr, transfer injected females to a mini cage.</li>
		<li>Add to the cage an equal number of uninjected young males, and label the cage.</li>
		<li>Add a weighing boat with cat food into the cage.</li>
		<li>Leave the cage in a 30 &#176;C incubator to allow mating. After 24 hr, females should start to lay eggs and embryos can be collected daily or at required time intervals.</li>
	</ol>
	</li>
</ol>
7. Phenotypic Analysis after RNAi
NOTE: At 30 &#176;C, it takes ~ 55 hr for eggs to hatch50.
<ol>
	<li>eRNAi Viability Analysis
	<ol>
		<li>Calculate hatch rate by comparing the number of hatched larvae to the total number of injected embryos. Be sure to include a negative control injection as embryos may be harmed or killed by the injection procedure. Typical hatch rates vary from 30% - 60%. Beware of hatched larvae eating unhatched eggs.</li>
	</ol>
	</li>
	<li>pRNAi Viability Analysis 
	NOTE: If female viability is impacted by the gene targeted by RNAi, females injected with specific dsRNA but not negative control dsRNA will begin to die. If egg production or egg laying is impacted, females will exhibit partial or complete sterility. Score female survival compared to negative controls or compare total number of eggs laid by experimental and control females (7.2.3).
	<ol>
		<li>Set up embryo collection following step 2.2.</li>
		<li>After 24 hr, collect embryos following steps 2.4 - 2.5.</li>
		<li>Count the total number of the embryos.</li>
		<li>Transfer the embryos to a 90 mm Petri dish. Label the Petri dish with the gene targeted, collection date, and number of embryos.</li>
		<li>Incubate the embryos in a 30 &#176;C incubator until they hatch. Since hatched larvae feed on unhatched embryos, check the Petri dish frequently and separate hatched larvae from unhatched embryos using forceps (see Discussion).</li>
		<li>Count the number of hatched embryos and calculate the hatch rate.</li>
	</ol>
	</li>
	<li>Examining Cuticle Defects in Hatched Larvae
	<ol>
		<li>Collect hatched larvae in a 1.5 ml microcentrifuge tube.</li>
		<li>In a fume hood, add 1 ml of Fixation Solution.</li>
		<li>Leave the microcentrifuge tube at 4 &#176;C at least overnight to let the solution penetrate the larval tissue.</li>
		<li>To visualize, remove as much fixative from the tube as possible.</li>
		<li>Rinse larvae at least three times with PBST.</li>
		<li>Transfer larvae to a multi-well glass plate with PBST using a P-1000 pipette tip with the end cut off to widen it.</li>
		<li>Under a dissecting microscope, stretch larvae using forceps to examine defects. It is critical to stretch out larvae to examine defects as their bodies contract in fixative.</li>
	</ol>
	</li>
	<li>Examining Cuticle Defects in Unhatched Larvae 
	Note: This procedure is useful for examination of early embryonic phenotypes, especially for embryos that do not survive to hatching.
	<ol>
		<li>For pRNAi, add PBST into the Petri dish (7.2.4) and transfer unhatched embryos to a 1.5 ml microcentrifuge tube using a P-1000 pipette tip with the end cut off. For eRNAi, add PBST into the Petri dish (5.5.4), brush unhatched embryos off the microscope slide and transfer them to a microcentrifuge tube.</li>
		<li>Fix the embryos with Fixation Solution and rinse them with PBST for visualization following steps 7.3.2 - 7.3.6.</li>
		<li>Using forceps, dissect embryos out of the eggshell under a dissecting microscope in PBST.</li>
		<li>Transfer the embryos back to 1.5 ml microcentrifuge tube (~ 200 &#181;l of embryos in each tube).</li>
		<li>Remove as much solution as possible.</li>
		<li>Add 1 ml of 90% lactic acid/10% EtOH. Leave the centrifuge tube in a 60 &#176;C incubator at least overnight. 
		NOTE: Embryos can be left at 60 &#176;C for several days.</li>
		<li>To mount the embryos, pipette them onto a microscope slide using a P-1000 pipette tip with the end cut off.</li>
		<li>Manually position the embryos with forceps to an ideal orientation.</li>
		<li>Cover the embryos with a cover-slip and visualize under microscope using DIC.</li>
	</ol>
	</li>
	<li>Examining Cellular and Molecular Defects 
	NOTE: This procedure is useful for investigating the underlying causes of cuticle phenotypes or lethality that is not accompanied by cuticle defects.
	<ol>
		<li>For pRNAi, separate embryos from cotton balls at appropriate development stage and transfer them into an embryo collecting basket. The collection basket can be the same or similar to that used for Drosophila embryo collections.
		<ol>
			<li>Make the egg basket from a 25 ml plastic scintillation vial with the bottom of the vial removed, and a circle cut out of the lid (roughly 1 cm diameter). Place a circle of super fine mesh about 2.5 cm in diameter inside the lid, and then screw the scintillation vial on and use upside-down.</li>
			<li>As the mesh can be easily removed once the embryos are washed, immerse the mesh in a microcentrifuge tube with water to recover any embryos sticking to the mesh.</li>
		</ol>
		</li>
		<li>For eRNAi, at appropriate stage, add PBST to the Petri dish. Brush the embryos off the microscope slide and transfer them to an embryo collecting basket. 
		NOTE: Fixation, in situ hybridization, antibody staining, and nuclear staining protocols can be found in Xiang et al. 201553.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose.

