Name: rearing and double-stranded rna-mediated gene knockdown in the hide beetle, dermestes maculatus
Uploaded: 2016-12-28T13:00:00
Description: Watch this Scientific Journal Video about Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus at JoVE.com

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 be...

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.

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	<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 ...

The overall goal of this procedure is to explore gene function in Dermestes maculatus, a new insect model system, which represents the diverse clade of beetles, and facilitates the study of basic and applied research questions. This method can help answer key questions in the fields of developmental biology, evolutionary biology and beyond, by making it possible to test gene function in Dermestes embryos and adults. The main advantage of this technique is that RNA interference can be used to target any gene, based only on sequence information, even in the absence of a full genome sequence.Methods introduced here not only provide an approach to study gene function in Dermestes. They can also be applied to develop new insect model systems, and potentially to control insect pests. To prepare a rearing cage for a D.maculatus, first spread a thin layer of wood shavings into a medium sized insect cage.Place a section of styrofoam into the cage to allow larvae an environment, in which to pupate. Add twenty to fifty adult beetles or late instar larvae to the cage. Place wet cat food into a Petri dish or weigh boat, as a food source.Cover the cage with a mesh cloth and place it into an incubator. Before starting embryo collection, first check the cage carefully for any embryos present and remove them. Once the cage is clear, place a stretched cotton ball into the cage and incubate the colony at 30 degrees Celsius.After the appropriate time window, collect the cotton ball, and remove any adults present, and place them back into the cage. Over a piece of black construction paper, pinch the cotton ball gently, and tear it slowly into thin filament, so that any eggs present fall onto the black paper. Take a standard Petri dish lid, and use a piece of black filter paper to cover the inside.Stick a piece of double-sided tape along the edge of a standard microscope slide. Place the microscope slide onto the black filter paper in the Petri dish lid. Taking the paper containing the collected embryos, gently tap to transfer the embryos to the Petri dish lid.Using a paintbrush, align the embryos on the tape perpendicular to the slide, with the anterior or posterior end towards the edge. To begin RNAi, first take up 2-4 microliters of dsRNA into a 20 microliter micro loader pipette tip. Insert the tip into a pre-pulled glass capillary, referred to as the needle, and gently pipette the solution into the needle.Fix the needle into a glass capillary holder and then assemble the capillary holder into the micromanipulator. Next, carefully transfer the slide with embryos attached onto the stage of the dissecting microscope. Move the slide to bring one embryo into the center of the field and focus the microscope.Whilst looking into the dissecting scope, position the tip of the capillary with the micromanipulator. Bring the tip close to the end of the first embryo in the row. Switch on the nitrogen or air supply, and then set the solenoid input selector switch to vacuum.Adjust the eject pressure regulator to 10-15 psi, depending on the apparatus. Open the capillary, and the colored dsRNA solution will fill the tip, ready for injection. Move the capillary tip forward, and gently puncture the embryo.Under ideal pressure settings, no embryonic fluid will flow into the capillary. Step on the foot switch to eject dsRNA into the embryo, until the appropriate amount is dispensed. Leave the tip inside the embryo for around two seconds, and then carefully remove it.Once all of the embryos have been injected, place the slide into a Petri dish. Add a wet cotton ball to the dish, taking care not to let it come into contact with the slide. Cover the Petri dish and wrap it with sealing film.Label it with the dsRNA type, concentration, date, time and number of injected embryos. Finally, place the dish into an incubator at 30 degrees Celsius until the embryos hatch. To perform pRNAi, first collect pupae from the colony.Sort male and female pupae into two separate dishes, and place these into a 30 degree Celcius incubator. Check the plates daily for eclosed beetles, and transfer these individuals to two separate Petri dishes for male and female adults. Feed these adults and maintain the plates every other day, until enough adults have eclosed to begin injection.Once the adult pool is complete, anesthetize a female beetle, and hold the individual ventral side up with one hand, holding the syringe in the other. Gently penetrate the segmacoria membrane between sternites 2 and 3. Check the syringe scale and slowly dispense around two microliters per female.After injection, hold the needle steady for at least 5 seconds and then carefully withdraw it. Transfer injected females to a Petri dish with cat food. Label the dish appropriately.Place the dish into a 30 degrees Celcius incubator. After 24 hours, transfer the injected females to a mini cage, and add an equal number of the uninjected young males. Label the cage, and add cat food.Place the cage into a 30 degree Celsius incubator, to allow mating. The resultant offspring can then be examined to determine the effects of the pRNAi. This image shows an offspring of a pRNAi individual injected with DS GFP, green fluorescent protein as a control.Here, as with wild-type individuals, the offspring hatched with one pigmented stripe per segment. Neighboring pigmented stripes were separated by a non-pigmented gap. Following paired pRNAi, affected offspring hatched with fused neighboring pigmented stripes, indicating defective segmental boundaries.Phenotypic severity was variable, but fusions consistently appeared at certain boundary regions, indicating pair rule like defects. Similarly, cuticle phenotypes of unhatched embryos after paired knockdown showed loss of abdominal segments, as well as shortened body length. Staining of early embryos with engrailed antibody to examine molecular defects showed stripes of expression of equal intensity in each segment in control individuals.In contrast, reduced expression was detected in alternate stripes in offspring of Dmac paired dsRNA injected females. This pattern of reduced expression is consistent with the defective cuticle pattern observed in the affected hatched larvae. Here, for the first time, we present our protocol for knocking down a gene of interest, during the early embryonic development in Dermestes maculatus, using RNA interference.While attempting these procedures, it is important to remember to use embryos or female adults at the appropriate stage. Once mastered, injecting about 100 embryos or 10 female adults can be done within half an hour. Following this procedure, other methods, like embryo staining or quantitative PCR can be performed in order to answer additional questions about gene regulatory mechanisms.After watching this video, you should have a good understanding of how to set up a Dermestes lab colony, and perform both embryonic and parental RNA interference in this beetle. Currently, only a small number of insect models are available for functional studies. The techniques presented here pave the way for researchers to use Dermestes as a model system, expanding the scope of insects amenable to molecular genetic analysis.

