This work was supported by the Institut National de la Sant&#233; et de la Recherche M&#233;dicale, the Centre National de la Recherche Scientifique, the Universit&#233; de Strasbourg, the H&#244;pital de Strasbourg, the Association pour la Recherche sur le Cancer, the Institut National du Cancer, the Agence Nationale de la Recherche and the R&#233;gion Alsace.
Pietro Laneve has been supported by the Fondation pour la Recherche M&#233;dicale. Currently, he is a recipient of a Istituto Italiano Tecnologia (IIT) fellowship. Publication costs are supported by the Neurex network (TriNeuron &#8211; Program Interreg IV Upper Rhine)

1. Sample Collection
<ol>
	<li>Detach semi-adherent Schneider&#8217;s 2 cells, (1-5 x 106 cells on average), by pipetting and harvest them by centrifugation (100 x g&#160;for 1 min). In case of in vitro cultured adherent cells, trypsinize them in standard conditions or directly collect them from plates into a convenient amount of RNA extraction reagent, by help of a cell scraper.</li>
	<li>For embryo collection, grow Drosophila strains in fly cages and let embryos accumulate on egg laying plates. Wipe embryos off by a paintbrush in distilled water (dH2O), discard debris by sieve filtering, rinse and recover embryos in a vial48.</li>
	<li>As an alternate technique, manually dissect tissues/organs. Isolate testes (a preferential system for piRNA analysis) from Drosophila abdomen in phosphate buffered saline (PBS) under a stereomicroscope (20X magnification), by using dissection needles or forceps as visualized in Sullivan et al49.</li>
	<li>Pestle&#160;homogenize samples upon addiction of 0.5 volumes of RNA extraction reagent.</li>
</ol>
2. RNA Extraction/Analysis
<ol>
	<li>Following manufacturers&#8217; instructions perform RNA isolation through commercial reagents. CAUTION! RNA extraction agents are usually toxic and caustic. Alternatively, apply a Proteinase K-based protocol for RNA isolation50, as follows:
	<ol>
		<li>Dilute samples into 400 &#956;l of Stop Mix and incubate at RT&#160;for 15 min.</li>
		<li>Recover hydrosoluble components by standard phenol:chloroform:isoamyl alcohol and chloroform extractions and by ethanol (EtOH) precipitation/washing. Air&#160;dry the samples and resuspend into diethylpyrocarbonate (DEPC)-treated double distilled (dd)H2O.</li>
	</ol>
	</li>
	<li>Assess RNA concentration by UV spectrophotometric measurement. Also ensure the RNA purity is high, close to 2.0, based on A260/A280 reading.</li>
	<li>Check RNA preparation on denaturing agarose gel as follows:
	<ol>
		<li>In a clean beaker, dissolve 1.2 g of Agarose in 82 ml of DEPC-ddH2O, under continuous stirring. Boil until completely dissolved.</li>
		<li>Allow the gel solution to cool down; when temperature reaches 60&#160;&#176;C add 10 ml of 10X 4-morpholinepropanesulfonic acid (MOPS) buffer to 1X final concentration (FC), and 8 ml of 37% formaldehyde to 3% FC. CAUTION!&#160;Formaldehyde is a carcinogen.</li>
		<li>Pour the gel into a horizontal electrophoresis cell. Allow the gel to solidify under a hood for at least 1 hr. Following solidification, submerge the gel into MOPS 1X.</li>
		<li>Aliquot 2 &#956;g of RNA sample and 1 &#956;g of Ladder (use an RNA marker when accurate&#160;evaluation is desired). Add 3 volumes of agarose loading dye (ALD) to RNA and to Ladder. Heat at 70 &#176;C for 5 min and immediately cool on ice.&#160;CAUTION!&#160;Contents of ALD (ethidium bromide and formamide, see Materials) are mutagen and corrosive, respectively.</li>
		<li>Load the samples and run the gel at 50 V for about 1 hr with occasional buffer recycling. Check the RNA fractionation pattern by UV visualization or gel imaging. Recognize discrete bands corresponding to 28S and 18S rRNAs and to tRNAs (see Figure 1).</li>