While a small number of sophisticated model systems (mice, flies, worms) were developed during the 20th century, the 21st century has seen a wave of new animal systems being developed in labs throughout the world. These new systems allow scientists to address comparative, evolutionary questions that cannot be probed using only the &#39;standard&#39; model systems. This deployment of new models requires the rapid development of methods for lab culturing, gene identification, and functional approaches...

In 1998,&#160;Fire and Mello reported that double-stranded RNA (dsRNA) can induce inhibition of gene function in Caenorhabditis elegans1. This response triggered by dsRNA was named RNA interference (RNAi), and such RNAi-mediated gene silencing was reported to be conserved in animals, plants, and fungi2-7. In plants and some animals, RNAi functions systemically, meaning that the effect can spread to other cells/tissues where dsRNA is not directly introduced (reviewed in8-10). Scientists have made use of this endogenous cellular RNAi response by designing dsRNAs to target genes of interest, thereby knocking down gene function without directly manipulating the genome (reviewed in11-14).
RNAi is a powerful tool for functional studies due to the following advantages. First, even with minimal gene sequence information, a gene can be targeted using RNAi. This is especially important for studies of non-model organisms lacking genomic or transcriptomic data. Second, in organisms where the RNAi response is robustly systemic, RNAi-mediated gene knockdown can be performed at almost any developmental stage. This feature is very useful for studying the function of pleiotropic genes. Third, in some cases, RNAi effects spread to the gonads and progeny, such that phenotypes are observed in offspring15,16. This phenomenon, known as parental RNAi (pRNAi), is especially advantageous for genes impacting embryonic development, as numerous offspring produced by a single injected parent can be examined without direct manipulation of eggs. For these reasons, pRNAi is the method of choice. However, if pRNAi is ineffective, for example for genes required for oogenesis, then embryonic RNAi (eRNAi) must be used. Fourth, RNAi can be used to generate the equivalent of an allelic series in that the amount of dsRNA delivered can be varied over a range to produce weak to strong defects. Such a gradation of phenotypes can be helpful for understanding gene function when the gene is involved in a complex process and/or complete loss of function is lethal. Fifth, delivery of dsRNA is generally easy and feasible, especially in animals showing robust systemic RNAi responses. dsRNA can be introduced by microinjection1,5, feeding/ingestion17,18, soaking,19,20 and virus/bacteria-mediated delivery21,22. Sixth, unlike some gene targeting/editing methods, there is no need to screen for organisms carrying the mutation or to carry out genetic crosses to generate homozygotes when using RNAi. Therefore, compared to many other techniques for studying gene function, RNAi is fast, inexpensive, and can be applied for large-scale screens23-25.
The broad utility of RNAi provides means&#160;to carry out functional studies in a wide range of organisms, expanding the range of species available for study beyond the traditional model systems for which genetic tools have been developed. For example, studies using non-model systems are required to give insights into the evolution of genes and gene networks by comparing the functions of orthologs from species representing different development modes or exhibiting distinct morphological features26-29. These types of studies will provide a better understanding of biological diversity, with impacts for both applied and basic research.
