The authors thank NSF (MCB-1157876) for funding. We also thank Alberto Civetta, John Roote, and two anonymous referees for their comments.

Small-scale experiments should be performed to become familiar with the whole procedure.
1. Collecting Virgin Females and Na&iuml;ve Males
The simplest version of the outlined experiment consists of four types of initial crosses, which involve the following combination of adults: a) w1118 individuals in order to collect virgin females; b) Oregon-R individuals in order to collect na&iuml;ve reference males; c) P{sw} homozygous males and virgin females from a control line that carries the wild-type organization of Sdic in order to collect na&iuml;ve experimental males (Type I); and d) P{sw} homozygous males and Sdic-sw-deletion-carrying virgin females in order to collect na&iuml;ve experimental males which carry the Sdic-sw deficiency (Type II).
<ol>
	<li>Set up multiple vials containing 8-10 females and 5 males each. Allow the females to lay eggs and transfer the adults to new vials every 3-5 days. Use bottles rather than vials if necessary and always use fresh food. The number of vials required will depend on the number of experimental male types under study and the number of individuals estimated to be necessary to detect differences (Table 1). More than 10 females per vial might result in over-crowding during larval growth, which may cause variation in the fertility of the progeny. Store vials at 25 &deg;C in a temperature-controlled chamber.</li>
	<li>Start collecting virgin females and na&iuml;ve males on days 11th and 12th after setting up the initial crosses. The waiting period varies according to the temperature; lower temperatures result in longer developmental time delaying the collection of individuals. Another factor affecting the timing to eclosion is the type of medium. Nutritious food, such as cornmeal-yeast medium 29, assures proper development of the adult reproductive system, which facilitates mating. Collect unmated flies every 4-6 hr. When collecting, anesthetize flies by introducing CO2 into the vial, tap the flies down, and sex them under stereomicroscope (Figure 2). Place the desired flies into different vials by sex and phenotype and label the vials appropriately.</li>
</ol>
Note 1. Collection routine starts in the morning by discarding the adults that emerged during the previous night. Collect virgin females and na&iuml;ve males once or twice during the day. Typically, D. melanogaster males become sexually mature 8 hr after eclosion at 25 &deg;C 30. If flies are maintained under 12:12 hr light/dark cycles, two peaks of eclosion are expected: during the first 1-2 hr after the light is turned on, and during the 2 hr before the light is turned off. Eclosion occurs within 24 hr after the pupa darkens.
Note 2. No more than 10 females should be put into the same vial to prevent over-crowding. This also limits the loss of females in the case they have to be discarded because at least one of them is not virgin.
Note 3. In order to collect the appropriate number of virgin females and na&iuml;ve males in a short time period, the following measures can be adopted. Set up 15-20 vials of w1118 for experiments involving two types of experimental males (Table 1). Lightly sprinkle the surface of the media and add a few dried active yeast pellets to facilitate oviposition. Transfer parents at least 4 times on consecutive days. To increase the available surface for pupation in each vial, insert multi-folded paper (7 x 5 cm) during the 4th or 5th day after the parents are transferred into another vial (Figure 3). If the number of emerged adults from the initial crosses is not enough, wait for a few more days and collect individuals from vials set up at different dates. Plan to carry out the experiments over several consecutive days to even out the workload.
2. Double-mating Experiments
Figure 4 shows the main steps involved in double-mating experiments performed in 14.
<ol>
	<li>On the morning of Day 1, set up the first mating. The Oregon-R males are the first to mate in the offense assay.
	<ol>
		<li>Using the aspirator, set up vials containing 10 4-5-day-old white-eyed w1118 virgin females and 10 red-eyed Oregon-R na&iuml;ve males each. Two to three vials are set up daily for 5 consecutive days. The number of vials to be set up might vary depending on the number of available individuals. Allow the flies to mate for 2 hr.</li>
		<li>Discard Oregon-R males and place each female into a new vial using an aspirator (Figure 5). Sex identification can be achieved by visual inspection of a few morphological differences (Figure 2). Label each vial appropriately and leave the female in the vial for 2 days (hereafter "v1").</li>
	</ol>
	</li>
	<li>On Day 3, set up the second mating. Selection of males and cross set up should be performed at random in order to minimize any potential bias towards any of the experimental male types.