	</ol>
	</li>
	<li>Proceed to Northern Assay or store RNA at -80 &#176;C.</li>
</ol>
3. Denaturing Acrylamide Gel Preparation
<ol>
	<li>Use a small vertical electrophoretic apparatus for this fractionation step (plate size about 8 cm x 9 cm, comb thickness 0.75 mm).</li>
	<li>Assemble clean glasses together in a casting frame.</li>
	<li>Prepare 10 ml of 10% acrylamide/bis-acrylamide (19:1) solution in MOPS 1X, 7 M urea. Immediately before pouring, add 100 &#956;l of 10% ammonium persulfate and 10 &#956;l of TEMED and rapidly mix the solution. CAUTION! Acrylamide is neurotoxic and carcinogen.</li>
	<li>Pour the gel between glass plates and insert the well comb carefully to avoid bubbles. Let the gel polymerize at RT for at least 45 min before use. Alternatively, store the gel O/N, wrapped in water-soaked filter papers and sealed in saran wrap.</li>
</ol>
4. Sample Preparation
<ol>
	<li>Aliquot 0.5-20 &#956;g of total RNA for each sample. If volume exceeds 3-4 &#956;l, vacuum&#160;dry the sample.</li>
	<li>For size determination, run an aliquot (20-30 cps) of radiolabeled low-range ladder in parallel to samples, as a marker (see section 4.3 and 4.4 for preparation). Treat it in parallel to samples, as described in step 4.5.</li>
	<li>Ladder: set up a phosphate forward reaction using 0.1 &#956;g of a 10 bp-step ladder. Incubate at 37 &#176;C for 30 min and stop the reaction by adding ethylenediaminetetraacetic acid (EDTA) to 20 mM FC.</li>
	<li>Purify by EtOH precipitation/washing, air&#160;dry and resuspend in ddH2O&#160;(about 100 &#956;l). CAUTION! 32P is a radioactive source.</li>
	<li>To each sample add an equal volume of acrylamide blue dye (ABD). Denature by heating at 80 &#176;C for 5 min and cool on ice until loading. CAUTION! Formamide (content in ABD) is corrosive.</li>
</ol>
5. Electrophoretic Fractionation
<ol>
	<li>Remove the glass plates from the casting support and properly assemble them in the electrophoresis cell. Fill both inner and outer chamber with sufficient Running Buffer (RB).</li>
	<li>Gently remove the comb and pre-run the gel at 200 V&#160;for 10 min. &#160;Before loading samples, wash away urea deposits from the wells by squirting some RB through a syringe.</li>
	<li>Use flat tips to carefully stratify samples from step 4.5 at the bottom of each well. Check anode and cathode connections to avoid inverted running. After the samples enter the gel at 100 V&#160;for 5 min, increase the voltage to 200 V.</li>
	<li>For a fractionation length of about 7 cm, a 45 min-long run is sufficient. Avoid Bromophenol blue (a tracking pigment in ABD) to overflow the gel.</li>
	<li>Visualize on-gel the fractionated long RNAs by ethidium bromide (EtBr) staining, as an optional step:
	<ol>
		<li>Cut away 2 cm from the top of the gel and briefly rinse the slice in ddH2O.</li>
		<li>Incubate the gel slice at RT for 15 min with gentle shacking in RB, EtBr 0.5 &#956;g/ml FC.</li>
		<li>Destain the gel for 15 min in RB, visualize long rRNAs by UV irradiation or gel imaging. Check for RNA loading and quality (see Figure&#160;1, Panel 7). CAUTION! Ethidium bromide is a mutagen.</li>
	</ol>
	</li>
	<li>Meanwhile proceed blotting the lower part of the gel containing small RNA species.</li>
</ol>
6. Gel Blotting
<ol>
	<li>Cut 6 pieces of blotting paper to fit the blotting area and an equivalent piece of uncharged nylon membrane. Dampen paper sheets, membrane and 2 blotting pads in RB. Make sure to completely soak the pads by repeatedly squeezing them in RB.</li>