Being the largest animal group on the planet, insects provide a great opportunity to explore the mechanisms underlying diversity. Additionally, insects are generally small, have short life cycles, high fecundity, and are easy to rear in the lab. In the past two decades, RNAi has been successfully applied in insects spanning orders, including Diptera (true flies)5, Lepidoptera (butterflies and moths)30, Coleoptera (beetles)16,31, Hymenoptera (sawflies, wasps, ants and bees)32, Hemiptera (true bugs), Isoptera (termites)34, Blattodea (cockroaches)35, Orthoptera (crickets, grasshoppers, locusts, and katydids)36and Phthiraptera (lice)37. Successful application of RNAi has provided functional data for studies of patterning in early embryogenesis (anterior-posterior axis32, dorsal-ventral axis28, segmentation26,38), sex determination39,40, chitin/cuticle biosynthesis41, ecdysone signaling42, social behavior43, and more. RNAi methods developed for different insect species may be of additional benefit in that they are likely to be useful for pest control (reviewed in44-46). RNAi effects will be gene-specific as well as species-specific, as long as non-conserved regions are chosen for targeting. For beneficial insect species like honeybees and silkworms, targeting genes vital for the survival of viruses or parasites to control infection may provide a novel strategy to protect these species47,48.
Dermestes maculatus (D. maculatus), common name hide beetle, is distributed worldwide except for Antarctica. As a holometabolous insect, the D. maculatus life cycle includes embryonic, larval, pupal, and adult stages (Figure 1). Because it feeds on flesh, D. maculatus is used in museums to skeletonize dead animals and forensic entomologists can use it to estimate time of death49,50. D. maculatus feeds on animal products including carcasses, dried meat, cheese, and the pupae/cocoons of other insects and thus causes damage to households, stored food, and the silk, cheese, and meat industries 51,52. Applying RNAi in this beetle could provide an efficient and environmentally friendly way to minimize its economic impact. Our lab has used D. maculatus as a new model insect to study segmentation53. In addition to being amenable to lab rearing, D. maculatus is of interest for basic research as it is an intermediate-germ developer, making it a useful species to study the transition between short- and long-germ development.
<img alt="Figure 1" src="/files/ftp_upload/54976/54976fig1.jpg" /> 
Figure 1: Life Cycle of D. maculatus. Photographs of D. maculatus at different life stages, as indicated. The life cycle from egg to adult takes three weeks at 30 &#176;C but longer at lower temperatures. (A, F) Freshly laid embryos are white to light yellow and oval, approximately 1.5 mm in length. Embryogenesis takes ~55 hr at 30 &#176;C. (B, C and G) Larvae have dark pigmented stripes and are covered with setae. Larvae go through several instars depending on the environment and their length can extend up to over 1 cm. (D, H) Young pupae are light yellow. Pupation takes ~ 5 - 7 days at 30 &#176;C. (E, I) Shortly after eclosion, dark pigmentation appears over the adult beetle body. Adults can live up to several months and one female can lay hundreds of embryos over her lifetime. <a href="http://cloudflare.jove.com/files/ftp_upload/54976/54976fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Previously, we showed that RNAi is effective in knocking down gene function in D. maculatus53. Here our experience rearing D. maculatus colonies in the laboratory is shared along with step-by-step protocols for both embryonic and parental RNAi set-up, injection, post-injection care, and phenotypic analysis. The dsRNA-mediated gene knockdown and analysis methods introduced here not only provide detailed information for addressing questions in D. maculatus, but also have potential significance for applying RNAi in other non-model beetle/insect species.