	<ol>
		<li>Two hours before the light is turned off, transfer again the female into a new vial with an aspirator. Label the new vial appropriately (hereafter "v2").</li>
		<li>Introduce three 5-6-day-old experimental males of the same genotype into v2.</li>
		<li>Repeat 2.2.1 and 2.2.2 for each female.</li>
		<li>On Day 4, two hours right before the light is turned on (i.e. no more than 12 hr after 2.2.1), discard males by using the aspirator.</li>
	</ol>
	</li>
	<li>On Day 6, transfer each female again into another new vial. Label the new vials appropriately (hereafter "v3").</li>
	<li>On Day 10, discard the w1118 females.</li>
	<li>Examine the progenies in v2 and v3. The first inspection is performed after 13-15 days after the female is introduced into v2 (or v3), e.g. on Day 17 after the female first started ovipositing in v2. The second inspection is performed exactly after 17 days. This time frame poses a safe upper temporal boundary that guarantees no second generation in the same vial. Two inspections prevent over-crowding while facilitating progeny counting.
	<ol>
		<li>On Day 17, inspect v1 and ensure that there are red-eyed progeny present; if not mark the vial accordingly. Presence of white-eyed progeny indicates that the female was not virgin at the time of initial mating while no progeny indicates that mating with the reference or the experimental males did not occur.</li>
		<li>On Day 17, perform the first inspection of v2. Anesthetize emerged progeny in v2 with CO2 and sort them by eye color and sex. Record progeny numbers fathered by the first and second males. Figure 6 shows the expected phenotypes in the progeny of each mating scheme. In this particular experiment 14, only female progeny can be unambiguously assigned as sired by the reference or the experimental males and therefore only the information from daughters can be used to calculate P2. If with other markers the paternity of male offspring can be unambiguously assigned, progeny counts of both sexes can be used in downstream calculations. Discard the progeny but keep v2 for second inspection.</li>
		<li>On Day 20, proceed as in 2.5.2 with the newly emerged progeny in v2 and then discard the vial.</li>
		<li>On Day 20, inspect the emerged progeny in v3 for the first time and follow step 2.5.2.</li>
		<li>On Day 23, proceed as in 2.5.3 with the newly emerged progeny in v3.</li>
	</ol>
	</li>
</ol>
Note 1: Due to the potential inflating effect on P scores that multiple mating might have (2.2.2 and 2.2.3), mating can be limited to a particular time frame. The time frame will depend on the remating frequency associated with the genotype of the female and males used. The probability of multiple mating is specially reduced during night time 11.
Note 2: Do not add live yeast pellets because this might cause problems due to potential overgrowth of the yeast when the number of adult flies is low.
3. Data Analysis
<ol>
	<li>Recorded progeny counts should be organized appropriately for easy visual inspection and efficient analysis with a suitable statistical package (e.g. JMP from SAS Institute) or free web-based tools (e.g. <a href="http://www.vassarstats.net/" target="_blank">http://vassarstats.net/</a>).</li>
	<li>Sperm competitive ability. Add counts from v2 and v3. Female flies that have given rise to no or very limited progeny (e.g. &lt;10), died during the procedure, or were only successfully inseminated by one of the two males are considered as non-informative and are excluded from any downstream statistical analysis (2.5.1 and Table 2). Calculate the P2 score for each informative female.</li>
	<li>Test whether there are statistically significant differences in P2 values among the experimental males compared. This can be done using parametric (e.g. Tukey's HSD) or non-parametric (e.g. Steel-Dwass) tests depending on several factors including the skewness of the distribution of P2 values and the dependency between the variance and the mean. The angular transformation is commonly applied to proportions such as P2 prior to the use of parametric tests 31.</li>
	<li>Mating rate differences (optional). For each experimental male type, calculate the number of doubly mated females and those that only mated with the first male (the reference male in the offense assay and the experimental male in the defense assay). Using a two-tailed Fisher's exact test (available at <a href="http://www.langsrud.com/fisher.htm" target="_blank">http://www.langsrud.com/fisher.htm</a>), determine whether there are statistically significant differences between the two types of experimental males.</li>
	<li>Sex ratio (optional). For each experimental male, use the chi-square test to determine whether the sex ratio in the progeny from each female deviates from the expected 1:1 ratio.</li>
</ol>

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No conflicts of interest declared.