	<li>Remove the gel from the apparatus and open the glasses by mean of a spatula. Use a clean cutter to remove the wells. Place a sheet of wet Whatman paper on the gel and gently lift it up.</li>
	<li>Place the nylon membrane on the free face of the gel. Mark a corner to orient the filter according to sample loading. Complete the &#8220;sandwich&#8221; (3 paper pieces each side). Roll a plastic rod on the blot surface, to remove any possible air bubbles.</li>
	<li>Place the blot between two blotting pads and then in the blotting cassette. Assemble the cassette in the blotting module, fill the chamber with RB and check for the correct orientation (nylon membrane must stay between the gel and the positive terminus).</li>
	<li>Transfer at 20 V for 20 min. Note: to ensure homogeneous conditions, after 10 min open the blotting module and rotate the sandwich by 180&#176;&#160; before completing the transfer.</li>
</ol>
7. Membrane Crosslinking
<ol>
	<li>Prepare 6 ml of fresh Crosslinking Solution (XLS). Saturate a 10 cm&#160;x 10 cm (larger than nylon membrane) piece of blotting paper with XLS. CAUTION! HCl (a component of XLS) is highly corrosive.</li>
	<li>Dismantle the blot and place the membrane on the wet 3MM filter paper. Note: RNA must not be in direct contact with the saturated paper. Place the filter and paper between two glass plates and wrap in saran-wrap.</li>
	<li>Heat the membrane at 60&#160;&#176;C for 2 hr,&#160;followed by a ddH2O washing. Store at -20&#160;&#176;C or proceed to&#160;hybridization.</li>
</ol>
8. Membrane Prehybridization
<ol>
	<li>If RNA loading has been checked (optional step 5.5), directly proceed to hybridization. Or else, cut away 1 cm from to the top of the nylon filter, to avoid probe cross-hybridizations to&#160;fractionated long RNAs.</li>
	<li>Preheat 10 ml of Hybridization Solution (HS) at 37 &#176;C. Denature 1 mg of Salmon Sperm at 95&#160;&#176;C for 5 min, cool on ice and add to HS.</li>
	<li>Incubate the filter in HS at 37&#730;C for at least 1 hr, under rotation in a hybridization oven. Check that the RNA-side of the filter faces HS.</li>
</ol>
9. Probe Synthesis
<ol>
	<li>Set up a phosphate forward reaction on 10 pmol of specific antisense oligonucleotide, as described in 4.3.</li>
	<li>Add 5 volumes of Tris-EDTA-NaCl (TEN) buffer to the reaction and purify the labeled primer from unincorporated nucleotides by gel exclusion chromatography on G-25 Sephadex columns (or also by EtOH precipitation). Probe activity can be assessed by Liquid Scintillation Counter.</li>
</ol>
10. Membrane Hybridization
<ol>
	<li>Replace the exhausted HS with a fresh 10 ml aliquot (prepared and complemented as described above). Heat the probe at 95&#160;&#176;C for 10 min, cool on ice and add to novel HS. Incubate the filter at 37&#160;&#176;C ON.</li>
</ol>
11. Membrane Washing
<ol>
	<li>Recover the hybridization solution and store it at -20 &#176;C in a radioactivity-shielding box.</li>
	<li>Rinse the filter in Washing Buffer (WB) and wash it thoroughly for 10 min at RT.</li>
	<li>Use a Geiger Mueller Detector for checking residual radioactivity at the filter corners. When background emission is around 5 cps, proceed to membrane exposition.</li>
</ol>
12. Signal Detection
<ol>
	<li>Expose the filter on autoradiography films or on a molecular imager screen ON. Reveal signals by photographic processing or digital conversion.</li>
	<li>Analyze the band intensity by dedicated quantification software (ImageJ or Optiquant).</li>
</ol>
13. Membrane Stripping
<ol>
	<li>To remove primary signals, submerge the membrane in 500 ml of boiling Stripping solution, then incubate at RT for 1 min. Proceed to novel (pre)hybridization or store the filter at -20 &#176;C.</li>
</ol>