<table border="0" cellpadding="0" cellspacing="0" width="642">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Dermestes maculatus live beetles</td>
			<td style="width:140px;">Our lab or Carolina Biological Supply</td>
			<td style="width:119px;">#144168</td>
			<td style="width:167px;">Our lab strain was verified by COI barcoding; strain variation from Carolina cannot be ruled out</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Wet cat food</td>
			<td style="width:140px;">Fancy Feast</td>
			<td style="width:119px;"></td>
			<td>Chunks of meat with gravy. Can buy at most pet food and grocery stores</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Dry dog food</td>
			<td style="width:140px;">Purina Puppy Chow</td>
			<td style="width:119px;"></td>
			<td>Can buy at most pet food and grocery stores</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Insect cage (size medium, 30.5x19x20.3 cm)</td>
			<td style="width:140px;">Exo Terra</td>
			<td>PT2260</td>
			<td>For colony maintenance. Can use larger cage if needed</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Insect cage (size mini, 17.8x10.2x12.7 cm)</td>
			<td style="width:140px;">Exo Terra</td>
			<td>PT2250</td>
			<td>For embryo collection</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Petri dish</td>
			<td style="width:140px;">VWR</td>
			<td style="width:119px;">89038-968</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cotton ball</td>
			<td style="width:140px;">Fisher</td>
			<td style="width:119px;">22-456-883</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Megascript T7 transcription kit</td>
			<td style="width:140px;">Fisher</td>
			<td style="width:119px;">AM1334</td>
			<td>For 40 reactions</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Pneumatic pump</td>
			<td style="width:140px;">WPI</td>
			<td style="width:119px;">PV830</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Capillary holder</td>
			<td style="width:140px;">WPI</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Micromanipulator</td>
			<td style="width:140px;">NARISHIGE</td>
			<td style="width:119px;">MN-151</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Black filter paper (90 mm)</td>
			<td style="width:140px;">VWR</td>
			<td style="width:119px;">28342-010</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Food coloring (green)</td>
			<td>McCormick</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Borosilicate glass capillary</td>
			<td style="width:140px;">Hilgenberg</td>
			<td style="width:119px;">1406119</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Needle puller (micropipette puller)</td>
			<td style="width:140px;">Sutter Instrument Co.</td>
			<td style="width:119px;">P-97</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Microscope glass slide</td>
			<td>WorldWide Life Sciences Division</td>
			<td style="width:119px;">41351157</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sealing film (Parafilm M)</td>
			<td style="width:140px;">Fisher</td>
			<td style="width:119px;">13-374-12</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Model 801 Syringe (10 &#181;l )</td>
			<td style="width:140px;">Hamilton</td>
			<td style="width:119px;">7642-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Needle (32-gauge)&#160;</td>
			<td style="width:140px;">Hamilton</td>
			<td style="width:119px;">7762-05</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Fixation Solution (Pampel&#39;s)</td>
			<td>BioQuip Products, Inc.</td>
			<td style="width:119px;">1184C</td>
			<td>Toxic, needs to be handled in fume hood</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Forcep (DUMONT #5)</td>
			<td style="width:140px;">Fine Science Tools</td>
			<td style="width:119px;">11252-30</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Cover slip (24X50 mm, No. 1.5)</td>
			<td style="width:140px;">Globe Scientific</td>
			<td style="width:119px;">1415-15</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Eppendorf Femtotips Microloader pipette tip</td>
			<td style="width:140px;">Fisher</td>
			<td style="width:119px;">E5242956003</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Dissecting microscopy for embryo injection</td>
			<td style="width:140px;">Leica</td>
			<td style="width:119px;">M420</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Dissecting microscopy for larval phenotypic visualization</td>
			<td style="width:140px;">Zeiss</td>
			<td colspan="2">SteREO Discover. V12</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">DIC microscopy</td>
			<td style="width:140px;">Zeiss</td>
			<td style="width:119px;">AXIO Imager. M1</td>
			<td></td>
		</tr>
	</tbody>
</table>

rearing and double-stranded rna-mediated gene knockdown in the hide beetle, dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

The authors&#39; lab has used RNAi technology to study the functional evolution of genes regulating segmentation in insects53,55. While all insects are segmented, the genes regulating this process appear to have diverged during insect radiations26,38,56-63. Genetic screens in Drosophila identified a set of nine pair-rule segmentation genes that are responsible for promoting the formation of body segments64-70. Here, the ortholog of one of these ge...