We have described the experimental design to assess differences in the relative contribution of genetically distinct D. melanogaster males to the progeny in controlled double-mating experiments 7,8. This has been done in the context of a genetic factor hypothesized to influence sperm competitive ability and it has been illustrated within the offense assay although a similar procedure applies to the defense assay (Figure 1). This experimental design can be modified to test other...

Since Geoff Parker noted the prevalence of sperm competition in insects and its evolutionary implications 2, a surge of studies in Drosophila and other species have tried to shed some light on this phenomenon at many different levels. Some examples of areas of interest have been the survey of its variation in natural populations 9,10, its genetic architecture and relevance of underlying genetic factors 11-14, and its role in driving coevolution between the sexes 15,16. In D. melanogaster females, the limited capacity of the specialized sperm-storage organs, a pair of spermatheca and the seminal receptacle 6,17, contributes to the competition of the sperm from different males. Approximately 1,500 sperm are transferred during mating to the female but only ~500 can be accommodated in the mentioned organs 18,19. In the laboratory, controlled double-mating experiments involving a reference male and one or more males of interest have been extensively used for evaluating sperm competitive ability 7,8.	
	Sperm competitive ability is estimated as the proportion of progeny sired by the experimental male in double-mating experiments over the total progeny, i.e. that from both the experimental and reference males. Sperm competitive ability comprises two components, each of them evaluated in a separate assay. In the offense assay, the ability of the experimental male sperm to displace the sperm from the first male, i.e. the reference male, is evaluated. Conversely, in the defense assay, the ability of the experimental male sperm to resist displacement or to reduce the fertilization success of the reference male sperm is evaluated. Depending on the type of assay, defense or offense, sperm competitive ability is estimated through the scores P1 or P2, respectively. P1 and P2 can only take values between 0 and 1. Intermediate values are usually interpreted as indirect evidence of sperm mixing, which suggests a physiological scenario involving direct sperm competition. Following the same rationale, extreme values can be interpreted as evidence for strong differential sperm competitive ability. Early studies showed that P2 in D. melanogaster is over 0.8 increasing as the time elapsed between the two matings lengthens 7. This same experimental design has been used in other Drosophila species, P2 being the commonly used statistic in studies to evaluate sperm competitive ability 20. For most species, the P2 values of the strains tested is higher than 0.6 21. Nevertheless, several other mechanisms unrelated to the direct competition between sperm of different males can yield identical scores (see Discussion).	
	Distinguishing progeny sired by the first or second males is possible through the use of easily identifiable markers. In early studies, one of the males was irradiated at sublethal doses of, for example, X-rays such that virtually all ova fertilized by irradiated sperm failed to hatch 7. Subsequently, mutations altering eye pigmentation or wing shape have been the most commonly used markers. Examples of the former are the mutations bw (brown) 9, cn (cinnabar)22 and w (white) 23, while the mutation Cy (Curly) 24 corresponds to the second type of phenotypes; some of these mutations have been combined in the same individual, e.g. cn bw. To a lesser extent, allozymes 25 and microsatellites 26,27 with known inheritance patterns have also been used.	
	The experimental design to test for differences in sperm competitive ability described here follows essentially that of Clark et al. 9. Results derived from these experiments give information solely about the differential paternity of the experimental male types under scrutiny. Assays that also make allowance for post-fertilization differences in fitness 14,28 and sperm visualization techniques 24 enable differences in P1 (or P2) scores to be interpreted as differences in sperm competence.	
	Figure 1 outlines the rationale of both the offense and the defense assays. To illustrate the logistics of the process, an offense experiment carried out in D. melanogaster 14 will be explained in detail. This particular offense assay was used to test for a measurable effect of the multigene family Sperm-specific dynein intermediate chain (Sdic) on sperm competitive ability. All the members of this multigene family reside in tandem on the X chromosome. Knockout males were generated by deleting the Sdic cluster. Because the deleted segment also included the essential gene short wing (sw) and the purpose of the study was to evaluate the relevance of Sdic, males carrying the Sdic-sw deletion were rescued by a transgenic copy of sw (symbolized as P{sw}, which also carried a mini-white reporter gene) on chromosome 2. Eye color was used as a visible marker for paternity identification. All the flies were in a white mutant background with the exception of those from the strain Oregon-R, which were used as reference males.