The authors declare that they have no competing financial interests.

Although our appreciation of the sophisticate intertwining through which RNA regulates neurogenesis is continuously increasing, the mechanistic implications of RNA-based circuitries affecting neural stem cell biology, neuronal differentiation/functional integration, development of neural pathologies and cancers remain unexplored. The maintenance of such networks through kingdoms makes the potential of genetic systems as Drosophila melanogaster instrumental to unravel unexplored cellular pathways, in which short ...

RNA biochemistry has experienced spectacular progresses in the past years1. Our comprehension of RNA potential in the control of gene expression has been burst by the identification of powerful noncoding ribo-regulators2, by the discovery of novel RNA-based regulatory mechanisms3,4 and by a deeper characterization of already known post-transcriptional events5. All together these studies have allowed RNA biology to dramatically make the scene, becoming a major research subject in the current scientific landscape. In particular, over recent years we are getting a sense of the pervasive impact of the &#8220;RNA world&#8221; on molecular neurobiology6, one of the most dynamic research domains in modern life science. In the last decade of the past century, the overall scientific scenario has been revolutionized by the discovery of the RNA interference7,8 and of the small regulatory RNAs9,10 with particular regard to microRNAs11, endogenously expressed small noncoding RNAs implicated in the control of nearly all cellular functions as pleiotropic and combinatorial regulators of gene expression.
Almost 10 years after miRNA initial discovery in Caenorhabditis elegans by Ambros and Ruvkun&#8217;s labs, renewed attention was turned to the field when high numbers of miRNAs were identified in Drosophila and in human cells as well12-15. Since then, thanks to versatile transgenic approaches, Drosophila melanogaster has stood out as a valuable biological context for delving into miRNA biosynthesis and activity. Drosophila miRNAs have revealed distinct functions in insect-specific or evolutionarily conserved processes, spanning from aging to metabolism, signaling pathways, behavior and, of course, neurogenesis. Along this direction, we recently unveiled a novel link16 in the intriguing correlation occurring between the master gene gcm/glide and the RNA pathway. The fly transcription factor Gcm/Glide17-19 constitutes a unique example of cell fate determinant, which dictates the glial vs. neuronal fate choice in multipotent fly neural precursors20. Twenty-year long research on this topic has clearly underlined the occurrence of multiple and overlapping inputs of gene expression regulation converging over gcm/glide21-28 to establish the threshold levels required for balancing the delicate ratio between neuronal and glial counterparts during neural development.
We discovered that regulation via Dmel-miR279 targeting represents a further control level contributing to post-transcriptional fine-tuning of gcm/glide expression16. Globally, these research lines have required specific methodological improvements: along years, several technologies originally developed to analyze traditional RNAs have been converted for quantifying small non coding RNAs, like as RNAse protection assays, cDNA arrays29-31, real-time PCR methods32-35 and sequencing36,37. On the other side, recalibration of technical approaches has fostered continuous progresses in the field.
Northern Blot assay (NB, or RNA gel blot assay) constitutes a representative instance: it is largely employed to profile RNA accumulation, since it ensures both expression level quantitation as well as size determination. However, the intrinsic poor sensitivity of the method is limiting when it is to be applied to low-abundant gene expression fine tuners, like short RNAs. A detrimental consequence is the requirement of large amounts of total RNA, which makes difficult its application to specific biological samples. For such reasons, specific NB variants for small RNAs detection have been developed38-40: we took advantage of an improved NB procedure41 (ENB, Enhanced Northern Blot), while elucidating the abovementioned interplay between Dmel-miR279 and gcm/glide.
This method relies on a chemical crosslinking step based on the activity of a Carbodiimide derivative [1-ethyl-3-(3-dimethylaminopropyl)carbodiimide, EDC] to fix nucleic acid onto solid supports. Carbodiimide is a versatile cross linker known to catalyze the formation of amide bonds between activated carboxyl or phosphate groups and amine groups42. This property can be exploited to covalently couple small RNAs via their mono-phosphorylated 5&#8242;-hydroxyl group to amino groups at the surface of nylon membranes. The resulting attachment configuration increases the accessibility of the immobilized nucleic acid and, in turn, the efficiency of probe-target hybridization, which results in remarkable detection enhancement43.
The technique assumes particular relevance in Drosophila molecular studies, by which the occurrence of novel and distinctive classes of small noncoding RNAs is emerging44. Among these, rasiRNAs45,46 constitute a specific subtype of piwi-interacting RNAs (piRNAs47), involved in sequence-specific gene silencing. Operative details of this method are fully described and visualized hereinafter, relative to the analysis of the microRNAs Dmel-miR279 and Dmel-miR286 and, for the first time, of one rasiRNA, rasi4. We pushed to extremes this method, which allowed us revealing poorly expressed targets from minimal amounts of RNA (less than 1 &#956;g).