Watch this Scientific Journal Video about Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus at JoVE.com

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

rearing-double-stranded-rna-mediated-gene-knockdown-hide-beetle

Universidad San Sebastian

Research

JoVE Journal

Genetics

10.5K Views.  University of Maryland. Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Video: Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Parallel Measurement of Circadian Clock Gene Expression and Hormone Secretion in Human Primary Cell Cultures

Circadian clocks are functional in all light-sensitive organisms, allowing for an adaptation to the external world by anticipating daily environmental changes. Considerable progress in our understanding of the tight connection between the circadian clock and most aspects of physiology has been made in the field over the last decade. However, unraveling the molecular basis that underlies the function of the circadian oscillator in humans stays of highest technical challenge. Here, we provide a detailed description of an experimental approach for long-term (2-5 days) bioluminescence recording and outflow medium collection in cultured human primary cells. For this purpose, we have transduced primary cells with a lentiviral luciferase reporter that is under control of a core clock gene promoter, which allows for the parallel assessment of hormone secretion and circadian bioluminescence. Furthermore, we describe the conditions for disrupting the circadian clock in primary human cells by transfecting siRNA targeting CLOCK. Our results on the circadian regulation of insulin secretion by human pancreatic islets, and myokine secretion by human skeletal muscle cells, are presented here to illustrate the application of this methodology. These settings can be used to study the molecular makeup of human peripheral clocks and to analyze their functional impact on primary cells under physiological or pathophysiological conditions.

Here, we describe settings to monitor in parallel circadian bioluminescence and the secretory activity of human islet cells and primary myotubes. For this, we employed lentiviral gene delivery of a luciferase core clock reporter, followed by in vitro synchronization and collection of outflow medium by continuous cell perifusion.

Circadian clocks are functional in all light-sensitive organisms, allowing for an adaptation to the external world by anticipating daily environmental ...

Preparation of rAAV9 to Overexpress or Knockdown Genes in Mouse Hearts

Controlling the expression or activity of specific genes through the myocardial delivery of genetic materials in murine models permits the investigation of gene functions. Their therapeutic potential in the heart can also be determined. There are limited approaches for in vivo molecular intervention in the mouse heart. Recombinant adeno-associated virus (rAAV)-based genome engineering has been utilized as an essential tool for in vivo cardiac gene manipulation. The specific advantages of this technology include high efficiency, high specificity, low genomic integration rate, minimal immunogenicity, and minimal pathogenicity. Here, a detailed procedure to construct, package, and purify the rAAV9 vectors is described. Subcutaneous injection of rAAV9 into neonatal pups results in robust expression or efficient knockdown of the gene(s) of interest in the mouse heart, but not in the liver and other tissues. Using the cardiac-specific TnnT2 promoter, high expression of GFP gene in the heart was obtained. Additionally, target mRNA was inhibited in the heart when a rAAV9-U6-shRNA was utilized. Working knowledge of rAAV9 technology may be useful for cardiovascular investigations.

In this manuscript, a method to prepare recombinant adeno-associated virus 9 (rAAV9) vectors to manipulate gene expression in the mouse heart is described.

Controlling the expression or activity of specific genes through the myocardial delivery of genetic materials in murine models permits the investigation ...

Studying Cell Cycle-regulated Gene Expression by Two Complementary Cell Synchronization Protocols

The gene expression program of the cell cycle represents a critical step for understanding cell cycle-dependent processes and their role in diseases such as cancer. Cell cycle-regulated gene expression analysis depends on cell synchronization into specific phases. Here we describe a method utilizing two complementary synchronization protocols that is commonly used for studying periodic variation of gene expression during the cell cycle. Both procedures are based on transiently blocking the cell cycle in one defined point. The synchronization protocol by hydroxyurea (HU) treatment leads to cellular arrest in late G1/early S phase, and release from HU-mediated arrest provides a cellular population uniformly progressing through S and G2/M. The synchronization protocol by thymidine and nocodazole (Thy-Noc) treatment blocks cells in early mitosis, and release from Thy-Noc mediated arrest provides a synchronized cellular population suitable for G1 phase and S phase-entry studies. Application of both procedures requires monitoring of the cell cycle distribution profiles, which is typically performed after propidium iodide (PI) staining of the cells and flow cytometry-mediated analysis of DNA content. We show that the combined use of two synchronization protocols is a robust approach to clearly determine the transcriptional profiles of genes that are differentially regulated in the cell cycle (i.e. E2F1 and E2F7), and consequently to have a better understanding of their role in cell cycle processes. Furthermore, we show that this approach is useful for the study of mechanisms underlying drug-based therapies (i.e. mitomycin C, an anticancer agent), because it allows to discriminate genes that are responsive to the genotoxic agent from those solely affected by cell cycle perturbations imposed by the agent.