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		<tbody>
		 <tr>
			<td>Name of Reagent/Material</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		 </tr>
		 <tr>
			<td>Amber latex tubing</td>
			<td>VWR</td>
			<td>62996-473</td>
			<td>1/4 inch ID, 1/6 inch Wall</td>
		 </tr>
		 <tr>
			<td>1 ml graduated XL filter tips</td>
			<td>USA Scientific</td>
			<td>1126-7810</td>
			<td>Any 1 ml pipette tip can be used but semi-transparent and long tips are better as a fly receiver. If filter tips are used, poke out and discard the filter using a needle after the tip end is trimmed</td>
		 </tr>
		 <tr>
			<td>Parafilm</td>
			<td>Sigma</td>
			<td>P-7793</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Mesh Fabric</td>
			<td></td>
			<td></td>
			<td>Thin and smooth</td>
		 </tr>
		 <tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>S6D</td>
			<td></td>
		 </tr>
		 <tr>
			<td>CO2 Tank</td>
			<td>Airgas</td>
			<td>CD-50</td>
			<td>Use FlyNap (Carolina Biological Supply Company) as an alternative if CO2 is not available</td>
		 </tr>
		 <tr>
			<td>FlyStuff Foot Valve, complete system</td>
			<td>Genesee Scientific</td>
			<td>59-121C</td>
			<td>Not necessary if FlyNap is used</td>
		 </tr>
		 <tr>
			<td>Plastic vials</td>
			<td>Genesee Scientific</td>
			<td>32-109</td>
			<td>Come with carboard trays that can be reused for holding vials</td>
		 </tr>
		 <tr>
			<td>Cotton balls</td>
			<td>Fisher Scientific</td>
			<td>AS-212</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Active dry yeast</td>
			<td>Red Star</td>
			<td></td>
			<td>Found in general grocery store</td>
		 </tr>
		 <tr>
			<td>Sharpie markers</td>
			<td></td>
			<td></td>
			<td>Different colors may be used for marking different genotypes</td>
		 </tr>
		</tbody>
 </table>

assessing differences in sperm competitive ability in drosophila

Competition among conspecific males for fertilizing the ova is one of the mechanisms of sexual selection, i.e. selection that operates on maximizing the number of successful mating events rather than on maximizing survival and viability 1. Sperm competition represents the competition between males after copulating with the same female 2, in which their sperm are coincidental in time and space. This phenomenon has been reported in multiple species of plants and animals 3. For example, wild-caught D. melanogaster females usually contain sperm from 2-3 males 4. The sperm are stored in specialized organs with limited storage capacity, which might lead to the direct competition of the sperm from different males 2,5.
Comparing sperm competitive ability of different males of interest (experimental male types) has been performed through controlled double-mating experiments in the laboratory 6,7. Briefly, a single female is exposed to two different males consecutively, one experimental male and one cross-mating reference male. The same mating scheme is then followed using other experimental male types thus facilitating the indirect comparison of the competitive ability of their sperm through a common reference. The fraction of individuals fathered by the experimental and reference males is identified using markers, which allows one to estimate sperm competitive ability using simple mathematical expressions 7,8. In addition, sperm competitive ability can be estimated in two different scenarios depending on whether the experimental male is second or first to mate (offense and defense assay, respectively) 9, which is assumed to be reflective of different competence attributes.
Here, we describe an approach that helps to interrogate the role of different genetic factors that putatively underlie the phenomenon of sperm competitive ability in D. melanogaster.

Table 2 summarizes some salient features of two offense experiments (Assays 1 and 2) in which D. melanogaster experimental males with and without (Type I and II, respectively) a functional Sdic cluster are compared 14. After taking into account different incidences encountered with some replicates, 58-83% of the females were found to be informative and therefore the paternity counts of their progeny could be used for calculating P2. Results of non-parametric tests ...

Watch this Scientific Journal Video about Assessing Differences in Sperm Competitive Ability in Drosophila at JoVE.com

Assessing Differences in Sperm Competitive Ability in Drosophila

Differential sperm competitive ability among Drosophila males with distinct genotypes can be ascertained through double-mating experiments. Each of these experiments involves one of the males of interest and a reference male. Readily identifiable markers in the progeny allow inference of the fraction of individuals fathered by each male.