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

Research

JoVE Journal

Genetics

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 ...

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 ...

A Bioinformatics Pipeline to Accurately and Efficiently Analyze the MicroRNA Transcriptomes in Plants

MicroRNAs (miRNAs) are 20- to 24-nucleotide (nt) endogenous small RNAs (sRNAs) extensively existing in plants and animals that play potent roles in regulating gene expression at the post-transcriptional level. Sequencing sRNA libraries by Next Generation Sequencing (NGS) methods has been widely employed to identify and analyze miRNA transcriptomes in the last decade, resulting in a rapid increase of miRNA discovery. However, two major challenges arise in plant miRNA annotation due to increasing depth of sequenced sRNA libraries as well as the size and complexity of plant genomes. First, many other types of sRNAs, in particular, short interfering RNAs (siRNAs) from sRNA libraries, are erroneously annotated as miRNAs by many computational tools. Second, it becomes an extremely time-consuming process for analyzing miRNA transcriptomes in plant species with large and complex genomes. To overcome these challenges, we recently upgraded miRDeep-P (a popular tool for miRNA transcriptome analyses) to miRDeep-P2 (miRDP2 for short) by employing a new filtering strategy, overhauling the scoring algorithm and incorporating newly updated plant miRNA annotation criteria. We tested miRDP2 against sequenced sRNA populations in five representative plants with increasing genomic complexity, including Arabidopsis, rice, tomato, maize and wheat. The results indicate that miRDP2 processed these tasks with very high efficiency. In addition, miRDP2 outperformed other prediction tools regarding sensitivity and accuracy. Taken together, our results demonstrate miRDP2 as a fast and accurate tool for analyzing plant miRNA transcriptomes, therefore a useful tool in helping the community better annotate miRNAs in plants.

A bioinformatics pipeline, namely miRDeep-P2 (miRDP2 for short), with updated plant miRNA criteria and an overhauled algorithm, could accurately and efficiently analyze microRNA transcriptomes in plants, especially for species with complex and large genomes.

MicroRNAs (miRNAs) are 20- to 24-nucleotide (nt) endogenous small RNAs (sRNAs) extensively existing in plants and animals that play potent roles in ...