We report two cell synchronization protocols that provide a context for studying events related to specific phases of the cell cycle. We show that this approach is useful for analyzing the regulation of specific genes in an unperturbed cell cycle or upon exposure to agents affecting the cell cycle.

The gene expression program of the cell cycle represents a critical step for understanding cell cycle-dependent processes and their role in diseases such ...

A Standard Methodology to Examine On-site Mutagenicity As a Function of Point Mutation Repair Catalyzed by CRISPR/Cas9 and SsODN in Human Cells

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point mutations, which often are responsible for a variety of human inherited disorders. Using a well-established cell-based model system, the point mutation of a single-copy mutant eGFP gene integrated into HCT116 cells has been repaired using this combinatorial approach. The analysis of corrected and uncorrected cells reveals both the precision of gene editing and the development of genetic lesions, when indels are created in uncorrected cells in the DNA sequence surrounding the target site. Here, the specific methodology used to analyze this combinatorial approach to the gene editing of a point mutation, coupled with a detailed experimental strategy to measuring indel formation at the target site, is outlined. This protocol outlines a foundational approach and workflow for investigations aimed at developing CRISPR/Cas9-based gene editing for human therapy. The conclusion of this work is that on-site mutagenesis takes place as a result of CRISPR/Cas9 activity during the process of point mutation repair. This work puts in place a standardized methodology to identify the degree of mutagenesis, which should be an important and critical aspect of any approach destined for clinical implementation.

This protocol outlines the workflow of a CRISPR/Cas9-based gene editing system for the repair of point mutations in mammalian cells. Here, we use a combinatorial approach to gene editing with a detailed follow-on experimental strategy for measuring indel formation at the target site&#8212;in essence, analyzing onsite mutagenesis.

Combinatorial gene editing using CRISPR/Cas9 and single-stranded oligonucleotides is an effective strategy for the correction of single-base point ...

Sequence-specific and Selective Recognition of Double-stranded RNAs over Single-stranded RNAs by Chemically Modified Peptide Nucleic Acids

RNAs are emerging as important biomarkers and therapeutic targets. Thus, there is great potential in developing chemical probes and therapeutic ligands for the recognition of RNA sequence and structure. Chemically modified Peptide Nucleic Acid (PNA) oligomers have been recently developed that can recognize RNA duplexes in a sequence-specific manner. PNAs are chemically stable with a neutral peptide-like backbone. PNAs can be synthesized relatively easily by the manual Boc-chemistry solid-phase peptide synthesis method. PNAs are purified by reverse-phase HPLC, followed by molecular weight characterization by matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF). Non-denaturing polyacrylamide gel electrophoresis (PAGE) technique facilitates the imaging of the triplex formation, because carefully designed free RNA duplex constructs and PNA bound triplexes often show different migration rates. Non-denaturing PAGE with ethidium bromide post staining is often an easy and informative technique for characterizing the binding affinities and specificities of PNA oligomers. Typically, multiple RNA hairpins or duplexes with single base pair mutations can be used to characterize PNA binding properties, such as binding affinities and specificities. 2-Aminopurine is an isomer of adenine (6-aminopurine); the 2-aminopurine fluorescence intensity is sensitive to local structural environment changes, and is suitable for the monitoring of triplex formation with the 2-aminopurine residue incorporated near the PNA binding site. 2-Aminopurine fluorescence titration can also be used to confirm the binding selectivity of modified PNAs towards targeted double-stranded RNAs (dsRNAs) over single-stranded RNAs (ssRNAs). UV-absorbance-detected thermal melting experiments allow the measurement of the thermal stability of PNA-RNA duplexes and PNA&#183;RNA2 triplexes. Here, we describe the synthesis and purification of PNA oligomers incorporating modified residues, and describe biochemical and biophysical methods for characterization of the recognition of RNA duplexes by the modified PNAs.