Assessing Differences in Sperm Competitive Ability in Drosophila

Competition among conspecific males for fertilizing the ova is one of the mechanisms of sexual selection, i.e. selection that operates on maximizing the ...

assessing-differences-in-sperm-competitive-ability-in-drosophila

Research

JoVE Journal

Biology

14.5K 조회수.  University of California, Irvine. 사이에 차동 정자 경쟁 능력 초파리 남성은 이중 결합 실험을 통해 확인 될 수 있습니다. 이 실험의 각각은 관심과 기준 남성의 남성 중 하나를 포함합니다. 자손에서 쉽게 식별 마커는 각각의 남성에 의해 아버지가 개인의 분수 추론 할 수 있습니다.

비디오: Assessing Differences in Sperm Competitive Ability in Drosophila

Fluorescence in situ hybridization (FISH) Protocol in Human Sperm

Aneuploidies are the most frequent chromosomal abnormalities in humans. Most of these abnormalities result from meiotic errors during the gametogenic process in the parents. In human males, these errors can lead to the production of spermatozoa with numerical chromosome abnormalities which represent an increased risk of transmitting these anomalies to the offspring.
For this reason, the technique of fluorescence in situ hybridization (FISH) on sperm nuclei has become a protocol widely incorporated in the context of clinical diagnosis. This practice provides an estimate of the frequencies of numerical chromosome abnormalities in the gametes of the patients that seek for genetic reproductive advice.
To date, the chromosomes most frequently included in sperm FISH analysis are chromosomes X, Y, 13, 18 and 21.
This video-article describes, step by step, how to process and fix a human semen sample, how to decondense and denature the sperm chromatin, how to proceed to obtain sperm FISH preparations, and how to visualize the results at the microscope. Special remarks of the most relevant steps are given to achieve the best results.

Fluorescence in situ hybridization FISH Protocol in Human Sperm

This video-article describes, step by step, how to process a semen sample to achieve good-quality fluorescence in situ hybridization on human spermatozoa. Preparations obtained can be used for aneuploidy screening in the context of clinical diagnosis.

Aneuploidies are the most frequent chromosomal abnormalities in humans. Most of these abnormalities result from meiotic errors during the gametogenic ...

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and hermaphrodites. In the earlier phase of gametogenesis, the germ cells of hermaphrodites differentiate into limited number of sperm - around 300 - and are stored in a small 'bag' called the spermatheca. Later on, hermaphrodites continually produce oocytes1. In contrast, males produce exclusively sperm throughout their adulthood. The males produce so much sperm that it accounts for &gt;50% of the total cells in a typical adult worm2. Therefore, isolating sperm from males is easier than from that of hermaphrodites.
Only a small proportion of males are naturally generated due to spontaneous non-disjunction of X chromosome3. Crossing hermaphrodites with males or more conveniently, the introduction of mutations to give rise to Him (High Incidence of Males) phenotype are some of strategies through which one can enrich the male population3.
Males can be easily distinguished from hermaphrodites by observing the tail morphology4. Hermaphrodite's tail is pointed, whereas male tail is rounded with mating structures.
Cutting the tail releases vast number of spermatids stored inside the male reproductive tract. Dissection is performed under a stereo microscope using 27 gauge needles. Since spermatids are not physically connected with any other cells, hydraulic pressure expels internal contents of male body, including spermatids2.
Males are directly dissected on a small drop of 'Sperm Medium'. Spermatids are sensitive to alteration in the pH. Hence, HEPES, a compound with good buffering capacity is used in sperm media. Glucose and other salts present in sperm media help maintain osmotic pressure to maintain the integrity of sperm.
Post-meiotic differentiation of spermatids into spermatozoa is termed spermiogenesis or sperm activation. Shakes5, and Nelson6 previously showed that round spermatids can be induced to differentiate into spermatozoa by adding various activating compounds including Pronase E. Here we demonstrate in vitro spermiogenesis of C. elegans spermatids using Pronase E.
Successful spermiogenesis is pre-requisite for fertility and hence the mutants defective in spermiogenesis are sterile. Hitherto several mutants have been shown to be defective specifically in spermiogenesis process7. Abnormality found during in vitro activation of novel Spe (Spermatogenesis defective) mutants would help us discover additional players participating in this event.