enhanced northern blot detection of small rna species in drosophila melanogaster

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular biology has been greatly fueled by the discovery of noncoding RNAs as major players in post-transcriptional regulation. Such a revolutionary perspective has been accompanied and triggered by the development of powerful technologies for profiling short RNAs expression, both at the high-throughput level (genome-wide identification) or as single-candidate analysis (steady state accumulation of specific species). Although several state-of-art strategies are currently available for dosing or visualizing such fleeing molecules, Northern Blot assay remains the eligible approach in molecular biology for immediate and accurate evaluation of RNA expression. It represents a first step toward the application of more sophisticated, costly technologies and, in many cases, remains a preferential method to easily gain insights into RNA biology. Here we overview an efficient protocol (Enhanced Northern Blot) for detecting weakly expressed microRNAs (or other small regulatory RNA species) from Drosophila melanogaster whole embryos, manually dissected larval/adult tissues or in vitro cultured cells. A very limited amount of RNA is required and the use of material from flow cytometry-isolated cells can be also envisaged.

By following the overall procedure described in the text and schematically represented in Figure 1, we assessed small RNAs expression from complex RNA samples of different origins. In the experiment presented in Figure 1, RNA was isolated from Drosophila Schneider&#8217;s 2 cells by Proteinase K extraction, checked for integrity (optional step: denaturing agarose gel fractionation) and loaded in double. One half of gel was blotted on a neutral membrane and chemically cross-linke...

Watch this Scientific Journal Video about Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster at JoVE.com

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The aim of this publication is to visualize and discuss the operative steps of an Enhanced Northern Blot protocol on RNA extracted from Drosophila melanogaster embryos, cells, and tissues. This protocol is particularly useful for the efficient detection of small RNA species.

Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

The last decades have witnessed the explosion of scientific interest around gene expression control mechanisms at the RNA level. This branch of molecular ...

enhanced-northern-blot-detection-small-rna-species-drosophila

Research

JoVE Journal

Biology

23.9K Views.  Institut de Génétique et de Biologie Moléculaire et Cellulaire.  מטרתו של פרסום זה היא לדמיין ולדון בצעדים האופרטיביים של פרוטוקול צפון כתם משופר על RNA שחולץ מתסיסנית עוברים, תאים ורקמות. פרוטוקול זה שימושי במיוחד לאיתור יעיל של מיני RNA קטנים. 

Video: Enhanced Northern Blot Detection of Small RNA Species in Drosophila Melanogaster

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

A Noninvasive Hair Sampling Technique to Obtain High Quality DNA from Elusive Small Mammals

Noninvasive genetic sampling approaches are becoming increasingly important to study wildlife populations. A number of studies have reported using noninvasive sampling techniques to investigate population genetics and demography of wild populations1. This approach has proven to be especially useful when dealing with rare or elusive species2. While a number of these methods have been developed to sample hair, feces and other biological material from carnivores and medium-sized mammals, they have largely remained untested in elusive small mammals. In this video, we present a novel, inexpensive and noninvasive hair snare targeted at an elusive small mammal, the American pika (Ochotona princeps). We describe the general set-up of the hair snare, which consists of strips of packing tape arranged in a web-like fashion and placed along travelling routes in the pikas&#x2019; habitat. We illustrate the efficiency of the snare at collecting a large quantity of hair that can then be collected and brought back to the lab. We then demonstrate the use of the DNA IQ system (Promega) to isolate DNA and showcase the utility of this method to amplify commonly used molecular markers including nuclear microsatellites, amplified fragment length polymorphisms (AFLPs), mitochondrial sequences (800bp) as well as a molecular sexing marker. Overall, we demonstrate the utility of this novel noninvasive hair snare as a sampling technique for wildlife population biologists. We anticipate that this approach will be applicable to a variety of small mammals, opening up areas of investigation within natural populations, while minimizing impact to study organisms.