We report the protocols for the synthesis and purification of Peptide Nucleic Acid (PNA) oligomers incorporating modified residues. The biochemical and biophysical methods for the characterization of the recognition of RNA duplexes by the modified PNAs are described.

RNAs are emerging as important biomarkers and therapeutic targets. Thus, there is great potential in developing chemical probes and therapeutic ligands ...

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development of a customizable platform to rapidly generate gene modifications in a wide variety of organisms, including zebrafish. CRISPR-based genome editing uses a single guide RNA (sgRNA) to target a CRISPR-associated (Cas) endonuclease to a genomic DNA (gDNA) target of interest, where the Cas endonuclease generates a double-strand break (DSB). Repair of DSBs by error-prone mechanisms lead to insertions and/or deletions (indels). This can cause frameshift mutations that often introduce a premature stop codon within the coding sequence, thus creating a protein-null allele. CRISPR-based genome engineering requires only a few molecular components and is easily introduced into zebrafish embryos by microinjection. This protocol describes the methods used to generate CRISPR reagents for zebrafish microinjection and to identify fish exhibiting germline transmission of CRISPR-modified genes. These methods include in vitro transcription of sgRNAs, microinjection of CRISPR reagents, identification of indels induced at the target site using a PCR-based method called a heteroduplex mobility assay (HMA), and characterization of the indels using both a low throughput and a powerful next-generation sequencing (NGS)-based approach that can analyze multiple PCR products collected from heterozygous fish. This protocol is streamlined to minimize both the number of fish required and the types of equipment needed to perform the analyses. Furthermore, this protocol is designed to be amenable for use by laboratory personal of all levels of experience including undergraduates, enabling this powerful tool to be economically employed by any research group interested in performing CRISPR-based genomic modification in zebrafish.

Efficient Production and Identification of CRISPR/Cas9-generated Gene Knockouts in the Model System Danio rerio

Targeted genome editing in the model system Danio rerio (zebrafish) has been greatly facilitated by the emergence of CRISPR-based approaches. Herein, we describe a streamlined, robust protocol for generation and identification of CRISPR-derived nonsense alleles that incorporates the heteroduplex mobility assay and identification of mutations using next-generation sequencing.

Characterization of the clustered, regularly interspaced, short, palindromic repeat (CRISPR) system of Streptococcus pyogenes has enabled the development ...

Production and Purification of Baculovirus for Gene Therapy Application

Baculovirus has traditionally been used for the production of recombinant protein and vaccine. However, more recently, baculovirus is emerging as a promising vector for gene therapy application. Here, baculovirus is produced by transient transfection of the baculovirus plasmid DNA (bacmid) in an adherent culture of Sf9 cells. Baculovirus is subsequently expanded in Sf9 cells in a serum-free suspension culture until the desired volume is obtained. It is then purified from the culture supernatant using heparin affinity chromatography. Virus supernatant is loaded onto the heparin column which binds baculovirus particles in the supernatant due to the affinity of heparin for baculovirus envelop glycoprotein. The column is washed with a buffer to remove contaminants and baculovirus is eluted from the column with a high-salt buffer. The eluate is diluted to an isotonic salt concentration and baculovirus particles are further concentrated using ultracentrifugation. Using this method, baculovirus can be concentrated up to 500-fold with a 25% recovery of infectious particles. Although the protocol described here demonstrates the production and purification of the baculovirus from cultures up to 1 L, the method can be scaled-up in a closed-system suspension culture to produce a clinical-grade vector for gene therapy application.

In this protocol, baculovirus is produced by transient transfection of baculovirus plasmid into Sf9 cells and amplified in a serum-free suspension culture. The supernatant is purified by heparin affinity chromatography and further concentrated by ultracentrifugation. This protocol is useful for the production and purification of baculovirus for gene therapy application.