Isolation and In vitro Activation of Caenorhabditis elegans Sperm

A protocol for isolating and activating spermatids from male C. elegans is described here. Cutting the posterior end of male releases spermatids. The spermatids can be activated by addition of protease.

Males and hermaphrodites are the two naturally found sexual forms in the nematode C. elegans. The amoeboid sperm are produced by both males and ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Competitive Genomic Screens of Barcoded Yeast Libraries

By virtue of advances in next generation sequencing technologies, we have access to new genome sequences almost daily. The tempo of these advances is accelerating, promising greater depth and breadth. In light of these extraordinary advances, the need for fast, parallel methods to define gene function becomes ever more important. Collections of genome-wide deletion mutants in yeasts and E. coli have served as workhorses for functional characterization of gene function, but this approach is not scalable, current gene-deletion approaches require each of the thousands of genes that comprise a genome to be deleted and verified. Only after this work is complete can we pursue high-throughput phenotyping. Over the past decade, our laboratory has refined a portfolio of competitive, miniaturized, high-throughput genome-wide assays that can be performed in parallel. This parallelization is possible because of the inclusion of DNA 'tags', or 'barcodes,' into each mutant, with the barcode serving as a proxy for the mutation and one can measure the barcode abundance to assess mutant fitness. In this study, we seek to fill the gap between DNA sequence and barcoded mutant collections. To accomplish this we introduce a combined transposon disruption-barcoding approach that opens up parallel barcode assays to newly sequenced, but poorly characterized microbes. To illustrate this approach we present a new Candida albicans barcoded disruption collection and describe how both microarray-based and next generation sequencing-based platforms can be used to collect 10,000 - 1,000,000 gene-gene and drug-gene interactions in a single experiment.

We have developed comprehensive, unbiased genome-wide screens to understand gene-drug and gene-environment interactions. Methods for screening these mutant collections are presented.

By virtue of advances in next generation sequencing technologies, we have access to new genome sequences almost daily.  The tempo of these advances is ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell imaging studies. Methods for detecting RNAs have expanded greatly since seminal studies linked site-specific mRNA localization to gene expression regulation. Dynamic mRNA processes can now be visualized via approaches that detect mRNAs, coupled with microscopy set-ups that are fast enough to capture the dynamic range of molecular behavior. The molecular beacon technology is a hybridization-based approach capable of direct detection of endogenous transcripts in living cells. Molecular beacons are hairpin-shaped, internally quenched, single-nucleotide discriminating nucleic acid probes, which fluoresce only upon hybridization to a unique target sequence. When coupled with advanced fluorescence microscopy and high-resolution imaging, they enable one to perform spatial and temporal tracking of intracellular movement of mRNAs. Although this technology is the only method capable of detecting endogenous transcripts, cell biologists have not yet fully embraced this technology due to difficulties in designing such probes for live cell imaging. A new software application, PinMol, allows for enhanced and rapid design of probes best suited to efficiently hybridize to mRNA target regions within a living cell. In addition, high-resolution, real-time image acquisition and current, open source image analysis software allow for a refined data output, leading to a finer evaluation of the complexity underlying the dynamic processes involved in the mRNA&#39;s life cycle.
Here we present a comprehensive protocol for designing and delivering molecular beacons into Drosophila melanogaster egg chambers. Direct and highly specific detection and visualization of endogenous maternal mRNAs is performed via spinning disc confocal microscopy. Imaging data is processed and analyzed using object detection and tracking in Icy software to obtain details about the dynamic movement of mRNAs, which are transported and localized to specialized regions within the oocyte.

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Here, we present a protocol for the visualization, detection, analysis and tracking of endogenous mRNA trafficking in live Drosophila melanogaster egg chamber using molecular beacons, spinning disc confocal microscopy, and open-source analysis software.

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell ...