We present a noninvasive sampling approach to efficiently collect hair samples from elusive small mammals, as shown for the American pika. We demonstrate the utility of this method by extracting DNA from sampled hair and amplifying several types of molecular markers commonly used in studies of wildlife ecology and conservation.

Noninvasive genetic sampling approaches are becoming increasingly important to study wildlife populations. A number of studies have reported using ...

Substrate Generation for Endonucleases of CRISPR/Cas Systems

The interaction of viruses and their prokaryotic hosts shaped the evolution of bacterial and archaeal life. Prokaryotes developed several strategies to evade viral attacks that include restriction modification, abortive infection and CRISPR/Cas systems. These adaptive immune systems found in many Bacteria and most Archaea consist of clustered regularly interspaced short palindromic repeat (CRISPR) sequences and a number of CRISPR associated (Cas) genes (Fig. 1) 1-3. Different sets of Cas proteins and repeats define at least three major divergent types of CRISPR/Cas systems 4. The universal proteins Cas1 and Cas2 are proposed to be involved in the uptake of viral DNA that will generate a new spacer element between two repeats at the 5' terminus of an extending CRISPR cluster 5. The entire cluster is transcribed into a precursor-crRNA containing all spacer and repeat sequences and is subsequently processed by an enzyme of the diverse Cas6 family into smaller crRNAs 6-8. These crRNAs consist of the spacer sequence flanked by a 5' terminal (8 nucleotides) and a 3' terminal tag derived from the repeat sequence 9. A repeated infection of the virus can now be blocked as the new crRNA will be directed by a Cas protein complex (Cascade) to the viral DNA and identify it as such via base complementarity10. Finally, for CRISPR/Cas type 1 systems, the nuclease Cas3 will destroy the detected invader DNA 11,12 .
These processes define CRISPR/Cas as an adaptive immune system of prokaryotes and opened a fascinating research field for the study of the involved Cas proteins. The function of many Cas proteins is still elusive and the causes for the apparent diversity of the CRISPR/Cas systems remain to be illuminated. Potential activities of most Cas proteins were predicted via detailed computational analyses. A major fraction of Cas proteins are either shown or proposed to function as endonucleases 4.
Here, we present methods to generate crRNAs and precursor-cRNAs for the study of Cas endoribonucleases. Different endonuclease assays require either short repeat sequences that can directly be synthesized as RNA oligonucleotides or longer crRNA and pre-crRNA sequences that are generated via in vitro T7 RNA polymerase run-off transcription. This methodology allows the incorporation of radioactive nucleotides for the generation of internally labeled endonuclease substrates and the creation of synthetic or mutant crRNAs. Cas6 endonuclease activity is utilized to mature pre-crRNAs into crRNAs with 5'-hydroxyl and a 2',3'-cyclic phosphate termini.

CRISPR/Cas systems mediate adaptive immunity in Bacteria and Archaea. Many Cas proteins are proposed to act as endoribonucleases acting on crRNA precursors of varying length. Here we illustrate three different approaches to generate pre-crRNA substrates for the biochemical analysis of Cas endonuclease activity.

The  interaction of viruses and their prokaryotic hosts shaped the evolution of  bacterial and archaeal life. Prokaryotes developed several strategies to ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Maintenance of a Drosophila melanogaster Population Cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life cycle of Drosophila enables collection of large quantities of material from embryos, larvae, pupae and adult flies, in a synchronized way, at a low economic cost. A major strategy for propagating large numbers of flies is the use of a fly population cage. This useful and common tool in the Drososphila community is an efficient way to regularly produce milligrams to tens of grams of embryos, depending on uniformity of developmental stage desired. While a population cage can be time consuming to set up, maintaining a cage over months takes much less time and enables rapid collection of biological material in a short period. This paper describes a detailed and flexible protocol for the maintenance of a Drosophila melanogaster population cage, starting with 1.5 g of harvested material from the previous cycle.

Maintenance of a Drosophila melanogaster Population Cage

This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life ...