Baculovirus has traditionally been used for the production of recombinant protein and vaccine. However, more recently, baculovirus is emerging as a ...

A Method to Study the C924T Polymorphism of the Thromboxane A2 Receptor Gene

The thromboxane A2 receptor (TBXA2R) gene is a member of the G-protein coupled superfamily with seven-transmembrane regions. It is involved in atherogenesis progression, ischemia, and myocardial infarction. Here we present a methodology of patient genotyping to investigate the post-transcriptional role of the C924T polymorphism (rs4523) situated at the 3&#39; region of the TBXA2 receptor gene. This method relies on DNA extraction from whole blood, polymerase chain reaction (PCR) amplification of the TBXA2 gene portion containing the C924T mutation, and identification of wild type and/or mutant genotypes using a restriction digest analysis, specifically a restriction fragment length polymorphism (RFLP) on agarose gel. In addition, the results were confirmed by sequencing the TBXA2R gene. This method features several potential advantages, such as high efficiency and the rapid identification of the C924T polymorphism by PCR and restriction enzyme analysis. This approach allows a predictive study for plaque formation and atherosclerosis progression by analyzing patient genotypes for the TBXA2R C924T polymorphism. Application of this method has the potential to identify subjects who are more susceptible to atherothrombotic processes, in particular subjects in a high-risk, aspirin-treated group.

In the present study, we describe a methodology to analyze the C924T genotype. The protocol consists of three phases: DNA extraction, amplification by polymerase chain reaction (PCR), and analysis of the restriction fragment length polymorphism (RFLP) on agarose gel.

The thromboxane A2 receptor (TBXA2R) gene is a member of the G-protein coupled superfamily with seven-transmembrane regions. It is involved in ...

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to abiotic stresses and diseases. As the plant microbiome can be highly complex, low-cost, high-throughput methods such as amplicon-based sequencing of the 16S rRNA gene are often preferred for characterizing its microbial composition and diversity. However, the selection of appropriate methodology when conducting such experiments is critical for reducing biases that can make analysis and comparisons between samples and studies difficult. This protocol describes in detail a standardized methodology for the collection and extraction of DNA from soil, rhizosphere, and root samples. Additionally, we highlight a well-established 16S rRNA amplicon sequencing pipeline that allows for the exploration of the composition of bacterial communities in these samples, and can easily be adapted for other marker genes. This pipeline has been validated for a variety of plant species, including sorghum, maize, wheat, strawberry, and agave, and can help overcome issues associated with the contamination from plant organelles.

Here, we describe a protocol to obtain amplicon sequence data of soil, rhizosphere, and root endosphere microbiomes. This information can be used to investigate the composition and diversity of plant-associated microbial communities, and is suitable for the use with a wide range of plant species.

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to ...

Characterizing Histone Post-translational Modification Alterations in Yeast Neurodegenerative Proteinopathy Models

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives each year. Effective treatment options able to halt disease progression are lacking. Despite the extensive sequencing efforts in large patient populations, the majority of ALS and PD cases remain unexplained by genetic mutations alone. Epigenetics mechanisms, such as the post-translational modification of histone proteins, may be involved in neurodegenerative disease etiology and progression and lead to new targets for pharmaceutical intervention. Mammalian in vivo and in vitro models of ALS and PD are costly and often require prolonged and laborious experimental protocols. Here, we outline a practical, fast, and cost-effective approach to determining genome-wide alterations in histone modification levels using Saccharomyces cerevisiae as a model system. This protocol allows for comprehensive investigations into epigenetic changes connected to neurodegenerative proteinopathies that corroborate previous findings in different model systems while significantly expanding our knowledge of the neurodegenerative disease epigenome.

This protocol outlines experimental procedures to characterize genome-wide changes in the levels of histone post-translational modifications (PTM) occurring in connection with the overexpression of proteins associated with ALS and Parkinson&#39;s disease in Saccharomyces cerevisiae models. After SDS-PAGE separation, individual histone PTM levels are detected with modification-specific antibodies via Western blotting.

Neurodegenerative diseases, such as amyotrophic lateral sclerosis (ALS) and Parkinson&#39;s disease (PD), cause the loss of hundreds of thousands of lives ...