Medium-throughput Screening Assays for Assessment of Effects on Ca2+-Signaling and Acrosome Reaction in Human Sperm

Ca2+-signaling is essential to normal sperm cell function and male fertility. Similarly, the acrosome reaction is vital for the ability of a human sperm cell to penetrate the zona pellucida and fertilize the egg. It is therefore of great interest to test compounds (e.g., environmental chemicals or drug candidates) for their effect on Ca2+-signaling and acrosome reaction in human sperm either to examine the potential adverse effects on human sperm cell function or to investigate a possible role as a contraceptive. Here, two medium-throughput assays are described: 1) a fluorescence-based assay for assessment of effects on Ca2+-signaling in human sperm, and 2) an image cytometric assay for assessment of acrosome reaction in human sperm. These assays can be used to screen a large number of compounds for effects on Ca2+-signaling and acrosome reaction in human sperm. Furthermore, the assays can be used to generate highly specific dose-response curves of individual compounds, determine potential additivity/synergism for two or more compounds, and to study the pharmacological mode of action through competitive inhibition experiments with CatSper inhibitors.

Medium-throughput Screening Assays for Assessment of Effects on Ca2+-Signaling and Acrosome Reaction in Human Sperm

Here, two medium-throughput assays for assessment of effects on Ca2+-signaling and acrosome reaction in human sperm are described. These assays can be used to quickly and easily screen large amounts of compounds for effects on Ca2+-signaling and acrosome reaction in human sperm.

Ca2+-signaling is essential to normal sperm cell function and male fertility. Similarly, the acrosome reaction is vital for the ability of a human sperm ...

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to recognize a wide range of particles and destroy microbial pathogens. As organisms age, this process begins to decline, yet little is known about the underlying mechanisms or the genetic basis of immunosenescence. Here, an injection based in vivo phagocytosis assay is used to assess age related changes in different aspects of phagocytosis, such as binding, engulfment, and degradation of internalized particles, by quantifying phagocytic events in hemocytes in adult Drosophila. Drosophila melanogaster has become an ideal model to investigate age related changes in innate immune function for many reasons. For one, many genetic components and functions of the innate immune response, including phagocytosis, are evolutionarily conserved between Drosophila and mammals. Because of that, results obtained from using this protocol are likely to be widely relevant to understanding the age related changes in immune function in a variety of organisms. Additionally, we note that this method provides quantitative estimates of hemocyte phagocytic ability, which could be useful for a variety of research topics, and need not be limited to studies of aging.

Assessing the Age-Specific Phagocytic Ability of Adult Drosophila melanogaster Hemocytes using an In Vivo Phagocytosis Assay

This protocol describes an in vivo assay of phagocytosis used to assess and quantify the ability of young and aged Drosophila melanogaster hemocytes to phagocytose bacteria.

Phagocytosis is an essential function of the innate immune response. This process is carried out by phagocytic hemocytes whose primary function is to ...

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

The pupae of Drosophila melanogaster are immobile for several days during metamorphosis, during which they develop a new body with a thin transparent adult integument. Their immobility and transparency make them ideal for in vivo live imaging experiments. Many studies have focused on the dorsal epithelial monolayer of the pupal&#160;notum because of its accessibility and relatively large size. In addition to the studies of epithelial mechanics and development, the notum has been an ideal tissue to study wound healing. After an injury, the entire epithelial repair process can be captured by live imaging over 6-12 h. Despite the popularity of the notum for live imaging, very few published studies have utilized fixed notum samples. Fixation and staining are common approaches for nearly all other Drosophila tissues, taking advantage of the large repertoire of simple cellular stains and antibodies. However, the pupal notum is fragile and prone to curling and distortion after removal from the body, making it challenging to complement live imaging. This protocol offers a straightforward method for fixing and staining the pupal notum, both intact and after laser-wounding. With this technique, the ventral side of the pupa is glued down to a coverslip to immobilize the pupa, and the notum is carefully removed, fixed, and stained. The notum epithelium is mounted on a slide or between two coverslips to facilitate imaging from the tissue&#39;s dorsal or ventral side.

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

The present protocol details the preparation and visualization of fixed tissue of the Drosophila pupal notum. It can be used for either intact or wounded tissue, and the original architecture of the tissue is preserved. The procedures for dissecting, fixing, and staining are all described in this article.

The pupae of Drosophila melanogaster are immobile for several days during metamorphosis, during which they develop a new body with a thin transparent ...